Plant Physiol. (1974) 53, 171-175

Biosynthesis of Phosphatidylethanolamine by EnzymePreparations from Plant Tissues

Received for publication June 22, 1973 and in revised form September 24, 1973

B. A. MACHER AND J. B. MUDDDepartment of Biochemistry and Statewide Air Pollution Research Center, University of California, Riverside,California 92502

ABSTRACT DG + CDP-ethanolamine -. PE + CMP (6)

The enzymic utilization of cytidine diphosphoethanolaminein the synthesis of phosphatidylethanolamine is localized inthe microsomal fraction of spinach (Spinacia oleracea)leaves. The metal ion requirement can be satisfied by Mn+(saturation approximately 0.6 mM) or Mg2` (saturation ap-proximately 25 mM). The enzyme has a pH optimum of 8.0 inthe presence of Mn2' and 7.5 in the presence of Mg2'. AMichaelis constant of 20 /M was determined for cytidinediphos-phoethanolamine. Enzyme activity was stimulated by thiolcompounds and inhibited by thiol reagents. No inhibition wasobtained with cytidine monophosphate and Tween 80.The in vitro biosynthesis of phosphatidylethanolamine was

inhibited by cytidine diphosphocholine and the biosynthesis ofphosphatidylcholine was inhibited by cytidine diphosphoetha-nolamine. Activities of the two synthetic systems were in-distinguishable on the basis of susceptibility to Iyophilizationand inhibition by thiol reagents.

An alternative pathway in animal tissue has been described(9):

PE + serine PS + ethanolamine (7)

(3)

However, reaction 7 is clearly an exchange reaction and canonly lead to net synthesis of PE if the organism can make PS(e.g. by reaction 2) and ethanolamine.The cells of higher plants have both procaryotic and eucary-

otic characteristics exemplified by the nucleic acids of the nu-clei and other subcellular organelles. It is, therefore, of some in-terest to examine the biosynthesis of PE in higher plants to seewhether the mechanism of biosynthesis is by the prevalentmethod in animals, the method in bacteria, or both. The reac-tion studied in this paper is the formation of PE from CDP-ethanolamine (reaction 6).

PS -> PE + CO2

Our understanding of the biosynthesis of phospholipids inanimal tissues stems from the basic findings of Kennedy andco-workers (8, 9, 12, 19, 20). Subsequent work with bacterialsystems has shown similarities with, but also important differ-ences from, the animal systems. The most remarkable differ-ence is in the biosynthesis of PE.' In bacteria this compound ismade by a pathway concluding in the three final steps (28):

PA + CTP -- CDP-diglyceride + PPi (1)

CDP-diglyceride + serine -- PS + CMP (2)PS-PE + CO2 (3)

In animals the major pathway to PE appears to be (20):

PA-DG + Pi (4)phosphorylethanolamine + CTP

CDP-ethanolamine + PPi (5)

'Abbreviations: PE: phosphatidylethanolamine; CDP-choline:cytidine diphosphocholine; CDP diglyceride: cytidine diphosphodi-glyceride; CDP-ethanolamine: cytidine diphosphoethanolamine;DEAE: diethylaminoethyl; DG: 1,2 diacyl-sn-glycerol; DDT: di-thiothreitol; lyso PE: lysophosphatidylethanolamine; PA: phosphati-dic acid; PC: phosphatidylcholine; PS: phosphatidylserine; pmb:p. mercuribenzoate.

MATERIALS AND METHODS

Materials. Plant materials were purchased from local gro-cery stores. CDP-ethanolamine was prepared by the method ofKennedy (19) or purchased from Sigma Chemical Company,St. Louis, Missouri, CDP-choline was purchased from SigmaChemical Company. "4C-CDP-choline was purchased from In-ternational Chemical and Nuclear Corporation, Irvine, Cali-fornia. "4C-Phosphoryl ethanolamine was obtained from NewEngland Nuclear.

"4C-CDP-ethanolamine was prepared from "4C-phosphorylethanolamine using enzyme preparations from chicken liver asdescribed by Chojnacki and Metcalf (10). The product was iso-lated by ion exchange chromatography on Dowex-1-formateaccording to the method of Kennedy (19). Purity of the productwas evaluated by paper chromatography in solvent systemsrecommended by Scheider (31) and scanning of the paper stripwith a radioactivity scanner.

Methods. Plant extracts were prepared by homogenizing agiven weight of material with twice as much homogenizing me-dium. The homogenizing medium consisted of 0.25 M sucrose,10 mM tris-HCl pH 7.5, 1 mm EDTA. The plant material andcold medium were homogenized in a Waring Blendor (three5-sec bursts), and the homogenate was pressed through twolayers of cheesecloth. In the preparation of subcellular frac-tions, the filtrate was first centrifuged for 5 min at 1,000g. Thesupernatant was centrifuged for 30 min at 20,000g and cen-trifuged for 60 min at 100,000g. The first two pellets (1,OOOgand 20,000g) were suspended in 20 ml each 10 mm tris-HCIpH 7.5, 1 mM EDTA, and the l00,OOOg pellet in 10 ml of thesame medium. Reaction mixtures contained 2 mm DTT, 20 mM

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MACHER AND MUDD

2000

E

1000

0

FRACTIONIn

2.0

1.0

0.2

EC

0toin0.1 <

0.0

SOLVENT ° 10 20 30 40 50 60 70 80 90I---C----tC/M:19/l j--C/M:9/1--j.--C/M: 2/1 -------C/M: 2/1----

I(I/. NH40Ac)100 ML 100 ML 150 ML 150 ML 200 ML

FIG. 1. Separation of radioactive products. Products of theenzymic synthesis (13,000 dpm in 4.5 ml CHCI3) were mixed withlipids isolated from egg yolk (250 mg in 1.5 ml CHCl3). The samplewas applied to a column (2.2 X 17 cm) of DEAE-cellulose (acetate)and washed into the column with CHC13. The sequence of solventsis indicated. Fractions of 7.7 ml were automatically collected.Aliquots of 1.0 ml were taken and dried under an air stream inscintillation counter vials. Scintillation fluid was added and thesamples counted. Aliquots of 1.0 ml were also taken for measure-

ment of ester bonds by the hydroxamate method. Aliquots were

also chromatographed on thin layer plates of silica gel G usingthe solvent CHCla-CH3OH-HAc-H20, 170:25:25:6. After develop-ment the spots were visualized by spraying the plates with 1%iodine in methanol. Spots were identified by comparison withstandards. Of the radioactivity applied to the column, 88% was

recovered in the fractions.

MgCl2, 50 mM tris-HCl pH 7.5, 0.05 mm "4C-CDP-ethanolamine(56,000 dpm) and enzyme in a final volume of 2.0 ml. Reac-tions were started by addition of the enzyme. Incubations were

for 20 min at 30 C. At the end of the reaction, 6 ml of meth-anol-chloroform (2:1, v/v) were added, and the extraction oflipid was performed according to the method of Bligh andDyer (7). The chloroform layer was washed twice with waterand then an aliquot of the chloroform was evaporated in an

air stream and 10 ml of scintillation fluid were added (4 g/lPPO and 0.2 g/l POPOP in toluene). Radioactivity in the vialswas counted in a 720 series scintillation counter (Nuclear Chi-cago).The product of the assay system was characterized by three

methods. (a) The radioactive product was co-chromatographedwith egg yolk lipids on thin layers of Silica Gel G in the solventsystem chloroform-methanol-acetic acid-water, 170:25:25:6.The plate was air dried and then sprayed with a 1% solution(w/v) of iodine in methanol. The area of PE was marked andwhen the iodine had sublimed all areas of the chromatogramwere scraped into scintillation vials and counted. More than90% of the recovered counts were found in the PE area. An-other plate run in the same solvent system was exposed to x-rayfilm. In addition to PE a small amount of radioactivity was de-tected in a spot tentatively identified as lyso-PE. (b) The radio-active product was subjected to conditions of mild alkaline hy-drolysis as described by Benson (4). The water soluble productswere chromatographed on strips of Whatman 4 filter paperusing butanol-propionic acid-water, 70:35:50 as solvent. Ra-dioactivity was subsequently located by passing the stripthrough a radioactivity scanner. The radioactivity was coinci-dent with glyceryl phosphoryl ethanolamine (obtained fromdeacylated egg yolk lipid) as detected by a ninhydrin spray(0.2% in water saturated butanol). (c) The radioactive productwas mixed with egg yolk lipids and the lipids separated on a

column of DEAE-cellulose (acetate form) according to the di-rections of Allen et al. (1). Effluent fractions were assayed forester bonds by the hydroxamate method (14), by radioactivity,and lipid components were measured by TLC. The results olEsuch a column are shown in Figure 1. Some radioactivity was

found in peaks preceding the major peak of PE, but the lattercontained most of the radioactivity.

Other analytical methods included the method of Arnon (3)for Chl, and the method of Lowry et al. (24) for protein.

RESULTS

Subcellular Distribution. The subcellular distribution inspinach leaves of the capability to synthesize PE is shown inTable I. Recovery of protein and Chl was close to 100%, butthe recovery of enzyme units was close to twice that determinedin the homogenate. Although this can be rationalized as dueto the presence of an inhibitor in the homogenate or a com-

peting enzyme (such as a phosphatase breaking down CDP-ethanolamine), no evidence has been gathered either way. Themajor points are that most of the enzymic activity and thehighest specific activity was found in the 100,000g pellet (thisfraction was used in subsequent assays), and the data for Chldistribution makes it unlikely that PE is synthesized in thechloroplast.

Characteristics of the Enzyme from Spinach Leaves. Resultsof a time course experiment are shown in Figure 2. Althoughincorporation was maintained throughout the 40-min reactionperiod, the relationship with time was not linear even in theearliest stages. In subsequent experiments reactions were run

for 20 min or less.The effect of changing protein concentration is shown in

Figure 3. Increased amounts of protein caused increased in-corporation in the range tested, but the linearity of the relation-ship was evident only at the lower protein concentrations.

Both magnesium and manganese ions fulfill a metal ion re-

quirement. Determination of saturation curves for these two

metal ions showed that saturation was achieved by magnesiumions at about 25 mm and by manganese ions at about 0.6 mM(data not shown). Table II shows the effects of two differentconcentrations of metal ions.

Determination of the Michaelis constant for CDP-ethanol-amine is shown in Figure 4. In the inset, the data from thesubstrate concentration curve are plotted by the method recom-

Table I. Subcellular Distribution of Ethanolaminie-phosphotransferase

Subcellular fractions were prepared from spinach leaves as

described in "Materials and Methods." Reaction mixtures andassay were standard as described in "Materials and Methods";0.4 ml aliquots of the homogenate, 1,OOOg pellet, and l00,OOOg

supernatant, and 0.2 ml aliquots of the 20,000g and 100,OOOg pelletswere tested. Incubation was for 20 min at 30 C. Values of activitywere determined from duplicate assays. An enzyme unit is definedas that amount of enzyme catalyzing the incorporation of 1 nmoleof ethanolamine in 20 min.

Protein Chlorophyll EnzymeFraction

Total Total Specific activity Total

Homogenate1,OOOg pellet

20,000g pellet100,OOOg pellet100,OOOg supernatant

112033819378552

4025.910.90.451.84

units/mg protein

0.240.060.813.870.05

u(nits

26519

15630228

I

A 0~~~~~~~~~

0~~~~~

NL---PC P

172 Plant Physiol. Vol. 53, 197zi

mg

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PHOSPHATIDYLETHANOLAMINE BIOSYNTHESIS

mended by Dowd and Riggs (15). The Michaelis constant isestimated to be 20 /tM.The pH optimum for the incorporation of phosphoryl eth-

anolamine was determined in the presence of either manga-nese or magnesium ions (Fig. 5). The optimum in the presenceof manganese ions appeared to be 8.0, and in the presence ofmagnesium ions 7.5, although a sharp maximum was not ob-tained.

Thiol compounds were always included in reaction mixtures.When tested directly some microsome preparations were stim-ulated (DTT was superior to GSH), but others were unaffected.

8

-2

0

0

0na)0

6

4

2

0

0 10 20 30 40time, min.

FIG. 2. Time course of PE synthesis. Reaction mixtures andassay were as described in "Materials and Methods."

8

-0a)

L-

0I.-0C-)c

(C)0Ec

6

4

o

0.0 0.4 0.8mg protein

10

173

1 I I 1

m ~ ~~~~= 20/.LM

_/~~~~~~~~~~~-

/ .-12 -8 -4 0 4 S8 12 16 20

-I -_0 2 4 6 8 10 12

ICDP-ethanolamine] x 105M14

FIG. 4. Effect of substrate concentration of PE biosynthesis. Re-action mixtures contained 50 mM tris-HCl pH 7.5, 2 mM GSH, 10mM MgSO4, 1'C-CDP-ethanolamine (32,000 dpm), and cold 0.5 mmCDP-ethanolamine to give the final concentrations indicated, 0.2ml microsomal preparation (approximately 0.8 mg protein), in afinal volume of 2.0 ml. Incubation was at 30 C for 20 min. Re-actions were assayed as described in "Materials and Methods."Direct plot and linear transformation (inset) are shown.

6

C

C-

c 4

c

07.0 8.0 9.0

pH

FIG. 5. pH curves for PE biosynthesis in the presence of Mg2`and Mn2+. Reaction mixtures and assay were as described in"Materials and Methods."

i.2 1.6

FIG. 3. PE biosynthesis as a function of protein concentration.Reaction mixtures and assay were as described in "Materials andMethods."

Table II. Effect ofMetal Ions on EthanolaminephosphotransferaseReaction mixtures and assays were as described in "Materials

and Methods," except for the variation in metal ion concentration.Reaction mixtures contained 0.88 mg of protein. Data presentedare averages of duplicate assays.

Metal Ion Added PE Synthesized

nmoles0 1.1210 mM Mg2+ 16.825 mm Mg2+ 19.20.4 mM Mn2+ 29.30.6mMMn2+ 34.8

This variability probably reflects the degree of SH oxidationduring preparation of the enzyme. It was considered that thereaction may be inhibited by one of the products, CMP. Whenthis was tested directly, no effect was observed up to concen-trations 80% of the CDP-ethanolamine substrate.

Detergents may stimulate lipid biosynthesis catalyzed bymembrane bound enzymes, either by making enzymes more ac-cessible to substrate or by solubilizing exogenous substrate. Wehave found no effect of Tween 80 up to concentrations of0.08% in the reaction mixture. The lack of inhibition makes itpossible to add lipid substrates or cofactors to the reaction mix-ture. So far we have not been able to stimulate PE biosynthesisby addition of exogenous diglycerides in Tween 80 suspensions.Does the Same Enzyme Catalyze the Reaction of Ethanol-

aminephosphotransferase and Cholinephosphotransferase? Theresults reported above on the saturation levels of Mn' andMg2+, and the shift of pH optimum depending on which metalion was used, reminded us strongly of our earlier studies on PCsynthesis (13). We have tried a number of methods to try todistinguish between the cholinephosphotransferase and the eth-anolaminephosphotransferase.

Plant Physiol. Vol. 53, 1974

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0

0~~~

II

2 _

2

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MACHER AND MUDD

According to the earlier work of Kennedy (19) on phospho-lipid synthesizing enzymes from rat liver, the enzyme catalyzingthe synthesis of PC is more stable to lyophilization than thatsynthesizing PE. Attempts to discriminate between PE and PCsynthesis by spinach leaf microsomes were unsuccessful: bothenzymic activities were quite stable to lyophilization. These re-sults are shown in Table III which also includes data on inhibi-tion by pmb. The synthesis of both PE and PC are equally sus-ceptible to the mercaptide forming reagent.

Data in Tables IV and V also emphasize the similarities inthe two enzymic activities. Tables IV and V show that CDP-choline is a potent inhibitor of PE synthesis and CDP-ethanol-amine is a potent inhibitor of PC synthesis. The information inthese tables again suggests that the same enzyme may catalyzethe two reactions. Kinetic experiments indicate that inhibitionof PE synthesis by CDP-choline is competitive with respect toCDP-ethanolamine.PE Synthesis in Different Plants. Tissues of several plants be-

sides spinach were tested for the ability to incorporate phos-phoryl ethanolamine from CDP-ethanolamine into PE. On aprotein basis preparations from the fruit of the bell pepper(Capsicum annum) were as efficient as those from spinach leaf(Spinacia oleracea), preparations from onion stem (A lliumcepa) were about half as effective, and those from carrot root(Daucus carota) and parsley (Petroselinuin sativa) gave no de-tectable activity.

Table III. I,ihibitiouz of Etlaiaolamirie anld Cholinle-phosphotrantsferase

Reaction mixtures and assays were as described in "Materialsand Methods." Fresh microsomes were assayed the day of prepara-tion. Lyophilized microsomes were prepared overnight and sus-pended the following day in water to give a volume equal to thatlyophilized. The enzyme was incubated with pmb for 10 min at30 C prior to the addition of substrate, Mg2+, and thiol. Proteinconcentration was 1.12 mg/reaction vessel. Data presented areaverages of duplicate determinations.

Substrate pmb Fresh LvophilizedMlicrosornes Microsornes

III.\fm tsoles incorporated

CDP-choline 0 l.01 0.760.01 0.37 0.350.1 0.01 0.01

CDP-ethanolamine 0 4.41 4.610.01 1.69 2.390.1 0.02 0.06

Table IV. I,ilibitioii of Etlhaiolaminiephosphot-anisferase byCDP-Cholinie

Reaction mixtures and assays were as described in "Materialsand Methods" (50 ,uM CDP-ethanolamine), except for the additionof CDP-choline as indicated. Protein concentration was 1.12mg/reaction vessel. Data presented are averages of duplicateassays. This experiment was performed simultaneously with thatpresented in Table V.

CDP-Choline PE Synthesized

,.M nmoles

0 17.32.5 12.05 10.710 8.1

20 7.6

40 3.8

Inhibition

C-

0

3138535678

Table V. lithibitiont of Choliniephosphotransferase byCD P-Ethantolamin7e

Reaction mixtures and assays were as described in "Materialsand Methods," except that '4C-CDP-ethanolamine was replacedby '4C-CDP-choline (10 ,uM; 150,000 dpm). Cold CDP-ethanol-amine was added to give final concentrations in the reaction mix-tures as indicated. Protein concentration was 1.12 mg/vessel.Data presented are averages of duplicate assays. This experimentwas performed simultaneously with that presented in Table IV.

CDP-Ethanolamine PC Synthesized Inhibition

AYl.5f nmsoles %

0 2.26 012.5 0.96 5725 0.61 7350 0.42 81100 0.25 89200 0.23 90

DISCUSSION

Lipid analyses of the tissues of higher plants invariably re-port substantial amounts of PE (1, 5, 6, 16, 21, 27, 30, 34, 36).On the other hand, PS is less frequently reported (5, 6, 16, 34).If PS is the precursor of PE, as it is in bacteria (28), only smallamounts may accumulate before decarboxylation takes places.The alternative pathway to PE is by phosphorylation of eth-

anolamine followed by reactions 5 and 6. These reactions, aswell as the analogous reactions for PC, have been well au-thenticated in animal tissues (20).The evidence for the biosynthesis fo PC in higher plants is

very good. Morre et al. (26) have measured the activities of allthree of the relevant enzymes (choline kinase, phosphorylcho-line cytidyltransferase, and cholinephosphotransferase) in onionstem. Ramasarma and Wetter (29) have characterized the cho-line kinase from rape seed and they specifically stated that theenzyme had no activity on ethanolamine. The choline kinaseof yeast has a Km from choline of 20 ,uM and 10 mM for ethanol-amine (38). Choline kinase in plants has also been reported byTanaka et al. (33), but only low activities were found. Subse-quent reports of choline kinase in higher plants have been madeby Johnson and Kende (17) and by Lord et al. (23). The reportof Morre et al. (26) on the cytidyl transferase has been verifiedin barley endosperm (16) and in castor bean cotyledons (23).The cholinephosphotransferase in plants has been described indetail (13) after Morre's initial report (26).The analogous pathway for PE synthesis is not well authen-

ticated. Marshall and Kates (25) have reported the presence ofethanolamine kinase in spinach leaf preparation. These authorswere unable to detect the cytidyl transferase enzyme. We havebeen unable to detect the formation of CDP-ethanolamine fromphosphoryl ethanolamine in experiments with plant tissue inour laboratory. On the other hand, Kennedy and Weiss (20)reported that enzyme preparations from carrot root were ableto synthesize CDP-ethanolamine. The data reported in this pa-per on the final step of PE synthesis, the ethanolaminephospho-transferase, show that it is indistinguishable so far from cho-linephosphotransferase. Taken together, the above considera-tions cast doubt on whether PE synthesis in plants takes placeby the "cytidine" pathway, and gives good reason to considerdecarboxylation of PS as the predominant pathway.Kennedy and Weiss (20) concluded that the cholinephospho-

transferase and ethanolaminephosphotransferase were distinctenzymes on the grounds of different lability on lyophilization.In our experience both of the enzymes from rat liver are stableto lyophilization as are the enzymes from spinach leaf. On the

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PHOSPHATIDYLETHANO

other hand, the two enzymes from rat liver can be distinguishedby their difference in susceptibility to inhibition by acyl CoAesters, PC synthesis being susceptible and PE synthesis resistant(11). In discussing their observation that CDP-choline inhibitedethanolaminephosphotransferase in microsomal preparationsfrom rat brain, Ansell and Metcalf (2) listed as a possibilitythat cholinephosphotransferase could catalyze the transfer ofphosphorylethanolamine from CDP-ethanolamine to diglycer-ide. If the final step of PE and PC synthesis were catalyzed bya single enzyme, the relative amounts synthesized would be de-pendent on the availability of CDP-choline and CDP-ethanol-amine, an unusual regulatory mechanism, and the Km valuesof the two substrates (20 ,uM for CDP-ethanolamine, and 10,uLM for CDP-choline [13]).

Experiments with intact tissue may show whether PE is madeby way of PS or by the cytidyl pathway. Willemot and Boll(37) showed that excised tomato roots metabolized "4C-serinewith the production of "4C-PC and PE and poorly labeled PS.They concluded that PS was converted to PE by decarboxyla-tion and that PC was formed from PE by successive methyla-tion reactions. This conclusion is supported by results of Lerchand Stegemann (22) who found that many plants contain freeethanolamine, but that no conversion of serine to free ethanol-amine could be detected. One difficulty in accepting the conclu-sion of Willemot and Boll (serine -- PS -* PE -e PC) is thatpmb and diethylethanolamine inhibit the formation of PS andPE while PC is still synthesized from "4C-serine. Perhaps thereis an alternative pathway to choline. Experiments with radio-active phosphate also bear on the synthesis of PE in plants.Singh and Privett (32) found that maturing soybeans incor-porated radioactive phosphate into lipids and the maximumpercentages in PS was after 30 min incubation. PE did notreach a maximum until 4 hr. During germination of the mungbean, PS was most rapidly labeled in the hypocotyls and radi-cles with increasing proportions in PE as the incubation pro-ceeded (18). This result is also consistent with the conversionof PS to PE by decarboxylation. In one other case of phosphatelabeling, radioactivity in PS was negligible even though PE be-came substantially labeled (6). If PE is normally made in vivoby decarboxylation of PS, we might expect that free ethanol-amine would be a poor precursor of PE. In fact the oppositewas true (37). It is possible that ethanolamine was introducedby an exchange reaction (35) rather than de novo synthesis.

Returning to studies at the enzyme level, Marshall and Kates(25) have reported the decarboxylation of PS by a particulatefraction from spinach leaves. Thus the enzymic capability ispresent in these leaves just as it is in bacteria (28).

In conclusion, PE can be synthesized by enzyme prepara-tions from leaves either by decarboxylation of PS or by transferof ethanolamine phosphate from CDP-ethanolamine. The pre-dominant pathway in vivo is still unknown.

LITERATURE CITED

1. ALLEN-, C., F. GOOD, H. F. DAVIS, P. CHISUM, AND S. D. FOWLER. 1966.Methodology for the separation of plant lipids and application to spinachleaf and chloroplast lamellae. J. Amer. Oil Chem. Soc. 43: 223-231.

2. AN5SELL, G. B. AND R. F. METCALF. 1971. Studies on the CDP-ethanolamine-1, 2-diglyceride ethanolamine phosphotransferase of rat brain. J. Neuro-chem. 18: 647-665.

3. ARNON-, D. I. 1949. Copper enzymes in isolated chloroplasts. Polyphenoloxidasein Beta vulgaris. Plant Physiol. 24: 1-15.

4. BEN-SON, A. A. 1963. Neutron activation chromatography of phosphoruscompounds. Methods Enzymol. 6: 881-888.

5. BENSON, A. A., J. F. G. NI. WINTERMANS, AND R. WVISER. 1959. Chloroplastlipids as carbohydrate reservoirs. Plant Physiol. 34: 315-317.

6. BIELESKI, R. L. 1972. Turno-er of phospholipids in normal and phosphorusdeficient Spirodela. Plant Physiol. 49: 740-745.

7. BLIGH, E. G. AN-D W. J. DYER. 1959. A rapid method of total lipid ex-traction and purification. Can. J. Biochem. Physiol. 37: 911-917.

ILAMINE BIOSYNTHESIS 175

8. BORKENHAGEN, L. F. AND E. P. KENNEDY. 1957. The enzymatic synthesis ofcytidine diphosphate choline. J. Biol. Chem. 227: 951-962.

9. BORKENHAGEN, L. F., E. P. KENNEDY, AND L. FIELDING. 1961. Enzymaticformation and decarboxylation of phosphatidylserine. J. Biol. Chem. 236:PC28-30.

10. CHOJNACKI, T. A-ND R. F. METCALF. 1966. Enzymatic synthesis of labeledcytidine-5'-diphosphate ethanolamine. Nature 210: 947-948.

11. DE KRL-YFF, B., L. M. G. vAN GOLDE, AND L. L. M. vAN DEENEN-. 1970.Utilization of diacylglycerol species by choline phosphotransferase, ethanol-aminephosphotransferase and diacylglycerol acyltransferase in rat livermicrosomes. Biochim. Biophys. Acta 210: 425-435.

12. DENN--IS, E. A. AND E. P. KENNEDY. 1972. Intracellular sites of lipid synthesisand the biogenisis of mitochondrion. J. Lipid Res. 13: 263-267.

13. DEVOR, K. A. AND J. B. MMUDD. 1971. Biosynthesis of phosphatidylcholine byenzyme preparations from spinach leaves. J. Lipid Res. 12: 403-411.

14. DITTMER, J. C. AND M. A. WELLS. 1969. Quantitative and qualitativ-e analysisof lipid components. Methods Enzymol. 14: 482-530.

15. DOWD, J. E. AND D. S. RIGGS. 1965. A comparison of estimates of 'Michaelis-Menten kinetic constants from various linear transformations. J. Biol.Chem. 240: 863-869.

16. GALLIARD, T. 1968. Aspects of lipid metabolism in higher plants I. Identifica-tion and quantitative determination of the lipids in potato tubers. Phyto-chemistry 7: 1907-1914.

17. JOHNSON, K. D. AND M. KENDE. 1971. Hormonal control of lecithin synthesisin barley aleurone cells: regulation of the CDP-choline pathway bygibberellin. Proc. Nat Acad. Sci. U.S.A. 68: 2674-2677.

18. KATAYAMA, MI. AND S. FUNAHASHI. 1969. Metabolic pattern of phospholipidsduring germination of mung bean, Phaseolus radiatus var. typicus. I.The incorporation of 32P-orthophosphate into phospholipids. J. Biochem.66: 479-485.

19. KEN`NEDY, E. P. 1956. The synthesis of cytidine diphosphate choline, cytidinediphosphate ethanolamine and related compounds. J. Biol. Chem. 222:185-192.

20. KENNEDY, E. P. AND S. B. WEIss. 1956. The function of cytidine coenzymesin the biosynthesis of phospholipids. J. Biol. Chem. 222: 193-214.

21. LEECH, R. M., M. G. RL-MSBY, W. W. THOMSON, W. CROSBY, AND P. WOOD.1971. Lipid changes during plastid differentiation in developing maizeleaves. Proceedings IInd Int. Congr. Photosynth. pp. 2479-2488.

22. LERCH, B. AND MI. STEGEMANN. 1966. Xthanolamin, ein verbreiteter Inhaltstoffder Bliitter hoherer Pflanzen. Zeit. Naturforsch. 216: 216-218.

23. LORD, J. M., T. KAGAWA AND H. BEEVERS. 1972. Intracellular distribution ofenzymes of the cytidine diphosphate choline pathway in castor bean endo-sperm. Proc. Nat. Acad. Sci. U.S.A. 69: 2429-2432.

24. LOWRY, 0. 'M., N. J. RoSEBROrGH, A. L. FARR, A-ND R. J. RANDALL. 1951.Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265-275.

25. MARSHALL, M. 0. AND M. KATES. 1973. Biosynthesis of phosphatidyl ethanol-amine and phosphatidylcholine in spinach leaves. FEBS Lett. 31: 199-202.

26. MORR£, D. J., S. NYQuIST, AND E. RIVERA. 1970. Lecithin biosynthetic en-zymes of onion stem and the distribution of phosphorylcholine cytidyltransferase among cell fractions. Plant Physiol. 45: 800-804.

27. ONGUN, A., W. W. THO11SON, AND J. B. MUDD. 1968. Lipid composition ofchloroplasts isolated by aqueous and non-aqueous techniques. J. LipidRes. 9: 409-415.

28. RAETZ, C. R. M. AND E. P. KENNEDY. 1972. The association of phos-phatidylserine synthetase with ribosomes in extracts of Escherichia coli.J. Biol. Chem. 247: 2008-2014.

29. RAMASAMRA, T. AND L. WETTER. 1957. Choline kinase of rapeseed (Brassicacampestris L.). Can. J. Biochem. Physiol. 35: 853-863.

30. ROUGHAN, P. G. 1970. Turnover of the glycerolipids of pumpkin leaves. Theimportance of phosphatidyl choline. Biochem. J. 117: 1-8.

31. SCHNEIDER, W. C. 1969. Enzymatic preparation of labeled phosphorylcholine,phosphorylethanolamine, CDP-choline, deoxy CDP-choline, CDP-ethanol-amine and deoxy CDP-ethanolamine. Methods Enzymol. 14: 684690.

32. SINGH, 'M. AND 0. S. PRIVETT. 1969. Incorporation of 83P in soybean phos-phatides. Biochim. Biophys. Acta 202: 20-202.

33. TANAKA, K., N. E. TOLBERT, AND A. F. GOHLKE. 1966. Choline kinase andphosphoryl choline phosphatase in plants. Plant Physiol. 41: 307-312.

34. TEVINI, M. 1971. Der Einfluss von Phosphat-Mangel-Ernahrung auf dieSynthese der Phospho- und Glycolipide bei Impatiens. Zeit. Pflanzenphys.66: 64-72.

35. VANDOR, S. L. AND K. E. RICHARDSON. 1968. Incorporation of ethanolamine-1,2-14C into plant microsomal phospholipids. Can. J. Biochem. 46: 1309-1315.

36. WEBER, E. J. 1970. Lipids of maturing grain corn (Zea mays L.): II. Changesin polar lipids. J. Amer. Oil Chem. Soc. 47: 340-343.

37. WILLEMOT, C. AND W. G. BOLL. 1967. Incorporation of serine-14C andethanolamine-14C into nitrogen containing phosphatides and effects ofmedium containing ethanolamine on phosphatide biosynthesis in excisedtomato roots. Can. J. Bot. 45: 1863-1876.

38. WITTENBERG, J. AND A. KORNBERG. 1953. Choline phosplhokinase. J. Biol.Chem. 202: 431-444.

Plant Physiol. Vol. 53, 1974

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