Studies on the biosynthesis of cyclopropane fatty acids in Escherichia coli

B&chimica et Biophysics Acta, 348 (1974) 63-75 6 Elscvicr ScientificPublishing Company, Amsterdam - Printed in The Netherlands BBA &$z8

STUDfiEi3ON THE 310SYNTHESIS OF CYCLOPROPANEFATTY ACIDS IN ESCHERICHIA COLI

JOHN E. CRONAN, Jr, WILLIAM D. NUNN and JOHN G. BATCHELOR Lkparttmmrof Mokwdar Biopkysics and Biockgrn~~t~y,Yale University, New Haven, Corm. 065x0 (U.S.A.) (Received November nth, 1973)

SUMMARY

By use of various mutants either defective in unsaturated fatty acid biosynthesis or altered in the level of S-adenosylmethionine we have tested the effects of changes in the intracellular concentrations of these substrates on in vivo cyclopropane fatty acid synthesis. A I .5-fold change in un~turated fatty acid content produced an equivalent change in cyclopropane fatty acid content. A zo-fold difference in S-adenosylmethionine content had little or no effect on cyclopropane fatty acid formation. Studies with unsaturated fatty acid auxotrophs showed that unsaturated fatty acid synthesis is not required for cyclopropane fatty acid synthesis. Other studies demonstrated that cyclopropane fatty acid content had little effect on phospholipid stability, Finally, we have shown by three methods @s-liquid ~hromato~aphy, proton and “C nuclear magnetic resonance spectra) that the cyclopropane ring of these Escherichia coli &&jS has the cis configuration.

INTRODUCTION

Most of our knowledge of the biosynthesis of the cyclopropane fatty acids of Escherichiu cob is due to the work of Law and co-workers [I, 31. The scheme outlined by these workers involves addition of a methylene group from the methyl carbon of S-adenosylmethionine to the double bond of the unsaturated fatty acid moeity of a ~ospho~p~ molecule. Most of these conclusions were based on in vitro experiments using a cyclopropane fatty acid-s~~es~~ system from CZo~tri~~ ~~t~rj~~ [z]. However, in the ensuing years, the isolation of various E. coli mutants has made it possible to examine the mechanism and control of cyclopropane fatty acids synthesis in vivo. In this paper, we report in vivo experiments probing the regulation of synthesis the function, and the structure of these acids in E. coli. MATERIALS AND METHODS

Bacterid strains The bacterial strains used are all derivatives of E. coli K12. Their relevant genotypes and derivations are given in Table I. Strain CY78 was obtained by plating

64

X340 on a broth plate spread with IO” T6 phage and picking a nonmucoid resistant colony. Strain CY14 was constructed by transduction of strain CY78 to pro+ with P, phage grown on strain K1o6o. The pro+ recombinants were then scored for the fadE phenotype. About 5 % of thepro+ recombinants werefad- (unable to grow on medium containing oleate as sole carbon source.) A starved culture of CY 14, one of the pro+ fadE recombinants, was assayed [4] for /?-oxidation and had < 0.5 % of the normal activity. Strain CY35 was constructed by mating an F- phenocopy culture (generated by growth in broth for 20 h at 30 “C) of UCIog8 with strain CY14. After mating for TABLE I BACTERIALSTRAINS The genetic symbols used are those of Taylor and Trotter [28]. The allele numbers are those of the Coli Genetic Stock Center, Yale University. Strain

Sex

Relevant characters

UC1 UCIog3 WNI

F+ F+ F+

prototroph, str’ fabAz, Cvc-, strR fab+, Cvc-, strR

CY33 CY35

F+ F+

CYI4 CY78 x340 K1o6o KIZ(L) RG62 RG73 RGIo~ RGIOO CY2

HfrC HfrC HfrC FF+ F+ F+ F+ F+ F-

Source

Cronan, et al. [6] Cronan, et al. [6] fab+ transductant of UC1098 by PI (UCr) fabAz, metBr, fadE62, tsxR, Cvc-, strR this paper fab+, metBr, fadE62, tsxR, Cvc-, strR fab + transductant of CY33 by PI (UCr) fadE62, metBr, tsxR, rel-r this paper proBz8, metB1, rel-r, tsxR this paper R. Curtiss via B. Bachmann proBz8, metBr, rel-I P. Overath fabB5, fadE62 R. C. Greene [23, 241 prototroph R. C. Greene [23, 241 metK84 R. C. Greene [23, 241 metK85 R. C. Greene [23, 241 metK86 R. C. Greene [22] metJ Cronan and Batchelor 1161 aceFro, gltAg

15 min at 34 “C, the mating mixture was diluted Ioo-fold into broth containing streptomycin (200 pgglml), then agitated on a vortex mixer for 5 min. The diluted mating mixture was incubated overnight at 26 “C (for phenotypic expression of tsxR recombinants). The resulting stationary phase cells were plated at 30 “C on a broth plate spread with 10~~T6 phage and streptomycin, Nonmucoid tsa?, strR recombinants were then scored for growth on medium containing oleate as sole carbon source. About 50 % of the tsxR strR recombinants were also fadE -. Strain CY35 is one such recombinant which also obtained the metBr allele from CY 14. The methods and media for the genetic procedures outlined above are given in earlier papers [5,6].

Media and growth conditions Unless otherwise noted, the minimal salts medium used was medium E [7]. The low sulfate medium used in the experiments of Table IV and Fig. 2 was the medium of Davis and Mingioli [8] with each sulfate salt replaced by the appropriate chloride salt. Sulfate was then added at IO-~ M. The low-phosphate medium used in the 32Pilabeling experiments has been described previously [g]. Media were supplemented with

65

required growth factors as previously described [5]. Dextrose was the carbon source used except when noted. Fatty acids were added as potassium salts and were solubihzed with Brij 58 detergent [5}. Bacterial cultures were incubated in gyromtory water bath shakers with vigorous aeration. The growth temperature was 37 “C unless otherwise noted. The fatty acids used were > 99 % pure by gas-liquid chromatography. Fatty acid extraction and methylation The cellular phospholipids were extracted as previously described [IO], then saponified with IO % KOH in 50 % aqueous methanol for I h at 70 “C. After acidification with 6 M HCl, the fatty acids were extracted into ether. The isolated fatty acids were converted to their methyl esters by treatment with diazomethane [II] or by transesterification in the presence of 2,zdimethoxypropane [IZ]. These esterification methods were chosen since strong acid catalysts have often been reported to cleave cyclopropane rings [13]. The resulting methyl esters were dissolved in toluene and analyzed by gas-liquid chromatography. Gas-liquid chromatography Gas-liquid chromatography was usually performed on an Apiezon L column as previously described [14]. Identification of the steric configuration of the fatty acid cyclopropane ring was performed on a 1/8 inch x 6 ft column of I 5 % diethyleneglycol succinate on 6o/80 Gas Chrom P. This column was run at 180 “C. Phospholipid analysis The phospholipids were resolved into component species and quantitated as previously described [9]. NMR spectra Proton NMR spectra were obtained on a Joelco Minimar IOO spectrometer. Tetramethylsilane was used as the reference compound. 13C NMR spectra were obtained on a Bruker 2rK gauss spectrometer converted for Fourier transform operation. Hexamethyldisilane was added to samples as an internal reference. Hexafluorobenzene in a 5-mm tube mounted coaxially provided a field frequency lock. Data sets of 4 K or 8 K were used providing chemical shift measurements accurate to f 0.1 ppm or f 0.05 ppm, respectively. The nonenriched cis and trMs cyclopropane fatty acid standards were neat solutions for spectroscopy. The 13C-enriched E. coli methyl esters were dissolved in chloroform to approximately 15 mg/ml final concentration. The assignment of the resonances has been described [r51. Materials E. coli CY2 fatty acids enriched with 13C in the odd carbons were obtained as previously described [ 16]. The cyclopropane fatty acid fraction obtained by preparative gas chromatography [16]. The unsaturated fatty acid standards were purchased from the Hormel Institute, Austin, Minn. The cyclopropane fatty acid standards were the products of Supelco, Inc., Bellefonte, Penn. and Analabs, North Haven, Conn. The chromatographic materials were also obtained from these companies.

66 RESULTS

Is cyclopropane fatty acid synthesis dependent on the supply of either of the substrates (unsaturated fatty acid or S-adenosylmethionine) of the reaction? Our experimental design was to alter the concentration of these substrates in vivo and then examine the effects on cyclopropane fatty acid synthesis. Our first experiment involved the alteration of the content of phospholipids containing unsaturated fatty acid content in growing cells*. Lack of regulation of cyclopropane fatty acidsynthesis by unsaturatedfatty acid content Does E. coli convert a set number of molecules of unsaturated fatty acid to

cyclopropane fatty acid or is a set proportion of the unsaturated fatty acid converted to cyclopropane fatty acid? We examined these alternatives by changing the cellular level of palmitoleic acid using mutants defective in unsaturated fatty acid synthesis. Both mutations used were originally isolated in strain UC1098 These mutations are the fabAt lesion [6,19] and the Cvc- phenotype [14]. The fubA.2 allele produces a /?-hydroxydecanoylthioester dehydrase of abnormal thermolability [IS]. This enzyme is responsible for the introduction of the double bond moiety of unsaturated fatty acid molecules [zo]. Thus, UC1098 requires an appropriate unsaturated fatty acid for growth above 36 “C (ref. 19). However, at 30 “C, UC1098 grows normally although the unsaturated fatty acid content of these cells is only about 60% of the normal amount. The unsaturated fatty acid of these cells is almost entirely palmitoleic acid; very little cis-vaccenic acid is formed [x4]. This phenotype was shown to be unrelated to the f&A gene and appears due to the lack of an enzyme involved in elongation of palmitoleic acid to cis-vaccenic acid [14] (the Cvc- phenotype). FubA+ derivatives of UC1098 synthesize normal amounts of unsaturated fatty acid but virtually all this unsaturated fatty acid is palmitoleic acid. Thus, the palmitoleic acid content of such fubA+ Cvc- cells is higher than normal [14]. Therefore, we are able to manipulate the pahnitoleic acid content of otherwise normal cells and examine the effect of these manipulations on cyclopropane fatty acid synthesis. Table II shows the fatty acid compositions of stationary phase cultures of UC1o98, of WNI (a fabA+, Cvc- transductant of UC1098), and of their wild type parent, UCI. As shown, about 95 % of the cellular palmitoleic acid was converted to cyclopropane fatty acid even though the palmitoleic acid content of the phospholipid varied about 1.5-fold. It was thought that the kinetics of cyclopropane fatty acid formation during growth might be altered in either UC1o98 or WNI or in both strains. However, Fig. I shows that the time courses of cyclopropane fatty acid formation in these strains are similar in relation to growth. This experiment was performed with strains CY33 and CY35. CY33 is a metB_, fadE-, derivative of UC1098, CY35 is a fubA+ transductant of CY33. The metB- marker allows the quantitation of cyclol It should be noted that two species of cyclopropane fatty acid are present in the phospholipids of E. coli, cis-g,ro-methylene hexadecanoic acid and lactobacillic @is-r I,IZ-methyleneoctadecanoic) acid [17]. These acids are the cyclopropane analogs of pahnitoleic (cis-g-hexadecenoic) acid and cisvacc&c (cis-1 r-octadecenoic) acid, the major species of unsaturated fatty acid found in E. coli [I 7, I 81. In our experiments, we have usually measured only the synthesis of the Cl7 cyclopropane fatty acid Lactobacillic acid was not usually measured because we have used strains of E. coli deficient in the synthesis of cis-vaccenic acid [14] and thus deficient in the synthesis of lactobacillic acid. Therefore, except when noted, the term, cyclopropane fatty acid, will denote the C17 acid.

TABLE II PERCENTAGE (w/w) FATTY ACID COMPOSITION OF VARIOUS STRAINS Fatty acid ester

Los Laurie cis-7-tetradecenoic* Myristic Unidentified Pahnitoleic Palmitic Methylene hexadecanoic cis-vaccenic Stearic Lactobacillic Palmitoleic+ 17-CPA** CPA** ____ Palmitoleic+CFA

trtr 3.3 o-4 29.0 30.2

Stationary

tr tr 3.9 I.3 0.6 32.5 30.5

Log tr tr I.6 tr

31.0 0.4 3.6

IS.5

31.1

31.1

45.0 44.4 4.3 2.7 2.0 0 49.3

6.8

98.1

9.1

2.1

UC1098

WNr

Strain: UC1

14.7 1.0

Stationary

Log

Stationary

5.7 I.3 I.5 42.8 43.7 3.2 0.9 0.8 45.2

tr tr 7.9 0.7 32.5 54. I I.9 I.2 I.7 0 34.4

tr tr 10.7 1.0 1.0 54.9 28.2 2.0 2.2 tr 29.2

96.7

5.5

96.6

0 0

* For identification see Batchelor and Cronan 1291. * CFA denotes c%-g,Io- methylene hexadecanoic acid. *** tr denotes only a trace of the acid was present.

propane fatty acid synthesis by a simple dual isotope label method [zI]. As shown in Fig. I, CY35, the strain with the greater content of unsaturated fatty acid, formed its increased content of cyclopropane fatty acid by continuing the maximum rate of cyclopropane fatty acid synthesis for an extended time rather than by changing either the maximum rate or the time of onset of cyclopropane fatty acid formation. Jt should be noted that strains UCl,UC1q8,WNr,CY33, and CY35 all grow identically at 30 “C. Luck of reg~~~ion of cyclopro~ fufty acid synthesis by S~~no~yl~e~~~~~ content Since the amount of cyclopropane fatty acid formed appears not to be limited by the supply of unsaturated fatty acid, we undertook experiments to test if the amount of the other substrate, S-adenosylmethionine, had a direct effect on the amount of cyclopropane fatty acid. Greene et al. [22-241, have isolated several types of E. coli K12 mutants with altered intracellular concentrations of S-adenosyhnethionine. Dr Greene has provided us with these strains and we have examined cyclopropane fatty acid formation during growth. Fig. 2 shows typical kinetics for cyclopropane fatty acid formation and growth of these strains. These strains grow and make cyclopropane fatty acid with kinetics very similar to those of their wild type parent. Table JJJ gives the cyclopropane fatty acid and S-adenosylmethionine contents of these strains. The data indicate that the intracellular S-adenosylmethionine content has little or no effect on the rate and/or extent of cyclopropane fatty acid formation. Turnover of ~elo~ro~e fatty acid ~on~~in~gphosph~~~i~ Five years ago, the senior author observed that cyclopropane fatty acid content correlated with the metabolic stability of the various phospholipid species of E. coli [21]. Hence, he suggested that the presence of a cyclopropane fatty acid molecule in a

68

Y) C 5 =, 100 CT .c f b ;

70 30

30

IO

Hour Fig. I. Kinetics of cyclopropane fatty acid synthesis in strains CY33 and CY35. The strains were grown on medium E supplemented with glucose (0.4 %) and 25 fig/ml of the standard L-amino acids with the exception of methionine. The medium also contained a2Pi (13.5 mCi/mole) and L-[Me-14C]methionine (0.375 Ci/mole, 50 fig/ml final concentration). The medium was inoculated with approx. IO’ cells/ml and incubated at 30 “C. At various time intervals, samples were taken and thejratio of cyclopropane fatty acid molecules to phospholipid molecules was determined by the radioactive dual labeling procedure described earlier [zI]. At the conclusion of the experiment samples of each culture were plated on the appropriate medium to score for fub+, and met+ revertants. Revertants of any of the markers were present in < IO-~ of the viable cells plated. Symbols: the open symbols signify CY33; the solid symbols signify CY35. Square symbols denote growth measurement; the circles denote ratios of cyclopropane fatty acid to phospholipid.

o,L~sL

1

1

4

1

’

6

’

’

’

’

io

Hours6

’

’

IL

%t”

Fig. 2. Kinetics of cyclopropane fatty acid formation and growth in S-adenosylmethionine content altered strains. A culture of strain RGIo~ was grown in glucose supplemented low-sulfate medium. Samples were taken at various intervals and the amount of seventeen carbon cyclopropane fatty acid quantitated by gas-liquid chromatography. Symbols: Open circles, turbidity; solid circles, cyclopropane fatty acid content. Strains KIZF +, RG62, RG73, and RGIOO displayed very similar kinetics of growth and cyclopropane fatty acid formation. Some of these data are given in Table IV.

TABLE XIX The bacteria were grown on glucose-low s&fate (IO-~M) minimal medium and were chocked for ethionine resistance [zzz at the conch&on of the experiment. Bacterial strain

Wild type RG62(metK-) RG73(metK-) RG 1 oo(meff -)** RGrog(metK-)

Intracelhdar S-adenosylmethionine concentration (nmoles/g cells)*

54.6 f 4.2 l 0.4 20.5 f 1.4 8.7 f 0.6

21.0

Cr7 C16:r+C17

at culture age-

2-4

6.5

12.0

0.32 0.24 0.56 0.37 0.30

0.65 o-33 0.80 0.78 0.52

0.88 0.33 0.80 o-93 0.85

* These values are the data of Greene, et al. [24]. ** Strain RG1oo has about twice the normal S-adenosylmethionine Greene, personal commu~i~tion). c** See Fig. 2 for correlation of culture age with growth phase.

(h) of: 24.0 0.93 0.72

0.95 0.96 0.92

concentration (Dr R. C.

phospholipid might somehow stabilize that phospholipid molecule during growth. With the isolation of unsaturated fatty acid auxotrophs, we were now equipped to test this suggestion. An unsaturated fatty acid auxotroph was grown on either a cyclopropane fatty acid or the homologous ~~~rat~ fatty acid and the phospholip~s were labeled with “Pi. The 32Pi was then removed from these cultures and the stability of the various phospholipid species examined during further growth. As shown in Fig. 3, the stabilities of each of the phospholipid species in either culture were quite similar. Therefore, the amount of cyclopropane fatty acid in a given phospholipid molecule does not directly affect the stability of that molecule during growth. Observations on*cyciopropanefarty acid synthesis and structure The mechanism of cyclopropane fatty acid synthesis proposed by Law and coworkers [I-31 would not require unsaturated fatty acid synthesis in order for cyclopropane fatty acid synthesis to occur in vivo. However, an alternative hypothesis put forth by Thorne 1257 suggested that concurrent unsaturated fatty acid synthesis must occur in order for cyclopropane fatty acid to be synthesized. We have tested these alternatives in vivo and find that cyclopropane fatty acid synthesis does not require concurrent unsaturated fatty acid synthesis. This experiment was done using strain K1o6o which contains mutations in both unsaturated fatty acid synthesis (j2M-) and j&oxidation (fadE_). This strain was grown to early log phase with palmitoleic acid as the required supplement. The supplement was then switched to oleic acid and the culture grown to stationary phase. Since no synthesis of palmitoleic acid containing phospholipids conld occur (due to the fubB mutation) at the time of cyclopropane fatty acid formation (entry into stationary phase), the only source of pahnitoleic acid was the unsaturated fatty acid moeities of the phospholipid (virtually all the unsaturated fatty acid in growing cells is found in phospholipid [IO, I 71) Since the phospholipid bound pahnitokzic acid was converted to cyclopropane fatty acid normally (Table N this indicated that ~0~~0~~s containing unsaturated fatty acid were indeed the substrate (or the source of the substrate) for cyclopropane fatty acid synthesis. A similar experiment in which strain CY33 was grown to log phase at 30 “C, shifted to 42 “C in the presence of oleate, then grown to stationary phase gave similar results.

60 htinutes

of

chase

120

160

240

Minutes of chase

Fig. 3. Turnover of the phospholipids of strain UC1098. Two cultures of UC1098 were grown at 42 “C. One culture was supplemented with potassium palmitoleate (left panel). The other culture was supplemented with potassium cis-9,ro-methylenehexadecanoate (right panel). The medium used was the low-phosphate medium supplemented with fatty acids at 80 ag/ml, Brij 58 detergent and glucose. After several generations of growth the cultures were labeled with szP, (20 ,uCi/ml) for 60 min during early log phase. The cultures were then centrifuged, washed once, and resuspended in the same medium containing SomM potassium phosphate buffer (pH 7.2). These cultures were incubated at 42 “C and samples were removed at the times indicated. The phospholipids were extracted and analyzed as described in Methods. Less than IO-~ of the viable cells in either culture were f&4+. The doubling times of the cultures were 75 min and 105 min for palmitoleate and cis-g,ro-methylenehexadecanoate, respectively. Symbols are: open circles, total phospholipid; PE (phosphatidylethanolamine), solid squares; PG (phosphatidylglycerol), solid circles; CL (cardiolipin), open triangles.

Configuration of the cyclopropane ring Crystallographic evidence [26] demonstrated that lactobacillic acid isolated from lactobacilli has the cis configuration. It has been assumed that the cyclopropane fatty acids of E. coli also have cis cyclopropane rings although the evidence in this organism was indirect [17]. We have accumulated three types of direct evidence that indicate the configuration of both the E. coli cyclopropane fatty acids is cis. The methods used were gasliquid chromatography, 13C NMR and proton NMR. The chromatographic data given in Table V indicate that < I % of either cyclopropane fatty acid found in E. coli has the tram configuration. This is confirmed by the 13C and proton NMR spectra of the seventeen carbon acid. 13C NMR clearly distinguishes between cis- and trans-cyclopropane fatty acids [r5]. As shown in Fig. dab, the natural abundance spectra of authentic transand cis-cyclopropane fatty acids differ markedly in the resonance of the carbons a to the ring. In a truns-cyclopropane fatty acid, these carbons resonate 36.9 ppm downfield from hexamethyldisilane, whereas these resonances are found 5.8 ppm further

TABLE IV Strain K1o6o was grown at 34 “C in medium E supplemented with glycerol (0.4”%), easein hydrolysate (I “%)Brij 58 (0.2 yd and potassium pahnitoleate (ioo rg/ml). A 3oo-ml culture was grown to early log phase (approx. 4 * IO’ cells/ml), then centrifuged. The cells were washed with medium E and resuspended in medium E at the original concentration. Then 250 ml of the culture was harvested and the lipids of the cell pellet were immediately extracted. The remaining 50 ml of cells in medium E was supplemented exactly as given above except potassium oleate (200 pg/ml) instead of pahnitoleate was the fatty acid added to support growth. The culture was then grown 16 h to late stationary phase, harvested, and the lipids were extracted. Upon lipid extraction a known amount of pentadecanoic acid was added to serve as an internal standard for quantitation of mass. The isolated phospholipids were saponified, esterified and analyzed by gas-liquid chromatography as described in the text. The culture was plated on the appropriate media to test for reversion of thefobB and the fadE characters of K1o6o. Less than IO-’ of the cells in the stationary phase culture was either@+ or fad+ or both. CFA, cis-9,ro-methylene hexadecanoic acid. Growth phase at harvest

Log phase (4 * ro* cells/ml) Stationary phase (2 . 10’ cells/ml)

Supplement

Fatty acid content (pg/5o ml culture) C16:1 C16:r +C17 Cr7

Percent CFA Cr7 C16:1+C17

palmitoleate palmitoleate to 4 * Ios cells/ml then oleate

45.6

11.9

57.5

20.7

2.7

50.8

53.5

94.9

TABLE V GAS CHROMATOGRAPHIC

ANALYSIS

OF E. COLZ CYCLOPROPANE

FATTY ACIDS

The methyl esters of the fatty acids were obtained and analyzed by gas chromatography as described in Materials and Methods. The retention times (accurate to about 5 “A are given relative to that of palm&ate (=~.oo). The Cl9 acids were the con8gurational isomers of g,ro-methyleneocta decanoic acid. The 9,10- and I r,r2-cis-methylene octadecanoates have essentially identical gas chromatographic behavior [30]. Since the acid was not commercially available, the retention time for the transC1, acid (as the methyl ester) was calculated from the data for the Cl9 compounds by standard procedures [31]. A similar calculation for the cis isomer gave a value of 1.61. The signal to noise ratio and the relative peak areas indicate that
Number of carbon atoms

ConiIguration

‘16:o

Synthetic Calculated Synthetic Synthetic E. coli B E. coli B E. coli K12 E. coli K12

17 17 19 19 17 19 17 19

cis trans cis tram

1.61 I.34 2.82 2.34 I .64 2.88 I .62 2.83

upfield for a cis acid. This difference is attributed to a steric interaction between the hydrogens on the carbons a to a cis cyclopropane ring. The substituted ring carbon resonance of trans-cyclopropane ring is found 3. I ppm downfield from the equivalent resonance of the cis isomer. Since we cani distinguish these two isomers, we have obtained the “C NMR spectrum of C,, cydopropane fatty acid from E. coli. In Fig. qc the spectrum of a sample enriched with ‘PC in the odd carbons is given. The spectrum is clearly that of a cis- rather than of a kanscyclopropane fatty acid.

a COOH

ppm downfield from HMDS Fig. 4. rJC NMR spectra of (a) frans-9, ro-methyleneoctadecanoic acid, (b) cis-9, ro-methyleneoctadecanoic acid and (c) methyl ester of the seventeen carbon cyclopropane fatty acid isolated from E. coli K12 strain CY2. Spectra (a) and (b) are of natural abundance material while spectrum (c) is of material 63 % enriched with W in the odd-numbered carbons. The enriched carbons are denoted by an asterisk. The peak marked a is the resonance of the substituted ring carbon of the cyclopropane ring (see Batchelor et al. [IS] for assignment of resonances). The comparison of free acids in panels (a) and (b) with a methyl ester in panel (c) is valid because only the resonance of carbon number 3 (in spectrum c) is affected by the substitution.

Proton NMR shows two characteristic peaks for a cis-cyclopropane fatty acid The peaks are 0.61 ppm downfield and 0.29 ppm upfield from tetramethylsilane, respectively. In contrast, a trans acid has two peaks (at 0.35 and approx. 0.14 ppm) downfield from tetramethylsilane. The spectrum of the cyclopropane fatty acid from E. coli K12 is clearly that of a cis cyclopropane fatty acid (Fig. 5). Therefore, all three methods indicate that the cyclopropane ring of the cyclepropane fatty acids found in E. coli have the cis configuration. [13].

DISCUSSION

We believe we are now able to discard several of the more obvious mechanisms proposed for the control of cyclopropane fatty acid synthesis. We previously showed that cyclopropane fatty acid synthetase activity does not appear to be induced or derepressed at the time when (entry into stationary phase) cyclopropane fatty acid synthesis is at a maximum in vivo [21]. The data presented in this paper indicate that cyclopropane fatty acid synthesis is not controlled by the levels of S-adenosylmethionine or of unsaturated fatty acid in the cell. These data can lead to the following conclusions. First, it seems unlikely that cyclopropane fatty acid molecules have specific cellular roles. We have shown that the amount of cyclopropane fatty acid formed is a

73

J

I

I

3

2

IO

I

I

I

I

I

2

IO

I

pm3 -

I 2.0

._ ._.-

I

I.5

I

I

I

1.0

I

I

0

-0.5

I

PPmQ Fig. 5. Proton NMR spectra of (left panel) cis-g,ro-methyleneoctadecanoic acid (right panel) truns-9, xo-me~ylen~tadec~oic acid, and (bottom panel) methyl esters of the ~~r~ionat~ fatty acids obtained from the phospholipids of E. coli KIZ. The resonances are given in reference to the resonance of tetramethylsilane (TMS). See Christie [131 for assignment of resonances.

direct function of the unsaturated fatty acid content (Fig. z, Table II). Also, Silbert, et al. [27] have shown that the formation of cyclopropane fatty acid is not required for cell viability since unsaturated fatty acid auxotrophs grow (albeit poorly) on trmrs unsaturated fatty acid supplements although no cyclopropane rings are formed. Another unlikely ~~ib~i~ is that cyclopro~e fatty acid fo~a~on is a mechanism to rid the cell of excess activated methyl groups. During exponential growth, S-adenosylmethionine is needed for the synthesis of the methyl moieties of several molecules including nucleic acids. Since most of these molecules are largely synthesized during exponential growth, cells in stationary phase might have excess S-adenosyhnethionine which could be consumed in cyclopropane fatty synthesis. However, this hypothesis would predict that cyclopropane fatty acid synthesis would show a strong dependence on S-adenosylmetbionine content. This dependence was not observed (Table ID) and thus this hypothesis also appears void. Since E. co& is able to grow witbout cyclopropane fatty acid (when the unsaturated fatty acids of the cells have the truns configuration 127J)perhaps some of the cyclopropane fatty acid in normal cells hers the trm configuration and thus could be substituted for by truns unsaturated molecules. This possibility seemed unlikely in view of the previous literature; however, since the previous experiments were indirect and qualitative, we examined the configuration of these acids by three different physical methods. We found the acids to be >gg % cis (Tables V, Figs 4 and 5). We also found

74

that cyclopropane fatty acid content does not affect phospholipid stability (Fig. 3) which probably eliminates another proposed function [21]. Why does E. coli form large amounts of cyclopropane fatty acid only in stationary phase? Is the presence of large numbers of cyclopropane fatty acid molecules detrimental to log phase growth? We examined the log phase rates of two cultures of an unsaturated fatty acid auxotroph. One culture was supplemented with an unsaturated fatty acid. The other culture was supplemented with a cyclopropane fatty acid homologous to the unsaturated fatty acid. Since the cyclopropane fatty acid supplemented culture grew with a rate only 65 % that of the unsaturated fatty acid supplemented culture (Fig. 3), high cyclopropane fatty acid content does appear to impede exponential growth. A previous experiment [27] reported that another unsaturated fatty acid auxotroph grew equally well on an unsaturated fatty acid or its homologous cyclopropane fatty acid. However, the cyclopropane fatty acid sample used by these workers was only 90 % pure and thus the test was probably not sufficiently stringent. Most of the above discussion assumes that cyclopropane fatty acids are synthesized by the mechanism delineated by the in vitro studies of Law et al. [I, 3,321. We have attempted to test some aspects of this mechanism in vivo. We had previously shown that the unsaturated fatty acid moeities of the various phospholipid species of IZ. coZiwere converted to cyclopropane fatty acid at differing rates [21]. Peypoux and Michel [33] later confirmed these findings. We now have shown that unsaturated fatty acid synthesis is not required for cyclopropane fatty acid synthesis (Table IV). These data are consistent with the mechanism of Law and co-workers [I-3, 321 and oppose the mechanism suggested by Thorne [25]. ACKNOWLEDGEMENTS We thank Drs. G. Powell and R. Bell for their advice on the manuscript. We thank Dr R. C. Greene for his kind gift of his mutants and preprints concerning his work. We thank John Clapp and Edward Gelmann for technical assistance. This investigation was supported by U.S. Public Health Service Grant AI 10186 and by National Science Foundation Grant GB-32063. The first two authors were supported by an N.I.H. Career Development Award (I-K~-GM-~o~II) and an N.I.H. Postdoctoral Fellowship (I Fez-A155, 327), respectively. REFERENCES I Law, J. H., Zalkin, H. and Kaneshiro, T. (1963) Biochim. Biophys. Acta 70, 143-151 z Law, J. H. (1967). Specificity of Cell Surfaces, (Davis, B. D. and Warren, L., eds) pp. 87-105, Prentice Hall, Englewood Cliffs, N. J. 3 Zalkin, H., Law, J., and Goldfine, H. (1963) J. Biol. Chem. 238, 1242-1248 4 Klein, K., Steinberg, R., Fiethen, B., and Overath, P. (1971) Eur. J. Biochem. 19, 442-450 5 Cronan, Jr, J. E. and Godson, G. N. (1972) Mol. Gen. Genet. 116, 199-210 6 Cronan, Jr, J. E., Silbert, D. F. and Wulff, D. L. (1972) J. Bacterial. 112, 206-211 7 Vogel, H. J. and Bonner, D. M. (1956) J. Biol. Chem. 218, 97-106 8 Davis, B. D. and Mingioli, E. S. (1950) J. Bacterial. 60, 17-28 9 Tunaitis, E. M. and Cronan, J. E., Jr. (1973) Arch Biochem. Biophys. 155, 420-427 IO Cronan, Jr, J. E. and Wulff, D. L. (1969) Virology 38,241-246 II Schlenk, H. and Gellerman, J. L. (1960) Anal. Chem. 32, 1412-1414 12 Karkas, J. D., Turler, H. and Chargaff, E. (1965) Biochim. Biophys. Acta I I I, 96-109

75 13 Christie, W. W. (1970) Topics in Lipid Chemistry, (Gunstone, F. D., ed.) Vol. I, pp. 1-89,WileyIn&science, N. Y. 14 Gelmann, E. P. and Cronan, Jr. J. B, (19721 f. Bacterial. 112,38I-387 I5 Batch&r, J. 6.. Cki&ley* R. J., Pru&qj& J. H., and Lip&y, B. R. (1973% J. Or& ti, in press 16 Ctonan, Jr, J. E. and Batchelor, J. G. (1973) cham. Phys. Lipids II, 196-202 17 Cronan, Jr, J. E. and Vagelos, P. R. (1972) Biochim. Biophys. Acta 265, 25-60 18 Cronan, Jr, J. E., (1967) Biochim. Biophys. Acta 144, 695-697 19 Cronan, Jr, J. E. and Gelmann, E. P. (1973) J. Biol. C&m. 248, 1188-1195 20 Blocb, K. (1971) The Enzymes (Boyer, P. D., ed.) 3rd ed, Vol. 5, pp. 441-464, Academic Press, N. Y. ZI Cronan, Jr., J. E., (1968) J. Bacterial. 95, 2054-2061 22 Su, C. H. and Greene, R. C. (1971) Proc. Natl. Acad. Sci. U. S. 68,367-371 23 Greene, R. C., Su, C-H., and Holloway, C. T. (1970) B&hem. Biophys. Res. Commun. 38, I rzo-1126 24 Greene, R. C., Hunter, J. S. V., and Coch, E. S. (1973) J. Bacterial. 115, 57-67 z5 Thorne, K. J. (1964) Biochim. Biophys. Acta 83, 35o-353 26 Craven, B. and Jeffrey, G. A. (1960) J. Am. Chem. Sot. 82,3858-3861 27 Silbert, D. F., Ruth, F. and Vagelos, P. R. (1968) J. Bacterial. 95, 1658-1665 28 Taylor, A. L. and Trotter, C. D. (1972) Bacterial. Rev. 36, 504-524 29 Batchelor, J. G. and Cronan, Jr, J. E. (1973) Biochem. Biophys, Res. Commun. 52,x374-1381 30 Christie, W. W., Gunstone, F. I)., Ismail, I. A. and Wade, L. (1968). Chem. Phys. Lipids 2, 1g6202 31 Ackman,R. 0. (1969) Methods Enzymol. 14, 329-381 32 Thomas, J. and Law, J. H. (1966) J. Biol. Chem. 241,5013-5018 33 Peypoux, F. and Michel, G. (1970) Biochim. Biophys. Acta 218,453-462