Dialogue between membranes and their lipid-metabolizing enzymes. WILLIAM E. M. LANDS. Department of Biological Chemistry, The University of. Michigan ...
Membrane Enzymes in their Lipid Environment Colloquium organised on behalf of the Lipid Group and the Membrane Group and edited by P. J. Quinn (London)
Dialogue between membranes and their lipid-metabolizing enzymes WILLIAM E. M. LANDS Department of Biological Chemistry, The University of Michigan, Ann Arbor, MI 48 109, U S A . Biological chemists have often regarded the fluidity or plasticity of a membrane as a property that may be adaptively altered to accommodate changes in environmental temperature. The implication that cells exhibit an intelligent (or at least, appropriate) response provides us with several questions to examine. (1) Is temperature or some aspect of fluidity perceived by the responsive cellular element? (2) How is the detected difference from ‘ideal’ in temperature or fluidity communicated to the synthetic enzyme(s)? (3) How can the selective actions of the enzyme(s) of phospholipid synthesis be changed to alter the fatty acid content of membrane lipids? A primary intellectual challenge in examining these responses is to discern the degree to which the cells adapt because of signals from an inadequate membrane fluidity in contrast with an independent alteration of synthetic selectivity that coincidentally alters the membrane fluidity. In a sense, one might ask whether the intracellular signalling is a dialogue or a monologue, or in other terms, whether the manner of membrane adaptation is purposefully successful (‘Lamarckian’) or fortuitously beneficial (‘Darwinian’). Certainly cells containing either adaptive mechanism could exhibit better survival in a limiting environment than would those cells without. A complicating factor in evaluating adaptive changes in the fatty acid composition of membrane phospholipids is that the synthetic enzyme(s) may act on only single fatty acid molecules by recognizing their chemical structural features rather than fluidity, which is a co-operative aspect of the interaction of many molecules. Thus we must be sensitive to the language in which messages are translated and communicated within cells. Few of the enzymes handling fatty acid derivatives have shown the ability to discriminate appreciably among the different acyl chains, and selective actions seem likely for only the fatty acid synthases, desaturases and acyltransferases. The factors used by these enzymes in selecting preferred acyl chains seem important elements in the ‘language’ of membrane lipid adaptation. Fatty acid biosynthesis in response to temperature One of the more completely interpreted membrane adaptations is that of Bacillus megaterium described by A. J. Fulco (Fulco, 1972; Fujii & Fulco, 1977). These bacilli increase the degree of unsaturation in their membrane lipids when grown at lower temperatures. This seemingly intelligent response of providing membrane lipids of greater fluidity appears due to the thermal instability of an oxygen-dependent phospholipid desaturase (Fulco, 1972). The enzyme activity decays slower at lower temperatures and thus provides more unsaturated membrane lipids. An added control feature is that the rate of synthesis of the enzyme is attenuated by a temperature-sensitive modulator protein (Fujii & Fulco, 1977). All the responses examined appear to be changes in levels of activity that are directly mediated by temperature and not by the state of membrane fluidity (Fulco & Fujii, 1979). Another well examined system is that of Escherichia coli. Marr & Ingraham (1962) showed that 2-fold more cis-vaccenate (cis-
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octadec-1 1-enoate) was present in cultures grown at 30°C lower temperature (21% at 10°C against 38% at 40°C). Seldom emphasized by other workers is the authors’ claim, given also in a subsequent report (Shaw & Ingraham, 1965), that the adjustment of fatty acid composition did not appear to be a prerequisite for growth at 10°C, and that the degree of unsaturation of the fatty acids did not appear to set the minimal temperature for growth. The increased portion of unsaturated acids at the lower temperature has been regarded as due to an adaptively altered specificity of the cellular acyltransferases (Sinensky, 1971; Cronan, 1975; Kit0 el al., 1975; Nishihara el al., 1976). However, Okuyama et af. (1977) showed that in vitro the fatty acid synthetase of E. coli produced more unsaturated fatty acids (85% against 40%) when the incubation temperature was lowered to 10°C from 40°C. Parallel studies showed that esterification by the acyltransferase(s) incorporated oleate and palmitate in accord with the ratio of available CoA thiol esters and was independent of incubation temperature. Thus the adaptive response to low temperature was not being communicated to the esterifying acyltransferases by altering their relative specificity, but rather by changing the relative availability of the different substrates for esterification. The results point to a thermal effect on the synthesis of cis-vaccenate that coincidentally results in elevated fluidity at low temperatures. The experiments gave no evidence that the increased synthesis of unsaturated fatty acids was in response to an ’inappropriate’ membrane fluidity. In fact, results provided by Gelmann & Cronan (1972) indicated that a mutant which forms negligible amounts of cis-vaccenate grows at the same rate as the wild-type cells at all temperatures tested, with no alteration in the fatty acid content of its membrane lipids. It seems that an increased amount of cis-vaccenate is not required for growth at the lower temperature, and that the ‘corrective adaptation’ was a fortuitous event unrelated to any measurable need for more fluid cellular membranes. Such a conclusion was forwarded 10 years earlier by Marr and Ingraham in their pioneering measurements of fatty acid alterations. Thus one again finds no firm evidence for a signal originating in ‘inadequate’ membrane fluidity to alter synthetic specificities. The report of a ‘homeoviscous adaptation’ (Sinensky, 1974) describes a provocative phenomenon that merits more careful interpretation. The signal that causes decreased biosynthesis of C 16: and increased biosynthesis of C18:I fatty acid, with little change in CI6: fatty acid is not yet clear. Recognition of the differences in two forms of ,4ketoacyl-acyl-carrier protein synthetase (D’Agnolo et al., 1975) allowed a tentative assignment of synthetase I as important in forming palmitoleate (C16:I ) and synthetase I1 in forming cis-vaccenate (C ,). Somehow these two enzymes also serve in the synthesis of saturated fatty acids and play a role in forming C : and C 14 : fatty acids, as well as C ,6 : fatty acid.
,
,
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Doesfatty acid biosynthesis respond tojuidity? Another adaptive response to lower fluidity might be to increase the ratio of C14:o/C16:ofatty acids, as indicated by Esfahani et al. (197 1). The shift of the ratio of C ,/C 16 : fatty acids from 0.1 with exogenous oleate (cis-octadec-9-enoate) to 1-3 with trans-octadec-9- and - I I-enoate or cis-eicos-1 I-enoate
,,:
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26 was regarded as a ‘correcting mechanism’ to attain the ‘proper’ fluidity. Such a corrective event was not observed during temperature-induced adaptations, and it may not be a general adaptive response. Later work by Silbert et al. (1973), Cronan (1975) and Okuyama et al. (1977) reported no accumulation of shorter-chain fatty acids. In fact, Marr 8c Ingraham (1962)and Silbert et al. (1973) showed decreased accumulation of C,2:o and C1,:o fatty acids at lower temperatures. The manner and extent of ‘adaptation’ to exogenous fatty acids needs much more careful interpretation, since previous claims of regulatory adaptation may have been unfounded (Polacco & Cronan, 1977). A somewhat different concept was forwarded for the eukaryotic Tetrahymena pyriformis. A rapid increase in unsaturated acids with decreased temperature was interpreted to be in response to decreased fluidity in the microenvironment of the microsomal desaturase(s) (Martin et al., 1976). The desaturase may be affected to provide an altered availability of acyltransferase substrates by fluidity effects without requiring a thermal influence (Kasai et al., 1976). Such an adaptive response might be significant in helping to maintain the cellular membrane integrity of homeotherms when stressed by agents that lower membrane fluidity. Presumably the decreased fluidity associated with increased membrane sterols could also enhance desaturase activity. This concept differs somewhat from that explored in detail by Strittmatter & Rogers (1975), in which the fluidity of the microenvironment of the desaturase was facilitating activity, and decreased fluidity was claimed to decrease activity in a manner not correlated with microviscosity (Enoch et al., 1976). No clear concept indicates how desaturase activity may be enhanced by lower fluidity.
Acyl-chain composition andfluidity In describing the possible effects of fluidity on cellular function we need better descriptions of the dimensions of fluidity and a clearer focus on the manner in which the unsaturated fatty acids quantitatively contribute to fluidity. Although we know some features of membrane lipids that occur in normal circumstances, we have imprecise evidence of what is required by cells, and we should be cautious in declaring the generally existing state to be a beneficial and desired state. For example, lipid heterogeneity gives a broad range for the solid-liquid phase transition and may provide abundant lateral phase separations regarded by some as desirable for normal cellular function (e.g. Shimshick & McConnell, 1973;Linden et al., 1973). Nevertheless, results with Acholeplasma laidlawii cells suggest that lateral segregation of lipids into gel state domains is not essential for proper membrane function (Silvius & McElhaney, 1978). There is sufficient evidence that the fluid state of lipids gives better membrane function than does the solid or gel. We must consider whether temperature-dependent variations in the apparent suitability of membrane lipids are due to a changing fluidity of the fluid state or to changing proportions of gel state within the fluid bilayer. Careful quantitative examination of specific enzyme activities in the more homogeneous fluid membrane matrix (Silvius 8c McElhaney, 1978) will be very helpful in assessing the relative contribution of fluidity and of specific unsaturated acyl chains to those activities. Also, we will need further designation of the lipid’s contribution in terms of its action as boundary (Jost et al., 1973)or annular (Warren et al., 1974) lipid in contrast with bilayer lipid. The relative contribution of acyl chains to fluidity per se still requires more quantitative assignment. Additional general indices of fluidity such as mole-% unsaturated fatty acids, the ratio of saturated to unsaturated fatty acids, or the ‘double-bond index’ do not give a clearly quantitative assignment of the level of fluidity in the lipid bilayer. The use of mole-% unsaturated acid fails to account for a greater contribution by polyunsaturated fatty acids. On the other hand, the use of a double-bond index, 1(mole-% unsaturated acid), x (double bonds per acyl chain),, may overestimate the relative fluidity because the most
BIOCHEMICAL SOCIETY TRANSACTIONS significant increase in area per mole is observed with the first double bond and is less with subsequent introductions of double bonds (Demel et al., 1972). We have attempted to assign a molar contribution by different unsaturated fatty acids to membrane function (Holub 8c Lands, 1975). and this has been useful in letting us describe differences among the many acyl chains studied. Nevertheless, all predictive molar indices currently used are inadequate. This inadequacy is in part due to the fact that fluidity is a conceptual feature of liquids that has diverse aspects (Hildebrand, 1971;Lakowicz et al., 1979).
Selectivitiesfor esterification Some years ago I found no significant alteration of acyltransferase specificity with temperature (Jezyk & Lands, 1978) or lipid matrix (Ellingson et al., 1970). At present, we still have no clear evidence of an ability to alter the selectivity of the esterifying enzymes beyond changes in the supply of unsaturated fatty acid substrates. Further search for examples of altered acyltransferase selectivity might be constructively pursued in cells unable to synthesize unsaturated fatty acids. Such cells grow well when the required type of acid is nutritionally available. We have observed high amounts of the unsaturated acid in cellular membrane lipids at early phases of culture growth that decreases as it is apparently redistributed to cells in subsequent generations. The high levels obtained indicate that a high membrane fluidity does not appreciably restrain the entry of more fluidizing fatty acids. Indeed, the possibility of a controlling response preventing too much fluidity has not been widely discussed or demonstrated, making a major void in our understanding of what may represent ‘too much’ fluidity for cells. During growth of auxotrophic cultures in limited supplies of nutrient unsaturated fatty acid, the cellular enzymes in the isothermal cultures are exposed to progressively lower fluidity with its presumed stimulus for an adaptive response. In studies of E. coli and Saccharomyces cerevisiae mutants have so far provided no evidence for an acyltransferase adaptation as the cells insert greater amounts of saturated fatty acids into membrane lipids and cell division and growth cease. It appears that the cells are unable to ameliorate the steady loss of membrane fluidity as the acyltransferases continue to make progressively lesscompetent membranes. Although it is premature to make firm conclusions, the fact that the cells continue to incorporate saturated fatty acids into the less fluid membranes suggests that little intelligent adaptive communication is acting in cells lacking the enzyme(s) for synthesizing the unsaturated fatty acids. If esterification of medium- or short-chain fatty acids into membrane lipids were a suitable adaptive response (Cronan, 1978), the proper occasion seemed present, but it did not trigger a response to alter the selectivity of the synthetic enzymes in time to save the cells from a self-inflicted disaster (Henning et al., 1969). We have no conclusive evidence that membrane fluidity alters the selectivity of the biosynthesis or the esterification of fatty acids. We have observed several unusual growth responses of auxotrophic mutants to certain exogenous nutrient fatty acids (Ohlrogge et al., 1976;Lands et al., 1977). Such results led us to consider that some effects that have been attributed to fluiditydependent signalling may actually be more dependent upon aspects of metabolic regulation (Lands et al., 1978). These effects reflect highly specific chemical features of the nutrient fatty acids. For instance, although trans-octadec-9-ynoate was not an effective nutrient supplement for the E. coli mutant grown in glucose, it was as effective as the cis-isomer for the S . cerevisiae mutant (Lands el al., 1977). More surprisingly, the trans-8- and - 10-ynoate isomers were more effective than the corresponding cis-isomers with both types of cell. Since all the acetylenic acids have high melting points, we need a major reappraisal of the way we believe acyl chains will affect membrane fluidity. A clear example is that of trans-octadec-9-enoate, which is often regarded to decrease the fluidity of membrane lipids. Growth of
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an E. coli auxotroph is glucose with the cis C,, , fatty acid was about 43 cells/fmol whereas growth with the trans-isomer was negligible. We found that the effectiveness of the trans acid in supporting growth increased from 1 to 19 cells/fmol when cyclic AMP was added (Tsao & Lands, 1979). Thus, the trans acyl chain can provide sufficient fluidity to support nearly normal cell growth and function, but its other interactions involving cyclic nucleotide and metabolic regulation dominate the cellular response. Our results help emphasize the need to designate in a more quantitative manner the contribution that an acyl chain makes to membrane fluidity and to assign separately the influence of the acyl chain upon metabolic events within the cell. Additionally, we need more quantitative evidence of the precise level of fluidity that can cause detectable influence of specific membrane-bound enzymes. The membrane fludities obtained by cells may be a fortuitous result of several events not causally linked to the suitability of the product. It remains possible that the selectivity of our acyltransferases cannot be altered, and that, isothermally, there is no signal to the desaturases so that cells which acquire inadequate lipid compositions die. The persistent desire of many individuals to perceive beneficial ‘corrective’ responses in Nature may not provide sufficient force to alter the nature of reality. ~
Cronan, J. E. (1975)J. Biol. Chem. 250,7074-7077 Cronan, J. E. (1978)Annu. Rev. Biochem. 47, 163-189 D’Agnolo, G., Rosenfeld, I. S. & Vagelos, P. R. (1975)J. Biol. Chem. 250,5289-5294 Demel, R. A., Geurts vanKessel, W. S. M. & van Deenen, L. L. M. (1972)Biochim. Biophys. Acta 266,2640 Ellington, J. S., Hill, E. E. & Lands, W. E. M. (1970)Biochim. Biophys. Acra 196,176-192 Enoch, H. G., Catala, A. & Strittmatter, P. (1976)J. Biol. Chem. 251, 5095-5 I03 Esfahani, M., Ioneda, T. & Wakil, T. J. (1971) J. Biol. Chem. 246, 5056 Fulco, A. J. (1972)J.Biol. Chem. 247,3511-3519 Fulco, A. J. & Fujii. D. K. (1979)fnr. Cong. Biochem. XIrh, Toronto Fujii, D. K. & Fulco, A. J. (1977)J.Biol. Chem. 252,3660-3670 Gelmann, E.P. & Cronan, E., Jr. (1972)J. Bacreriol. 112,381-387 Henning, U.,Dennert, G., Rehn, K. & Deppe, G. (1969)J. Bacteriol. 98,784-796
Hildebrand, J. H. (1971)Science 174,490493 Holub, B. J. & Lands, W. E. M. (1975)Can. J. Biochem. 53. 12621277 Jezyk, P. & Lands, W. E. M. (1968)J.LipidRes. 9,525-531 Jost, P., Griffith, 0. H., Capaldi, R. A. & Vanderkooi, G. (1973) Biochim. Biophys. Acra 311,141-152 Kasai, R., Kitajima, Y., Martin, C. E., Nozawa, Y., Skriver, L. & Thompson, G. A., Jr. (1976)Biochemistry 15,5228-5233 Kito, M.,Ishinaga, M.,Nishihara, M., Kato, M., Sawada, S. & Hata, T. (1975)Eur. J. Biochem. 54,55-63 Lakowicz, J. R., Prendergast, F. G. & Hogen, D. (1979)Biochemistry 18,508-519 Lands, W. E. M., Ohlaogge, J. B., Robinson, J. R., Sacks, R. W., Barve, J. A. & Gunstone, F. D. (1977)Biochim. Biophys. Acta 486, 451461 Lands, W. E. M., Sacks, R. W., Sauter, J. & Gunstone, F D. (1978) Lipids 13,878-886 Linden, C. D., Wright, K. L., McConnell, H. M. & Fox, C. F. (1973) Proc. Natl. Acad. Sci. U.S.A. 70,2271-2275 Marr, A. G. & Ingraham, J. L. (1962)J. Bacteriol. 84,1260-1267 Martin, C. L.. Hiramitsu, K., Kitajima, Y.,Nozawa, Y., Skriver, L. & Thompson, G. A., Jr. (1976)Biochemistry 15,5218-5227 Nishihara, M., Ishinaga, M., Kato, M. & Kito, M. (1976) Biochim. Biophys. Acta 431,5661 Ohlrogge, J. B., Barber, E. D., Lands, W. E. M., Gunstone, F. D. & Ismail, I. A. (1976)Can. J. Biochem. 54,736-745 Okuyama, H., Yamada, K., Kameyama, Y., Ikezawa, H., Akamatsu, Y . & Nojima, S. (1977)Biochemistry 16,2668-2673 Polacco, M. L. & Cronan, J. E., Jr. (1977)J. BioL Chem. 252, 54885490 Shaw, M. K.& Ingraham, J. L. (1965)J.Bacteriol. 106,449-455 Shimshick, E. J. & McConnell, H. M. (1973)Biochemistry 12, 235 12359 Silbert, D. F., Ladenson, R. C. & Honegger, J. L. (1973) Biochim. Biophys. Acra 31 I, 349-361 Silvius, J. R. & McElhaney, R. N. (1978)Nature (London) 272, 645647 Sinensky, M. (1971)J. Bacteriol. 106,449-455 Sinensky, M. (1974)Proc. Narl. Acad. Sci. U.S.A. 71,522-525 Strittmatter, P. & Rogers, M. J. (1975)Proc. Natl. Acad. Sci. U.S.A. 72,2658-266 1 Tsao, Y . K. & Lands, W. E. M. (1979)Science, in the press Warren, G. B., Toon, P. A., Birdsall, N. J. M. & Metcalf, J. C. (1974) Biochemistry 13,5501
The control of phosphatidylinositol turnover m cell membranes ROBIN F. IRVINE and REX M. C. DAWSON Biochemistry Department, A.R.C. Institute of Animal Physiology,Babraham, Cambridge CB2 4A T, U.K. A rapid and apparently specific turnover of phosphatidylinositol occurs in many tissues, which increases in response to physiological stimulation (see Mitchell, 1975; Fain & Berridge, 1979, for references). The percentage of cellular phosphatidylinositol involved in the heightened turnover appears to be minimal, although a net loss of the phospholipid can be observed in certain circumstances. This points to an enzyme system (or systems), specifically designed to hydrolyse this phospholipid, which is (or are) very tightly controlled. A variety of functions for stimulated phosphatidylinositol turnover have been suggested (for example, Michell, 1975; Hawthorne & Pickard, 1979; lrvine & Dawson, 1979), but they are as yet unproven. As a means of elucidating the physiological role of phosphatidylinositol turnover, we have investigated the ways in which animal cells can catabolize this phospholipid and how this catabolism may be controlled; we shall therefore here consider these various catabolic enzymes and their properties in uirro (see Fig. 1). First there is a Ca2+-independentphosphatidylinositol phosphodiesterase that is lysosomal in location (Irvine et al., 1977, 1978). We have discussed elsewhere the properties of this Vol. 8
enzyme and the possible role it may play in stimulated phosphatidylinositol turnover in brain (Irvine & Dawson, 1979) and some other tissues (Richards el al., 1979). Suffice to say that where stimulated turnover of the phosphoinositol moiety of phosphatidylinositol occurs in association with increased catabolism of other phospholipids, there is a strong possibility of increased lysosomal activity being responsible. Lysosomes also have a phosphatidylinositol deacylating system. To our knowledge these are the only enzymes deacylating phosphatidylinositol assigned to a specific cellular location. White et al. (1971) have described a phosphatidylinositol total-deacylating activity in homogenates of guinea-pig pancreas with a pH optimum around 6.0. We have found a similar activity in sheep pancreatic juice with a more acid pH optimum (5.0) and with no requirement for Ca2+ (R. M. C. Dawson & R. F. Irvine, unpublished work). These secreted enzymes are therefore very similar to those in liver lysosomes and may represent a secretion by specialized pancreatic 1ysosomes. Despite the ability of lysosomes to deacylate phosphatidylinositol, the major route of phosphatidylinositol degradation by rat liver lysosomes either given a membrane or pure phospholipid substrate is by the phosphodiesterase to liberate phosphoinositol rather than deacylation to give glycerophosphoinositol (Irvine et al., 1978; Richards et al., 1979). The
