7YBS - .~4av 19,~;3

older tissues are generall 3 more resistant to cyanide than younger tissues: it is well established that senescence is associated with an increased production of free radi- cals in cell membranes 2°,

Finally, such a process could also take place in any cell membrane that contains both free unsaturated fatty acids and quinones. It could well operate in plastid membranes. In this context, it is interesting to note that there is a recent report 2~ on the occurrence of a respiratory chain in the chloroplasts (chlororespiration). Curiously enough, the oxidases involved in chlororespiration vary from one organism to another and appear to be analogous to the oxidase of the alternative pathway of mitochondrial respiration. Would this not be the operation of a sequence of reactions analogous to those of Fig. 1 ?

References I Kelly. G. J. (1982l lrends Biocheal. Sci. 7,233 2 Palmer. J. M. (1982) lrends Biochem. Sci. 7,

357 3 Goldstein, A. H, Anderson, J. O. and McDaniel,

R. G. (198(I) Plant Physiol. 66,488-493 4 Siedow, J. N. and Girvin, M. E. (198(I) Plant

PhysioL 65,669--674 5 Parrish. D. J. and Leopold, A. C. (1978) Plant

Physiol. 62, 47(I--472 6 Bendall, B. S. and Bonner, W. D. Jr (19711 Plant

Physiol. 47. 236-245 7 Bonnet. W. D. Jr and Rich, P. R. (19781 in Plant

~litochondria (Ducet, G, and Lance. C., eds). pp. 241-247, Elsevier/North-Holland Biomedical Press

8 Huq, S. and Palmer, J. M. (1978) FEBS Lett. 92, 317-320

9 Rich, P. R. (1978)FEBS Lett. 96,252-256 1(1 Dizengremel, P. and Lance, C, (1976) Plant

Physiol. 58, 147-151 11 Laties, G. G, (1982) Annu. Rev. Plant Physiol.

33, 51%555

157

12 Edwards, D. L., Rosenberg, E. and Maroney, P. A. (1974)J. BIOl, Chem. 249, 3551-3556

13 Erecinska, M. and Storey, B. T. (197(I) Plant Physiol. 46,618-624

14 Dupont, J., Rustin, P. and Lance, C. (1982)Plant Physiol. 69, 1308-1314

15 Rustin, P., Dupont, J. and Lance, C. (19821 Physiol. Veg. 20,721-727

16 Grossman, S. and Za.kut, R. (19781 ~4ethods Biochem. Anal. 25,303-327

17 Miller, M. G. and Obendorf, R. L. (19811 Plant Physiol. 67,962-964

18 Rychter, A., Janes, H. W. and Frenkel, C. (1979) Plant Physiol. 63, 149-151

19 Beutelman, P. and Kende, H. (19791 Plant Physiol. 63,888-893

2(I Leshem, Y. Y. and Barness, G. (19821 in Biochemistry and Metabolism of Plant Lipids (Wintermans, J. F. G. M. and Kuiper, P. J, C., eds), pp. 275-278, Elsevier/North-Holland Biomedical Press

21 Bennoun, P. (1982)Proc. Nad Acad. SoL U.S.A. 79, 4352-4356

Emerging Techniques

Intracellular translocation and proteins. Although the enzymes respon- sible for lipid biosynthesis reside primarily

metabolism of fluorescent o. the rough and smooth endoplasmic reticulum, lipids are found throughout the lipid ~e,, with different intracellular membranes analogues in cultured often having different lipid compositions 3. Additionally, each leaflet of a given mere-

m a m m a l i a n cells brane bilayer (e.g. the plasma membrane) may have a markedly different lipid corn-

Richard E. Pagano and Kenneth J. Longmuir position from the other ~. Thus the synthesis and translocation of lipids present sorting and targeting problems analogous to those

Fluorescent analogues o f phosphol ip ids add a new d imens ion to studies o f lipid encountered with the assembly and/or se- metabol i sm by prov id ing in format ion about the distribution o f lipid metabolites in cretion of protein molecules. The purpose of living cells. Using this approach we present evidence for the 'sorting' o f lipids during this article is to present new technology

the irs ynthesis, transport and assembly into intracellularmernbranes, which we feel may be useful for future studies of the metabolism, translocation

One of the most exciting and rapidly much a part of membrane biogenesis and and assembly of cellular lipids into mem- developing areas of cell biology today is membrane differentiation as membrane branes. the study of the translocation of protein .... : molecules from their sites of synthesis and t i ~ - ~ ( ENOOPC SMIC RET COLt~ ) ~

, DfGLYCERiDE ] ~ f ~ ~i • glycosylation in the endoplasmic reticulum I ~ / To I -__ r . . . . ~ t_J - - . . . . , ~ ,/ TRIG LYCE R$ DE and Golgi apparatus of the cell, to various : :1 ; (~ & P.osP.ot,,0 [ ~ . ~ ~ - ~ - , : . . . . . ~ ) I~ Ill CONVERTER. J ~"k ~ J ~ - - ~ _ . t , ~ t ( / % ~ ~ / intracellular compartments or the cell sur- ~ IIIIKEEP AT aT*C! i.t.--" "<£.,J' -A4@/(q ~ ......

face. The mechanism(s) by which these :1 --3~ ~ ~__~ y~-~r- ~ , a ~ 3 q ~ ~ ' ~ , / ~ ~ , - - ( =(at3 5 molecules are apparently sorted out and , n "

'targeted' to their appropriate destination(s) ~ SOa~,NG within the cell or the extracellular environ- ~ ~ ~ , ~ ~ x'~-~l] ~ ~

biolog are under active investigati°n by molecules, which amount to : 1 ~ ~ ~ t ~ ' - - ~ ~ I " 10-90°/, of the dry weight of a cell, a reas ~ ~ ~ " ~ ! .~,

Richard E. Pagano is at the Department o f ~ ~ . . ~ , . . , i 'i~ ):, i Embryology, Carnegie Institution of Washington, ~ " . _ x 115 West University Parkway, Baltimore, AID 21210, U.S.A. Kenneth J. Longmuir was at the Department of Embryology. Carnegie Institution oJ' Washington, and is now at the Department of "~','~ ~'~-ccc.:cc~c,',:cc~¢ c ~ c c ~ , ' ¢ , c v , ' j Physiology and Biophysics, College of Medicine, ~ ' University o]" CaliJornia, lrvine. CA 92717, U.S.A. ~7 ...........

~; 1983, Elsevier Science Publishers B ~ . Amsterdam 0376 5007/83/$01 on

158 TIBS - May 1983

Until now, limited information has been obtained about the mechanisms of lipid translocation within eukaryotic cells. This is because the study of the distribution and movement of lipid molecules is hampered by various shortcomings of the traditional methods used to study metabolism at the subcellular level. For example, in the 'pulse-chase' experiment, which involves the uptake of radiolabeled lipid precursors followed by cell fractionation, labeled lipids may exchange between isolated membrane fractions, either by the action of soluble exchange proteins 5'6, or by the tend- ency of certain lipids to transfer spontan- eously between membranes. Thus, the appearance with time of a particular lipid species in a given membrane fraction may or may not be indicative of what actually occurs in situ. The purity of subcellular fractions, an important issue in all mem- brane biochemical studies, is particularly critical when dealing with lipids. The plasma membrane of eukaryotic cells, for example, contains only about 1-2% of the total cell lipid mass, making contamination with even small amounts of intracellular lipids a serious problem. Finally, a major problem faced by the cell biologist or lipid biochemist interested in the movement of lipids within cells is that except for electron microscopic autoradiographic techniques, which are tedious and require special atten- tion to lipid retention during sample proces- sing 7'8, the location of lipids within intact cells cannot be readily determined. This is because, in contrast to studies of protein translocation, which can utilize tagged and specific antibodies or toxins for examining subcellular distribution, no specific probes for the major classes of lipids exists.

To circumvent some of the problems out- lined above we have developed a new approach employing fluorescent phos- pholipids which apparently can be used as true analogues of some of their natural counterparts. This approach is potentially very powerful since, as will be discussed, we can carry out traditional experiments in lipid metabolism, then correlate these biochemical data with 'positional' informa- tion on the intracellular location of the lipid metabolites in living cells, using conven- tional fluorescence microscopy. In this report, we highlight the unique properties of a fluorescent derivative of phosphatidic acid, 1 - acyl - 2 - (N - 4 - nitrobenzo - 2 - oxa- 1,3 - diazole) - aminocaproyl - phos- phatidic acid, or C6-NBD-PA (Fig. 1 ). This lipid appears to he particularly well suited for examining those regions of the cell where lipid biosynthesis takes place.

In our experiments, small unilamellar vesicles are used as a vector for introducing the fluorescent phosphatidic acid into cul-

tured Chinese hamster fibroblasts 9. In a typ- ical experiment, vesicles are first prepared from a mixture of dioleoyl phosphatidyl- choline, C6-NBD-PA, and a non- exchangeable fluorescent lipid 1°. The vesi- cles are then incubated with cells in mono- layer culture or in suspension for 60 rain at 2°C in a simple, protein-free, balanced salt solution. Under these incubation condi- tions, -->90% of the cell-associated C~- NBD-PA is taken up by preferential lipid transfer ~.1° from vesicles to cells, rather than by alternate mechanisms of uptake ~ ~4 such as vesicle-cell adsorption, fusion, or endocytosis. Following this incubation, the cells are washed, and either examined by fluoresence microscopy, or the lipids are extracted and analysed by conventional analytical procedures.

Intracellular distribution and translocation of fluorescent iipids

In early studies we found that if vesicle- cell incubations are carried out at 2°C using 1 - acyl-2.-[NBD-aminocaproyl]- phosphatidyl-choline or phosphatidyl-

H I

H2C C f t

0 0

I I O:C C-:O

I i R (CHa)

5

H

I 0

I O=P-O

I 0 I CH 2

NH

~ ~ N \ O

%N /

NO 2 Fig . l . S l r u c l u r e o f ! - a c y l - 2 - ( N - 4 - n i t r o -

b e r ; z o - 2 - o x a - 1 , 3 - d i a z o l e ) - a m i n o c a p r o y l p h o s -

p h a l i d i c a c i d ( C ~ - N B D - P A ) .

ethanolamine, these lipids are selectively incorporated into the plasma membrane '°. Furthermore, no internalization of these fluorescent lipids takes place as long as the cells are kept at 2°C. It was therefore quite surprising to find that under the same incu- bation conditions, C6-NBD-PA is not only transferred into the cell at 2°C, but all of the cell-associated fluorescence is internalized to cytoplasmic membranes. Two prominent cytoplasmic features are seen by fluores- cence microscopy (Fig. 2). First, a portion of the fluorescence is localized to a reticular network, which we have identified as the endoplasmic reticulum. Second, bright dots of fluorescence, approximately 1-2 /zm in diameter, are distributed throughout the cytoplasm. These organelo les are mitochondria.

Identification of the reticular network was accomplished by staining with rhodamine-conjugated lectins, which pre- ferentially label the endoplasmic reticulum of eukaryotic cells ~5. Comparison in the same cell of the reticular network stained by Cs-NBD-PA with the endoplasmic reticulum stained by the lectin demon- strated that both probes were localized to the same membranes. Similarly, identification of the bright fluorescent dots in the cyto- plasm was accomplished by showing that the Cs-NBD-PA co-localized with the fluores- cent probe Rhodamine 3B, a cationic dye that is specifically accumulated by the func- tioning mitochondria of living cells t". These co-localization studies allowed us to conclude that when cells are incubated with Ce,-NBD-PA at 2°C, the mitochondria and the endoplasmic reticulum are the principal intracellular sites of fluorescence localiza- tion.

A new pattern of intracellular fluores- cence labeling emerges if cells which have been treated with C~-NBD-PA are washed and then warmed to 37°C (a pulse-shift experiment) 17. As seen in Fig. 3, the fluorescence of the endoplasmic reticulum is diminished, and spherical regions of fluorescence appear in the cytoplasm. Using a variety of techniques, we have demonstrated that these spherical organ- elles correspond to intracellular lipid drop- lets, which are present in many cell types grown in serum-containing culture medium TM. It should be noted that although these droplets were present at all times dur- ing our experiments, they were not fluores- cent after incubation with C6-NBD-PA at 2°C, and became fluorescent only when the temperature was shifted to 37°C.

Cellular metabolism of fluorescent lipids Specific events in lipid metabolism

accompany both the initial uptake of C~- NBD-PA at 2°C, and its subsequent redis-

T I B S - May 1983

tribution at 37°C. During incubation at 2°C, most ( -80-90%) of the C6-NBD-PA is converted to NBD-diglyceride, with the remaining lipid consisting principally of intact C~NBD-PA. We do not yet know whether conversion of the phosphatidic acid to diglyceride is necessary for transport into the cell, or whether the two events, transport and hydrolysis to diglyceride, are independent of one another. We have, however, established that uptake does not

proceed by hydrolysis to the fluorescent fatty acid, followed by its transport into the cell, and re-esterificationlL Instead, the glycerol backbone and fatty acid portions of the molecule remain together during trans- port.

If the cells are wanned to 37°C, the NBD-diglyceride is converted largely to NBD-triglyceride and NBD-phosphatidyl- choline, with small amounts of other NBD-phospholipids also formed. This can

159

be seen in the thin-layer chromatogram pre- sented in Fig. 4, in which the fluorescent lipids obtained from cell extracts after dif- ferent periods of time at 37°C are shown. Thus, the redistribution of intracellular fluorescence seen during a pulse-shift experiment (Fig. 3) is accompanied by a marked change in the composition of cell- associated NBD-lipids.

Subcellular fractionation studies follow- ing the pulse-shift experiment demonstrate

Fig. 2. bluorescence micrographs o f ( "hinese hamster fibroblast rnonolayer cultures incubated witt C8-N B D-PA containing vesicles ]or 60 rain at 2 ( ". ( a) Bar, 10 lain. (b) The same cell at higher magnification. Bar is 2 tam. Reproduced from Pagano et al. ~ by permission of The R~x:keleller U niversit,¢ Pre~,s.

160

that primarily NBD-triglyceride is asso- ciated with the intracellular lipid droplets, although substantial amounts of NBD- labeled phosphatidic acid and phos- phatidylcholine are also present in the intact cells. This finding was confirmed in a sep- arate experiment in which the fluorescent lipids associated with the intracellular lipid droplets were selectively and irreversibly photobleached in si tu. Extraction and analysis of the fluorescent lipids revealed that only NBD-triglyceride was photo- bleached. From these results we conclude that NBD-triglyceride is selectively trans- ported to the intracellular lipid droplets, while other NBD-lipids remain in other intracellular membranestL Hence the cell recognizes the different classes of fluores- cent glycerolipids as they are lormed, and can transport or "sort' them to different cytoplasmic locations. Ahhough the molecular mechanism(s) underlying this process is not yet known, one possibility is that some NBD-triglyceride is synthesized directl? al the intracellular lipid droplet sites. Alternauvel~,,, NBD-triglyceride may be It)treed elsewhere in the cell (e.g. at the endoplastntc reticulum) and translocated to thc mtracellular lipid droplets by lateral dillu~,ic, n'", vesicular transporP ~. l ip id excl~ange" ", or emulsification ~'.

• \1 present u,c do not klloVx, ht:,w precisely Ihc metabolism and mtracellular transloca- (lOll OI C,~-NBD-PA represent that OI

I'2g. 3. bluorescence micrograph o f Chinese ham- ~tcr fihrohlast monohzver cuhure after a ptds'e-shtft c~ twriment with ( ",~NBD-FA. ( "ells were incubated with ( ',~ N B I )- I'A ('ontaining veswles ]or 60 rain at 2 ( , washed, and fitrther incubated in a balanced ~ah ~ol.tion jor 30 rnin at 37 ( ' prior to photo- grapItv. .Arrows indicate ~everal sites ~] fluor- e~cen/(~ laheh'd inlraceUular liFid droplets. Bar is 5 h~ltl

I I ' O

e o o i O i

I I B S - A.htv 19,'¢3

- - T 6

- D 6

- FA

- PG

- P C

- - PS - PA

0 5 I 0 15 2 0 2 5 3 0 4 5 6 0 I'tg. ~ l . rm ,mon oJ NBl)-Iipid product~Jrom ( . - N B D - t ' A . C 'elL~ were treated wilh ( ' ,~NBD-/ 'A eotllaitl- in,q l c~wA'~ jor 60 rain at 2 (', washed, and sahsequent(v tncuhaled in a balanced sah solutum at 37 ( 'Jor t'arlotLs" tittles. 771e cells were then washed, scraped from the ell[lure dish, and the lipids extracted. 7tl¢' lipid erlracts were subjected to thin-layer chromatography and the chromatogram was photographed under U V hght. Note the decrease in N B D-diglyceride ( N B D-D( ; ) and the apl)earam e o f N B D-pho whatidvh'h¢dine ( N BD-P( ' ) and NBD-trig(vceride (NBD-TG) with time (mini at 37 ('.

endogenous phosphatidic acid. However, the tollowing should be noted. First, based on the known metabolic pathways for glycerolipid biosynthesis (reviewed in Refs 21,22), C6NBD-PA is apparently metabol- ized through the diglyceride pathway to the fluorescent analogues of diglyceride, tri- glyceride, phosphatidylcholine, and small amounts of other lipids. Hence, the enzymes of lipid metabolism which cata- lyse these reactions are not inhibited by the fluorescent NBD-moiety on the fatty acid chain. Second, the NBD-labeled lipids do not randomly disperse throughout the cyto- plasm, but rather assume a unique distribu- tion within the cell during the metabolism of Cs-NBD-PA. Since it is generally believed that intracellular lipid droplets in cultured cells are storage depots for cellular triglyceride TM, the intracellular 'sorting' of fluorescent triglyceride is further evidence that the NBD-lipids behave in a manner similar to the normal products of lipid metabolism. Finally, preliminary experi- ments using exogenously supplied di-(9,10-[3H]-oleoyl phosphatidic acid indicated that this compound, like C~- NBD-PA, was hydrolysed to diglyceride and further metabolized to phosphatidyl- choline and neutral lipids (unpublished observations).

We conclude that fluorescent lipid analogues such as C~-NBD-PA hold much promise for correlating biochemical inves- tigations of lipid metabolism with fluores- cence microscope studies of the intracellu- lar localization and transport of lipid metabolites during their assembly into cellular membranes. In addition, this compound provides an elegant means |br ob~rving the endoplasmic reticulum in living cells, previously possible only after fixation, permeabilization and staining procedures.

Acknowledgements The authors thank Dr N. G. Lipsky |br

suggesting the accompanying cartoon. This work was supported by USPHS Grant GM-22942 to R. E. Pagano. K. J. Longmuir is supported by a USPHS Pulmonary Faculty Training Award (5-KOS-HL00456) sponsored by the Department of Physiology and Biophysics, College of Medicine, University of California, Irvine.

References I Palade, G. E, (1975)Science 189, 347-358 2 Farquhar. M. J. and Palade, G, E. (1981)J. (~'//

Biol. 91,77s-106s 3 White, D. A, (1973) in Porto and Function oJ

Phospholipids (Ansell. G. B., Hawthorne, J. N

T I B S - May 1983

and Dawson, R. M. C., eds), Ch. 16, Elsevier/North-Holland Biomedical Press, Ams- terdam

4 0 p den Kamp, J. A. F. (1979) Annu. Rev. Biochem. 48, 47-71

5 Bloj, B. and Zilversmit, D. B. (1981)Mol. Cell. Biochem. 40, 163-192

6 Wirtz, K. W. A. (1982)inLipid-Protein Interac- t/ons (Jost, P. C. and Griffith, O. Hayes, eds), Vol. 1, Ch. 6, John Wiley and Sons, New York

7 Stein, O. and Stein, Y. (1971)Adv. LipidRes. 9, 1-72

8 Gould, R. M. and Dawson, R. M. C. (1976) J. Cell Biol. 68,480--496

9 Pagano, R. E., Longmuir, K. J., Martin, O. C. and Struck, D. K. (1981)J. CellBiol. 91,872-877

10 Struck, D. K. and Pagano, R. E. (1980)J. Biol. Chem. 255, 5405-5410

11 Pagano, R. E. and Weinstein, J. N. (1978)Annu. Rev. Biophys. Bioeng. 7,435---468

12 Pagano, R. E., Schroit, A. J. and Struck, D. K. in Liposomes: From Physical Structure to Therapeutic Applications (Knight, C. G., ed.), Ch. 11, Elsevier/Noah-Holland Biomedical Press, Amsterdam

13 Huang, L. in The Liposomes (Ostro, M. J., ed.), Ch. 3, Marcel Dekker, New York (in press)

14 Szoka, F., Jacobson, K., Derzko, Z. and Papahad- jopoulos, D. (1980) Biochim. Biophys. Acta 600, 1-18

15 Virtanen, I., Ekblom, P. and Laurila, P. (1980) J. Cell Biol. 85,429-434

16 Johnson, L. V., Walsh, M. L., Bockus, B. J. and Chen, L. B. (1981)J. Cell Biol. 88, 526-535

17 Pagano, R. E., Longmuir, K. J. and Martin, O. C. (1983)J. Biol. Chem. 258, 2034-2040

161

18 Spector, A. A., Mathur, S. N., Kaduce, T. L. and Hyman B. T. (1981) Prog. Lipid Res. 19, 155-186

19 Edidin, M. (1981) in New Comprehensive Biochemistry (Finean, J. B. and Michell, R. H., eds), Vol. 1, Elsevier/North-Holland Biomedical Press, Amsterdam

20 Small, D. M. (1981) in Proceedings of the Inter- national Conference on Biological Membranes (Bloch, K., Bolis, L. and Tosteson, D. C., eds), Ch. 2, pp. 11-34, PSG Publishing Company, Boston

21 Van Golde, L. M. G. and Van den Bergh, S. G. (1977) in Lipid Metabolism in Mammals (Snyder, F., ed. ), Vol. 1, pp. 1-33, Plenum Press, New York

22 Bell, R. M. and Coleman, R. A. (1980) Annu. Rev. Biochem. 49, 459-487

Textbook Errors

The stoichiometry of the citric acid cycle

T. Norman Palmer and Mary C. Sugden

Students are often asked to describe the pathway whereby palmitate (or related long-chain fatty acids containing an even number of carbon atoms) is converted to glucose in mammalian liver. Needless to say, this is a 'trick question'. Mammalian liver and kidney are unable to convert pal- mitate into glucose. This fact is related to the stoichiometry of the citric acid cycle. The entry of the two-carbon acetyl unit of acetyl CoA (the product of/3-oxidation of palmitate) into the cycle is balanced by the loss of two carbon atoms by decarboxyla- tion within the cycle (catalysed by isocitrate dehydrogenase and the 2-oxoglutarate dehydrogenase complex). Therefore the acetyl unit is oxidized in the cycle and can- not form oxaloacetate de novo.

The fact that the stoichiometry of the citric acid cycle commits the acetyl unit of acetyl-CoA to oxidation (to two molecules CO2) is frequently dealt with in some depth in undergraduate textbooks. An equally important facet of the stoichiometry of the citric acid cycle appears, however, to be frequently ovedooked. This is that the acetyl unit of acetyl-CoA is the only sub- strate that the cycle can oxidize completely. Goldstein and Newsholme x and Vinay et al. 2 have made this point previously but it warrants reiteration given its fundamental importance to intermediary metabolism.

Textbooks, in consequence, are in error when they imply that conversion of the car- bon skeletons of certain amino acids to citric acid cycle intermediates results in complete oxidation per se. These amino acids are glutamate, glutamine, histidine, proline and arginine (cycle entry at 2-oxoglutarate), isoleucine, methionine, threonine and valine (cycle entry at succinyl-CoA), phenylalanine and tyrosine (cycle entry at fumarate), and aspartate and asparagine (cycle entry at oxaloacetate) (Fig. 1). For example, entry of 1 mol of glutamate into the cycle as 2-oxoglutarate

T. Norman Palmer and Mary C. Sugden are at the Department of Biochemistry, Chafing Cross Hos- pital Medical School, London, W6 8RF, U.K.

and its metabolism through one revolution of the cycle results in:

oxoglutarate + 3NAD + + FAD + GDP + Pi + 2I-hO + acetyl-CoA ---,

oxoglutarate + 2CO~ + 3NADH + FADI-h + 2H ÷ + GTP + CoA.

It is clear that, provided no intermediates leave the cycle, oxoglutarate is regenerated whilst 1 mol acetate (as acetyl-CoA) is oxidized. Needless to say, the entry of 2-oxoglutarate into the cycle, whilst result- ing in no net oxidationperse, would lead to a continuous increase in the concentration of cycle intermediates.

The amino acids specified above are obviously capable of being oxidized in vivo. The question is how this is achieved. Given that the acetyl unit of acetyl-CoA is the only substrate oxidized quantitatively by the citric acid cycle, logic dictates that

Acetyl CoA

Asparagine ~ - Aspartate ] / • Oxaloacetate

Malate Citrate

[ rosine 1 l 1 Phenylalanine I ~- Fumarate Isocitrate

Succinate C02 [

Isoleucine ~ Succinyl CoA 2-Oxoglutarate ~ Methionine ,,F-~ ~ - - - ~ " Threonine CO 2 Valine

Fig. 1. Fates of the carbon skeletons of relevant amino acids.

Glutamate Glutamine Histidine Proline Arginine

,~: 1983, Elsevier Science Publishers B.V.. Amsterdam 0376 - 5067/83/$01.00