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I ,IIII ‘I II I,I II II III‘I'I’III IIII III IIIII'IIIIIII" W IIII'II‘I'I‘IIIII IIIIIIIIIIII'IIIIIIIIIIIIIIII‘ IIIIII IIIIIIIIIIIII IIIIIII IIIIIIIII‘I‘ PART I . PART II. This is to certify that the thesis entitled ISOLATION AND PARTIAL CHARACTERIZATION OF AN INACTIVATING FACTOR FOR FATTY ACID SYNTHETASE ISOELECTRIC FOCUSING IN DENSITY GRADIENTS IN THIRTY-FIVE MINUTES presented by GARY ARDEN NEUDAHL has been accepted towards fulfillment of the requirements for M.S. degree inBIOCHEMISTRY fixfl 1/1442: ($274160; Major professor Date QLKQLZ /Z7ZC/‘ PALMITATE + 14 NADP+ + 8 CoA + 7 co2 + 6 H20 . There is no evidence that the enzymes catalyzing this reaction are as- sociated in yigg, although a specific acyl carrier protein (ACP) has been localized near the surface of the cell membrane (Volpe and Vagelos, 1975). This acyl carrier protein and six individual enzymes (Goldman g£,gl., 1961; Lennarz gg‘gl., 1962) are required for the overall reac- tion. Each of these enzymes, when purified, catalyzes a specific reac- tion in the biosynthetic sequence. The acyl carrier protein has a cen- tral role in the overall reaction in that all intermediates are cova- lently bound to it through its prosthetic group, 4'-phosphopantetheine. The enzymes involved in the overall fatty acid-synthetic reaction are: (l) scetyl-CoA-ACP transacylase (Alberta 55 31., 1964), which catalyzes the transfer of the acetyl group from CoA to ACP, (2) malonyl-CoA-ACP transacylase (Alberta st 21., 1964), which catalyzes the transfer of the malonyl group from CoA to another molecule of ACP, (3) B-ketoacyl ACP synthetase (Alberta 35 31., 1965), which catalyzes the synthesis of B-ketoacyl thioesters of ACP from malonyl-ACP and fatty acyl-ACP with concomitant loss of C02 and release of free ACP, (4) B-ketoacyl ACP re- ductase (Majerus 23 31., 1965), which catalyzes the NADPH-dependent re- duction of B-katoacyl-ACP to B-hydroxyacyl-ACP, (5) B-hydroxybutyryl- ACP dehydratase (Majerusletigl., 1965, 1969), which catalyzes the dehy- dration of short chain B-hydroxyacyl-ACP to 5533343,B unsaturated acyl- ACP, and (6) enoyl-ACP reductase (Weeks and Wakil, 1968), which cats- 4 1yzes the NAD(P)H-dependent reduction of enoyl-ACP to fatty acyl-ACP. Many of these reactions have been studied using model substrates (Lynen, 1961; Trams and Brady, 1960) for the physiological ones, in- cluding acylthioester derivatives of CoA, pantetheine, and N-acetyl- cysteamine. Some of the enzymes have very rigid specificities, so the model substrates could not be used (Yang 35 31., 1967). Since 4'-phos- phopantetheine is a component of both ACP and CoA, the importance of the apoprotein structure of ACP in confering increased reactivity to the bound intermediates is clear (Majerus, 1967). Model substrate and other studies on these enzymes have shown that a sulfhydryl residue on ACP, other than that of the 4'-phosph0pantetheine group, is involved in both B-ketoacyl-ACP synthetase (Vagelos 35 31., 1969) and acetyl-CoA-ACP transacylase (Williamson and Wakil, 1966) reactions. Type III fatty acid synthetases are microsomal or mitochondrial mem- brane-associated fatty acid elongation systems (Bloch and Vance, 1977). The microsomal enzymes modify fatty acids formed by Type I enzyme (see below) and also modify the essential unsaturated fatty acids (Nugteren, 1965; Seubert and Podack, 1975). The desaturation (Marsh and James, 1962) and elongation (Guchhait 2E 31., 1966) functions performed by these enzymes yield polyunsaturated fatty acids such as arachadonic acid. Brain is a particularly good source of these Type III fatty acid synthe- tases; it has at least three separate Type III FASs which are apparently involved in myelinogenesis (O'Brien, 1965). The mitochondrial fatty acid synthetases elongate short chain-length fatty acids by the reverse of B-oxidation, except an enoyl-CoA reductase replaces acyl-CoA dehydrogenase (Seubert and Podack, 1975; Wakil, 1961; Hinsch 35 31., 1976). These enzymes are most active when the 12.2222. ATP level is high and the oxygen level is low (Hinsch and Seubert, 1975; Whereat, 1971). The type 1 fatty acid synthetases are those which contain multiple catalytic activities on single polypeptide chains (Stoops £3 21., 1978; Mattick 35 51., 1981). They are found in mammals, birds, insects (Mun- icio st 31., 1977), yeast, and some fungi (Giompres and Packter, 1978) and bacteria (Kawaguchi and Okuda, 1977). The fatty acid synthetase complex from yeast (Lynen, 1961) was the first of the Type I enzymes to be extensively studied. The reactions required for overall fatty acid- 5 synthetic activity are identical to those catalyzed by the dissociable Type II enzymes, except the acyl intermediates in Type I fatty acid synthetase remain bound to a single 4'-phosphopantetheine group within the complex until synthesis of the fatty acid is completed. The "acyl carrier protein" of Type I fatty acid synthetase is thus an integral part of a multifunctional polypeptide chain in the complex. Also, in yeast, the synthesized fatty acids are released as acyl-CoAs, rather than as free fatty acids. The yeast fatty acid synthetase complex is a high molecular weight (2.5 x 106) substance of subunit composition 0656 and overall dbmen- sions 21 x 25 nm (Lynen, 1969). The nonidentical subunits have been shown by genetic mapping to be coded by two unlinked gene loci desig- nated £331 and £332 (Schweizer and Bolling, 1970; Schweizer 35 31., 1975; Kuhn 35 51., 1972; Knobling £5 31., 1975). The £331 locus codes for the 8 subunits, which has enoyl reductase, dehydratase, acetyl trans- ferase, and malonyl (palmitoyl) transferase activities. The £352 locus codes for the a subunit, which has condensing enzyme (B-ketoacyl synthe- tase) and B-ketoacyl reductase activities and the acyl carrier protein (Schweizer £5 31., 1975). The assignment of activities to one or the other subunit has been verified by protein chemical studies (Wieland ,g£.gl., 1979; Engeser 22.21:: 1977; Kresze gg‘gl., 1976). The molecular weights of the a and 6 subunits are 215,000 and 205,000, respectively (Stoops and Wakil, 1978). The complex will utilize NADH in the reduc- tive steps, but NADPH is preferred; also, FMN serves as a hydrogen car- rier in the reaction catalyzed by the enoyl reductase activity (Lynen, 1969). The complex has optimal activity at pH 6.5 to 7.0. The Km values for acetyl-GOA, malonyl-GOA, and NADPH are 28, 8, and 67 uM, re- spectively. Many of the partial reactions,which together comprise the overall fatty acid-synthetic activity, have been studied using model substrates (Lynen, 1961, 1962, 1967), and the structural-functional re- lationships in the complex have been examined in detail (Lynen, 1980; Stoops and Wakil, 1981a). Type I fatty acid synthetases from mammalian and avian sources differ somewhat from those of yeast and various bacterial sources: (1) the overall fatty acid-synthetic reaction is the same as for the Type II enzyme; that is, palmitate and other free fatty acids are produced, 6 rather than acyl-CoAs, (2) FMN is not utilized by the complex, (5) NADPH specificity is more stringent, (4) the complex has subunit structure a2 or aEB, and (5) there is a palmitoyl (long chain acyl) transacylase ac- tivity separate from the malonyl transacylase activity. Genetic studies similar to those performed with yea-st have not been possible with mammalian and avian fatty acid synthetases, so identity or nonidentity of the two subunits which comprise the native FAS complex in animal tissues has instead been suggested on the basis of physica- chemical and catalytic properties of the subunits. Messenger RNA for fatty acid synthetase is of sufficient size to code for complete subunits (Lau st 31., 1979; Nepokroeff and Porter, 1978), and in cell-free trans- lation systems it produces proteins of the apprOpriate size and immuno- logical characteristics (Flick st 31., 1978; Mattick 35 31., 1981). Two types of subunits have been obtained from pigeon liver and have been separated and characterized (Lornitzo st 31., 1974, 1975). Each subunit catalyzed some, but not the same, partial reactions; this suggested that the subunits were different. The evidence for nonidentical subunits in fatty acid synthetase complexes from other tissues and species has been based on finding only one mole of 4'-phosphopantetheine per mole of com- plex (Smith and Abraham, 1975a; Yun and Hsu, 1972; Nepokroeff gg‘gl., 1975). Other studies have given 522 moles of 4'-phosphopantetheine per mole of complex, however (Paskin and Mayer, 1976; Albertslgg‘gl., 1975; Buckner and Kolattukudy, 1976; Hardie and Cohen, 1978), and have sug- gested that limited proteolysis of identical subunits is responsible for both the low 4'-phosphopantetheine content and differing subunit activities reported by some workers (Stoops 35 31., 1978). In any case, the subunits, if not identical, are of very similar molecular weight and amino acid content for complex from any given source. Also, the subunits cannot be genetically coded in the same manner as those of yeast, as both subunits lack B-ketoacyl synthetase activity; this activity re- quires domains of both subunits (Kumar gg 31., 1970; Stoops and Wakil, 1981b). Mammalian and avian fatty acid synthetases have been isolated from a large number of species and tissues (Bloch and Vance, 1977), including red blood cells (Jenik and Porter, 1981), pigeon liver (Ken 35 31., 1969), chicken liver (Arslanian and Wakil, 1975), rat liver 7 (Nepokroeff £3 21., 1975), pig liver (Kim:££.El-: 1977a), lactating rat mammary gland (Smith and Abraham, 1975b), and rabbit mammary gland (Dils and Carey, 1975; Hardie and Cohen, 1978). These complexes all have molecular weights between 488,000 and 555,000 in their native forms. The specific activities of the purified enzymes, although determined under varied conditions, are all between 200 and 1400 nmoles NADPH oxi- dized per minute per milligram protein, with optimal pH for the overall activity assay between 6.5 and 7.0. Substrate Km values for the com- plexes are between 2.5 and 25, 10 and 50, and 20 and 70 “M for acetyl- CoA, malonyl-GOA, and NADPH, respectively. The major product of fatty acid synthesis is palmitic acid for all except the mammary gland enzyme, which also produces large amounts of butyryl-CoA (Strom 25.31., 1979), apparently the result of high catalytic efficiency in the reduction of acetoacetylfCoA and trans-crotonyl-COA (Kumar and Dodds, 1981). All the complexes are malonyl-GOA dependent, but some prefer butyryl-CoA over acetyl-GOA as a primer (Maitra and Kumar, 1974; Lin and Kumar, 1972: Smith and Abraham, 1971). Methylmalonyl-CoA was viewed by most workers as a competitive inhibitor of fatty acid synthetase (Frenkel and Kitchens, 1977), but when it replaces malonyl-00A in an assay, branched chain fatty acids are formed, although at a slow rate (Buckner gt 31, 1978). Most studies using methylmalonyl-CoA as a substrate were performed with goose urOpygial gland fatty acid synthetase (Buckner and Kolattukudy, 1976). Several mechanisms for control of fatty acid synthetase activity have been proposed. Although acetyl-GOA carboxylase was originally thought to be the critical regulatory enzyme in fatty acid biosynthesis, it now seems likely that under certain dietary conditions (Porter 23 21., 1971; Guynn 35 31., 1972; Butterworth 33 31., 1966; Burton 35.31., 1969), and especially in developing tissues (Vagelos, 1974), fatty acid synthetase is the rate limiting enzyme (Chang 55 31., 1967; Smith and Abraham, 1970; Volpe and Kishomoto, 1972). Indeed, the capacity of developing mouse liver to synthesize fatty acids from acetate correlates best with changes- in fatty acid synthetase activity (Ahmad £5 31., 1979). Allosteric regulators of Type I fatty acid synthetases are unknown at this time. Because these enzyme complexes have bound intermediates, it is unlikely that any one step is rate—limiting (Bloch and Vance, 1977); the overall fatty acid-synthetic rate is not dependent on the Km.va1ues of individual intermediates, but rather on the rate of 8 transfer between sites (Bloch, 1977). Nevertheless, reagents external to the complex can affect product distribution and the overall reaction rate. Lower malonyl-GOA to acetyl-GOA ratios yield shorter chain- 1ength fatty acids (Smith and Abraham, 1975a, 1975b; D113 and Carey, 1975), as do specific chain length modifying proteins from mammary tis- sue (Smith and Abraham, 1975a; Carey, 1977). Palmitoyl-CoA inhibits the complex (Hsu gg'g1., 1969; Tubbs and Garland, 1965; Lynen £5 21,, 1964; Lust and Lynen, 1968; Robinson 31 21., 1965), apparently by a nonspecific detergent-like denaturation of the complex (Dorsey and Porter, 1968; Taketa and Pogell, 1966). Recently CoA was shown to ac- tivate fatty acid synthetase at very low concentrations, so low that the activation was not believed to be of physiological significance (Linn g._t_ 91., 1980). Control of the activity of fatty acid synthetase by covalent'modifi- cation of several types has been reported. Qureshi SE 31. (1975,1978) gave evidence for a phosphorylation/dephosphorylation control mechanism, but the presence of a phosphorylated form of the complex has not been verified (Rous, 1974; Hardie and Cohen, 1978). Also, excessive malonyla- tion of the enzyme has been shown to inactivate its overall activity (Kumar and Srinivasan, 1981). The complex has two catalytic sites for the binding of malonyl-GOA. When exposed to high levels of malonyl-GOA (more than 575 moles per mole complex), six moles of malonate are bound to the complex. Binding is apparently near a catalytic site, thereby inactivating the enzyme. Inactivation of fatty acid synthetase by this mechanism is not universal, however (Roncari, 1975). For at least some fatty acid synthetases (e.g., those from rat and pigeon liver), inter- conversion of the apo- (no 4'—phosph0pantetheine group) and holo- (co- valently attached 4'-phosphopantetheine) forms of the complex has been put forth as a mechanism for the 12_!132 control of enzyme activity. Indeed, the turnover rate of the 4'-phosphopantetheine group is more rapid than that of the remainder of the FAS complex (Tweto 35.31., 1971; Tweto and Larrabee, 1972). Also, the enzymes responsible for the trans- fer of this group from CoA to the apocomplex (Yu and Burton, 1974a,b) and for the hydrolytic release of this group from the holocomplex (Roncari, 1975; Kim 25 31., 1977b) have been partially characterized (Sobhy, 1977, 1979). 9 Another proposed mechanism for control of fatty acid synthetase ac- tivity involves dissociation of the complex into subunits. This has been demonstrated 12’31552, especially at temperatures around 4° (Muesing and Porter, 1975; Kumar 35 31., 1972; Yun and Hsu, 1972; Smith and Abraham, 1971b). Invariably, the purified enzyme is cold labile in low ionic strength solutions, especially in the absence of reducing agents (Strom and Kumar, 1979; Dils and Carey, 1975). There is no evidence for dissociated subunits 12:3132, although inactive FAS has been reported (Lion and Donaldson, 1977; Straweer and Larrabee, 1976). The long term (adaptive) control of mammalian and avian fatty acid synthetases 15:3112 (control by variation in the rates of protein syn- thesis and degradation) occurs through both dietary and hormonal channels. Dietary effects on FAS levels are greatest in liver, small in lung (Kumar, 1977), and nil in brain (Volpe and Vagelos, 1975). A diet high in saturated fatty acids lowers hepatic FAS levels (Romsos and Leveille, 1974), as does fasting (Lion and Donaldson, 1973: Allmann and Gibson, 1965). The lower levels of FAS are the result of decreased translation rather than an increased degradation rate for the complex (Volpe and Vagelos, 1975; Craig 25 31., 1972; Fischer and Goodridge, 1978). The only dietary condition thus far shown to significantly affect the de- gradation rate of FAS is transfer from a fat-free to a linoleste-rich diet; the transfer causes a decrease in the half-life of rat liver FAS from 5.8 to 1.8 days (Flick 25 21., 1977). Upon refeeding with a fat- free diet following a fast, a sharp rise in the amount of fatty acid synthetase-messenger RNA, and then FAS peptide associated with poly- somes,is observed (Nepokroeff 55 31., 1979; Flick £5 31., 1978; Taylor and Schimke, 1974). After 48 hours, the maximal supranormal level of fatty acid synthetase activity is reached; the enzyme level then draps over about a week to normal levels (Alberta g£_g1,, 1975). Weaning of rat and mouse pups (Volpe and Kishomoto, 1972) and hatching of chicks (Zehner 55.31., 1977) increase liver FAS levels as the result of the involuntary change from a high fat diet to a high carbohydrate diet. Hormonal regulation of FAS levels 1g‘x1zg_is quite organ specific. Prolactin raises mammary gland FAS levels (Speake SE 31., 1976), and estrogen raises oviduct FAS levels (Aprahamian 35.21., 1980) with little effect on liver FAS levels. Prednisolone increases FAS levels in brown 10 fat but has no effect on hepatic FAS levels (Hahn 35 31., 1978). Ad- .9 renal steroids in general lower adipose tissue FAS levels, but have no effect on liver FAS levels (Flick 31.21., 1977). Insulin, the most wide- ly studied hormonal effector of FAS levels, increases FAS levels in liver, especially upon administration to diabetic (naturally or chemically in- duced) animals or their cultured explants (Joshi and Aranda, 1979; Nepokroeff 31,31., 1979; Lakshmanan 31 31., 1972; Dahlen 22 31., 1968; Gibson and Hubbard, 1960). The absolute requirement for insulin in raising fatty acid synthetase levels 12:3132 has been questioned (Kukulansky and Yagil, 1979). FERRITIN Iron is a required nutrient for cells in almost all phyla (Munro and Linder, 1978). Because ferrous ion in aqueous solution at physiologi- cal pH tends to oxidize and form insoluble ferric hydroxide or oxyhy- droxide, and then precipitate, a means for intracellular storage and transport of iron is needed. Although low molecular weight polyphos- phates have been implicated in the small scale transport and storage of iron (Jacobs, 1977; Konopka and Romslo, 1980; KonOpka 25,31., 1981), the protein ferritin has been found to be the major intracellular iron storage depot. Ferritin maintains the iron for cellular metabolic needs in a soluble and nontoxic form. In addition to its function as an iron storage protein, it participates directly in the transfer of iron from plasma membrane to the mitochondria (Ulvik 31 31., 1981; Mazur and Carleton, 1965; Nunez 21 21,, 1980; Ulvik and Romslo, 1978, 1979), where the iron is transferred to a ferrochelatase (Ulvik and Romslo, 1981) and then inserted into protoporphyrin IX to complete heme biosynthesis (Ulvik, 1981) . The first preparation of an iron-containing protein was obtained from pig liver. This poorly defined, denatured "ferratin" contained about 5% iron by weight (Schmiedeberg, 1894). It was in deference to this "fer- ratin" that the first pure iron-containing protein, isolated from horse spleen, was named. Horse spleen ferritin (Laufberger, 1957) is the best characterized of the subsequently isolated ferritins. Among the other tissues and organisms from which ferritin has been isolated are 11 Escherichia coli (Yariv 35.31., 1981), fungi (David and Easterbrook, 1971; Bozarth and Goenaga, 1972), legumes (Crichton 35 31., 1978), ‘ plants (Seckbach, 1972), dog, cat, jackal, Guinea pig, rabbit, tuna- fish, dolphin, and human spleen (Kuhn 25 31., 1940; Granick, 1945; Muraoka 55,31., 1966; Kato 35.31., 1968; Kato and Shimada, 1970; Wag- staff 25 51., 1978), rat and human heart (Valimiri 35 21., 1977; Wag- staff 35 a_l., 1978), and amphibian red blood cells (Thiel, 1973). A summary of the relative occurrences of ferritin in various animal tis- sues has also been published (Michaelis, 1947). Ferritin consists of an approximately spherical hollow shell of pro- tein with a core containing up to 4500 iron atoms (Crichton, 1975). The core consists of iron in the same crystalline form as the mineral ferri- hydrite (5Fe203-9H20), with apprOpriately adsorbed phosphate (Towe, 1981; Chukhrov g£_g1,, 1975; Harrison, 1965; Michaelis 35 11., 1945). The iron is present.in a layered structure with flattened octahedral arrangement of oxygen about each iron atom (Heald 25 21., 1979), with iron in the ferric state (Blaise 31.31., 1965). The phosphate is pri- ‘marily on the surface of the core (Granick, 1942; Fishbach 21 21., 1969) and typically constitutes l to 1.5% of the total mass of the core (Granick and Hahn, 1944), although as the core increases in size, the relative amount of phosphate present decreases (Treffry and Harrison, 1978). The roughly spherical protein shell about the core consists of 24 subunits, arranged such that the shell belongs to the F452 space group (1980b). In mammalian ferritin, the shell is about 2.4 nm thick and has an outer diameter of 12-15 nm and inner diameter of 6-7 nm. Six channels, located at the fourfold symmetry axes of the shell, provided for the entry and exit of small molecules about the interior (Banysrd ‘g£.g1., 1978). These channels are 0.9-1.2 nm wide at the outer surface of the shell and 1.7-2.0'nm at the inner surface (Jones 25 31., 1978. The subunits comprising the shell are of two types (Linder 35.31., 1974; Urushizaki 21,21,, 1971) designated H and L (Russel and Harrison, 1978; Adelman 25 31., 1975; Ishitani 55 51., 1975). The respective molecular weights of these subunits are approximately 21,000 and 19,000 daltons, giving an overall molecular weight of the shell of 440,000 to 480,000 daltons, depending on the proportion of L and H subunits (Hoare 12 ‘35‘31., 1975; Banyard SE 31., 1978; Crichton and Bryce, 1970; Bjork and Fish, 1971; Crichton 55 31., 1975s; Drysdale gt; 51., 1975). These sub- units have distinct immunological properties despite their many common structural features. Each subunit is of cylindrical shape and consists of a bundle of four long helices lying parallel (or antiparallel) to one another, together with two much shorter helices (Clegg 55 31., 1980a) and a long 100p over the outside surface of the subunit which forms a short section of B-pleated sheet with the loop from a neighboring sub- unit (Clegg 35 31., 1980b). The L subunits have a more highly ordered secondary structure than the H subunits; the L subunit is about 50% or helix and the H subunit is about 50% ahhelix (Otsuka 35 51., 1981). In either case, the subunit conformations bear resemblance to those of myo- hemerythrin, hemerythrin, tobacco mosaic virus, and cytochromes b-562 and c' (Clegg 55 51., 1980b). The amino acid sequence of a subunit from horse spleen ferritin has been determined (Heusterspreute and Crichton, 1981). _ The ratio of L to H subunit in ferritin shells varies with species and tissue (Crichton 25 31., F975b; Msssover, 1978; Kohgo‘gg‘g1., 1980). Iron poor tissues, such as heart, intestine, and pancreas, have a lower ratio of L subunit to H subunit in their ferritin shells than iron-rich tissues (Bomford‘ggig1., 1981; Arosio 35151., 1978). Ferritin shells with higher L to H subunit ratios have been shown to have longer turn- over times in the cell, and also to have a lesser surface charge (Kohgo, 3531., 1980). Under physiological conditions, iron is transferred to and from far- ritin in the ferrous state, although it is stored in the ferric state (Niederer, 1970‘. Deposition of iron in the shell involves oxidation of ferrous ion to the ferric state followed by hydrolysis (Treffry and Harrison, 1980a; Macara 25 31., 1972; Crichton 25.31., 1977). Initia- tion of deposition apparently occurs as follows: two ferrous ions bind to catalytic sites on adjacent polypeptide chains, followed by bonding of dioxygen by the ferrous ions. The iron is oxidized to the ferric state, reducing the oxygen to a peroxo- complex. Incoming ferrous ion then displaces the peroxo- complex, resulting in the hydrolysis and de- position of the ferric ion (Crichton 25 31., 1980). After initiation of the core, iron is then deposited directly onto the preexisting core; consequently, the deposition rate is greatest in ferritin shells which 13 are half-filled, containing about 1400 atoms of iron (Ulvik 21,31., 1981). Certain carboxyl groups, located in residues 55—59 of the 174 amino acid subunit of horse spleen ferritin (wetz and Crichton, 1976; Vandamme, 1980) have been implicated in the binding of iron and cataly- sis of oxidation. The reductive mobilization of iron from ferritin 12 2132 by oxidoreductases in the cytosol and microsomes (Zaman and Verwil- ghen, 1977), by an NAD(P)H: flavin oxidoreductase (Sirivech gg‘g1., 1977), and by a mitochondrial ubiquinol-FMN oxidoreductase (Ulvik 21.31., 1981; Ulvik and Romslo, 1981) have been suggested. A variety of bio- logical reducing agents also release iron from ferritin; these include cysteine, ascorbate, glutathione (Dognin and Crichton, 1975) and di- hydroflavin (Jones 35 31., 1978; Sirivech 25 31., 1974). The apparent mediators of transfer of iron from ferritin in biological systems are flavins, since FMNH2 generated at the mitdchondria by s ubiquinol-FMN oxidoreductase seems to reduce iron in ferritin to the ferrous state, and then transport it to a ferrochelatase in the inner mitochondrial membrane (Jones and Jones, 1969; McKay SE 31., 1969). An increase in flow of iron into cells stimulates ferritin synthesis in all cells studied to date with the exception of lymphocytes (Lema and Sarcione, 1981). Ferritin levels in the cell can be increased up to ten-fold through the use of iron-rich media or diet (Hoy and Jacobs, 1981). Distinct messenger RNAs for the H and L subunit types (Watanabe and Drysdale, 1981b) are produced in disproportionate amount under iron- rich conditions: the greater the iron concentration, the greater the pro- portion of L subunit formed (Otsuka 21 31., 1981). Under normal dietary conditions, ferritin messenger RNA is the most common messenger RNA in polysomes and mRNP particles. It is present in sufficient quantity to be translated immediately upon entry of iron to the cell (Watanabe and Drysdale, 1981a). As iron concentration rises in the cell, the amount of polysome-associated ferritin messenger RNA increases even further (Zahringer gg‘g1., 1976) and translation of the ferritin messenger RNA occurs at an increased rate (Schaefer and Thiel, 1981). The ferritin produced, as well as having a higher L to H subunit ratio, also has a lower isoelectric point, apparently due to its greater incorporation of iron (Treffry and Harrison, 1980c; Harrison, 1959; Banyard 25 31., 1978; Bomford 55 31., 1981; Kohgo 51 31., 1980) rather than its higher L to H subunit ratio (Treffry and Harrison, 1980b; Hoy and Jacobs, 1981). CHAPTER 2 IMPROVED PARTIAL REACTION ASSAY AND PURIFICATION PROCEDURE FOR FATTY ACID SYNTHETASE ABSTRACT This chapter presents improved procedures for assay and for purifi- cation of fatty acid synthetase from pig liver. The decalone reductase activity assay used in this laboratory at the beginning of this work had limited applicability because of precipitation of the substrate, pggggg-l- decalone, at high salt and protein concentrations. Increasing the ethanol concentration and decreasing the 15233-1-decalone concen- tration in the assay now allows the assay to be used at any step in the purification of fatty acid synthetase. This assay is about fifteen times more sensitive than the overall activity assay. The procedure for the purification of fatty acid synthetase used in this laboratory at the beginning of these studies provided enzyme of high purity in good yield, but required five days for completion. Ad- dition of 5% isopropanol to the homogenization buffer, modification of the procedure for DEAR-cellulose chromatography, and elimination of the calcium phosphate gel treatment have significantly reduced the time and materials required for purification of the enzyme. The enzyme is now purified in three days without compromise of purity or yield. The purified enzyme is highly susceptible to air-oxidation. The oxidation is readily reversed by the addition of reducing agent to the enzyme. 14 15 INTRODUCTION Dutler 31,31. (1971) described several substrate analog model com- pounds specific for the 5-oxo-acyl carrier protein-reductase component of fatty acid synthetase. The Km and Vmax values for (9R)f£ESEE' decal-1,4-dione were 1.2 mM and 165 sec'l, respectively. The Km and Vmax values for (9R)-££gng-l-decalone were 0.17 mM and 51.5 sec'l, re- spectively. They used the (i) gasps-decal-l,4-dione dissolved in di- ethyleneglycol dimethyl ether to assay for the B-ketoacyl reductase com- ponent of fatty acid synthetase, apparently because of its higher Vmax value. A partial reaction assay for fatty acid synthetase using (i)- ‘EggEg-l-decalone was reported by this laboratory (Kim, Unkefer, and Deal, 1977). The commercially available Esgng-l-decalone was dissolved in absolute ethanol because, unlike diethyleneglycol dimethyl ether, ethanol is stable indefinitely. Although the value for Vmax 0f.£££22' l-decalone is only a third of that for Egggs-decal-l,4-dione, the deca- lone reductase assay for fatty acid synthetase is still about fifteen times more sensitive than the overall assay. The decalone reductase assay reported earlier by this laboratory worked well for relatively pure preparations of enzyme at low ionic strength. However, samples with high salt or protein content, e. g., crude liver fractions and fractions with added ammonium sulfate, could not be assayed for decalone reductase activity because 53353-1-decalone precipitated upon its addition to the incubation mixture. Part of this chapter describes an analysis of the precipitation problem and an im- proved assaying procedure which eliminates precipitation of the 55323-1- decalone. The purification procedure for fatty acid synthetase described by Kim, Neudahl, and Deal (1981) yielded high purity enzyme in good yield, but required a large investment of time (five long days) and gave enzyme which was not as stable as desired. Therefore, studies aimed both at obtaining enzyme that was more stable and at obtaining high purity en- zyme in good yield in a lesser period of time were performed. The re- sult was an improved procedure for purification of the enzyme. 16 MATERIALS AND METHODS Reagents. Acetyl-CoA, malonyl-GOA, and NADPH were obtained from P-L Biochemicals. Dithiothreitol was a product of Eastman-Kodak and £5322- l-decalone of Aldrich Chemical Co. Absolute ethanol was obtained from Aaper Alcohol and Chemical Co. Bovine serum albumin, gum arabic, brom- phenol blue, and Coomassie Brilliant Blue R were obtained from Sigma Chemical Co. Tannic acid was obtained from Baker, and DEAE-cellulose (standard, 0.91 meq/g) was obtained from Schleicher and Schuell. Am- monium persulfate, N,N,N',N'-tetramethy1ethylenediamine was a product of Canalco. Riboflavin was a product of Calbiochem, and 2-mercaptoetha- nol was a product of Mallinckrodt. All other reagents were of analytical reagent grade. 1132;. Pig livers were obtained from Peet Packing Company (Chesaning, MI) within an hour of slaughter and were frozen on dry ice. The livers were stored at -50°. Protein Determination. Protein was determined by the method of Massey and Deal (1975). The stock solutions were as follows (distilled, deion- ized water was used throughout): 1. Tannin reagent: Liquified phenol (2 ml, 88%) was dissolved in 98 ml of 1 M HCl. Tannic acid (10 g) was then dissolved in the solu- tion. The reagent was stored at 4°. Before use, the reagent was brought to room temperature and was stirred until any gum present had dissolved. After filtering through Whatman No. 1 filter paper, the reagent was ready for use. 2. Arabic reagent: Gum arabic (100 mg) was dissolved in 100 ml water. The reagent was stored at 4° and was brought to room temperature prior to use. 5. Protein standard: Crystalline bovine serum albumin (10.0 mg) and sodium chloride (900 mg) were dissolved in 100 ml water. This solution was stored at 4°. 4. Diluent: Sodium chloride (900 mg) was dissolved in 100 ml water. The protein determinations were carried out as follows: 1. A set of six 15 x 125 mm test tubes, which contained 0, 10, 20, 40, and 50 ug bovine serum albumin in 0.9% NaCl, were prepared. 2. Several dilutions of the unknowns were prepared, such that at least one dilution for each unknown had between 10 and 100 ug 17 protein per ml sample. The samples, of 0.50 ml volume, were placed in 15 x 125 mm test tubes. Estimates of protein concen- tration were made by measuring absorbsnce at 280 nm; an absor- bance of 1 represents a protein concentration of about 1 mg/ml. 5. Tannin reagent (0.50 ml) was added in a defined sequence to each tube at regular intervals of 45 seconds. Immediately after adding the tannin reagent, the solutions were thoroughly mixed. 4. Twenty minutes after adding tannin reagent, arabic reagent (1.00 ml) was added to each tube. The solutions were thoroughly mixed after addition of the arabic reagent. 5. The apparent absorbances of the turbid suspensions were measured at 650 nm. The sample containing only NaCl, tannin reagent, and arabic reagent was the reference solution. A standard curve was constructed using the absorbances of the protein standards. Pro- tein concentrations for the unknowns were obtained using this curve. Since this protein determination is a turbidimetric one, absorbance measurements should be made as soon as possible in order to minimize er- rors due to settling of the turbid suspension. The standard curve was linear between 5 and 50 mg protein per assay. Above 50 ug protein per assay, absorbsnce per ug protein rapidly decreases. For this reason, the testing of several different dilutions of unknown is required. Conductivity and pH Measurements. Conductivities of solutions were meas- ured using a Radiometer Gm2e conductivity meter with a CDC114 cell . The values of pH were determined at room temperature using a Radiometer PHM4c pH meter equipped with a GK2521C electrode. Native Po1yacry1amide Gel Electrophoresis. Native polyacrylamide gel electrophoresis was performed using the method of Clark and Switzer (1977) with minor modification. The solutions required for PAGE were the fol- lowing (Staining and destaining solutions were stored at room tempera- ture; ammonium persulfate solution was prepared fresh; all other solu- tions were stored at 4°; deionized, distilled water was used throughout): 1. Solution A: Tris(hydroxymethyl)aminomethane (56.68), 48 m1 of 1M HCl, and 0.25 ml TEMED were taken to 100 ml with water. The pH of this solution was approximately 8.9. 2. Solution B: Tris(hydroxymethyl)aminomethane (1.5 g), 11.7 ml of 1 M HCl, and 0.15 ml TEMED were taken to 25 ml with water. The pH of this solution was 6.9. 5. Solution C: Acrylamide (56.0 g) and N,N'-methylenebis(acrylamide) (1.47 g) were taken to 200 ml with water. The solution was fil- tered through Whatman No. 2 filter paper before use. 4. Solution D: Acrylamide (5.0 g) and N,N'-methylenebis(acrylamide) (1.25 g) were taken to 50 ml with water. The solution was filtered 18 through Whatman No. 2 filter paper prior to use. 5. Solution E: Riboflavin (1.25 mg) was dissolved in 25 ml water. 6. Solution F: Sucrose (40 g) was taken to 100 ml with water. 7. Solution G: Ammonium persulfate (1.12 mg/ml) was prepared and de- aerated by aspiration immediately prior to use. 8. Electrophoresis buffer: Tris(hydroxymethyl)aminomethane (6.0g) and glycine (28.8 g) were taken to one liter with water. This buf- fer is diluted ten-fold with water prior to use. 9. Tracking dye solution: Bromphenol blue (0.5 mg) was dissolved in 5 ml water and 5 m1 glycerol was then added. 10. Staining solution: Coomassie Brilliant Blue R (1.5 g) was dis- solved in a solution containing 265 ml 95% ethanol, 50 m1 glacial acetic acid, and 255 ml water. The staining solution was reused indefinitely. ll. Destaining solution: Glacial acetic acid (280 m1) and 95% ethanol (500 ml) were taken to four liters with water. Gel tubes were prepared by cutting 7 mm (o. d.) Pyrex tubing into 15 cm lengths and then fire-polishing both ends. Marks were etched 10 cm for one end of the tubes. These markings indicate the filling positions for the resolving gel solution. Polyacrylamide gels were formed one to seven days prior to electro- phoresis. The tubes were sealed at the ends opposite the markings with Parafilm and were mounted vertically. Resolving gel solution (abOut 1.8 ml per tube) was then prepared on ice by mixing one volume Solution A, two volumes Solution C, and five volumes Solution G. The gel tubes were filled to the etched markings using a Pasteur pipet. The resolving gel solution was overlayered with about 0.1 ml water. During polymerization of the resolving gel, the stacking gel solution was prepared. One volume Solution B, two volumes Solution D, one volume Solution E, and four volumes Solution F were placed in an aluminum foil- wrapped test tube. A volume of 0.2 m1 of the resulting solution is re- quired for each tube. After polymerization of the resolving gel, the water overlayer was re- moved using a Pasteur pipet. Stacking gel solution (0.2 ml per tube) was then applied. Again, a water overlayer was applied; this overlayer promotes the formation of a sharp, flat boundary upon polymerization of the gel solutions. The tubes were placed under a fluorescent lamp until the stacking gel polymerized. Immediately before a PAGE experiment, the water overlayer was removed 19 from the gels. After moistening the tops of the tubes with glycerol, the tubes were inserted in the electrophoresis apparatus. The two reservoirs and the space in the tubes above the stacking gels were then filled with the ten-fold diluted electrophoresis buffer. Samples were prepared for electrophoresis by adding at least a tenth- volume of tracking dye solution. Sample size was limited to about 200 ul by the dimensions of the gel tubes. Removal of salt from the samples prior to electrophoresis was not required, but high salt concentrations increased the amount of time required for electrophoresis. The samples (with tracking dye) were applied just above the stacking gels using an Eppendorf pipet. After all samples were applied, the chambers were attached to a Heath- kit Model IP-52 regulated power supply. The negative lead was attached to the upper reservoir, the positive lead to the lower reservoir, and a 200 volt potential was applied. Electrophoresis was continued until the dye band was about 5 mm from the bottom of the resolving gels. After turning off the power supply, the tubes were removed from the apparatus. Gels were removed from the tubes using a stream of water pointed at the bottom of the tube, between the gel and the wall of the tube. The stream of water was produced by an 18 gauge needle attached to a water faucet by a length of Tygon tubing. The gels were placed in 16 x 150 mm screw- cap culture tubes containing staining solution. Until the staining solu- tion had been used eight times, gels were stained for about two hours; later, overnight staining was used. After staining, the staining solution was removed and the gels were rinsed with distilled water. Destaining solution was then added to the tubes, and the tubes were left on a mechanical shaker overnight. The destaining solution was then discarded and fresh destaining solution was added to the tubes. The destaining solution changes were continued un- til stain was no longer released from the gels. Assays for Fatty Acid Synthetase. The assays described below (Kim, Un- kefer, and Deal, 1977) were the standard assays for fatty acid synthe- tase in use in this laboratory when this work was begun. They therefore were the starting point for the development of improved assays. Initial Standard Decalone Reductase Assay. This partial reaction as- say was routinely used in partially purified preparations of fatty acid 20 synthetase. The stock reagents for this assay (final concentrations in parentheses) are given below: potassium phosphate, 0.2 M, pH 6.10 (0.1 M) trans-l-decalone, 200 mM, in ethanol (10 mM) NADPH, 2 mM, in 1 mM NaOH (50 M) enzyme (about 4 ug/ml) The reaction was carried out at 25°. It was initiated by the addi- tion of Eggng-l-decalone. The final assay volume was 0.4 ml. Activity values were measured from the initial slopes of the absorbance curves. One unit of activity was the amount of enzyme that catalyzed the oxida- tion of one meI NADPH per minute. Initial Standard Overall Assay. The overall reaction assay was used primarily to determine the specific activity of purified fatty acid syn- thetase. The stock reagents for this assay (final assay concentra- tions in parentheses) are given below: potassium phosphate, 0.4 M, pH 6.89 (0.1 M) acetyl-00A, 2 mM (50 uM) malonyl-CoA, 2 mM (50 an) NADPH, 4 mM, in 1 mM NaOH (200 an) bovine serum albumin, 20 mg/ml (1 mg/ml) enzyme (about 60 ug/ml) The reaction was carried out at 25°. It was initiated by the addi- tion of malonyl-GOA. The final assay volume was 0.4 m1. Activity values were measured from the initial slopes of the absorbsnce curves. Activity units are as described for the decalone reductase assay. Spec- ific activity is expressed as units per milligram of protein. RESULTS The Decalone Reductase Assay. The initial standard decalone reduc- tase assay was about fifteen times more sensitive for fatty acid synthe- tase than the overall assay. This sensitivity would make it the pre- ferred assay for crude preparations of the enzyme; smaller sample sizes would be required, and any contaminating enzyme activities would be pro- portionately reduced. Also, the decalone reductase assay required fewer and less costly reagents than the overall assay. Unfortunately, £5235- 1-decalone precipitated when samples with high concentrations of protein or of salt were assayed for decalone reductase activity. Studies aimed at extending the usefulness of the decalone reductase assay to samples 21 of high protein or of high salt concentration were therefore undertaken. Development of an Improved Assay. Apparently negative activities were obtained when the initial standard decalone reductase assay was used with samples of high salt or protein concentration. The cause was the addition of trans-l-decalone, which produced precipitation (Tablel). Table 1. Determination of the Optimal Concentrations of trans-l-decalone and Ethanol in the Decalone Reductase Assay for Samples Containing Up To 1.4 M Ammonium Sulfate. igithzlcgsztszgs Sample Solution Added Precipitation?b (210 I1) (50 :11) (1+0 L11) 1004, NADPH, n20 H20 200 mM TDEC in ethanol yes KPO4, H20 H20 200 mM TDEC in ethanol yes KPO4, NADPH, H20 H20 ethanol no KPO4, H20 H20 ethanol no 1004, NADPH, H20 H20 100 mM TDEC in 95% ethanol yes 100,, NADPH, 1120 ago 100 mM TDEC in ethanolc no KPO4, NADPH, H20 1.4 M (1~m.,,)zso4 100 mM TDEC in ethanol yes 1004, NADPH, H20 1.4 M (1411412304 so mM TDEC in ethanold no aConcentrations of NADPH and potassium phosphate buffer (KP04) were as indicated in Materials and Methods for the decalone reductase assay. bPrecipitation was measured as an increase in apparent absorbence at 540 nm upon addition of the substrate (ethanol-containing) solutions; absorbance normally decreases as the enzyme oxidizes NADPH. cThis is a previous stock solution of trans-l-decalone (TDEC) for the assay. dThis is the present stock solution of trans—l-decalone for the assay. Addition of ethanol alone to the assay mixture did not cause precipita- tion. The precipitation upon addition of the Egggg-l-decalone stock so- lution was prevented by increasing the ethanol concentration in the as- say mixture to 10%. This was accomplished by dilution of the 15333-1- decalone stock solution to 100 mM Eggng-l-decalone in ethanol. Precipi- tation again occurred, however, when samples other than water were used in this modified assay Two alternatives were considered for the elimination of the precipi- 22 tation: increasing the ethanol concentration, or decreasing the 15222-1- decalone concentration. Since the Km for Egggs-l-decalone is 0.17 mM (Dutler 25,31., 1971), the concentration of Erggg-l-decalone in the as- say was reduced. At concentrations of up to 5 mM EEgEs-l-decalone (with 10% ethanol) in the assay, one-eighth volume of 0.55 saturated ammonium sulfate (the highest ionic strength reached in the purification of fatty acid synthetase) did not induce precipitation (Table 1 ). Consequently, a stock solution of 50 mM Eggng-l-decalone in ethanol was selected for the assay. This stock solution yields final concentrations of 5 mM Eggng-l-decalone and 10% ethanol in the assay. Enzyme Concentration and Activity. Proportionality tests between enzyme concentration and reaction rate in both the decalone reductase and overall assays were performed. The overall activity of the enzyme was linearly proportional to enzyme concentration over at least a ten- fold range of enzyme concentration. This proportionality was retained even when bovine serum albumin was eliminated from the assay and the NADPH concentration in the assay was reduced to 100 mM. For this reason, the present standard overall activity assay has the following final con- centrations of reagents in the assay: potassium phosphate, 0.1 M (pH 6.89) acetyl-GOA, 50 uM malonyl-00A, 50 “M NADPH, 100 “M enzyme (about 0.006 units/ml) All other details of the assay are as for the initial standard overall assay. The specific activity of fatty acid synthetase as measured by decalone reductase activity increased significantly with enzyme concentration up to 0.015 units of activity in the assay (Figure 1). Above this level, activity is proportional to the concentration of enzyme over more than a two-fold range of enzyme concentration. The decreasing specific ac- tivity at lower enzyme concentrations is apparently due either to dis- sociation of the enzyme into inactive (or less active) subunits or to inactivation of the enzyme by Eggggfl-decalone. The improved decalone reductase assay has the following stock reagents (final concentrations in the assay in parentheses): -potassium phOSphate, 0.2 M, pH 6.10 (0.1M) -trans-l-decalone, 50 mM, in ethanol (5 mM) 23 .mxmmmm one am pom: coca ouo3 cowusaom menu «0 moesfio> maceum> "Ao~.® :AV acumen oumsnmosn Enwmmmuoa z m~.o cums pfiowicou pounfiao ems season are mo :Owus~0m xooum < .ommu nocucxm pwom zoumm commence mo meanest osowum> momma Ou mom: was mudsmom mo wcwacawon one up poucomoua zmmmm ommuonwou ocofimooo pumpcmum one .momm<.wuw>wuo< ommuonpom ocodmooa one wean: cogumuucoocoo oEchm mo :oHuocsm o no oomuocucxm owo< awash mo zuw>wuo< owwwoomm .d ounwfim 24. Figurel. .9 Ta . C C *2 C ‘ a on m to C is m < Li. Jun . EMX (IV/51W”)- MIMIQV OHIOHdS 25 -NADPH, 2 mM, in 1 mM NaOH (so on) -enzyme (0.04-0.12 units/m1) All other details of the assay are as for the initial decalone reductase assay. FattygAcid Synthetase Purification. Purification of fatty acid syn- thetase according to the method of Kim, Unkefer, and Deal (1977) re- quired about five days and yielded enzyme which was not as stable as desired. It was thought that the instability of the enzyme might be due to proteolysis during its purification. Therefore, the enzyme was puri- fied in the presence of a protease inhibitor, phenylmethanesulfonyl- fluoride. This compound is not water soluble, so it was dissolved in isopropanol. A stock solution of 20 mM PMSF in isOpropanol was prepared, yielding concentrations of 1 mM PMSF and 5% isopropanol throughout the enzyme purification. These substances stabilized the decalone reductase activity in the early stages of the purification. It also substantially decreased the amount of contaminating protein precipitated with fatty acid synthetase in the first ammonium sulfate fractionation step of the purification procedure. To determine whether these effects were due to PMSF, parallel purifications of fatty acid synthetase were performed in the presence of PMSF and isopropanol, or isopropanol alone. IsoprOpanol proved responsible for the effects noted above. So 5% isOpropanol was incorporated into the homogenization buffer. The initial method for purification of fatty acid synthetase included a DEAE-cellulose chromatography step which required about 18 hours to complete. By replacing this step with DEAE-cellulose pad chromatography, using stepwise rather than gradient elution, the time required for chrom- atography was reduced to three hours. Elimination of the calcium phos- phate gel treatment further reduced the amount of time required for puri- fication of the enzyme. These changes in the purification procedure decreased the time required for preparation of the enzyme to three days, without effect on overall yield or purity. The details of this improved purification procedure are presented below. Improved Purification Procedure for Fatty Acid Synthetase. Unless otherwise indicated, centrifugation was performed in the Sorvall GSA rotor at 0° and 11,000 rpm for 20 minutes, or in the Sorvall SS-54 rotor at 0° and 18,000 rpm for 20 minutes. Step 1 and Step 2 were performed at 26 4°; Step 5 and Step 4 were performed at room temperature. Step 1. Clarified centrifuged extract. Frozen pig liver (800 g) was cut into small pieces with a knife and was suspended in two volumes of cold homogenization buffer. This buffer contained 50 mM tris(hydroxy- methyl)aminomethane (pH 7.4), 1 uM EDTA, and 5% (v/v) isOpropanol. The suspension was homogenized for 50 seconds at low speed, 50 seconds at medium speed, and 60 seconds at high speed in a Waring commercial blen- dor. Homogenization at each speed was interrupted at the midpoint in time for 15 seconds; these interruptions were also performed when chang- ing speeds. During homogenization, the temperature of the suspension typically increased from 0 to 6°. The homogenate was centrifuged in a GSA rotor for 40 minutes at 11,000 rpm and 0°. The supernatant was fur- ther centrifuged in polyallomer tubes in a Beckman Type 21 rotor at 0° for 150 minutes at 21,000 rpm. The dark red, faintly turbid supernatant from this centrifugation was designated the clarified centrifuged ex- tract. Step 2. Ammonium sulfate fractionation. The clarified centrifuged extract was brought to 0.25 saturation ammonium sulfate by adding satu- rated (4°) ammonium sulfate solution (previously adjusted to pH 7.4 at 25° and 0.1 saturation) with magnetic stirring. Stirring was continued for at least 15 minutes; the suspension was then centrifuged in a GSA rotor. The small amount of precipitated material was discarded, and the 0.25 saturated supernatant was brought to 0.56 saturation by adding more, ammonium sulfate solution (saturated at 4°). Again, stirring was con- tinued for at least 15 minutes. The suspension was then centrifuged in a GSA rotor. The pellets were collected and were suspended in homogeni- zation buffer to a volume equal to that of the original clarified cen- trifuged extract. After the pellets dissolved, saturated ammonium sul- fate solution was added with stirring to 0.56 saturation. After stir- ring at least 15 minutes, the suspension was centrifuged in a GSA rotor. The pellets from this centrifugation were resuspended in TED buffer (100 ml) and were then dialyzed overnight against a total: of four liters of TED buffer, with one buffer change. Dialysis typically began 12 hours after homogenization of the liver. After dialysis, the dialyzate was centrifuged in an 88-54 rotor to remove denatured (precipitated) protein. The supernatant from this centrifugation was warmed to room 27 temperature. Step 5. DEAF-cellulose pad chromatography. An 11.5 cm (d) x 5.0 cm (h) DEAE-cellulose pad was prepared in a Buchner funnel over Whatman No. 1 filter paper. Firm packing was achieved by preparing the pad un- der reduced pressure with the aid of an aspirator and a filtering flask. The pad was washed with TED buffer (one liter), the aspirator was de- tached, and the dialyzate supernatant from Step 2 was applied to the column. After the supernatant was absorbed by the pad, the aspirator was again attached. The pad was washed with TED buffer (one liter) and 50 mM KCl in TED buffer (one liter), drawn through the pad at a rate of about 50 ml per minute. The TED buffer wash fraction was extremely tur- bid. The KCl in TED buffer wash fraction was pale yellow-green and con- tained measurable catalase activity. Following these washes, fatty acid synthetase was eluted with 500 mM KCl in TED buffer. The bright yellow eluate was taken to 4° and was brought to 0.55 saturation with saturated (4°) ammonium sulfate solution. After stirring for 15 minutes, the sus- pension was centrifuged in a GSA rotor. The pellets from this centrifu- gation were combined and were suspended in a minimal volume (less than 20 ml) of PED buffer. The suspended pellets were dialyzed at 4° against PED buffer (500 ml) for at least two hours. The dialyzate was then can- trifuged in an 88-54 rotor. The clear yellow-orange supernatant was warmed to room temperature. Step 4. Sucrose density gradient centrifuggtion. Six linear sucrose density gradients (5 to 20%) containing PED buffer were prepared. The gradients (54 ml) were prepared in 1 x 5% inch polyallomer tubes using an ISCO Model 570 gradient former. One-sixth volume of the dialyzate supernatant from Step 5 was applied to each gradient. The tubes were then placed in the buckets for a Beckman SW27 rotor and centrifuged for 18 hours at 27,000 rpm and 20° in a Beckman L5-50 ultracentrifuge. U1- tracentrifugation typically began seven hours after the overnight dialy- sis at the end of Step 2. After centrifugation, the gradients were frac- tionated,using an 1800 Model UA-5 absorbance monitor to measure absor- bance at 280 nm and an ISCO Model 185 gradient fractionator to fraction- ate. The fractionation rate was 1.5 ml per minute, and 1.5 ml fractions were collected. Fractions having significant FAS activity were combined and taken to 4°. Saturated ammonium sulfate solution was then added to 28 0.50 saturation with stirring. After stirring for at least 15 minutes, the suspension was centrifuged in an 88-54 rotor. The pellets were com- bined and were taken up in a minimal volume of PED buffer. The suspended pellets were dialyzed for at least two hours against PED buffer at 4°- If satisfactory removal of the 7 S contaminant(s) had been achieved, dialysis was continued overnight, and the purified enzyme was frozen at -50° for storage. If further removal of the contaminant was desired, ultracentrifugation of the dialyzate was repeated. Comparison of the Original and Improved Purification Procedures for Peggy Acid Synthetase. Briefly, the changes introduced in the purifi- cation procedure for fatty acid synthetase were the following: 1. Isopropanol (5%) was incorporated in the homogenization buffer. 2. The first and second ammonium sulfate fractionations were changed from 0.2-0.55 and 0-0.55 saturation to 0.25-0.56 and 0-0.56 satu- ration, respectively. 5. DEAE-cellulose column chromatography with gradient elution was re- placed by DEAE-cellulose pad chromatography with stepwise elution. 4. The calcium phosphate gel treatment step and the associated Sepha- dex G-25 desalting procedure were eliminated. The effects of these changes in the purification procedure are detailed below. Effects of changes in homogenization buffer and ammonium sulfate frac- tionations. Table 2 presents a summary of data from parallel, small scale purifications of fatty acid synthetase. The procedure of Kim, Un- kefer, and Deal (1977), with and without 5% isopropanol in the homogeni- zation buffer, was used. The same pig liver was used for both purifi- cations. Essentially identical decalone reductase activity recoveries were obtained in the early fractions. It is informative to compare the 0.55 saturated supernatant fractions, which were discarded in the purifi- cation procedure. The supernatant fractions which were prepared with 5% isopropanol in the homogenization buffer contained about 2% times more decalone reductase activity than the supernatant fractions prepared with- out isOprOpanol. Thus, fatty acid synthetase was more soluble in 0.55 saturated ammonium sulfate when 5% isopropanol was present. Other pro- teins were also more soluble under these conditions. This was demon- strated by the greater amount of protein in the 0.55 saturated superna- tants and by the smaller pellets obtained from the 0.55 saturated 29 Table 2. Comparison of Preparations of Fatty Acid Synthetase Carried Out With and Without IsOpropanol (5%) in the Homogenization Buffer? Homogenization Buffer Homogenization Buffer Without Isopropanol With IsOpropanol Total Total DRb Total Total DR Volume Activity‘ Yield SAMPLE Volume Activity Yield (m1) (units) (%) (m1) (units) (3%) 210 194 100 Gsfi'centrifuged 200 168 100 omogenate clarified centri- 155 130 67 fuged extract IhS 116 69 0.2 saturated 179 125 65 supernatant 168 108 64 c 1st 0.55 satu- 200 12 ( 6) rated supernatant 198 30 (18) rated supernatant aThe purifications were performed by the method of Kim, Unkefer, and Deal (1977) with the exceptions of starting with 100 g pig liver and, for one purification, adding 5% isopropanol to the homogenization buffer. bDR is an abbreviation for decalone reductase. cThese values are in parentheses because these fractions are discarded. suspensions when isopropanol was present (protein content was measured by both the tannin turbidimetric method and by relative intensity of banding on native polyacrylamide gels). To minimize the loss of decalone reductase activity during the ammon- ium sulfate fractionations with isopropanol present, the upper cut was raised from 0.55 to 0.56 saturation. The lower cut was raised from 0.2 to 0.25 saturation because fatty acid synthetase did not precipitate at 0.25 saturation, regardless of the presence or absence of isopropanol. Effects of changes in DEAE-cellulose chromatography. A comparison of the original and improved procedures for DEAE-cellulose chromatography (on the dialyzates obtained at the end of Step 2 in the purification pro- cedures) is presented in Table 5. The dialyzate supernatant obtained using the original procedure of Kim, Unkefer, and Deal (1977) was turbid dark green and contained 7400 mg protein. That obtained using the im- proved procedure was also turbid, but was a much lighter golden-tan color and contained only 420 mg protein from the same amount of liver. 30 Table 5. Comparison of Yields and Degrees of Purity of Fatty Acid Syn- thetase Fractionated on DEAE-cellulose by Gradient Elution from a Column or by Stepwise Elution from a Batch-scale Pad. COLUMN CHROMATOGRAPHY“ PAD CHROMATOGRAPHYb Total Total DRc Total Total Total DR Total Volume Activity Protein Yield Volume Activity Protein Yield (m1) (units) (ms) (t) SAMPLE (n1) (units) (mg) (as) Step 2 130 236 7400 100 Dialyzate 129 652 820 100 FAS eluted 1000 54 -e- 14 from DEAE- 960 508 - e 78 cellulose a Columns were prepared and eluted by the method of Kim, Unkefer, and Deal (1977). Pads were prepared and eluted by the method given in Results. DR is an abbreviation for decalone reductase. Enzyme for column chromatography was obtained from 800 g pig liver by the method of Kim, Uhkefer, and Deal (1977); enzyme for pad chromato- graphy was prepared from 1600 g liver by the procedure in the Results. Protein concentration was not determined for these fractions. Also, its specific activity was twenty-five times that of the dialyzate supernatant at the corresponding point in the original purification pro- cedure. The difference in the purity of these dialyzate fractions is due to a solubilizing effect of 5% iSOpropanol upon many contaminating proteins which precipitated with fatty acid synthetase in the 0.55 satu- rated ammonium sulfate suspensions in the original procedure. The amount of protein in the dialyzate fraction from the original procedure was so great that about half the protein did not bind to the DEAE-cellulose column. This apparently accounts for most of the activity lost during DEAE-cellulose chromatography in the original procedure; the remainder was apparently lost due to irreversible binding to, or dena- turation on, the column. The amount of protein in the dialyzate ob- tained using the present procedure is sufficiently low that the quantity of DEAE-cellulose used was reduced to one-third of that used in the ori- ginal procedure. The 500 cm3 of DEAE-cellulose in the pad is sufficient to bind all the fatty acid synthetase in the dialyzate. Therefore, loss of activity during pad chromatography is apparently due only to 31 irreversible binding or denaturation of the enzyme on the column. One major purpose of the DEAE—cellulose chromatography step was to remove catalase, since it is similar in size to FAS and cannot be readily separated from FAS by ultracentrifugation. Both procedures were largely successful in removing catalase; however, FAS purified by either procedure has some catalase activity. An entire day was required for column chromatography in the original procedure; pad chromatography in the present procedure requires three hours. Yield and Purity of the Purified Enzyme. Both purification procedures yielded between 15 and 30% of the fatty acid synthetase in the clarified centrifuged extract as pure enzyme. Lower total yield and lower percent yield of enzyme was obtained from livers which had been stored for long period of time. Also, less enzyme was recovered from the DEAF—cellulose pad in Step 3 when more than the minimum amount Of ion-exchanger necessary for binding of the enzyme was used. The specific activity of the enzyme purified by the present procedure is comparable to that Of the enzyme purified by the original procedure (typically 1.6 units per milligram protein, based on the decalone reductase assay). The contaminating proteins in the purified enzyme, however, were distinctly different for the two procedures. Since the present procedure did not use the calcium phosphate gel treatment step, a small quantity of hemoprotein H-450 was present in the purified enzyme. This hemoprotein, with its visible absorption maxima, gave the FAS purified by the present procedure its pale yellow color. Fatty acid synthetase purified by the original procedure, although presumably free of hemoprotein H—450, had several other measurable enzymic activities. Relatively high levels of pyruvate kinase, malate dehydrogenase, and glutamate dehydrogenase activities were found, as well as lesser phosphoglyceromutase and catalase activities. Only catalase was found in FAS purified by the present procedure. All these contaminating activities were located primarily in the first 0.33 saturated supernatant of Step 2 in the original purification procedure for fatty acid synthetase. Apparently, in the present 32 purification procedure, even more of these enzymes are eliminated during the ammonium sulfate fractionations; that is, they are more soluble in 0.36 saturated ammonium sulfate with isopropanol present than in 0.33 saturated ammonium sulfate. Stability of the Purified Enzyme. The overall FAS activity was extremely sensitive to air-oxidation. When the enzyme was stored at 4° in PED buffer, 70% of the overall activity was lost after 24 days, and 95% was lost after 45 days. Over the same 45 day period less than 5% of the decalone reductase activity was lost. Rejuvenation of the air-oxidized enzyme was attempted by the addition of reducing agent at several different concentrations (Figure 2). Dithiothreitol was far superior to 2-mercaptoethanol in rejuvenating the enzyme. Enzyme stored in high concentrations of reducing agent (20—50 mM), however, lost all activity within 24 hours. Dithiothreitol at lower concentration (3 mM) reactivated the enzyme at a slower rate, but it also allowed the increase in overall activity to be retained for a much longer period of time. Therefore, overnight dialysis of the enzyme against PED buffer was tested; it was perhaps the most effective way of restoring overall activity to air-oxidized enzyme. The most effective means of preventing air-oxidation of the enzyme was to store the enzyme frozen at -30° in PED buffer. Less than 20% of the overall activity was lost after the enzyme had been frozen for three months. Most of the lost overall activity could be recovered by overnight dialysis against PED buffer. The remaining activity was apparently lost as irreversibly denatured enzyme, which was present in the thawed solution as a fine flocculent precipitate. DISCUSSION The initial decalone reductase assay would be the assay of choice for crude preparations of FAS. Unfortunately, 55333-1—decalone precipitated in the presence of these fraction because of their high salt and protein concentrations. By increasing the ethanol concentration in the assay from 5% to 10% and decreasing EEEEE'I’de' calone concentration from 100 mM to 50 mM, the assay can now be 33 .ucoewuoaxo osu usoswsousu acmumcoo opus muOdedo any mo mmauw>wuow omouoscou ocoamoon one .muso: mo noumw Houfiouzuownuflv :8 mg ou :oxuu was douaoucancqu SE n wsaafimucoo 395m 9:. .25 ADV Om no av om ou HocwcuoOuawouoEum mo in. AIV m: no «A4V S “ADV m ou Houwouzuonuwc "osmuao onu.wo muoDVwfio ou wovvo cosu ma3 ucowo wcwoanom .xuw>«uou mama -osvou mcofimuov mo mugs: muoa x N.~ was Auw>fiuom Admuo>o mo mafia: vuofi x m.o to: oesflo> fiasco mo uosowfio :a mega sowLB um “axon om ~w:o«uwvvo an 0: an concum cozu mos mamuso use .u0mmsb 9mm umcfiwwm unwwcuo>o vmumdmwv was aw «want mm now 0: um mahuao onu wcguoum noum< .muw>wuom -wuo>o mo mugs: «nod x w.n vac «sauce vowwfiusn xasmouw ecu mo uosvfidm cc .mucow< wowoocom an dmmuo>om muH vow omwuosucmm wfio< muumm mo coaumvfixouufi< .m ouswfim 34 3E5: v ”2: 22:282. 9 m c 9: no 2. mm a . d E _ J _ 825 ._.._b J/ql haul-IV 0 A E h (I max 2... a ‘1 a z m .\.' , Us: «2:. a R. w. o 4. . \ r \\ \. v 11111.. IIHJHHV fiJ "' .IIII-I-Ilu * \H\ I N m Iol /- . ta 2.: m \w m + In # m to SE 2 to 2.: me I m m m Figure 2. 35 used at any step in the purification of fatty acid synthetase. The total decalone reductase activity in the assay should be above 0.015 units, however, as below this level the specific activity of the enzyme is a function of its concentration. The cause is apparently a partial inactivation of the enzyme by trans-l-decalone, although dissociation of the enzyme into subunits with lesser (or without) decalone reductase activity is also possible. The initial procedure for the purification of fatty acid synthetase yielded enzyme of good purity, but required much time and yielded enzyme which was not as stable as desired. Using the present purifi- cation procedure, the time required for purification of the enzyme has been reduced from five days to three days. Most of the reduction in time comes from using a DEAR-cellulose pad and stepwise elution in place of gradient elution from a column. Some time is also saved by eliminating the calcium phosphate gel treatment step. This step can be incorporated for removal of the hemoprotein H-450 when desired. Comparable yields and specific activities of FAS are obtained using either procedure. The number of contaminating enzymic activities is apparently much less, however, in FAS purified by the present procedure. Storing FAS either frozen in PED buffer or in dialysis at 4 0against PED buffer minimizes air-oxidation of the enzyme and subsequent loss of overall activity. Freezing is preferable for long—term storage. As a final note, the fractional saturations with ammonium sulfate in Step 2 of the present purification procedure assume that isopropanol does not affect the solubility of ammonium sulfate. CHAPTER 3 Inactivation of Fatty Acid Synthetase in the Presence of Various Pellet Fractions from Pig Liver ABSTRACT A search for activators and inactivators of fatty acid synthetase in pig liver fractions has led to the discovery of inactivating capacities in the 6,000 x 3, 30,000 x g, and 124,000 x g pellet fractions from the crude homogenate. The 124,000 x g pellet fraction has the greatest inactivating capacity per gram wet weight. The rate of inactivation of fatty acid synthetase in the presence of suspension of these pellet fractions is dependent upon a number of factors. The rate of inactivation is increased at higher temperature and at higher concentration of pellet fraction. The rate of inacti- vation is also increased when air-oxidized enzyme is used. The presence of 1 mM dithiothreitol or 200,pM NADPH decreases the rate of inactivation. Continued addition of reducing agent to pellet fraction suspensions during incubation with fatty acid synthetase completely prevents the inactivation. The inactivation as measured by loss of overall FAS activity follows first-order kinetics. A preliminary study was performed to determine whether the inactivating capacities of the various pellet fractions could be extracted from the particulate material. The 124,000 x g pellet fraction released most of its inactivating capacity in soluble form immediately upon resuspension in TE buffer at room temperature; the other pellet fractions released only a small portion of their inactivating capacities, even after extended periods of incubation at 37°. Release of inactivating capacity from the pellet fractions was accompanied by a small but reproducible decline in pH of the suspensions which was proportional to the amount of inactivating capacity in the unincubated suspension supernatants. 36 37 The mechanism of inactivation of fatty acid synthetase in pellet fraction suspension was investigated using several techniques. In pellet suspensions containing FAS and incubated at 37°, overall FAS activity was lost more rapidly than decalone reductase activity. Analysis of these suspensions by sucrose density gradient centrifugation revealed that the native FAS peak decreased in magnitude as the extent of inactivation increased. Native polyacrylamide gel electroph- oresis of these suspensions also demonstrated the loss of the native form of the enzyme upon inactivation. These observations suggest that a nonspecific aggregation of FAS to a very high molecular weight material occurs shortly following inactivation. Since inactivation of FAS causes loss of the native form of the enzyme from native polyacrylamide gels, and native forms of other proteins extracted from the pellet fractions during incubation are not changed, the inactivating capacity may be specific for FAS. At 4°, isopropoanol (5%) decreased the rate of loss of decalone reductase activity in crude extracts of pig liver, and also decreased the rate of loss of overall activity of the purified enzyme in TE buffer. At 37°, however, both FAS in TE buffer and FAS in pellet fraction suspensions lost overall activity more rapidly in the presence of 5% isopropanol. At 49, sodium octanoate (0.5%) decreased the ' rate of overall activity loss by the purified enzyme in TE buffer. At 37°, the rate of overall activity loss by the pure enzyme in TE buffer containing 0.1% octanoate was greater than that in TE buffer; however, the rate of inactivation of FAS in pellet suspensions containing 0.1% octanoate was less than in suspensions without oct- anoate. These observations suggest that the inactivation of FAS by pellet fraction suspensions cannot be expressed solely in terms of nonspecific changes in the structure of the enzyme. 38 INTRODUCTION Much work has been done on the isolation and characterization of fatty acid synthetase from a variety of sources. Although much is now known about many of its physical and chemical properties, little is known about short-term control of the enzyme by factors other than its substrates and products. Consequently, studies were undertaken to determine whether activators or inactivators of the enzyme are present in the liver tissue from which the enzyme is isolated. It has long been known that the microsomal fraction of liver from most species tested inactivates fatty acid synthetase. The enzyme is unstable in liver fractions prior to the removal of the microsomal fraction. This is the initial report of an inactivating capacity in the 6,000 x g (10 min), 30,000 x g (15 min), and 124,000 x g (70 min) pellet fractions obtained from pig liver homogenates. This chapter describes the preparation of several inactivating pellet fractions and a partial characterization of the inactivation process. MATERIALS AND METHODS Reagents. Caprylic (octanoic) acid and dithiothreitol were obtained from Sigma Chemical Co. Malonyl—CoA, acetyl—GOA, and NADPH were products of P-L Biochemicals. Transyl-decalone was obtained from Aldrich Chemical Co. Acrylamide and N,N'-methylenebis(acrylamide) were products of Bethesda Research Laboratories, and N,N,N',N'-tetrame- thylethylenediamine of Canalco. All other reagents were of analytical reagent grade. Livers. Pig livers were obtained from Peet Packing Co. (Chesaning, MI) within two hours of slaughter and were frozen on dry ice. The livers were stored at -30°. Fatty Acid Synthetase. Fatty acid synthetase was prepared as described in Chapter 2. Assays for Fatty Acid Synthetase. The decalone reductase activity and overall FAS activity assays as described in Chapter 2 were used. 39 Polyacgylamide Gel Electrophoresis. A modification of the method of Clark and Switzer (1977), as described in Chapter 2, was used to perform native polyacrylamide gel electrophoresis. pflyMeasurement. Values of pH were determined at room temperature using a Radiometer PHM-4c meter equipped with a GK2321C electrode. Sucrose Denisty Gradient Centrifugation. Linear 5-20% sucrose gradients were prepared by the procedure of Martin and Ames (1961). The 5.1 ml gradients, which contained TED buffer, were formed in % x 2 inch cellulose nitrate tubes. After sample application, the tubes were centrifuged in a Beckman SW50.1 rotor at 200 and between 10,000 and 47,000 rpm for between 15 and 150 minutes. After centrifugation, the gradients were examined visually and then the absorbance profile at 280 nm was determined. Fractionation was performed using an ISCO Model 183 density gradient fractionator and an 1800 Model UA—S absorbance monitor. During fractionation the flow rate was 0.37 ml per minute, and 0.185 ml fractions were collected. RESULTS The aim of this research was to determine whether activators or inactivators of fatty acid synthetase were present in pig liver. So various fractions were prepared from pig liver homogenates by differential centrifugation. Initial tests for inactivation were performed at 4°, but inactivation was very slow. Subsequent tests were conducted at 37° to obtain faster inactivation. Preparation of Liver Fractions. Pig liver was cut into small pieces and suspended in two volumes of TE buffer. The suspension was homogen- ized for two minutes in a Waring blendor and then separated into several fractions by differential centrifugation by one of two methods. In method I (summarized in Figure 3), which was used primarily to obtain extract fractions, the homogenate was centrifuged at 11,000 rpm in a Sorvall GSA rotor at 00 for 40 minutes. The turbid supernatant from this centrifugation was called centrifuged extract. Upon centrir fugation of the centrifuged extract at 21,000 rpm in a Beckman Type 21 rotor at 00 for 150 minutes, the microsomal pellet fraction was 40 HOMOGENATE (2 ml TE buffer per g liver) 16,000 x g, 40 min pellet CENTRIFUCED [unstable]«“ EXTRACT (contains microsomal material) microsomal 45,000 x g, 150 min pellet fraction . CLARIFIED ‘”CENTRIFUCED (almost microsome-free) EXTRACT remaining parti- (NH4)2804 to 0.2 saturation; culate material 16,000 x g, 30 min 0.2 SATURATED SUPERNATANT [less unstable] [indefinitely stableJ-uw. (clear) Figure 2. Preparation of Supernatant Fractions from Pig Liver Homo- ggnate and the Stability of the Decalone Reductase Activities in the Fractions. The conclusion is that an inactivator of fatty acid syn- thetase is present in the particulate fraction of pig liver. obtained. The faintly turbid supernatant from this centrifugation was called the clarified centrifuged extract. In Method II (summarized in Figure 4), which was used to obtain pellet fractions, three such fractions were obtained from the liver homogenate. In this method, the homogenate was first centrifuged at 6,700 rpm in a Sorvall GSA rotor (6,000 x g) at 00 for 10 minutes. The nuclear ppd'gpll debris pellet fraction was obtained from this centri- fugation. The supernatant from the 6,000 x g centrifugation was then centrifuged at 18,000 rpm in a Sorvall 88-54 rotor (50,000 x g) at 0° for 15 minutes. The pellet obtained was called the " remicrosomal" pellet fraction. The turbid supernatant from this centrifugation was then centrifuged at 0° in either a Beckman Type 21 rotor at 21,000 rpm (44,000 x g) for 150 minutes or a Beckman Type 42.1 rotor at 40,000 rpm (124,000 x g) for 70 minutes. The pellet obtained was the microsomal pellet fraction. Each of the pellet fractions was further purified by washing at 4° with TE buffer. Washing consisted of resuspension of the pellet frac- tions followed by centrifugation. The pellet fractions were resuspended 41 HOMOGENATE (0°) "NUCLEAR AND CELL DEBRIS" PELLET FRACTION 5:000 x 8, 10 min supernatant l "PREMICROSOMAL" PELLET FRACTION 30,000 2: g, 15 min supernatant 2 MICROSOMAL PELLET FRACTION 1215000 x 3’ 70 “in clarified supernatant Figpre 4. Preparation of Pellet Fractions from Pig Liver Homogenate by Differential Centrifugation. All pellet fractions were washed before use to remove any remaining soluble material from the homogenate. Wash- ing consisted of resuspending the pellets and repeated centrifugation. in a volume of buffer equal to the volume of suspension from which they originally came. Shaking was used to resuspend the fractions obtained by centrifugation in GSA rotors. Fractions obtained by centrifugation in Beckman rotors were resuspended by a single 30 second burst of homo- genization in a glass waring blendor. The pellet fractions (resuspended by shaking or by homogenization) were then centrifuged under conditions identical to those used to initially obtain the particular fractions. Washing was continued until soluble proteins present in the crude homo- genate had been removed (as demonstrated by native polyacrylamide gel electrophoresis. Three or four washes were typically required. Some protein was extracted from the pellet fractions during washing. These proteins gave banding patterns in native polyacrylamide gels which were characteristic for each pellet fraction (Figure 5). 42 ”NM {135K tvqbza'r Figure 5. Native polyacrylamide Gel Electrophoresis Banding Patterns of Solubilized Proteins of Purified Pellet Fractions. After the pellets were washed, they were resuSpended in ten volumes of water. The suspen- sions were centrifuged at 0° and 40,000 rpm for 70 minutes in a Beckman 42.1 rotor. Native polyacrylamide gel electrophoresis was performed on the supernatants using the procedure given in Chapter 2, yielding the banding patterns shown for the nuclear and cell debris (6.7K tan), pre- microsomal (12.75K), and microsomal (56Kf10c) pellet fractions. 43 Tests for Inactivation of the Decalone Reductase Activity of Fatty Acid Synthetase by the Supernatant Fractions. Endogenous decalone reductase activity was measured as a test for inactivation Of the decalone reductase activity of FAS by the centrifuged extract and the clarified centrifuged extract fractions. There was a slow loss of activity in both extracts at 4° (Figure 6 and Figure 7). Half of the activity was lost in two days in the centrifuged extract and in three days in the clarified centrifuged extract. The clarified centrifuged extract was faintly turbid. To determine whether the turbid material was responsible for the inactivation a clear supernatant was obtained from the clarified centrifuged extract by taking the extract to 0.2 saturation with ammonium sulfate and centrifuging at 11,000 rpm in a Sorvall GSA rotor at 00 for 25 minutes. The decalone reductase activity in this supernatant was stable for more than ten days (Figure 8). Taking the clarified centrifuged extract to 0.2 saturation with ammonium sulfate is part of the standard procedure for the purification of FAS. Tests for inactivation of fractions in the purification procedure for FAS beyond the 0.2 saturation ammonium sulfate fractionation were negative; the fractions were stable in- definitely. Hence, the turbid material seems responsible for the inactivation of FAS. Tests for Inactivation of Overall FAS Activity by Pellet Fractions. In order to determine the factors upon which the rate of inactiva- tion of overall FAS activity in pellet fraction suspensions might depend, a standard procedure for inactivation experiments using pellet fractions was defined. This procedure was as follows: a suspension of one gram pellet fraction in ten milliliters TE buffer was prepared. The pellet fraction was suspended using a 15 ml Dounce homogenizer with a "large" clearance pestle. Fatty acid synthetase in PED buffer was then added to an aliquot of the suspension to a concen- tration of 0.16 units (overall activity) per milliliter of FAS-con- taining suspension. Between one and two volumes of FAS in PED buffer for each ten volumes of pellet fraction suspension was typically re— quired. After measuring the overall activity of a flD‘pl aliquot of the FAS-containing suspension, the suspension was placed in a water 0 . . bath at 37 to incubate. Aliquots were removed at regular intervals 44 .%~o>wuooammu ads “on mums: om.o was mm.~ mums fiocmaouaomw fim Aev unocuHB was AOV saws wowmsn :« cowumuwcowoeoz up monsoon; uo>wd wan mo muomuuxo camoufluusoo ecu a“ mOwuH>Huom ammuoavou aco~mooc fimwuwcw use .HochOHAOmH fiw mo uoowmm Aw0>an mam mo uouuuxw mowSmHuucoo ecu cw Mua>wuo< Omwuosvom ocofimooa .m ouswwm 45 Figure 6. 585 is: 20:28:. a. R. a. 8. a a a a IeOIIeeleee.o I Iota as m O 19:: o: 46 .>~m>wuooom0p «#5 non mums: on.~ com om.~ mums «pmwwsp coHumuficmono: ecu cw LoamQOHQOmH fim Aev unocuwz pom on Laws auo>wd wan Eouw nonmaoua mucouuxo mowsm -wuucoo pemeumHO now memuw>wuom Ommuoswou Ocofiwoou fimfiuHcH one .HOCwaouQOmH mm mo uoommm "uo>wq Man «O uompuxm wowswwuunoo pmwwwumdo ecu cw zuw>wuo< smouospmm ocodmooo .5 suswwm mum com .285 us: 22382. 47 Figure 7 ‘4 q a - u d d it I 19:: c: .Ha won mugs: No.0 on: uceuecuonsa ozu mo hua>auue oeeuospou anodeoou AewuaCu any .uo>«A mam mo uosnuxm vowsmwuucoo pawmwuofio Eoum unnumcuomam enougsw anacossm vausuauum N.o on» Ca wwfi>uuo< oauuuspom ocodwosa .w shaman A285 “2: 22:382. mum 8N m: :3 m2 9: fl . om mu c u - J¢ u , a u u u d N .c: 8 4 0000.. cc 49 and assayed for overall activity. The first inactivation tests on the pellet fraction suspensions were temperature dependence studies. Overall FAS activity was measured in all tests. Effect of temperature on the inactivation rate. Washed pellet fractions were suspended at 4° in TE buffer at a concentration of 0.1 gram pellet per milliter suspension using Potter-Elvehjem homogenizer. Pellet fraction suspension (325 pl) was then mixed with FAS in PED buffer (SOpl, 0.13 units). The mixture was incubated at 4° and assayed intermit- tently (Figure 9). The most rapid loss of activity was in the mixture containing microsomal pellet fraction. The loss of activity at 4° was very slow, however. Pellet fraction suspension containing FAS and incubated at 37° had relative inactivating capacities similar to those at 4°, but about seventeen-fold greater (Figure 10). For this reason, inactivation studies using pellet fraction suspensions were performed at 37°. Effect Of pellet concentration on the inactivation rate. The rate of inactivation of FAS in pellet fraction suspensions is directly propor- vtional to the concentration of pellet fraction. Studies at several con- centrations of pellet fraction demonstrated the inverse prOportionality between pellet fraction concentration and the half-life of overall FAS activity (Figure 11). Effect of air—oxidation of the enzyme on inactivation rate. The rate of inactivation of FAS in pellet fraction suspensions was greatly af- fected by the relative oxidation state of the enzyme. Purified FAS stored at 4° eventually loses a considerable portion of its overall activity, but only much later loses any of its decalone reductase ac- tivity (Figure 2). The loss of overall activity was apparently caused by a reversible air-oxidation of the enzyme, as the overall activity could be restored by the addition of, or dialysis against, a reducing agent. Consequently, accurate elucidation of the effects of potential inactivators on FAS requires a knowledge of the relative oxidation state of the enzyme, particularly when the pellet fraction suspensions contain a reducing agent. The most reliable and easily interpretable results were obtained with fresh enzyme in its fully reduced, most active state. Generally, therefore, tests were conducted with enzyme which had been dialyzed overnight against PED buffer. 50 .xuwownmo wcwuo>wuoocw o>wum~ou Ou ~o:0wuuoaoua aduoouwo mum muoHa Omega «0 moaon ecu vow aoefiu mamuo> w< mod: on oeuuofia one mwmozu meu :« mama ecu ACOmmou mwnu pom .op mafia msmuo> w< mod: mo msowuo .zuwooamo wowum>auomcw cu zuaawcoHuuoaoua uoouwo .oHaEwm w o>ww you on acnmuw nose ao>wumwoc mum moaodm dfim conga usm .Azufiooamo wcHum>quQCw ecu usuwouw on» aoaofiu ozu mo opsuwcwme Ou3~0mnm any usumouw osu a. o .wv mufiowamo wsaum>wuowa Ou Hm:Owuuoa noun ops mopsuwcwoe mmmwmmmm.0mo:3 momofim Laws mocfid mo>ww oEHu moauo> m< on+ mo camuw one .OEHu saws mommouooo auw>wuom Oaxaca meowm OQOAm O>aumwoc m on: o>n=o a Loam .Oewu mo cOHuocsw a as neuuofia ma Am< wofiv oewu cm>ww a um wcwcwmsou >ua>fiuoo Oaxaca Hmwuacw ecu mo COwuomuw on» no Ezuwnmwod ecu coc3 pocwmuao a“ mafia unwfimpum n «.o.a Hosanna ou uooam use sows nopuo umnwm -ov3Oma no nopuo umuww a“ ammooua Ono vow «mascaauoaxo coduo>wuomcfi mcausp mean cums ammoouoop muw>wuow -mpo>o o5H .oewu mo :ofiuonsm o as auw>wuum Oaxaca onu ou pousdou zfiwumswuds ma coHLOOHM a mo zuaowowo wcwuo>wuomcfi one .>~o>wuooauou amusos mm was «mm «mm sums mconnoamsm acauowuw pegged deOmouuHE can adeOmouOHEOHQ amwunov game can Hosanna ecu a“ >ug>wuom mo mo mo>wduw~mz .Auxou oomv >uw>wuom -muo>o ecu monwawnmuo xawuouonEOu ucowm wcwospou one .~OuwounuoH:uwv :5 m pocfioucoo :menoamSa some can 0: us mm3 cowuopso new ”msoHuoooxo omosu Lows acofiuwvcoo unaccoum pops: vouwnaocw was vousnoun owes oconcoaosm och .03 um mconcwmmsm cOLuooum aegfiomlwm ommuocucxm wwo< Nuumm mo nofiuw>AuumaH .NWMMMMNM 51 8 Figure 9. 1 585 as: 22:382. T £58 :3 use .862: o: N... .r 3. 3. ad 1 o._ ’v 901 - 52 1.0 f 0.8- _ / / ' premicrosomal <- microsomal A - LOG Af <—nuclear and cell debris 4. A. n 1 J J 0 100 200 300 400 500 INCUBATION TIME (MINUTES) Figure 10. Inactivation of Fatty_Acid Synthetase by Pellet Fraction Suppensions at 313. The suspensions were prepared and incubated under standard conditions with the exception of the suspensions containing 2 mM dithiothreitol. The reducing agent temporarily stabilizes the overall activity (see text). Half-lives of overall FAS activity in the nuclear and cell debris (A), premicrosomal (C), and microsomal (I) pellet fraction suspensions were 320, 245, and 76 minutes, respectively. The slope of each curve is proportional to the inactivating capacity of the corresponding suspension (see Figure 9 legend for further details. 53 0.15291ml 1.0 ,- 9 <'0.076 Iml 0.8 - ' g ¢£"' 0.6 - ‘. 8 0.0389lml 7' I 0.4 - ‘/ 0.2 - II lll"; k l l l I I 0 100 200 300‘ 400 500 INCUBATION TIME (MINUTES), Figure 11. Inactivation of Faggy Acid Synthetase by7Varioua Concen- trations of Pellet Fraction. Premicrosomal pellet fraction and fatty acid synthetase were combined and analyzed under standard conditions with one exception: pellet fraction concentrations were 0.038 (A), 0.076 (Q), and 0.152 (I) grams per m1 suspension. The relative speci- fic inactivating capacities (the slape of the plot of -1og Af versus time divided by the pellet fraction concentration) for these suspen- sions were 0.026, 0.049, and 0.051 min'ml/g, respectively. So at least at higher pellet fraction concentration, inactivating capacity is pro- portional to the slope of curve (see Figure 9 legend for further de- tails. 54 Effect Of reducing agent on the inactivation rate. When 1 mM dithio- threitol was added to pellet fraction suspensions containing FAS prior to incubation, the inactivation rate was decreased (inactivation was prevented) for a time. Nuclear and cell debris, premicrosomal, and microsomal pellet fraction suspensions containing FAS and 1 mM dithio- threitol had inactivation half-lives for FAS of 90, 245, and 325 min- ute§,respectively. Addition of dithiorthreitol to FAS-containing pel- let fraction suspensions at a rate of 1 mM/hr prevented the inactiva- tion (Figure 12). Thus, the redox environment in the incubation mix- ture significantly affects the inactivation rate. Effect of NADPH, malonyl-GOA, and acetyl-GOA on the inactivation rate. Incubation of FAS in pellet fraction suspensions containing a saturat- ing level of NADPH (200 pM) reduced the rate of inactivation of the enzyme (Figure 13). Half-lives of inactivation fdr FAS in the premicro— somal and microsomal pellet fraction suspensions were 88 and 62 minutes in the absence of NADPH, and 167 and 88 minutes in the presence of NADPH, respectively. Addition of 50 pM acetyl—COA and 50 pM malonyl-CoA to FAS-containing pellet fraction suspensions slightly destabilized the enzyme (Figure 13). Half-lives of inactivation for FAS in the prem- icrosomal and microsomal pellet fraction suspensions in the presence of acetyl-GOA and malonyl-GOA were 77 and 52 minutes, respectively. The Inactivation of Overall FAS Activity by Pellet Fraction Suspensions Follows First-Order Kinetics. The inactivation of FAS in pellet frac- tion suspensions containing no dithiothreitol fellows first order kin- etics (Figure 11). Therefore, the relative inactivating capacities of the suspensions can be determined from the slope of a plot of the log- arithm of the fraction of the initial overall FAS activity in the sus- pension (logAf) as a function of time. Plotting -logAf versus time results in the slope of the curve being directly proportional to the amount of inactivating capacity present (see Figure 9 legend). As previously shown (Figure 9 and Figure 10), dithiothreitol at an initial concentration as low as 0.2 mM in pellet fraction suspensions containing FAS has a stabilizing effect on the enzyme. The duration of the stabilization is proportional to the amount of dithiothreitol in the suspensions. After the period of stabilization, inactivation of the enzyme again follows first-order kinetics . Consequently, relative 55 .comwuwano 90m :3onm aw cowuomuw uoHHoa mo commune osu aw Oaxaca ecu mo xuwawnwum OLE .cowmp was GOHDonaonw usumm mouscwe can man «can aomm um XE m an man «comes ems noHuw£=OCw nmumm sausage mw~ mam .ow~ ammH «mm «mm ‘on us SE a xo memmmuo Isa mwB commcoamsm mwcu cw coHumuucoosoo douwoucuoHLuwm use .coHumnsocA wcfiusm momma Houwounu IOHLuHm SE cod mo muosvwam mm: concoamzm Ono mam anofiumnaocw mo uumum ecu um AOuwousuoHsuam ZEN mongmunoo m:o«mcoam:m :uon "macauamoxm wawsoaaom osu sums m:0au«maou mummamum Moms: monxa Imam mam mmuwamud mums anewmcoamsm :oHuouum usagoa assemououz .msowmaoamsm newuomum usaaum we Ommummwcaw mwo< Nuuom mo :oHuw>wuomcH mucm>oum acuwousqucuwa mo nofiuwmm< owmofiuom .mfi shaman 56 Figure 12. 552.5 Us: 22582. 8e 8m :8 .8m 8w :2 o - IIdII q u u .d mugul 2.83:3 « q 4lull-ll '- IUWll|ll|||||l|l|AVHHHHHHHHHHH;I\\\\. LHHHHmemmvlJ _o I. n. \«q\ \ sears . 2 . ed . 9: Au .. a... SenatoE'v L .3 ’v 901 - 57 .%~O>Huooamou ammunawa Nod mam mm cw >ua>wuow Awwuacw osu mam; moum>wuomCH mmnwuoocw ~o>«uooamow ..2325.: mm mam we :« muw>wuom mo 13:5 one :6: meant/:02: snowmcoagm cofiuowuw oozed Adv amEOmouuHEmua mam A4V :EOm IouoHE muwmcmum use .mconcoamam ozu Ou fiuownH ecu :o muoommm HOuomwoo mam museumnam .m~ ouswwm 58 Figure 13, «$52.28 as: 22:382. :8 en =8 8" e - - - - a $222 53.382259... ll. ....... us as . 0.... 1%.. .. 9: JV 333.2359... .lnv . . 2 $22 5.; 32822.“... <8 .22 ES <8u< 5;. 3532252,. . . .. 3 4 <8 .22 En <82 5.; 353.82.: 38882:. 59 inactivating capacities of suspensions containing reducing agent can be determined from the slope Of the plot of -logAf versus time in the re- gion where inactivation has begun. One unit of inactivating capacity is defined as that amount of in- activating capacity required to produce the loss of one unit of overall FAS activity in one minute at 37° in the inactivation assay. One unit of overall FAS activity is as defined in Chapter 2; that is, one unit of overall FAS activity is the amount of FAS catalyzing the oxidation of 1 umole NADPH per minute in the standard overall FAS activity assay. Extraction in Aqueous Solution of the Inactivatinngapacity from Pellet Fractions and the Requirement of Intact Membranous Pellet Fraction for the Inactivation of FAS. The long range goal is the isolation and iden- tification of the inactivating componenets in the pellet fractions. The first step toward this goal was tO determine whether the inactivating capacity could be extracted and separated from the pellet fractions. Thus, the questions were (a) is intact pellet fraction membrane re- quired for inactivation, and (b) if not, can the inactivating compon- ent be extracted from the pellet fraction? To determine whether the inactivating capacity required intact pellet fraction, the pellet fractions were resuspended at room temper- ature in TE buffer containing 2 mM dithiothreitol in standard propor- tion. The suspensions were then divided into three portions. One por- tion was incubated with FAS and analyzed under standard conditions. An- other portion was centrifuged at 40,000 rpm for 70 minutes in a Beckman Type 42.1 rotor at 0°. Fatty acid synthetase was then added to the super- natant to a concentration of 0.13 units (overall activity) per m1 super- natant. The FAS-containing supernatant was then incubated at 37° and assayed intermittently for overall FAS activity. The third portion was incubated at 370 for 20 hours and then centrifuged at 0° for 70 minutes in a Beckman Type 42.1 rotor at 40,000 rpm. Fatty acid synthetase was added to the supernatant to a level of 0.13 units (overall activity) per m1 supernatant and then incubated at 37°. The FAS-containing supernatant was assayed intermittently for overall FAS activity. 60 The results from these inactivation tests are given in Table 4. Table 4. Rate of Solubilization of Inactivating Capacities from Pellet Fraction Suspensions at 57°. -------$esesizesiea-gseesisiseissisalsllslgf ......... a supernatant of un- supernatant of P6119t fraction suspension incubated suspensionb incubated suspension nuclear and cell debris 5‘8 0’9 1'7 premicrosomal 5.8 1.5 2.4 microsomal 7.9 7.8 7.4 aPellet fractions were suspended in TE buffer containing 2 mM dithio- threitol. Suspensions contained 1 g pellet fraction/ 10 ml suspension. bSupernatants were obtained by centrifugation of suspensions at 40,000 rpm for 70 minutes at 0° in a Beckman 42.1 rotor. cThe suspension was incubated at 57° for 20 hours. The inactivating capacity of the microsomal pellet fraction was released in soluble form.immediately after resuspension at room temperature. Only about one-fourth of the inactivating capacities of the other pellet frac- tions was released upon resuspension of the pellet fractions, and less than half of the inactivating capacity was released even after incubation at 57° for 20 hours. For this reason, the inactivating capacity of the microsomal pellet fraction seems to be different from.the inactivating capacities of the other pellet fractions. The pH values of the pellet fraction suspensions for these inac- tivation tests were measured. A decline in pH, which was proportional to the inactivating capacity of the unincubated suspension supernatants, was observed. The pH values for FAS-containing suspensions of the nu- clear and cell debris, premicrosomal, and microsomal pellet fractions were 7.52, 7.52, and 7.47, respectively. The pH of a solution of FAS in buffer alone was 7.55. Thus, the pH is lowered about 0.01 for each 10“ unit of inactivating capacity per ml unincubated suspension super- natant. The significance of this phenomenon is not yet known. Studies on the Mechanism of Inactivation of FAS by Pellet Fraction Suspensions. To determine whether a conformational change occurred 61 in fatty acid synthetase upon its inactivation by pellet fraction suspension, sucrose density gradient centrifugation was performed on pellet fraction suspensions containing FAS which had been incubated at 37° for various periods of time. As the overall FAS activity was inactivated, the magnitude of the native FAS peak in the sucrose density gradients, as measured by absorbance at 280 nm, decreased (Figure 14). At no time during the inactivation process was a peak other than that for native FAS observed in the sucrose density gradients. Consequently, in terms of gross physical structure, inactivation results in an aggregation of FAS to very high molecular weight material. The aggregation is apparently nonspecific, as no new discrete peaks appear in the sedimentation patterns obtained from sucrose density gradient centrifugation of the FAS-containing pellet fraction suspensions. A similar conclusion was drawn from comparable experiments using native polyacrylamide gel electrophoreisis instead of sucrose density gradient centrifugation. The native FAS band decreased in intensity as the enzyme was inactivated, and no other bands appeared in the polyacrylamide gels (Figure 15). Native polyacrylamide gel electrophoresis performed on FAS-contain- ing pellet fraction suspensions also suggested that the inactivation by the microsomal pellet fraction may be specific for FAS. Fatty acid synthetase, 0% or 98% inactivated by microsomal pellet fraction suspension, was clarified by centrifugation at 18,000 rpm in a Sorvall SS-rotor at 0° for 20 minutes, and the supernatant was subjected to native PAGE. The native FAS bands decreased markedly in intensity upon inactivation of the enzyme, while bands from contaminating proteins (solubilized during resuspension of the pellet fraction; see Figure 5) remained at about the same intensity, apparently un— changed by incubation with the inactivating capacity. Also, after 98% inactivation, there was a substantial amount of native FAS still present, indicating that aggregation follows inactivation. The relative rates at which decalone reductase activity and overall FAS activity were lost during incubation of FAS with pellet fraction suspensions were measured. Assuming that the aggregated enzyme has neither partial nor overall FAS activity, then if aggregation is concommitant with 62 Figure 14. Inactivation of Fatty Acid Synthetase and Loss of the Native FAS Peak During Sucrose Density Gradient Sedimentation Velocity Analysis. Microsomal pellet fraction was suSpended and incubated with fatty acid synthetase under standard conditions for 0, 96, 220, or 580 minutes. After these incubation periods, 0, 52, 74, and 95%, respectively, of the overall FAS activity was inactivated. After incubation, the sus- pensions (150 “1) were applied to sucrose density gradients (see Meth- ods). Centrifugation was initially at 5,000 rpm (for 5 minutes) in or- der to gently transport the particulate material into the gradients. Centrifugation was then continued for 20 minutes at 50,000 rpm. The elution profiles at 280 nm, obtained during fractionation of the grad- ients, are for (a) no incubation, (b) 96 minutes incubation, (c) 220 minutes incubation, and (d) 580 minutes incubation. Similar profiles were obtained when incubating fatty acid synthetase with premicrosomal pellet fraction. Figure 14. ABSOR BANCE w. l “I TOP ‘ T I 0. 2 A280 nm BOTTOM l 0 minutes r——-- 220 minutes 63 96 minutes 380 minutes 61+ m “ Ilil - I! I, M a} ‘4. “I Figure 15. Native Polyacrylamide Gel Electrophoresis Banding Patterns of Microsomal Pellet Fraction Suspension ContainingpFatty Acid Synthe- tase before and after Inactivation of the Enzyme. The pellet fraction suspension was prepared and incubated under standard conditions with one exception: the suspensions contained 1 mM dithiothreitol. Overall FAS activity was 0 (210) or 98% (21f) inactivated when polyacrylamide gel electrophoresis was performed on the clarified suspensions (centri- fuged at 0° and 18,000 rpm for 20 minutes in a Sorvall 88-54 rotor). The procedure for PAGE presented in Chapter 2 was used. 65 FAS inactivation, overall and decalone reductase activities should be lost at prOportionate rates. Experimentally, the decalone reductase activity was lost less rapidly than the overall activity of the enzyme (Table 5). Table 5. Relative Rates of Decalone Reductase and Overall Fatty Acid Synthetase Activity Loss During Inactivation of Fatty Acid Synthetase by Microsomal Pellet Fraction SuSpensions at 57°. Incubation time (minutes) 0 5O 94 160 250 510 Fraction of initial over- all activity remaining 1°°0 1'05 0°85 0-56 0-50 0.24 Fraction of initial decalone 1 reductase activity remaining ‘00 0'75 0‘78 0'69 0°72 0-51I Model Compounds for the Inactivatipg:0apacipy of the Pellet Fractions. Two model compounds for the inactivating capacity in the pellet frac- tions were examined to see how closely they mimicked the inactivation of FAS by pellet fraction suspensions. These model compounds were isopropanol and sodium octanoate; how they were chosen and their effects upon FAS are examined below. Inactivation of overall FAS activipyfpy isppropanol. IsOprOpanol (5%) was examined as a result of studies performed during the purification of fatty acid synthetase. In those experiments, isOprOpanol was used as a solvent for phenylmethanesulfonylfluoride, a widely used protease inhibitor. The goal of these eXperiments was to determine whether a more stable FAS preparation might be obtained by preparation of the enzyme in the presence of a protease inhibitor. A control purification of the enzyme was performed using isopropanol without the phenylmethanesulfonylfluoride. This 5% isopropanol control was found to significantly stabilize the enzyme at early stages in the purification (Figure 6 and Figure 7). The half-lives of decalone re- ductase activity in the centrifuged extract and clarified centrifuged extract at 4° with 5% isoprOpanol were 90 and 150 hours, and without isOpropanol were 45 and 60 hours, respectively. IsOprOpanol also had a 66 stabilizing effect on the ”natural" inactivation (air-oxidation) of the overall activity of the pure enzyme in TED buffer at 4° (Figure 16). The stabilization of overall FAS activity in the isopropanol containing buffer and extracts at 4° is apparently caused by isopropanol lowering the dielectric constant of the solution. Changes in the dielectric constant can cause changes in the stable conformations of proteins by altering hydrogen, electrostatic, and hydrophobic bonding of the proteins. Hence, it appears that the most stable conformation(s) for FAS in the presence of 5% isopropanol at 4° is different from that in its absence, and that the conformation of FAS induced in 5% isopropanol is less sensitive to both inactivation and air-oxidation. At 37°, isopropanol inactivated the pure enzyme in TED buffer (Figure 17). The half-lives Of overall FAS activity in the presence of 2% and 5% isopropanol in TED buffer (0.13 units/ml) were 300 and 46 minutes, respectively. Both native PAGE and density gradient sedimentation velocity analysis of the 5% isopropanol-containing FAS solution at 37° demonstrated the loss of the native form of the enzyme during or closely following inactivation. Therefore, the most stable conformer(s) of FAS in 5% isopropanol at 37° is apparently either inactive or much more susceptible to air-oxidation than the enzyme in the absence of isopropanol, as overall FAS activity was lost rapidly and the enzyme aggregated nonspecifically to high molecular weight material. The aggregation of FAS was also observed when FAS was inactivated by the pellet fraction suspensions. In the presence of both pellet fraction suspension and 5% isopropanol at 37° the inactivation rate of overall FAS activity was greater than that observed using either pellet fraction suspension or 5% isopropanol alone (Figure 18). Hence, 5% isopropanol and the pellet fraction suspensions, separately or combined, produced the same overall effects on FAS; the enzyme was inactivated and aggregated nonspecifically. However, isopropanol .zun>nuoo mo new anucouunsnuuan mozamua onw3 mom on up monOua one: mo~AEwm one .Alc nocaaondOan Rm wcncnwunoo new Imps nub no an£VIOumocauuo mm.o wcmcmmucou nuwwsn amp «Aev nowman Que nonuno mo #2 own an momma was A zun>nuun -wnm>o mo nuns: moo.o e onsuosucxm.mnoo zuuww muannam .aocu on OuH cannousoo nommsm OMB no sumo: .38 “me 2:53:00 5:3 8:. .583 can nofinm an o: as 83.2.3.3 32 ~32 no 333...; .8 38$ 585 us: 22:38,: 8N m: :2 m2 8" R 8 mm a .l . . . _ _ _ J c m I Nd I <5 I 0.: .555 am: e \ oe . I w c 68 1.0 - ' II / ‘ 2°10 iPrOH 0.2 ‘/+_ . o ’ 0.4 IoiPrOH /‘ K ‘m“"“ [Jafl“““, 0 ’ 0,0/ <4— 0.1% i PrOH -—--"'A“'7 352%. '\ no iPrOH -0.2 _ l l l l 0 100 200 300 400 INCUBATION TIME (MINUTES) Figure 11. Inactivation of Fatty Acid Synthetase in TED Buffer by Iso- propanol at Various Concentrations. Purified fatty acid synthetase (15 “1, 0.029 units of overall activity) was added to TED buffer (300 pl) containing 0 (e), 0.1 (A0, 0.4 63), 2.0 (A), or 5 % GI) isopropanol. The samples were incubated at 57° and assayed intermittently for overall fatty acid synthetase activity. 69 ' I:— microsomal . ALL WITH 5°10 1.2 - ISOPROPANOL premicrosomal 1.0 " l A 4 nuclear and 0.8 . I celldebris <"" . CD 0.6 - 9 buffer I . A l 0.4 - . (34" 0.2 - O l A 0 , 50 ‘ 100 INCUBATION TIME (MINUTES) Figure 18. Inactivation of Fatty Acid Synthetase at 57° by Pellet Frac- tion Suppensions in.TED Buffer Containingpjfi Isopropanol. Suspensions were prepared and analyzed under standard conditions with the exception of adding 5% isopropanol to all suspensions (and buffer). The half- lives of overall FAS activity in the nuclear and cell debris (A), pre- microsomal (e), and microsomal (I) pellet fraction suspensions were 25, 15, and 5 minutes, respectively. Inactivation of enzyme in TE buffer containing 5% isopropanol is shown for comparison. 70 is distinctly different from the inactivating capacity in the pellet fraction suspensions, in that at 4° isopropanol stabilizes the enzyme, while the pellet fractions inactivate it. Inactivating of overall FAS activity by octanoate. The pellet fractions consist of various membrane—enclosed organelles and thus have a rela— tively high lipid content. Therefore it seemed that one possible mech- anism for the inactivation of FAS in pellet fractions might involve a detergent-like denaturation of FAS by fatty acids either on the surface of the membranes, or released from them. For this reason, a representative fatty acid was tested for its inactivating capability. Octanoate was chosen because it is very soluble and, unlike palmitate, has little tendency to form micelles even at relatively high concen- tration. The hydrocarbon portion of octanoate, although short enough to minimize micelle formation, was long enough to be expected to pene- trate the hydrophobic regiOns of the enzyme. At 4°, octanoate (0.5%) had a stabilizing effect on FAS; it slowed the "natural" inactivation (air-oxidation) of the pure enzyme in TED buffer (0.13 units overall FAS activity/ ml; Figure 16). When the in- cubation was carried out at 37°, however, overall FAS activitwaas inactivated in octanoate-containing TED buffer (Figure 19). At 0.1 and 0.5% octanoate, the half-lives of overall FAS activity were 415 and 55 minutes, respectively. Sucrose density gradient sedimentation velocity analysis of the inactivation of FAS in solutions containing octanoate revealed that a peak with a higher sedimentation coefficient than that of native FAS was formed during inactivation of the enzyme (Figure 20). This faster sedimenting peak had neither decalone reduc- tase nor overall FAS activity. It was an intermediate in the denatur- ation of the enzyme as indicated by its relatively small size and by the appearance of pellets at the bottoms of tubes to which enzyme that was inactivated to large extents had been applied. These observations suggest that at 4° octanoate did not signifi- cantly alter the conformation of FAS. The stabilization of the enzyme may be due to octanoate forming a monolayer at the interface between the solution and air, thereby slowing the diffusion of oxygen into the solution, and so slowing air-oxidation of the enzyme. At 37°, the octanoate has clearly affected enzyme conformation, apparently yielding 71 1.0 F' 0.8 4 ./ ,_ 0.6 - <£ II 8 ‘I‘ 0.4 - o . *—— 05 lo octanoate ‘ 0.10/0 octanoate ', o. 02 ° 0‘ 3(— I0 octanoate . _—-o ,.." no octanoate I I I A 100 200 3(1) 400 INCUBATION TIME (MINUTES) Figure 19. Inactivation of Fatty Acid Synthetase at 57° in TED Buffer by Octanoate at Various Concentrations. Purified fatty acid synthetase (20 n1, 0.05 units of overall activity) was added to TED buffer (250Id) containing 0 (e), 0.02 (D), 0.1 (A), or 0.5% (I) sodium octanoate. The samples were incubated at 57° and were assayed intermittently for over- all FAS activity. 72 .mun>nuoo mo nos omouosmon oco~ooom nosuno: mounnncxo xooa mnuoc osu co nomnsosn manuaoanmoo noumow ose .csocm an uconmonw sumo mo COnuo:Onuoonm noon mocnouno En owm no onnwono :0nu:~o one .omm no mouscne mg now Eon oooaom no nos cOnuowawnnucoonuH: .mmOnnod GOHuonaocn omosu noumo umOa uoz >un> Inuoo mo ozu mo nano>nuooamon . moo moo no» aOm ao anouosnxonoa< .Amv mousana wbm mam «AOV men .Aev mm «boy 0 n0w own up acnuonsocn noumo mmozuoz an co>nw onamooono onu an monsoono muconmonw zunmcom omonoom on monaoao ono3 ALE mm.ov ouoaon~< .AnE Om.dv ouoOCouoo *m.o wcncnoucoo nowwsn owe cunz mo:n@Eoo moB Azun>nuoo damno>o mo nuns: :>.o «#8 m.ov ommu Ionucxm mnom zuuom monmnnsm .ouoOCouoo Rm.o Mm nommsm owe an ONW no omouosucmm mno< huuom mo coHum>nuomcH osu mo mnMNaoc< zunoo~o> :oHumucoEnmom usonmono Nunoaom omonosm .om onswnm 73 Figure 20. 5:55 wk fl 8:53. mh~l_ I r a . Ilflllllluh (1 SEE... N» W! I I 85550 20:8 _ 7. scam/V N... a a H r a2 ; "'— HONVS HOSSV 74 a confirmation of the enzyme which eventually aggregates to high mole— cular weight material. In the presence of pellet fraction suspension containing 0.1% octanoate, the octanoate protected the enzyme from inactivation to a small extent (Figure 21). Fatty acid synthetase in premicrosomal and microsomal pellet fraction suspensions containing 0.1% octanoate (0.13 units overall FAS activity/ ml) had half-lives of overall FAS activity of 92 and 68 minutes; in the absence of octanoate, half-lives were 68 and 49 minutes, respectively. The decreased rate of overall FAS activity loss in pellet fraction suspensions containing octanoate suggests that the conformational change(s) induced by octanoate, although slightly destabilizing to FAS, protects the enzyme to some extent from interaction with the inactivating capacity of the pellet fraction; that is, octanoate seems to block the site(s) of interac- tion between the pellet fraction and the enzyme. DISCUSSION These studies provide a preliminary characterization of the in- activation Of fatty acid synthetase by pellet fractions obtained from pig liver homogenates. The microsomal pellet fraction is dis- tinctly different from the other pellet fractions with respect to inactivating capacity. Microsomal pellet fraction suspensions have 40% and 110% more inactivating capacity (per gram) than the premicro— somal and nuclear and cell debris pellet fractions, respectively. Also, almost all the inactivating capacity of the microsomal pellet fraction is soluble after resuspension Of the fraction at room temperature. Only about one-fourth of the inactivating capacities of the premi- crosomal and nuclear and cell debris pellet fractions is soluble after resuspension at room temperature, and only one-half is soluble after incubation of the suspensions at 37° for 20 hours. The rate and extent of solubilization of the inactivating capacity in the various pellet fractions may be explained most easily by either or both of the following possibilities ; (a) the inactivating capacity of the microsomal pellet fraction is chemically different 75 .COmnnmano now csozm ono ouoocouoo *~.o mononoucoo nowmsp me an no nowwsn my on osxnco noM moumn conum>nuoocn ore .>Ho>nuooamon \@35an No man mm ono3 ouoosouoo &H.o wancnoucoo mnOnmcoamao cOnuoonm uoanoo Anr noEOmonOnEona moo An; ngOmononE ocu an mo>nn Iago: .z~o>nuooaoon amouscde mm mom a: ono3 mGOnmcoamom :Onuoonm uoaaoa AOV HmEomonOnEono mco Alv nmEOmonOnE mnomcoum one an xunzuuo mfg :onoa/o mo 3372mm .monnEoo mo monnom nonnmnoo m mo one On ouoocouoo &H.o wcnmmo mo cOnuaooxo ocu nun3 chnunmsoo mnomaoum nomc: mouanoco moo monoaona ono3 ocOHmcoamsm .ouoocouoo &H.O.wcn:noucou nommsm me an no nomwsm we on onenmcommsm COnuomnm uofifiomINn own no owouozucxm mno< quom mo :oHuo>nuoo:H .nm anamnn 76 355.5 5.: 222555 332.58 a... .6 55. 3589.35 I'Ivh 35898.59... I..v\ 358225 Ill- 4 a; 77 from that of the other two pellet fractions, or (b) the inactivating capacities are compartmented differently in the various pellet frac- tions. The pellet fractions were comprised of different cellular materials as indicated by analysis using native PAGE and sucrose density gradient sedimentation velocity experiments. The nuclear and cell debris pel- let fraction contained relatively few solubilizable proteins, and most of these proteins had smilar relative mobilities on native polyacryla— mide gels. The premicrosomal and microsomal pellet fractions had iden- tical banding patterns on native polyacrylamide gels, except the micro— somal pattern had a very strong band (at a relative mobility of 0.12 with respect to bromphenol blue) corresponding to the protein ferritin. Sucrose density gradient sedimentation velocity'analysis of these two pellet fractions revealed that some of the premicrosomal material sedi- mented at the same, slow rate as the microsomal material. It was also shown that freezing of the premicrosomal pellet fraction suspension causes the formation of a considerable amount of particulate material which sediments at the same rate as the microsomal pellet fraction. ' These observations suggest that the manner in which the inactivating capacities are compartmented is responsible for the different rate and extent of solubilization of the inactivating capacities. The preceding statement is also supported by the similar effect of the various factors tested upon the rate of inactivation of FAS, re- gardless of the type of pellet fraction used (e.g., dithiothreitol prevented inactivation of FAS by any of the pellet fractions, and NADPH slowed inactivation by any of the pellet fractions). CHAPTER 4 Further Purification and Characterization of an Inactivating Factor for Fatty Acid Synthetase ABSTRACT The previous chapter reported the discovery of several particulate fractions from pig liver which were capable of inactivating fatty acid synthetase. The process of inactivation in pellet fraction suspensions was partially characterized, and it was shown that the inactivating capacity could be extracted. This chapter describes further purification and characterization of the inactivating factor from the microsomal pellet fraction. The steps in the further purifica- tion of the inactivating capacity involve: incubation of the microsomal pellet fraction in water at 20° to release the factor in soluble form, ultrafiltration to separate high molecular weight compounds, DEAR-cellulose chromatography, and Sephadex G-10 gel filtration. The inactivating factor is a small substance (molecular weight less than 750 daltons) and is of net neutral or slightly negative charge at physiological pH. It is stable at 100° in aqueous solution for at least five minutes, and it contains no detectable free sulfhydryl or disulfide groups. The elution profile of the inactivating capacity from Sephadex G-10 suggests that it is composed of several distinct compounds, presumably of similar physical properties since they copurify through so many steps in the purification. Iron may be a component of the inactivating factor. Several factors affecting the inactivation of overall FAS activity by crude preparations of the inactivating factor (pellet fraction . suspensions) were described in Chapter 3. The effects of these factors,and some additional ones, were examined using the further purified inactivating factOr. The inactivation process still follows first order kinetics. Addition of 1 mM dithiothreitol at any time during the inactivation process temporarily prevents, but does not reverse, inactivation of FAS. Addition of dithiothreitol at a rate of 4 mM/hr prevents inactivation of the enzyme indefinitely. The rate of inactivation is significantly reduced by 20 mM EDTA and 78 79 and by 200 uM NADPH; the stabilizing effect of NADPH is increased when 50 pM acetyl—GOA and 50 pM malonyl-GOA are also present. The rate of inactivation is very slightly reduced by 50 pM acetyl-COA, by 50 pM malonyl-GOA, and by 40‘pM palmitate. The mechanism of inactivation is not yet known. Because EDTA decreased the rate of inactivation of FAS by the inactivating factor, suggesting the presence of a multivalent cation, and because ferritin (about 1.5 mg/ml) inactivates FAS, 1 mM ferric ion was investigated as a model compound for the inactivating factor. Ferric ion was an unsatisfactory model compound for the inactivating factor because the kinetics of inactivation were different for the two substances. In high concentrations of dithiothreitol (2-4 mM) inactivation of FAS by ferric ion occurred extremely rapidly after a brief period of time during which the overall FAS activity was stable; inactivation of FAS by the inactivating factor was prevented for a period of time, and then occurred at a rate similar to that observed when dithiothreitol was present only at a very low concentration (0.2 mM). 80 INTRODUCTION An inactivating capacity for fatty acid synthetase, derived from the particulate fraction Of pig liver, was reported in the previous chapter. A procedure for preparation of inactivating pellet fractions and a partial characterization of the inactivation process were given. It was also discovered that the inactivating capacity from the microsomal pellet fraction could be easily extracted in soluble form. This chapter describes the further purification of the inactiva- ting capacity from the microsomal pellet fraction and some further observations concerning the process of inactivation. MATERIALS AND METHODS . Reagents. Malonyl-COA, COA, and NADPH were products of P-L Biochemi- cals. Dithiothreitol was a product of Eastman-Kodak, and 2-mercaptoeth- anol of Mallinckrodt. Palmitic acid,5,5'-dithiobis(2—nitrobenzoic acid), and sodium nitroprusside were products of Sigma Chemical Co. Dowex 1 (OH-form, 200-400 mesh) and Dowex 50W-X8 (H+'form, 50-100 mesh)were obtained from Dow Chemical Co. CM-cellulose (standard, 0.67 meq/gm) and DEAE-cellulose (standard, 0.91 meq/gm) were products of Schleicher and Schuell, and Sephadex G—10 and Sephadex G-25 (coarse) Of Pharmacia Fine Chemicals. Acetyl-COA was prepared by a modification of the method of Simon and Shemin (1953), using continuous diethyl ether extraction of the acidified reaction mixture for four hours to remove reaction by-products. All other reagents were of analytical reagent grade. Livers. Pig livers were obtained from Peet Packing CO. (Chesaning, MI) within two hours of slaughter and were frozen on dry ice. The livers were stored at -30°. Fatty AcidySynthetase. Fatty acid synthetase was prepared from pig liver by the procedure given in Chapter 2. Assay for the Inactivation Factor. The inactivating factor was assayed by measuring the rate of overall FAS activity loss in the presence of the inactivating factor. The assay for overall FAS 81 activity is given in Chapter 2. The incubation mixture was fatty acid synthetase in PED buffer and the appropriately diluted sample to be tested for inactivating factor, combined on ice. Sufficient FAS was added to the incubation mixture to give an absorbance change of 0.05 per minute when 20 pl of incubation mixture was assayed. The incubation mixtures were then taken to 37°, and at periodic intervals, samples (201pl) of the incubatiOn mixture were removed and assayed for overall FAS activity. One unit of inactivating factor was defined as the amount of inactivating factor required to produce the loss of one unit of overall FAS activity in one minute at 370 in the inactivation assay. Measurement of pH and Conductivity. Values of pH were determined at room temperature with a Radiometer PHM4c pH meter equipped with a GK2321C electrode. Conductivity was measured with a Radiometer CDM2e conductivity meter equipped with CD0114 conductivity cell (cell constant 0.56 cm). Absorption Spectra. Ultraviolet absorption spectra were obtained with a Beckman DB spectrophotometer attached to a Sargent Model SRL recorder. Sulfhydpyl and Disulfide Analysis. Sulfhydryl content of samples was determined by denaturing any protein present with sodium dodecyl sulfate and then reacting the sulfhydryl groups with DTNB (Habeeb, 1971). Sample was diluted to 1.0 ml with a solution containing 2% sodium dodecyl sulfate, 80 mM sodium phosphate, pH 8.0, and 0.5 mg/ml EDTA. To 0.5 ml of the solution was added 0.02 ml DTNB solution (40 mg DTNB in 10 ml of 0.1 M sodium phosphate buffer, pH 8.0). The color was developed for 15 minutes and the absorbance at 410 nm determined against the remaining 0.5 ml of sodium dodecyl sulfate- treated sample, to which DTNB was not added. A reagent blank is subtracted from the apparent absorbance to give the net absorbance. For calculation of sulfhydryl content, the net absorbance is employed with a molar extinction coefficient of 13,600 m-1cm-1. Disulfide content was determined by the reduction of disulfide to free sulfhydryl in BM urea followed by the reaction Of the sulf- hydryl with DTNB (Cavallini, Graziani, and Dupre, 1966). Test tubes, 10 82 x 75 mm, with marks at 0.6 and 1.2 ml, were used. The following material was added to the tubes in the order shown; 0.29 grams solid urea, 0.02 ml of 0.1 M EDTA,.l-.2nu.sample, 0.2 ml of 2.5% sodium borohydride, water up to the 0.6 ml mark, and a drop of n-octyl alcohol as an antifoaming agent. The tubes were shaken at 38° to dissolve the urea and were then kept at this temperature for 30 minutes. At the end of the temperature treatment, 0.1 ml of 1M KH2P04 containing 0.2 M H01 was added. The wall of the test tube was carefully wetted with the solution in the tube in order to destroy traced of NaBflg adhering to it. Five minutes after the addition of the KH2?O‘-HC1 solution, the destruction of NaBH‘Iwas completed by adding 0.4 ml acetone. The mouth of the tube was closed with Parafilm and the tube inverted several times to bring the solution in contact with the wall. Nitrogen was bubbled slowly through the solution for five minutes. The DTNB solution (0.1 ml of 0.01 M DTNB in water) was then added to the solution, and the solution brought to 1.2 ml with water. Nitrogen was again bubbled through the solution for 2 minutes. The tube was then stoppered and allowed to stand at room temperature for 15 minutes. The absorbance at 412 nm was then determined against water in a Gilford 2000 spectropho- tometer. The absorbance of blank solutions, which contained all the reactants except the sample, were subtracted from the sample absorbances. A standard curve was constructed from absorbances. for samples which contained 0.25—40 nmoles dithiothreitol. Concen- trations of disulfides were determined from this standard curve. RESULTS Chapter 3 has described initial studies on the inactivating capacities from pig liver. The inactivating capacity from the micro— somal pellet fraction was shown to be extracted upon resuspension in TE buffer at room temperature. Inactivating capacities of the other pellet fractions were only partially extracted, even after incubation at 37° for 20 hours. Also, the total inactivating capacity 83 of the microsomal pellet fraction was greater than that of the other pellet fractions. For these reasons, the microsomal inactivating capacity was chosen for further purification and Characterization. Development of washingyand Extraction Procedures. The first step in purification of the inactivating factor was the purification of the microsomal pellet fraction. This was accomplished in Chapter 3 by repeated washing of the microsomal pellet fraction in TE buffer at 4°. This buffer, of moderate ionic strength, apparently immediately extracts the inactivating factor from microsomal pellet fractions at 200 (Table 4). Hence it is expected that some fraction of the inactivating capacity would be released from the pellet fraction during washing with TE buffer, even at 4°. So water at 0° was tested as a washing solution for the microsomal pellet fraction. The amount of inactivating capacity in the wash supernatants decreased with each washing. The first, second, and third wash supernatants had inactivating capacities of 1.3 x 104% 5.1 x 10‘6 capacity decreased to very low levels (the apparent inactivating -6 , and 1.5 x 10 -6 units/ml, respectively. The inactivating capacity of water is about 10 units/ml) as the remaining soluble components of the homogenate were removed (Figure 22). Thus, the inactivating capacity in the wash supernatants is apparently derived from the soluble components, not the pellet fraction. This is consis- tent with the discovery of inactivating capacity in the clarified centrifuged extract of pig liver homogenate which was reported in Chapter 3. Therefore, very little inactivating capacity seems to be released by the microsomal pellet fraction during washing with ice-cold water. The washed microsomal pellet fractions were extracted by suspen- sion in water for two hours at 20°. The amount of inactivating factor extracted was equal to the inactivating capacity in the pellet fraction suspension. The extract, which was separated from the extracted pellet fraction by centrifugation, was of sufficiently low ionic strength to be applied to ion exchange columns (see below). Also, the absence of added buffer in the extract reduced the amount of ionic substances eluted from Sephadex columns with the inactivating 84 Figure 22. Purification of Microsomal Pellet Fraction by Washing with Water at 0°. Pig liver was homogenized in water at 0° and the homogen- ate was centrifuged at 18,000 rpm for 15 minutes at 0° in a Sorvall 88-54 rotor. The supernatant was centrifuged again in the 88-54 rotor under identical conditions. The supernatant from the second centrifu- gation was then centrifuged at 00 for 60 minutes at 42,000 rpm in a Beckman 42.1 rotor. The supernatant from this centrifugation is the first wash supernatant (pH 6.54, conductivity 2.15 mmho). The pellets from this centrifugation were taken up to 20 ml with water at 0° and were resuspended using a Potter-Elvehjem homogenizer. The suspension was taken up to 150 ml with more water at 0° and was then centrifuged once again in the Beckman 42.1 rotor as described above. The super- natant from this centrifugation is the second wash supernatant (pH 6.62, conductivity 520 nmho). Further wash supernatants were obtained by repeating the resuspension and centrifugation steps. 85 Figure 22. (angrily... 86 factor (see below). It is useful to minimize ionic strength for an- other reason as well; fatty acid synthetase is inactivated at high ionic strength (Table 6). Table 6. Spontaneous Inactivation of Fatty Acid Synthetase in Solutions of TE Buffer of Various Ionic Strengths. Fold normal TE buffer concentration8 1 5 9 13 17 Half-life of overall FAS activity (min) 900 510 335 265 218 aNormal TE buffer contains 50 mM tris(hydroxymethyl)aminomethane, pH 7.4, and 1 mM EDTA. Incubation was carried out at 57°. The inactivation rate is not proportional to TE buffer concentration. A more highly inacti- vating solution gives a shorter half-life for overall FAS activity. Preliminary Characterization of the InactivatingyFactor and the Develop- ment of Separations Based on Molecular Size and on Net Chgpgg. The goals of (1) development of a purification procedure for the inactiva- ting factor, and (2) characterization of the inactivating factor, were closely interrelated. Studies in either area yielded information use- ful in the other. For example, examination of prospective purification steps based on the size of the inactivating factor (gel filtration, dialysis, ultrafiltration) yielded some knowledge of its size. Simi- larly, examination of separations based on the net charge prOperties of the inactivating factor (Dowex and DEAE- and CM-cellulose chromato- graphy) yielded information about its net charge. Early experiments were undertaken to determine the size of the inac- tivating factor. The initial test was based on gel filtration. 'Micro- somal pellet fraction suspension in TE buffer (1 g pellet fraction/ 10 m1 suspension) was centrifuged at 25° in a Beckman 42.1 rotor for 70 ‘minutes at 40,000 rpm. The clear yellow supernatant from this centri- fugation (20 ml) was applied to a Sephadex G-25 column (coarse, 2.2 x 50 cm) and eluted with TE buffer; 0.051 of 0.054 units of inactivating capacity (90%) was eluted in the low molecular weight region. There- fore, the molecular weight of the inactivating factor was less than the exclusion limit of the gel (5,000 daltons). 87 The next test of size for the inactivating factor consisted of dialyzing the microsomal pellet fraction in aqueous suspension (4 g pellet fraction in 16 ml water) against 500 ml water at 20° for two hours. This was followed by overnight dialysis at 4° against the same 500 ml water, to allow diffusion of low molecular weight components (less than 14,000 daltons) across the dialysis membrane to reach equilib- rium. A considerable amount of inactivating capacity was found outside the dialysis casing, substantiating the small size of the inactivating factor as determined by gel filtration. Subsequently, ultrafiltration was used to separate the solubilized inactivating factor from the microsomal pellet fraction and proteins. Ultrafiltration was performed using an Amicon PM-30 ultrafiltration mem- brane. Molecules of molecular weight more than 30,000 daltons were re- tained, while the inactivating factor passed through (as greater than 80% of the inactivating capacity applied). Tests for separation of the inactivating factor from contaminants by gel filtration on Sephadex G-10 columns also provided evidence that the inactivating factor was very small, since most of the inactivating capa- city (greater than 802 of that applied) was eluted in the low molecular weight region (see below). Hence, the molecular weight of the inacti- vating factor is less than 750 daltons (the exclusion limit of the gel). Tests for separation of the inactivating factor from contaminants by ion exchange chromatography showed that the inactivating factor has neu- tral or slightly negative net charge near neutral pH. Ultrafiltered ex- tract of microsomal pellet fraction suspensions was applied to columns (0.6 x 16 cm) of Dowex BOW-X8 (H+ form), Dowex 50W-X8 (Na‘+ form), and Dowex 1 (OH. form), and the columns were eluted with water (Table 7). The eluate from the Dowex SOW-XB (Na+ form) column contained all the applied inactivating capacity, and apparently more. So the inactivating factor is apparently chemically altered when passing through this column, and is not bound. The eluate from the Dowex 50W-X8 (H+ form) column con- tained about 10% of the applied inactivating capacity. Again, the inac- tivating factor appears to have been chemically modified, this time the modification decreased the effectiveness of the inactivating factor in inactivating fatty acid synthetase. The eluate from the Dowex 1 (0H- form) column had essentially no inactivating capacity. Since the inac- tivating factor was bound by this column, it apparently has a net 88 negative charge. Table 7. Binding of Partially Purified Inactivating Factor to Dowex Resins. Resin useda Total Inactivating Total Inactivating Capacity of Applicate Capacity of Eluate (units) x 103 (units) x 103 Dowex sow-xs (Na+ form) 1.5 4.0 Dowex 50W-X8 (H+ form) 8.2 0.82 Dowex 1 (OH' form) 8.2 <0.1 aInactivating factor taken through Step 2 in its purification procedure was applied to 0.6 x 16 cm columns of the resins. The columns were eluted with water at 23°. The eluate was taken to pH 7.4 with 6M NaOH or 6M HCl prior to assaying for the inactivating factor. To confirm the net charge on the inactivating factor, DEAE-cellulose and CMecellulose chromatography were performed on the extracted, ultra- filtered inactivating capacity from the microsomal pellet fraction. Binding of the inactivating capacity to CM—cellulose could not be demon- strated. At sufficiently low ionic strength, however, the inactivating capacity was bound quantitatively to DEAE-cellulose. When eluted with a 0 to 0.25 M NaCl gradient, the inactivating capacity peaked at 50 mM NaCl (Table 8), again suggesting a slightly negative net charge on the inac- tivating factor. Since the binding of a nonionic surfactant, Triton X-100,tx>DEAE-cellulose has been demonstrated (unpublished results from Table 8. Inactivating Capacities of Fractions Obtained from.DEAE-cellu- lose Chromatography During Purification of the Inactivating Factor. Fraction number8 37 38 39 40 43 Inactivating ca acity (units) x 10° 0.56 1.09 0.44 0.17 0.44 aFractions 38 and 43 correspond to the peaks at 3.3 and 6.1 mmho conduc- tivity, respectively, in Figure 24. The first peak has ten times the absorbance of the second at 254 nm. 89 this laboratory), the binding of the inactivating factor to DEAR-cellulose may be due either to a negative charge, or to nonelectrostatic interaction between the inactivating factor and DEAE-cellulose. The studies just described led to the following standard purification procedure for inactivating factor from microsomal pellet fraction. This procedure is summarized in Figure 23. Standard Procedure for the Partial Purification of Inactivating Factor from the Microsomal Pellet Fraction of Pig Liver. Distilled deionized water was used throughout. Step 1. Microsomal pellet fraction isolation. Frozen pig liver (800 g) was cut into small pieces and homogenized for 90 seconds with 1600 ml water at 4° in a waring blendor. The homogenate was centrifuged at 6,700 rpm for 10 minutes in Sorvall GSA rotors at 0°. This centrifugation re- moved larger organelles. connective tissue, and whole cells. The super- natant was centrifuged at 18,000 rpm for 15 minutes in Sorvall SS-34 ro- tors at 0°. The supernatant from this centrifugation was centrifuged a- gain under identical conditions to complete removal of mitochondria, lyso- somes, and peroxisomes. The supernatant from the repeat centrifugation was then centrifuged in a Beckman Type 21 rotor at 21,000 rpm and at 0° for 150 minutes. The microsomal pellet fraction was obtained from this centrifugation. The pellet fraction was washed by resuspending the pel- lets in ice-cold water to a volume of 150 ml using a "large" clearance Dounce homogenizer, and then centrifuging the suspension at 40,000 rpm in a Beckman Type 42.1 rotor at 0° for 70 minutes. Three washes were com- pleted before continuing to the next step. Step 2. Extraction of the inactivating capacity and separation from .pigpymolecular weight contaminants. Four grams of washed microsomal pellet fraction were suspended in thirty-six milliliters of water by Dounce homogenization at room temperature. After incubation at room tem- perature for two hours, the suspension was centrifuged in a Beckman Type 42.1 rotor at 20° for 70 minutes at 40,000 rpm. The clear yellow super- natant (36 ml) was applied to an Amicon Mbdel 52 pump with a PM3O ultra- filtration membrane and was filtered at 15 psi. The pale yellow fil- trate (32 ml) was collected. The solution retained in the filtration ap- paratus was golden yellow and turbid. Step 3. DEAE-Cellulose chromatography. A DEAF-cellulose column (1.25 x 12 cm) was prepared and equilibrated with TE buffer. The column was 90 .nOuoem.anuo>nuoeoH noes-omonn mo segueonmnnsm can now unono scum .NN onsunm an noun m50v A: noum mcov onaaoonn mooamon up senuonooe> Io ansuon an nauoow wonuu>nu xooo 3: Am noum mcov on: I n mouauo mo COnuenusoocoo acnuonunnw now ouonmoenoucn on *mn 1'. . H<¢HAHmcman= «acoononoeonzo occaonnoonmcmn onnu xomosoom 7‘1 xuom 3: so. on mow An doom mcov unso: osu noH com um cannonoonn zo~so.|, .I soon: $.55: ONES—5km do: eon—39, on 3 50:52:58.. A<20momouzflmmm<3 poses a as 3: “w”; Axnmu ouooowoeo: oau zeahummm Hannah mo oucocoaeoo enaonoe o no nouns an mcoousuon Aosen cu seasons or .w w ooo.:m_ monmnnodo :Onuuwounnucoo noonon III IInzo5on o» ona.m~ aw x ooo.on acnuowounnucoo unooon az<5n~ Eon» non cm: as my maezmoozo: E 555 or. :9... more: "05.22.35 no 282355... «on 5.889... V .‘l 91 then washed with water until the conductivity of the eluate was less than 5 nmho. The conductivity of the ultrafiltrate from the previous step was adjusted to less than 15 pmho and was then applied to the column, after which the column was washed with 30 ml water. The column was eluted with a linear 0 to 0.25 M NaCl gradient (104 ml, 52 ml each reservoir). A characteristic elution profile at 254 nm was obtained (Figure 24). Twen- ty drop fractions (about 0.8 ml) were collected using an ISCO Model 328 fraction collector. The peak of inactivating capacity eluted at 3 mmho conductivity (about 50 mM NaCl). Fractions with inactivating capacity were combined and concentrated by rotary evaporation under vacuum at 40° to a volume of less than 2 ml. Step 4. Sephadex G-lOpgel filtration. The concentrated inactivating factor from the DEAE-cellulose chromatography was applied to a Sephadex G-10 column (1.25 x 99 cm) and was eluted with water at a flow rate of 6.6 ml per hour. Twenty drop fractions (about 0.7 ml) were collected. A typical elution profile at 254 nm is shown in Figure 25. More than 80% of the inactivating capacity eluted in the NaCl (low molecular weight) peak. Additional Properties of the InactivatingyFactor. The studies described at the beginning of the Results indicated that the inactivating factor is small (less than 750 daltons) and has a net neutral or slightly negative charge near neutral pH. Some additional tests were performed on the inac- tivating factor in order to further elucidate its physical and chemical structure. The results of these tests are presented below. Stability in aqueous solution at 100°. Inactivating factor extracted from the microsomal pellet fraction, but not yet ultrafiltered, was taken to 100° in a boiling water bath and was allowed to incubate for five minutes. After incubation, inactivating capacities of both unincubated and incubated extracts were determined. No inactivating capacity was lost as a result of the incubation. ‘ Sulfhydryl and disulfide content. All tests for sulfhydryl groups and disulfide bonds in the inactivating factor were negative. The inacti- vating factor for these experiments was obtained by performing Sephadex G-25 gel filtration on the extracted inactivating capacity from the micro- somal pellet fraction. High and low molecular weight eluates from the column were combined separately and were concentrated to levels twenty times those in the applied samples. When assayed for free sulfhydryl 92 .onswnm ozu co oucnoo on co>nw onm moOHuomnw ~mno>oo mo monunooaoo wanuo>nuoo5n one .zun>nuosmcoo onEE m.n no moxooa zunooooo wanum>nuoocH .ounomom osu an co>nw onomooona ocu 5n onouonoosou Soon on mouano moo ceanoo ocu on mongooo conu mos wannou oeu omnmuoo 50nu=~Om one .onSuonoQEou Soon as sum o now nouns unonowo cOnuoonm uofinoo noEOoonons wannanonm an mocnouno mos Ana wcnv oHQEom monnaoo one .nOuoom onuo>nuooc mo conwmowmnnsm nonunmm now 5: :mm no oanmonm cOnusaw >£oonMOuoEonno ooo~=~nooumom mo mmwuwuwamo wcfium>wuumcw use .mufismmm ca cm>ww musvmooua msu Cu wawouooom upsumumanu Econ um noum3 saws amusflw mm3 :E:~ou use .m:E:~oo unawquumEouno omodaadmoummu Scum unawuomuw wcwum>wuomcw mcu mo wuwmoaeoo wmumuucmocou a «ma wdaawo wwwaaan use .uouowmwwcauu>fiuowaH mo cofigmofimwpsm Hmwuumm you a: :mm um ofiwmoum aofiusfim cofiuwuufiwm duo o~-u xonmnmmm .wm ouswfim 95 EDI x (sum) movavo SNILVAILOVNI 3 °° ‘° V N :3 '(Owaf mmfincmoo a 8 w a a g f I 1 . 41f , J5 E *3 2 E =3 8 S .— U \ 8 . 8 #8 lax—+4 < la ! c. m c: m c N .3 .: =5 UJU vszv FRACTION NUMBER 96 and disulfide content, less than lO‘pM free sulfhydryl and 20‘pM di- sulfide were found in the low molecular weight concentrate. The protein- containing high moelecular weight concentrate had 200 pM disulfide and less than ZOIpM free sulfhydryl. Heterogeneity:several copurifyinggcompounds. The inactivating factor purified using the standard procedure apparently contains at least two major components of different molecular weight (Table 9). Most of the Table 9. Inactivating Capacities of Fractions Obtained from Sephadex G-10 Gel Filtration During Purification of the Inactivating Factor. Fractiona Conductivity Inactivating Capa- (mmho) city (units) x 103 54 (leading edge of high MW peak) 0.15 0.31 57 (high MW peak) 0.10 0.27 63 (intermediate MW region) 0.61 1.59 68 (leading edge of low MW peak) 6.45 0.34 73 (low MW peak) 24.0 710 79 (trailing edge of low MW peak) 0.01 0.15 8The fractions are from the gel filtration whose elution profile is shown in Figure 25. The portions of the elution profile from which the frac- tions were taken are indicated in parentheses. inactivating factor eluted in the low molecular weight (NaCl) peak (80%), but an additional 152 eluted in the fractionation range for the gel (the 'intermediate' molecular weight range). The remaining inactivating capa- city was spread throughout the eluted fractions. The inactivating capa- cities had associated with them an absorbance at 226 nm, 270 nm, or both (Figure 26). ‘ Extractibility in several organic solvents. The extractibility of inac- tivating factor in several organic solvents was determined. Addition of hydrochloric acid to 0.15 M to the inactivating factor did not affect its absorption spectrum for at least 24 hours, so the extractions were per- formed on inactivating factor (with associated absorption maxima at 226 nm and 270 nm) in 0.15 M HCl. Extraction with either cyclohexane or methyl- ene chloride gave similar results; the material with absorption maximum at 226 nm.was not extracted, and that with absorption maximum at 270 nm was 97 .1 . .,| l+§ 73 ABSOR BANCE -—-—- zoo no 60200 ' ' 250‘ an WAVELENGTH (NM) WAVELENGTH (NM) Figure 26. Ultraviolet Absorption Spectra of Fractions from Sephadex G-10 Gel Filtration for Partial Purification of Inactivating Factor. These fractions are from the same gel filtration column described by Table 9 and Figure 25. Fraction 7} contained the greatest amount of inactivating capacity. Absorption spectra of fractions 5h, 57, and 65 were measured at thirteen-fold dilution; those of the other frac- tions were measured at thirty-one -fold dilution. Absorption spectra were obtained using a Beckman DB spectrOphotometer. 98 only partially extracted. When taken to dryness by rotary evaporation under reduced pressure, the extracts yielded oily orange solids. Extrac- tion with diethyl ether gave a turbid colorless solution. Ultraviolet analysis of the extract was not possible because of the very high back- ground absorbance of the remaining ether. Complexed ferric ion. The presence of ferric ion was consistent with several observations made during the purification and characterization of the inactivating factor. The inactivating factor gave a translucent orange, noncrystalline solid when taken to dryness by rotary evaporation. It gave no absorption maxima in the visible absorption spectrum. Also, at several steps in the purification of the inactivating factor, orange. precipitates occasionally formed; formation of the precipitate was assoc- iated with the loss of inactivating capacity. The redissolved precipi- tates gave ultraviolet-visible absorption spectra without maxima, with absorption increasing monotonically with decreasing wavelength. These absorption characteristics are consistent with those for ferric ion. The inactivation of fatty acid synthetase by the inactivating factor is inhibited by EDTA (see below); however, EDTA did not eliminate the inactivation of the enzyme. Also, DEAE-cellulose will not bind a cationic metal ion. Thus, if a metal ion is present in the inactivating factor, it must be in a complexed form which has a net neutral or negative charge. The behavior of the inactivating factor on Dowex resins can be easily ex- plained if the inactivating factor has s complexed metal ion. 0n Dowex l, the inactivating factor would be degraded to metal ion, which binds to the column, and free ligand, which is released and eluted from the column. 0n Dowex 50W-X8 (H+ form), the inactivating factor would be degraded in part, so that a small amount of free metal ion might elute from the column, and so account for the small inactivating capacity observed in the eluate. On Dowex SOW-XB (Na+ form), the ligands might be more completely stripped, or the complex changed in some other manner, so that the free metal ion would be eluted quantitatively from the column. The effect of ferric ion on fatty acid synthetase will be presented shortly. Further Characterization of the Inactivation Process. Tests of the inac- tivation process using the further purified inactivating factor gave re- sults similar to those obtained in Chapter 3 using the inactivating capa- city in microsomal pellet fraction suspensions. Continuous addition of 99 dithiothreitol to the incubation mixture at a rate of 4 mM per hour was sufficient to almost halt the inactivation of fatty acid synthetase, even in extremely high concentrations of inactivating factor (Table 10). Addi- tion of 1 mM dithiothreitol was sufficient to halt, but not reverse, the Table 10. Periodic Addition of Dithiothreitol to Incubation Mixtures Pre— vents Inactivation of Fatty Acid Synthetase by Inactivating Factora. Incubation time (minutes) 0 38 78 152 250 Fraction of no DTT overall FAS added 1'00 0°03 0 0 0 __ activity DTT remaining addedb 1.00 0.91 0.90 0.76 0.60 8The sample was the concentrated inactivating capacity obtained at the end of Step 3 in the purification of the inactivating factor. bDithiothreitol concentration in this sample was increased by 1 mM every 15 minutes with a 100 mM stock solution of dithiothreitol. The incuba- tion was carried out at 37°. inactivation process. The rate of inactivation of the enzyme was also sig- nificantly reduced in the presence of 20 mM EDTA, 200 pM NADPH, and especi- ally 200 pM NADPH with 50‘pM acetyl-CoA and 50‘pM malonyl-GOA. Malonyl-CoA, acetyl-CoA, or palmitate alone only slightly stabilized the enzyme (Table 11). Table 11. Effectors of the Inactivation of Fatty Acid Synthetase by Inacti- vating Factor at 37°. Additional component in NOne Palmitate AcCoA MalCoA NADPH EDTA MalCoA (SO‘pM), the incubation (40 M) (50 pH) (50 M) (200 pH) (20 mM) AcCoA (50 pM), Inixtureél NADPH (200 ,uM) Relative inactivating 1.0 0.68 0.74 0.88 0.43 0.34 0.21 capacity aIncubation mixture contained fatty acid synthetase and Amicon ultrafil- trate from the purification of the inactivating factor (latter at two- thirds its initial concentration). 100 Inactivation of Fatty Acid Synthetase by Ferric Ion Differs Greatly from That by Inactivating Factor. Several observations (see previously) were consistent with the presence of iron in the inactivating factor. Also, ferritin, which is present in large amounts in the soluble extract from microsomal pellet fractions, inactivates fatty acid synthetase. Freshly purified, fully reduced ferritin at about 1.5 and 8 mg per m1 inactivated half the overall enzyme activity in a standard incubation mixture con- taining 1 mM dithiothreitol in 750 and 350 minutes, respectively. This suggested that ferritin might be the source of the inactivating factor; ferrous ion might be released from ferritin, oxidize, and then react with fatty acid synthetase. The capacity of ferric ion to oxidize free sulf- hydryl groups at concentrations as low as 0.1 mM is well established. Thus, the inactivation of fatty acid synthetase by ferric ion would be ex- pected to closely model that by inactivating factor if an oxidative modi- fication of the enzyme is a part of the inactivation process. For this reason, ferric ion was tested as a candidate for the inactivating factor. Ferric ion was obtained from ferric chloride brought to neutral pH with l M NaOH solution. Inactivation of fatty acid synthetase by ferric ion (1 mM) occurred at a rate comparable to the inactivation of the enzyme by inactivating factor when using the extracted material at Step 2 in the purification procedure as the source of inactivating factor (Figure 27). The kinetics of inacti- vation were the same using either ferric ion or inactivating factor when the dithiothreitol concentration was low (less than 0.2 mM). At higher dithiothreitol concentrations, however, dithiothreitol protected the en- zymg from inactivation by inactivating factor, but accelerated greatly the inactivation of the enzyme by ferric ion. For this reason, ferric ion proved to be a poor model for the inactivating factor. DISCUSSION These studies provide a partial characterization of the inactiva- ting factor derived from the microsomal pellet fraction of pig liver. The data show that the inactivating factor is a low molecular weight sub- stance (less than 750 daltons) of net neutral or slightly negative net charge. It is stable in aqueous solution at 100° for at least 5 minutes 101 A» ? V 1 mM Fe+3 and 1.2 :7“ "'4 mM DTT 1 mM Fe*3 and *‘2 mM DTT 1.0 F 0.8 - ,1 I <“" 1 mM Fe"’3 and g 0-6 0.2 mM on .—| 0.4 F """" 0.2 mM DTT L o/ 0.2 / A/ 2 mM DTT __A ......... o P’ \ 4 mM on o 100 200 300 INCUBATION TIME (MINUTES) Figure 21. Inactivation of Fatg Acid Synthetase by 1 mM Fe+5 in TE Puffer Containigg:Various Amounts of Dithiothreitol. Fatty acid syn- thetase and Fee were combined in a standard incubation mixture con- taining either 0.2 (O), 2 (A), or 4 mM (I).dithiothreitol. (DTT). The effect of TE buffer containing 0.2 (o), 2 (A), or 4 (0) mM dithio- threitol upon overall FAS activity is shown for comparison. 102 and apparently contains no free sulfhydryl nor disulfide groups. Its elution profile from Sephadex G-10 indicates that it may be comprised of a series of closely related compounds. Its visible absorption spectrum, with absorption monotonically increasing as wavelength decreases, is con- sistent with that of ferric ion. The ultraviolet absorption spectrum, which exhibits peaks at 226 nm and at 270 nm in varying proportion, sug- gests the presence of one or more organic compounds. Since the chemical structure of the inactivating factor has not yet been elucidated, further studies in this area are indicated. Oxidation of fatty acid synthetase as a step in its inactivation seems likely since dithiothreitol, a reducing agent, has an inhibitory effect on the inactivation. Because EDTA also has a (lesser) inhibitory effect on the inactivation, and because ferritin is a major component in the soluble extract from microsomal pellet fraction, ferric ion was sug- gested as a candidate for the inactivating factor. Tests revealed that ferric ion was not the inactivating factor. The data also showed that it was not a good model compound for the inactivating factor; at high dithio- threitol concentrations the inactivation of fatty acid synthetase by inac- tivating factor was inhibited, while inactivation of the enzyme by ferric ion was greatly accelerated. Although ferric ion was eliminated as the inactivating factor, oxida- tion was not eliminated as a mechanism for the inactivation. Detergent- like denaturation of fatty acid synthetase by the inactivating factor is very unlikely because an intermediate conformer was formed during incuba- tion of the enzyme with octanoate, and no such intermediate was observed during incubation of the enzyme with inactivating factor. The total mass of the inactivating factor is sufficiently small (considerably less than 0.12 of the mass of the microsomal pellet fraction, although not exactly known) that a nonspecific effect on hydrogen or electrostatic bonding is also unlikely (e. g., 1% isopropanol was needed to achieve inactivation of the enzyme at a rate far enough above that in buffer alone to be de- tected). Consequently, the best mechanism for the inactivation involves an oxidation, or other covalent modification, of the enzyme which re- sults in the enzyme having an increased tendency to aggregate. Further studies are needed to elucidate the exact mechanism. CHAPTER 5 ISOELECTRIC FOCUSING IN THIRTY-FIVE MINUTES ABSTRACT Certain unstable macromolecules require rapid analytical techniques for fractionation and analysis. This chapter describes procedures which greatly shorten the time required for isoelectric focusing in density gradients. Focusing times as short as thirty-five minutes were obtained for hemoglobin; the shortest time previously reported was two hours. Key characteristics of the present system are: (l) a short solution col- umn (50 mm): (2) incorporation of dilute K01 (0.03%) in the gradient column, the gel plug soak solution, and both reservoir solutions; (3) a uniform initial ampholyte concentration in the density gradient, at the lowest concentration consistent with the formation of a linear pH grad- ient (0.50%); and (h) incorporation of sucrose in the gel plug. In addition, this chapter also describes various conditions to further maxi- mize the linearity of the pH gradient, though at the eXpense of longer focusing times. Standard polyacrylamide gel electrophoresis apparatus is used except for the electrofocusing tubes, which are specially prepared from standard glass tubing. 103 104 INTRODUCTION Isoelectric focusing of macromolecules in density gradients has sev- eral major advantages over isoelectric focusing in polyacrylamide gels. These include: (1) rapid and quantitative recovery of sample for fur- ther analysis, (2) easier determination of pH values at each point in the tube, and (3) faster electrofocusing for comparable column heights. For these reasbns, there has been an interest in development and re- finement of faster and better techniques for isoelectric focusing in density gradient solutions (for references to reviews, see Behnke st 31., 1975). Massey and Deal (1973) described an apparatus and techniques for rapid isoelectric focusing. The key feature of this procedure was the use of a short (110 mm) density gradient column formed over an agar plug. Technical difficulties with the agar plug were overcome by re- placing it with a polyacrylamide gel plug (Behnke st 31., 1975). In addition, it was demonstrated that multiple tubes could be easily run simultaneously. This procedure for isoelectric focusing was a great improvement over that using the commercially available apparatus, which required up to 48 hours for focusing, but did not give focusing times short enough to accommodate certain unstable samples. For this reason, the factors affecting focusing time, as well as those affecting the ac- curacy and reliability of isoelectric point determinations, were sys- tematically studied. A number of factors affecting focusing time and linearity of the pH gradient have been identified. Under conditions where focusing time is minimized, the focusing time of hemoglobin has been reduced by a factor of almost four, to about 30 minutes. Procedures which yield greater resolution and maximal pH gradient linearity at the expense of longer focusing times have also been developed. MATERIALS AND METHODS Ethanolamine (95%) was obtained from Aldrich Chemical Co. Ampholyte (Biolyte 3/10) was purchased from BIORAD; it had a pH range of 5 to 10 and a concentration (w/v) of 40% solids. Bovine hemoglobin was obtained from Sigma Chemical Co. Ammonium persulfate, N,N'-methylenebis(acryla‘ mide), and acrylamide were products of EM Laboratories. Canalco 105 supplied N,N,N',N'-tetramethylethylenediamine. All other reagents were of analytical reagent grade. A standard polyacrylamide gel electrOphoresis apparatus was modified by replacing the grommets holding the PACE tubes in the upper reservoir with grommets to hold 10 mm (o. d.) glass tubing. Two sizes of electro- focusing tubes were used, a 10 x 90 mm tube and a 10 x 150 mm tube. Each was made from 10 mm (o. d.) glass tubing; a slight bulb was blown at the bottom of each tube as previously described (Behnke g£_gl,, 1975). The shorter tube was used for the ultimate test of minimum fo- cusing time. The longer tube was used in selecting Optimum electro- focusing conditions with respect to other variables because focusing time differences were greater, and deviations from linearity in the pH gradient were easier to detect, in the longer column. In preparing for an experiment, the electrofocusing tubes were pro- vided with a polyacrylamide gel plug in the bottom to support the den- sity gradient solution. The bottoms of the tubes were tightly sealed with Parafilm.and the tubes were filled to 3 mm above the tops of the bulbs with aqueous gel plug solution. This solution contained 7.5% acrylamide, 0.18% N,N'-methylenebis(acrylamide), and 25% sucrose. Af- ter addition of the gel plug solution, a layer of water was applied. Upon polymerization of the gel plug solution, the water overlayer was removed and the tubes were filled with, and immersed in, a gel plug soak solution for a minimum of seven hours. The soak solution con- tained 3% acetic acid, 0.03% KCl, and 25% sucrose. Soaking periods of longer than seven hours did not enhance or diminish performance of the plugs. Just prior to use, the soak solution was removed, the electrofocusing tube was rinsed with distilled water, and the tube was blotted dry with absorbent tissue. In the short focusing tube, a 2.50 ml (50 mm height) density gradient containing 5 to 12% sucrose was formed. In addition to sucrose, the gradient contained 0.50% ampholyte, 0.03% KCl, and 50 to 200 pg protein, all distributed uniformly throughout the gradient. When the long focusing tube was used, a 5.6 ml (112 mm height) 5 to 20% linear sucrose gradient with 0.86% ampholyte was formed; all other so- lution components were as for the short tubes. The lower electrOphoresis chamber was filled with 3% (v/v) acetic 106 acid solution containing 0.03% KCl. The electrofocusing tubes contain- ing the gradients were then moistened near their tops with glycerol and were inserted into the empty upper electrophoresis chamber. Upper re- servoir solution, 3% (v/v) ethanolamine containing 0.03% K01, was care- fully layered over each gradient until each tube was filled, and the upper chamber was then filled. The positive lead of the power supply was attached to the lower reservoir and the negative lead was attached to the upper reservoir. Electrofocusing was begun at 95 volts for the short tubes or at 200 volts for the long tubes, yielding an initial cur- rent of about three milliamperes per tube. Using this procedure, hemo- globin focused within thirty-five minutes in the short tube. After electrofocusing, the tubes were removed and mounted vertically on a stand; the upper reservoir solution was then removed from the tubes with a Pasteur pipet. The top of a tube was stoppered with a cork through which the barrel of an 18 gauge needle extended; the needle was attached to plastic tubing leading to the inlet of an ISCO Model UA-5 spectrOphotometric flow cell with an attached fraction collector. An- other 18 gauge needle, attached by plastic tubing to a syringe pump which contained 50% sucrose, was inserted upward through the gel plug until its tip was just above the gel plug. This Operation was carried out with the pump on slowly so that no air bubbles were introduced at the bottom of the density gradient. The 50% sucrose was pumped at a rate of 0.37 ml per minute by an 1800 Model 183 density gradient frac- tionator. Fifteen second (0.09 ml) fractions were collected for the short tube; thirty second fractions were collected for the long tube. The pH values of the fractions were determined using a Radiometer PHM4c pH.meter equipped with a GK2321C combination microelectrode. Focusing times were determined using hemoglobin as the test sample since its color allowed visual monitoring of the focusing process. Sharp protein boundaries were formed near the extremes of each column initially; they migrated toward each other and ultimately coalesced into a single, focused boundary. This process was described in detail in a previous paper from this laboratory (Behnke, st 31., 1975). RESULTS The major goal of this study was to determine the conditions which would minimize the focusing time for macromolecules subjected to iso- 107 electric focusing in sucrose density gradients. However, because of a sharp decline in pH in the lower third of the sucrose density gradients using the previous method (Behnke 35 31., 1975), it was first necessary to improve the linearity of the pH gradient. For these studies, as well as for studies on the effects of various factors on focusing time, the original, longer (112 mm, 5.6 ml) density gradient column was used. This allowed greater sensitivity in detecting differences in both focus- ing times and shapes of pH gradients. Unless otherwise indicated, the experiments described below were performed with the longer (5.6 ml) gradients. Sucrose in the Gel Plug: Effect on pH Gradient Linearity. ‘Working on the assumption that convection caused the sharp decline in pH in the bot- tom third of the density gradient, 25% sucrose was incorporated in the gel plug. This greatly improved the linearity of the pH gradient in the bottom portion of the density gradient, but at the expense of a slightly increased focusing time (Figure 28). Nevertheless, the improvement in linearity of the pH gradient was judged more important than the increase in focusing time, so inclusion of sucrose in the gel plug was adopted as part of the standard procedure. Acetic Acid as the Lower Reservoir Electrolyte: Effect on pH Gradient Linearity. Since the pH gradient was still steeper than desired at the bottom of the density gradient, a weaker acid was substituted for the strong acid generally used in the acid reservoir solution. The use of acetic acid instead of sulfuric acid substantially improved the linear- ity of the pH gradient by raising the pH at the bottom of the density gradient about two units (Figure 29). Hence, acetic acid (3%) was in- corporated in the standard procedure. Uniform Initial Ampholyte Distribution: Effect on pH Gradient Linear- ity, In the previous electrofocusing system.(Behnke g£_gl,, 1975), an ampholyte concentration gradient was used to complement the sucrose gradient, thereby providing a greater initial density gradient. Com- parison of this system with one having a uniform initial ampholyte con- centration showed the latter system to yield a more linear pH gradient (Figure 30). So uniform initial ampholyte concentration is a part of the standard electrofocusing procedure. 108 .%Ho>wuoodm uou .mouscwe 0mg can mam mums mEoumzm oBu omoco ca swoodono; pow moawu wcwmooom mafia flow ecu cw omouosm fimm wowumuoduoocw mo cowumoawaooa mech ecu cues no “A 6---; V 23: .MM mm oxcsom mo pocuoe use an ooeuowuoo mos wcwmooOM owuuooaoomH .MMwmsoom owuuooaoomH a“ ucofioouo xuwmcoo omouosm one wcwuuoamwm wsdm How ecu cu omouosm wcwmo< mo uoommm .wm ousmwm 109 Eaten Figure 28. Ems—dz 22.55.... cm 3.2535 5s: 83.: 32.2; .. €2,685 an: 83$ :22? 0— $3.52 . 110 A .>Ho>wuoodmou anous—SE 0mg can QNH mums mucoEwquxo osu omocu ca canofiono: How omega wcwmsoom .Auuunnv owes oauooo mm no mood owuom~sm m Hugues mp3 cowuodom awo>pomop-u03o o u an «omouosu mm confluence a n o mooou mcwmsoomouuoodo ago as awofid How ecu "moowumowwwpos omens cuas Ambmfiv .mm.mm.oxc£om mo cosmos one an poeuomuod was wowoooom oouuoodoOmH .wcfimooom ofiuuooaoomw you cowusfiom awo>uomom peso; ecu cw wwo< oquMHsm HON vwo< owuooHo>wuooamou “awesome ww~ woo wm~ ouoB mucoewumaxm omocu cw swoofiono: MOM moEHu wcwmsoom .>~o>wuoodmou «muovmooud as m- ucomoud ocu cw pom: umcu moswu m.# use Amwmfiv .mm.mm.oxc:mm mo cosmos ago a“ com: omozu V fiwm.~ mo cowumuucoocoo snowman m um no Anuuynv ouxdosoEm $:~.m ou n:.o «0 ucowoouw m on nonuwo ucowomuw zufimcoo omouo=m ecu ow ucomoua was ouzfiochm ecu udooxo «apogee: pom ufiowuouoz ecu cw ponwuomov ousvoooua uswfioz seafloo ucofiomuw EE mdfi mumocmum ecu ou wowvuooom coeuowuoa mos wcwoooom ofiuuooamomw .wmmmmmmm owuuoodoomw cw assumes“; ucowomuo mm com: cowuoowuumfin oomfiocma< HmfiuwcH mo uoowwm .Om ouowwm 113 Figure 30. E5222 20.55: on ON 0— mam—2&2 . - _ u no— N— 114 Decrease of Focusing Time by Decreasing Ampholyte Concentration. As shown in Figure 31, the time required for electrofocusing decreased as ampholyte concentration decreased. However, as ampholyte concentration was decreased, the pH gradient became nonlinear and the magnitude of the slape of the pH gradient increased. Presumably this is a reflection of the loss of ampholyte buffering capacity at the extremes of the sucrose gradient, where the gradient is in direct contact with acid and base. For the long and short columns, ampholyte concentrations of 0.86 and 0.50%, respectively, were chosen as the best compromise between focusing time on the one hand and linearity and slope of the pH gradient on the other. Decrease in Focusing:Time by Including_0.03% KCl in the Gel Plug Soak Solution, Density Gradient, and Reservoir Solutions. The presence of a low concentration of electrolyte in the density gradient and in the re- servoir solutions significantly shortened the electrofocusing time (Table 12). The optimal concentration of K01 is about 0.03%. When no Table 12. Effect of KClfi(0.03%) in Various Components of the Isoelec- tric Focusingggystem upon Hemoglobin Focusigg Time. Components to which KCl was added8 Focusing time (minutes) none 122 gel plug 122 gradient 103 gradient and gel plug 103 gradient, gel plug, and reservoirs 100 8The standard 112 mm column height procedure for isoelectric focusing was used with the exception of adding KCl only to the solutions indi- cated. electrolyte is added, focusing time is increased by about 20%. If the sample to be focused contains electrolytes, best results are obtained with nonfocusable electrolyte equivalent to about 5 mM (0.035%) KCl. Higher concentrations cause significant heating. Decrease in FocusingyTime by Acid Soaking the Gel Plug. In the previous electrofocusing system (Behnke £5 31., 1975), soaking the gel plugs in lower reservoir solution was required for good performance. If the gel plugs were not acid-soaked, focusing times were increased by 20 to 25%. In the present procedure, soaking of the gel plugs in lower reservoir solution containing 25% sucrose decreases focusing time about —. I— v 115 .: ma soaon doauo>o mmw.o can n:.o mo mcowumuucoocoo ouzfiozdaw cows mucoewuooxm now muaowcauw so ace .ouswfim ago as cacao one mocoapmuw omonu cg ofioofiwoao: pom moses wcfimooom .pooooapca on ones aucowpouw one a“ mcofiumuucoocoo ouzaochm udooxo aeponuoz one mfimfiuouoz one ca confluence ouoooooud mamas: csaaoo ucowpmuw 55 m- vuooooum one means omeuompod mos wcwmsoom ofiuuoofioomH .waHmSoom ewuuooaoomw cw xuwuoocHA ucomomuc :m‘ocm wage wowmooom com: cowumuucoocoo muzfiocm5< mo uoomwm .Hw ouswfim 116 Figure 31. mumszz 20.55.... E cm 3 3 m=Qm_zm_.2 . 352.5. ”.2: $2. mac”. 3: 20.53.55 209 552$ 52 d‘ 2 N. 117 five percent. The different soaking effects in the two procedures are due to the incorporation of conducting electrolyte in the gradient and to the uniform initial ampholyte distribution in the gradient. Decrease in Focusinngime by Decreasing the Gradient Column Height. With the improvements made in pH gradient linearity, it was possible to shorten the gradient column height. This is the most effective way of decreasing focusing time. Figure 32 shows the effect of decreasing the column height from 112 mm in the previous procedure (Behnke st 31., 1975) to 50 mm in the present procedure. The time required for focusing in the shorter column is about one-third that in the longer column. The pH gradient, although not as linear as in the longer column, is still satisfactory. DISCUSSION Two major advances are provided by the present method of isoelectric focusing in sucrose density gradients: (1) the time required for elec- trofocusing has been reduced to its limit, to as little as thirty-two minutes for hemoglobin, and (2) the linearity of the pH gradient has been greatly improved, yielding more accurate and reliable isoelectric point values. Figure 33 summarizes the differences between the initial and improved electrofocusing procedures. Figure 34 summarizes the changes made in the initial procedure during the development of the cur- rent procedure and the effects on focusing time and pH gradient linear- ity of those changes. Many factors affected both focusing time and pH gradient linearity, so changing one factor at a time was required in determining the effects of each factor on the electrofocusing system. The following factors always produced relatively large effects: (1) reducing the ampholyte concentration, which always decreased focusing time, (2) reducing the gradient column height, which always reduced focusing time, and (3) incorporation of sucrose in the gel plug, which always yielded more linear pH gradients. The magnitude of the effects of the following fac- tors often varied, depending on the other conditions of the experiment: (1) the presence and concentration of KCl, (2) the acid used in the lower reservoir solution, (3) the initial ampholyte distribution, and (4) the soaking of gel plugs prior to isoelectric focusing. 118 .x.o>wuooommu amouocHE 00 one mm mums masofioo SE ma. one 55 Om any a. canoawosoc How omega wcwmsoom .>.o>wooodmou .mceoaoo Auunnuv 88 NA. 0:» can A V as cm 9: now. .5 806 can mzo.o mo moE:.o> om: wouomfiaoo mooHuomuw ecu mom amchHoo soon a. *Om.o mp3 cofiuouucmocoo oquoszm ecu "udooxo apocomz pom mfimwpoumz a. ponwuomoo mm mouaomooua ucwwm; candoo ucmfipmuw as m.. new as Om compound ecu moan: noeuomuoa mus wcaosoom owuuooaoooH .wmamsoom ofiuuooaoomH a“ muwumoCWA ucowwmuu mm pom oEaB wdwmsoom co ufiwwom CESHOU pompouomn mo uoowmm .m on: am 119 Figure 32. 22.5.. $2222 22.55.“. . m8m.2m..2 & 855:. 8. z 300 as m: A3552 mm. 2.22.50 5:. cm 120 INITIAL PROCEDURE (FOCUSING TIME 115 MINUTES FOR HEMOGLOBIN): -3% SULFURIC ACID IN LOWER RESERVOIR, 5% ETHANOLAMINE IN UPPER RESER- VOIR. -POLYACRYLAMIDE GEL PLUG IN ELECTROFOCUSING TUBE (TO SUPPORT SUCROSE DENSITY GRADIENT), SOAKED OVERNIGHT IN LOWER RESERVOIR SOLUTION. -112 mm SUCROSE DENSITY GRADIENT COLUMN (3- -20%) CONTAINING 0. A3- 2.1h% AMPHOLYTE GRADIENT. -200 VOLTS. IMPROVED (PRESENT) PROCEDURE (FOCUSING TIME 32 MINUTES FOR HEMOGLOBIN): -3% ACETIC ACID IN LOWER RESERVOIR, 3% ETHANOLAMINE IN UPPER RESER- VOIR, BOTH RESERVOIRS ALSO CONTAINING o. 03% KCl. -POLYACRYLAMIDE GEL PLUG IN ELECTROFOCUSING TUBE (TO SUPPORT SUCROSE DENSITY GRADIENT), PLUG CONTAINING 25% SUCROSE, SOAKED AT LEAST SIX HOURS IN LOWER RESERVOIR SOLUTION CONTAINING 25% SUCROSE. -50 mm SUCROSE DENSITY GRADIENT COLUMN (5-12%) CONTAINING 0.50% AMPHOLYTE AND 0.03% KCl. -95 VOLTS. Figure 33. Components of Initial and Improved Isoelectric Focusing Pro- cedures. The initial procedure is that of Behnke et a1. (1975). The improved procedure is that presented in the Materials and Methods. The 50 mm gradient column height electrofocusing procedure represents the best combination of the factors studied for Obtaining minimal focus- ing time with good pH gradient linearity. The 112 mm gradient column height procedure represents the best combination of the factors studied for obtaining maximal pH gradient linearity and maximal resolution with satisfactory focusing time. The reduction in time to reach equilibrium during isoelectric focus- ing which was achieved by shortening the gradient column height is not as pronounced as might be expected based on sedimentation equilibrium theory, where the time to reach equilibrium is proportional to the square of the solution column height. The difference in time reduction in isoelectric focusing and sedimentation is due to the fact that in sedimentation equilibrium experiments, the centrifugal field may be main- tained as solution column height is decreased; in isoelectric focusing experiments, the electrical field must be reduced as the gradient column height is decreased. Maintaining the electric field strength as the gradient column height is reduced in isoelectric focusing results in heating due to the lower resistance and higher current flow across the shorter column. 121 EFFECT ON CHANGE FOCUSING TIME EFFECT ON pH GRADIENT ADDING SUCROSE TO THE GEL PLUG +10% LINEARITY IMPROVED ACETIG ACID REPLACING SULFURIC ACID +20% MINIMUM pH RAISED ADDING KCl T0 GRADIENT.AND RESERVOIRS 20% MINIMUM pH LOWERED INITIALLY DISTRIBUTING AMPHOLYTE UNI- _ 5% FORMLY RATHER THAN AS A GRADIENT REDUCING AMPHOLYTE CONCENTRATION -10% " REDUCING COLUMN HEIGHT -63% avLINEAR (pH 10.3 to 3.0) LINEAR (pH 10.5 to 2.5) Figure 34. Summary of the Steps in the Development of an Improved Pro- cedure for Isoelectric Focusinggin Sucrose Density Gradients. Presented are the changes made in the initial procedure for isoelectric focusing (Behnke 55H=1.; D975) and their effects upon focusing time and pH grad- ient linearity. CHAPTER 6 Purification and Partial Characterization of Pig Liver Ferritin ABSTRACT A yellow material copurified with fatty acid synthetase when using the procedure of Kim, Unkefer, and Deal (1977) to purify~ fatty acid synthetase. A portion of this material was removed at several steps in the purification, but some remained with the enzyme even after sucrose density gradient centrifugation. This material has been shown to be the iron storage protein, ferritin. Since ferritin copurifies with fatty acid synthetase, it was of interest to determine whether there was any interaction between them. Also, ferritin is the major protein extracted in water from the microsomal pellet fraction of pig liver described in Chapter 3 and Chapter 4. SO it was also of interest to see whether ferritin inactivates fatty acid synthetase. Consequently, a purification procedure for ferritin was developed. The purification procedure for pig liver ferritin is a modifica- tion of the method of Kim, Unkefer, and Deal (1977) for the purifica— tion of fatty acid synthetase. It includes ammonium sulfate fraction- ation of the centrifuged crude extract, DEAF-cellulose pad chromato- graphy, and sucrose density gradient centrifugation. An extremely pure preparation of the protein is Obtained: the banding pattern of the purified ferritin on native polyacrylamide gels is practically identical to that of commercially prepared horse spleen ferritin. The subunit molecular weight of pig liver ferritin is about 20,000 daltons based on sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The isoelectric point of the protein is 5.5 in 6 M urea and 5.8 in 2 M urea. The sedimentation coefficient of the protein on sucrose density gradients is independent of concentration. In sucrose density gradient sedimentation velocity experiments, the peak for protein as measured by the tannin turbidi- metric method and the peak for iron as measured by absorbance at 480 nm were not coincident; the iron peak was further down the gradient than the protein peak. The solubility Of ferritin 122 123 depends strongly upon its oxidation state: in TE buffer containing 0.33 saturated ammonium sulfate, ferritin will air-oxidize and precipitate over a period of weeks; in TE buffer containing 10 mM dithiothreitol, 0.60 saturated ammonium sulfate is required to precipitate the protein. Native polyacrylamide gel electrOphoresis showed that the native conformation Of fatty acid synthetase was not changed by incubation with ferritin. However, inactivation of fatty acid synthetase by ferritin was demonstrated. 124 INTRODUCTION A yellow-colored substance cOpurified with fatty acid synthetase during the latter's purification by the method Of Kim, Unkefer, and Deal (1977). This substance has been identified as the iron storage protein, ferritin. Ferritin is comprised Of an approximately Spheri- cal hollow protein shell containing up to 4500 atoms Of iron in the form of an oxyhydroxidephosphate. It is also the major protein ex- tracted from the microsomal pellet fraction from pig liver; this frac- tion was shown in Chapter 3 and Chapter A to inactivate fatty acid syn- thetase. SO it was of interest to determine whether there was any in- teraction between fatty acid synthetase and purified ferritin. This chapter describes a purification procedure for pig liver ferritin and the results of tests for interaction between it and fatty acid synthetase. Some prOperties of the purified ferritin are also given. MATERIALS AND METHODS Reagents. Dithiothreitol was a product of Eastman-Kodak. DEAF-cellulose (standard, 0.91 meq/g) was Obtained from Schleicher and Schuell. Coo- massie Brilliant Blue R, horse spleen ferritin, gum arabic, 2-mercapto- ethanol, sodium dodecyl sulfate, and tris(hydroxymethyl)aminomethane were purchased from Sigma Chemical CO. Ampholyte (Biolyte 3/10) was Obtained from BIORAD; it had a pH range of 3 to 10 and a concentration (w/v) of h0% solids. N,N,N',N'-tetramethylethylenediamine was a pro- duct of Canalco. Ammonium persulfate, N,N'-methylenebis(acrylamide), and acrylamide were Obtained from EM Laboratories. Phenol (88%, 11- quified) was a product Of Mallinckrodt. Tannic acid was purchased from Baker Chemical CO. All other reagents were Of analytical reagent grade. Absorbance Measurements. Iron content of ferritin was estimated by measuring absorbance at ABC nm using a Gilford Model 2000 absorbance recorder with water as the reference. Protein Determinations. Protein concentrations were determined by the tannin turbidimetric method described in Chapter 2. Native Polyacrylamide Gel ElectrOphoresis. Native polyacrylamide gel 125 electrOphoresis was performed using the modified method of Clark and Switzer (1977) as described in Chapter 2. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis. Sodium dodecyl sulfate- polyacrylamide gel electrophoresis was performed using a modification of the method of Clark and Switzer (1977) as described below. Distilled deionized water was used throughout. The stock reagents for the procedure were the following: 1. SDS gel buffer: 7.80 g NaH2P04-H20, 20.44 g NaeHPO4, 2.00 g sodium dodecyl sulfate, and water to two liters. 2. BIS-AC solution: 22.2 3 acrylamide, 0.50 g N,N'-methy1ene- bis(acrylamide), and water to 100 m1. This solution is filtered with Whatman NO. 1 filter paper and is stored at 4°. 3. Persulfate solution: 15 mg ammonium persulfate per ml water; this solution is prepared just before use. 4. SDS stock solution: 1.00 g sodium dodecyl sulfate in 10 ml water. 5. Dye solution: 0.2 mg bromphenol blue in 10 ml 50% (v/v) glycerol in water. 6. Staining solution: 120 mg Coomassie Brilliant Blue R is dissolved in 125 ml isOprOpanol; 50 ml glacial acetic acid and 325 ml water are then added. 7. Destaining solution 1: 500 m1 isOprOpanOl and 200ml glacial acetic acid to two liters with water. 8. Destaining solution 2: 300 ml 95% ethanol and 280 ml glacial acetic acid to four liters with water. 9. Protein standard solution: In these experiments, this solution contained each Of the following proteins at 1 mg/ml: Bovine serum albumin, yeast hexokinase, yeast enolase, and rabbit muscle glycerol-3-phosphate dehydrogenase- triose phosphate isomerase; the molecular weights of which are 68,000, 51,000, 44,000, 34,000, and 26,500, respectively. The electrOphoretic procedure is as follows: Gel tubes of dimensions 7 mm (0. d.) x 13 cm were sealed at one end with Parafilm. The tubes were then mounted vertically and the aqueous gel solution was prepared on ice. The gel solution contained 5.00 ml SDS gel buffer, 4.50 ml BIS-AC solution, 0.5 ml persulfate solution, and 12 ul TEMED for each six tubes. Immediately after preparing the gel solution, it was added to the gel tubes (1.8 ml per tube) using a Pas- teur pipet. A thin layer of water was then carefully added to the top 126 Of the gel solution in each tube. Polymerization was allowed to occur for at least one hour. While the gel solution was polymerizing, the protein samples were prepared by adding (for each sample in its separate tube) protein (10-50 pl Of a 1-2 mg/ml solution), SDS stock solution (5 pl), and 2-mercaptoethanol (5 ml). The samples were then heated in a boiling water bath for ten minutes. After cooling, dye solution (50 pl) was added to each sample. After polymerization Of the gel solution in the gel tubes, the water overlayers were removed. The tubes were then placed in the upper elec- trOphoresis chamber with the unfilled end of the tube upward. Both reservoirs were filled with SDS gel buffer and any air bubbles at either end of the gel tubes were removed. The boiled samples were then applied to the gels using an Eppendorf pipet; each tube received one sample, applied just above the gel to minimize mixing with the reservoir solu- tion. The positive lead from the power supply was attached to the lower reservoir and the negative lead to the upper reservoir. Electrophoresis was then performed at 90 volts until the dye front reached the bottom Of the tubes. The power supply was turned Off and the gel tubes were re- moved. The gels were removed from the tubes by gently rimming the lower end of the tube with a 22 gauge needle supplied with a stream Of water from a faucet. The gels were cut at the front edge Of the tracking dye and were then placed in 16 x 150 mm screw-top test tubes which contained staining solution. Staining was allowed to occur overnight. The staining solution was then decanted from the gels and was replaced with destaining solution 1. Destaining in this solution was allowed to occur overnight. The solution was then decanted and replaced with destaining solution 2. Destaining with this solution was continued until the gels were adequately destained. Isoelectric Focusing. Isoelectric focusing was performed using the standard 112 mm gradient height procedure described in Chapter 5 ex- cept the gel plugs contained 30% sucrose and the system contained no KCl. Also, for isoelectric focusing in 2 M urea, the sucrose density gradient contained 2‘M urea and 750 pg ferritin protein, the upper re- servoir contained 2 M urea in 1.8% ethanolamine, and the lower reser- 127 voir contained 2 M urea in 1.8% sulfuric acid. For isoelectric fo- cusing in 6 M urea, the sucrose density gradient contained 6 M urea and 350 pg ferritin protein, the upper reservoir contained 6 M urea in 1.2% ethanolamine, and the lower reservoir contained 6 M urea in 1.2% sulfuric acid. Duplicate gradients were prepared for each sam- ple; one gradient was fractionated after two hours at 200 volts and the other tube was fractionated after four hours at 200 volts. The position of ferritin in the gradients was determined both by measuring the absorbance of the fractions at 280 nm and by performing native polyacrylamide gel electrophoresis on the fractions. Sucrose Density Gradient Centrifugation and Sedimentation Velocity Analysis. An 1800 Model 570 gradient former was used to form 36 m1 linear 10 to 25% sucrose density gradients buffered with TED buffer in 1 x 3% inch polyallomer tubes. For purification of ferritin, cen- trifugation was at 20° for four hours at 25,000 rpm in a Beckman SW27 rotor. After centrifugation, fractions (1.5 ml) were collected using an ISCO Model 183 gradient fractionator with a flow rate of 3 m1/min. An 1800 Model UA-S absorbance monitor with a 10 mm pathlength flow cell was used to detect absorbance at 280 nm during fraction. For analysis of the relative protein-to-iron ratio in purified ferritin, the above procedure was used except 20 mg ferritin protein in 2‘ml PED buffer was applied to the gradient, centrifugation was performed at 27,000 rpm for three hours, and the gradient was fractionated into 1.00 ml fractions. For sucrose density gradient sedimentation velocity analysis Of ferritin, 5.3 ml linear 5 to 20% sucrose density gradients containing PED buffer were prepared in'fi x 2 inch cellulose nitrate tubes. Puri- fied ferritin (0.15 ml) at a concentration between 0.1 and 10 mg protein per milliliter was applied to the gradients. Centrifugation was per- formed at 47,000 rpm.for 60 minutes in a Beckman SW50.1 rotor at 25°. The gradients were then fractionated, monitoring the absorbance at 280 nm with an 1800 Model UA-5 absorbance monitor. Fractionation was performed at 0.75 ml per minute, and 0.25 ml fractions were collected. 128 RESULTS Pig liver ferritin was initially isolated as a byproduct in the purification of fatty acid synthetase by the method of Kim, Unkefer, and Deal (1977). The initial steps in the purification of ferritin described below are therefore modified from the procedure for the purification of fatty acid synthetase. Ammonium sulfate fractionation. The method of Kim, Uhkefer, and Deal (1977) for the purification of fatty acid synthetase was followed through the first dialysis step with these modifications: the first ammonium sulfate fractionation was changed from.0.2 to 0.33 saturation to 0.30 to 0.40 saturation, and the second ammonium sulfate fractiona- tion was taken to 0.40 saturation rather than 0.33 saturation. These changes reflect the fact that ferritin is slightly more soluble than fatty acid synthetase in ammonium sulfate solution at this stage in its purification. DEAE-cellulose chromatoggaphy. The dialyzate obtained following the ammonium sulfate fractionations was centrifuged at 20° for 30 minutes at 18,000 rpm in a Sorvall 88-34 rotor to remove a soft-packing precipi- tate. The supernatant was then applied to a 16 (d) x 4 cm (h) DEAE- cellulose pad which had been formed over a piece of Whatman NO. 1 fil- ter paper in a Buchner funnel. After the applied sample was absorbed, the pad was washed with one liter Of TED buffer and one liter 50 mM KCl in TED buffer; the eluates were discarded. Ferritin was then eluted as a bright orange solution using 1.5 liters of 300 mM KCl in TED buffer. The eluted ferritin was cooled to 6° and ammonium sulfate was added to 0.30 saturation. After stirring for 15 minutes, the suspension was centrifuged at 11,000 rpm for 25 minutes at 0° in a Sorvall GSA rotor. The supernatant was brought to 0.50 saturation ammonium sulfate with stirring to precipitate the ferritin. The suspension was centrifuged at 11,000 rpm for 25 minutes at 0° in a Sorvall GSA rotor. The pellets were collected and suspended in a minimal volume of TED buffer (less than 20 ml). The ferritin suspension was then dialyzed against one liter of TED buffer at 4° until the ferritin was solubilized. Sucrose density ggadient centrifugation. The dialyzate from the DEAE- cellulose chromatography step was centrifuged in a Sorvall 88-34 rotor at 16,000 rpm for 20 minutes at 20°. The supernatant was divided into 129 six equal portions and was applied to six SW27 gradients. Centrifugation was performed as indicated in Methods. Following fractionation of the gradients, the bottom two-thirds of the gradients were combined, taken to 40, and brought to 0.50 saturation with saturated ammonium sulfate solution. After stirring for 15 minutes, the suspension was centri- fuged in a Sorvall SS-34 rotor at 0° for 20 minutes at 16,000 rpm. The pellets were collected, suspended in a minimal volume of TED buffer, and dialyzed against TED buffer until the ferritin dissolved. Once solu- bilized, the ferritin is ready for a second SW27 centrifugation. Den- sity gradient centrifugation is repeated until native polyacrylamide gel electrophoresis no longer reveals contaminants. The contaminants are visible after the first SW27 centrifugation as a slower moving shoul- der in the gradient fractionation profile at 280 nm (Figure 35). Characteristigg_g§_£§g Protein: Identity. The purified protein was identified as ferritin by comparison of its electrOphoretic behavior with that Of commercial horse spleen ferritin. In our native poly- acrylamide gel system, both produced bands with relative mobilities of 0.04 and 0.12. The band with higher relative mobility was always pre- sent in greater quantity (Figure 36). Subunit molecular weight. The molecular weight of pig liver ferritin subunits was about 20,000 daltons, based on sodium dodecyl sulfate- polyacrylamide gel electrophoresis. When ferritin was incubated with 2-mercaptoethanol and sodium dodecyl sulfate, the solution changed in color from orange to rose to colorless. When the sample was applied to the gels and subjected to electrOphoresis, an orange material (pre- sumably an iron oxyhydroxide) remained at the tap of each gel; the ferritin subunits, which migrated into the gel, were colorless. Isoelectric 22335. Ferritin precipitated when it was subjected to isoelectric focusing in the absence of urea. Although some of the pro- tein precipitated in 2‘M urea, a preliminary value of 5.8 was obtained for the isoelectric point of ferritin. In 6 M urea, the protein re- mained soluble, and a value of 5.5 was determined for its isoelectric point. Nonuniform.;522 distribution. The preparation of pig liver ferritin was polydisperse as determined by sucrose density gradient sedimenta- tion velocity analysis; that is, the protein shells Of ferritin enclosed 130 .Adv codename edaoadsd Assam suw3 oceaomuw OLD you So cam on ofiwwoua :OHOOHOMQM ecu Boom Daemon a. nomasocm mace .Amv xmoa :«uwuHOM ego mo ovum wcwocoEHpOm uo3o~m may do hoodsonm o no mfinamfi> mum mucm:HEmuaoo use .moosuoz ow co>ww ouowoooud ecu moan: coauomuom ems cofiuomsmauucoo .caufiuuom powwwusm Wagon pom >22mfiuumd mo mamammc cowumucoEHpom uncapmuo xmwmcom Omouoom .Mm ouswwm Figure 37. In F! 135 wuosvv 1.0 l-fi F4 (1w/9w) NOIIVEINIONOO mama FRACTION NUMBER 136 .1 F— a - b 0 2 0.2mg/ml 1'2 4mg/ml 2 e 0.08 - 2 0-3 ' 1?: c: o o 3 0- |: G- a: «a O o E < "" m 0.04 '¢ ‘ 0.4 - \A 0 .51... e _ \LJ 0 ' l 1 1 10 20 1 10 20 FRACTION NUMBER FRACTION NUMBER Elgpre 38. Sucrose Density_Gradient Sedimentation Velocity Analysis of Ferritin at Two Concentrations. Centrifugation was performed using the procedure given in Methods. Ferritin in TED buffer (0.15 ml) at either 0.2 mg/ml (a) or at 4.0 mg/ml (b) was applied. AbsOrbance at 280 nm was measured during fractionation of the gradients. 137 synthetase was sufficient to yield an initial absorbance change of 0.05 per minute when 10 pl of incubation mixture was assayed for overall FAS activity. Two different ferritin concentrations were tested. Half- lives of overall FAS activity in incubation mixtures with 0.8 mg/ml and 4 mg/ml ferritin were 750 and 350 minutes, respectively, after taking into account the initial stabilization of overall FAS activity by dithio- threitol. DISCUSSION A procedure for the purification of ferritin has been presented. The purified protein is practically identical to the commercially prepared horse spleen ferritin in banding pattern on native polyacrylamide gels. The most interesting Observations about the protein are its oxidation- state dependent solubility in ammonium sulfate solutions and its inac- tivation of fatty acid synthetase. Ferritin in,a more highly oxidized state has been shown to be less soluble in ammonium sulfate solutions. One explanation for this pheno- menon is a change in conformation of the protein when its exposed sulf- hydryl groups are oxidized. As was demonstrated by sucrose density gradient sedimentation velocity analysis and native polyacrylamide gel electrophoresis, this conformational change does not detectably alter the sedimentation or electrOphoretic behavior of the protein. 80, apparently, oxidation causes movement of nonpolar amino acid residues to the outer surface of the shell or movement Of polar residues to the interior Of the shell, or both. Inactivation of fatty acid synthetase by ferritin can be explained as a change in the oxidation state of the enzyme. When fatty acid synthe- tase was incubated in high concentration of reducing agent (e. g., 45 mM dithiothreitol or 50 mM 2-mercaptoethanol), overall FAS activity initial- 1y increased, but then declined rapidly until it was lost completely (Figure 2). When fatty acid synthetase was incubated in TE buffer, air-oxidation of the enzyme occurred, resulting in inactivation of the enzyme. SO there is apparently a narrow range of redox environments in which fatty acid synthetase is stable. Ferritin may alter the environ- ment sufficiently to destabilize (inactivate) the enzyme. LIST OF REFERENCES LIST OF REFERENCES Adelman, T. G., Arosio, P., and Drysdale, J. W. (1975) Biochem. Biophys. Res. Conlnun. 63, 1056. Ahmad, P. M., Russel, T. R., and Ahmad, F. (1979) Biochem. ,1. 182, 509. Alberts, A. w., Majerus, P. W., Talamo, 13., and Vagelos, P. R. (1964) Biochemistry 3, 1563. Alberta, A. W., Majerus, P. W., and Vagelos, P. R. (1965) Biochemistry 4, 2265. Alberta, A. W., Strauss, A. W., Henessy, S., and Vagelos, P. R. (1975) Proc. Natl. Acad. Sci. USA 2, 3956. Allmann, D. W., and Gibson, D. M. (1965) ,1. Lipid ge_s_. _6, 51. Aprahamian, S., Arslanian, M. J., and StOOpa, J. K. (1980) Lipids 34, 1015. Arosio, P., Adelman, T. G., and Drysdale, J. W. (1978) ,1. Biol. Chem. 233, 4451. Arslanian, M. J., and Wakil, S. J. (1975) Meth. Enzml. 33, 59. Banyard, S. H., Steamers, D. K., and Harrison, P. M. (1978) Nature 271, 282. Behnke, J. N., Dagher, S. M., Massey, T. 11., and Deal, W. C. (1975) Anal. Biochem. 39, l. Bjork, I., and Fish, W. w. (1971) Biochemistry 12, 2844. Blaise, A., Chappert, J., and Giradet, J. (1965) g. _R_. Acad. Sci. 261, 2310. Bloch, K., and Vance, D. (1977) Annu. Rev. Biochem. 4_6_, 263. Bomford, A., Conlon-Hollingshead, 0., and Munro, H. N. (1981) 3. Biol. Chem. 236, 948. Bozarth, R. F., and Goenaga, A. (1972) Q93. ,1. Microbiol. 38, 619. Bloch, K. (1977) Adv. Enzymol. 43, 1. Brady, R. 0., and Gurin, s. (1952) ,1. Biol. Chem. 199, 421. Buckner, J. S., and Kolattukudy, P. E. (1976) Biochemistry 13, 1948. Buckner, J. S., Kolattukudy, P. E., and Rogers, L. (1978) Arch. Biochem. BiOphys. 186, 152. Burton, D. N., Collins, J. M., Kennan, A. L., and Porter, J. W. (1969) 3. Biol. Chem. 244, 4510. 138 139 Butterworth, P. H. W., Guchhait, R. B. Baum, H., Olson, E. B., MMrgolis, S. A., and Porter, J. W. (1966) Arch. Biochem. Bioghyg. 116, 453. Carey, E. M. (1977) Biochim. Bioghys. Acts 486, 91. Cavallini, D., Graziani, M. T., and Dupre, S. (1966) Nature 212, 294. Chang, H. G., Seidman, 1., Teebor, G., and Lane,‘M. D. (1967) Biochem. BiOphys. Res. Commun. 26, 682. Chukhrov, F. V., Zvyagin, B. B., Gorschkov, A. I., Yermilova, L. P., and Balashova, V. V. (1973) Int. Geol. Rev. 16, 1131. Clark, J. M., and Switzer, R. L. (1977) i3 Experimental Biochemistry, 2nd Ed., p. 47, W. H. Freeman and Co., San Fransisco. Clegg, G. A., Stansfield, R. F. D., Bourne, P. B., and Harrison, P. M. (1980a) Nature 288, 298. Clegg, G. A., Stansfield, R. F. D., Bourne P. B., and Harrison, P. M. (1980b) Biochem. Soc. Trans. 6, 65 . Craig, M. C., Nepokroeff, C. M., Lakshmanan, M. R., and Porter, J. W. (1972) Arch. Biochem. Bioghys. 152, 619. Crichton, R. R. (1975) Angew. Chem. Int. gs. lg, 57. Crichton, R. R., and Bryce, C. F. A. (1970) FEBS Lett. g, 121. Crichton, R. R., Collet-Cassart, D., Ponce-Ortiz, Y., Warters, M., Roman, F., and Paques, E. (1977).iE Proteins of Iron Metabolism (Eds. Brown, E. B., Aisen, P., Fielding, J., and Crichton, R. R.), Grune and Stratton, New York. Crichton, R. R., Eason, R., Barclay, A., and Bryce, C. F. A. (19733) Biochem. g. 131, 855. Crichton, R. R., Millar, J. A., Cummings, R. L. G., and Bryce, C. F. A. (1973b) Biochem.g. 151, 51. Crichton, R. R., Ponce-Ortiz, Y., Koch,‘M. H. J., Parfait, R., and Sturhman, H. B. (1978) Biochem. g. 111, 349. Crichton, R. R., Roman, F. Roland, F., Paques, B., Paques, A., and Vandamme, E. (1980) g, Mbl. Catal. I, 267. Dahlen, J. V., Kennan, A. L., and Porter, J. W. (1968) Arch. Biochem. Bioghzs. 124, 51. David, C. N., and Easterbrook, K. (1971) J, Cell Biol. 465 15. Dils, R., and Carey, E. M. (1975) Meth. Enzymol. 35, 74. Dorsey, J. A., and Porter, J. W. (1968) g, Biol. Chem. 243, 3512. Drysdale, J. W., Arosio, P., Adelman, T., Hazard, J. T., and Brooke, D. (1975) 13 Proteins of Iron Storage and Transport in Biochemistry and'Medicine (Ed. R. R. Crichton), North-Holland, Amsterdam. Dutler, H., Coon M. J., Kull, A., Vogel, H., Waldvogel, G., and Prelog, v. (1971) Eur. g, Biochem. 22, 205. Engeser, H., Wieland, F., and Lynen, F. (1977) FEBS Lett. gg; 139. 140 Fischer, P. W., and Goodridge, A. G. (1978) Arch. Biochem. Biophys. 190, 332- Fishbach, F. A., Harrison, P. M., and Hoy, T. G. (1969) 1. M01. Biol. 22.: 255- Flick, P. K., Chen, J., Alberts, A. W., and Vagelos, P. R. (1978) Proc. Natl. Acad. Sci. USA 75, 750. Flick, P. K., Chen, J., and Vagelos, P. R. (1977) g. Biol. Chem. 252, 4242. Formica, J. V., and Brady, R. 0. (1959) g, Amer. Chem. Soc. 61, 752. Frenkel, E. P., and Kitchens, R. L. (1977) Proc. Soc. Expgal. Biol. Med. 156, 151. Gibson, D. M., and Hubbard, D. D. (1960) Biochem. Biophys. Res. Commun. Eb 531. Giompres, P., and Packter, N. M. (1978) Biochim. Bigghy_. Acta‘5g9, 189. Goldman, P., Alberts, A. W., and Vagelos, P. R. (1961) Biochem. Bioghzs. Res. Commun. 5, 280. Granick, s. (1942) 1. Biol. Chem. 146, 451. Granick, s. (1945) 1, Biol. Chem. 149, 157. Granick, 8., and Hahn, P. F. (1944) 1. Biol. Chem. 155, 661. Guchhait, R. R., Putz, G. R., and Porter, J. W. (1966) Arch. Biochem. Bioghzs. 111, 541. Guynn, R%3:§, Veloso, D., and Veech, R. L. (1972) 1, Biol. Chem. 241, Habeeb, A. F. A. s. (1972) Meth. Enzzggl. 25, 457. Hahn, P., Seccombe, D., Kirby, L., Girard, J., Assan, R., and Kervran, A. (1978) Endocrinolggy{1g5, 1417. Hardie, D. G., and Cohen, P. (1978) Egg. 1. Biochem. 92, 25. Harrison, P. M. (1959) ,1. 1191. B131. _1_, 69. Harrison, P. M. (1963) g, M91. B131. 6, 404. Heald, S. M., Stern, E. A., Bunker, B., Holt, E. M., and Holt, S. L. (1979) ;. Amer. Chem. Soc. 101, 67. Heusterspreute, M., and Crichton, R. R. (1981) FEBS Lett. 129, 322. Hinsch, W., Klages, C., and Seubert, W. (1976) E25. g. Biochem. 64, 45. Hinsch, W., and Seubert, W. (1975) Egg. 1. Biochem. 55, 437. Hoare, R. J., Harrison, P. M., and Hoy, T. G. (1975) Nature 255, 653. Hoy, T. G., and Jacobs, A. (1981) Biochem. g, 195, 87. Hsu, R. Y., Butterworth, P. H. W., and Porter, J. W. (1969) Meth. Enzyggl. 1E, 33. Ishitani, K., Niitsu, Y., and Listowsky, I. (1975) 1. Biol. Chem. 250, 31h2. 141 Jacobs, A. (1977) 3211131313. m. 51, 91. Jenik, R. A., and Porter, J. W. (1981) M. Enzmol. 719, 97. Jones, M. 8., and Jones, 0. T. G. (1969) Biochem. J. _l_11, 507. Jones, T., Spencer, R., and Walsh, C. (1978) Qiochemistry 11, 4011. Joshi, v. C., and Aranda, L. P. (1979) 1. 13191. 91:59. 254, 11785. Kato, T., and Shimada, T. (1970) _._I_. Biochem. 68, 681. Kato, T., Shinjo, s., and Shimada, 1'. (1968) 1. Biochem. 65, 170. Kawaguchi, A., and Okuda, S. (1977) 11:93. _N_a_§_1_._. Acad. Sci. USA 73, 3180. Kennedy, E. P., and Lehninger, A. L. (1948) g. 15131. M. 172, 847. Kim, I.-C., Neudahl, G., and Deal, W. C. (1981) 112g. Enzml. 119, 79. Kim, I.-C., Unkefer, C. J., and Deal, W. C. (1977a) Arch. Biochem. Bioghzs. 118, 475. Kim, M., Qureshi, A. A., Jenik, R. A., Lornitzo, F. A., and Porter, J. w. (1977b) Arch. Biochem. Bigphzs. 181, 580. Knobling, A., Schiffmann, D., Sickinger, H.-D., and Schweizer, E. (1975) §_u_r_. .1. Biochem. 5_6_, 559. Kohgo, Y., Yogota, M., and Drysdale, J. W. (1980) :1. Biol. Chem. 255, 5195. KonOpka, K., Mareachale, J.-C., and Crichton, R. R. (1981) Biochim. Bigghfl. Acta 661, 417. KonOpka, K., and Romslo, I. (1980) Egg. ,1. Biochem. 101, 433. Kresze, G.-B., Oesterhelt, D., Lynen, P., Castorph, H., and Schweizer, E. (1976) Biochem. Bigghfi. 13.33. Comun. 69, 893 Castorph, H., and Schweizer, E. (1972) £1115. g. Biochem. 24, £92. Kuhn, R., Sorenson, N.A., and Birkofer, L. (1940) 9.2!.- 13, 823. Kukulansky, T., and Yagil, G. (1979) H_o_rl_n_. m. .1333. 11, 14. Kumar, s. (1977) 2191- Biochem. Bioghys. 1_85, 625. Kumar, 8., and Dodds, P. F. (1981) M. Enzymgl. I_]._C_, 86. Kumar, 8., Dorsey, J. A. Muesing, R. A., and Porter, J. W. (1970) g. Biol. Chem. 245, 4752. Kumar, 8., Phillips, G. T., and Porter, J. W. (1972) 1115. 1. Biochem. L 15- Kumar, 8., and Srinivasan, K. R. (1981) Biochemistry g9, 3393. Lakshmanan, M. R., Nepokroeff, C. M., and Porter, J. W. (1972) Proc. Natl. Acad. m. gs_A, 1.1.. 889. Langdon, R. G. (1955) g. Amer. Chem. §gg. fl, 5190. Langdon, R. G. (1957) 1. Biol. Chem. 22_6, 615. Lau, H.-P., Nepokroeff, C. M., and Porter, J. W. (1979) Biochem. Bigghyg. Res. Commun. 69, 264. Kuhn, L. 142 Laufberger, v. (1957) Bull. Soc. Chim. Biol. 19, 1575. Lema, M. J., and Sarcione, E. J. (1981) Comp. Biochem. Physiol. 5 69B, 287. Lennarz, W. J., Light, R. J., and Bloch, K. (1962) Proc. Natl. Acad. Sci. USA 116, 840. Liou, G. I., and Donaldson, W. E. (1973) Can. 1. Biochem. 51, 1029. Liou, G. 1., and Donaldson, W. E. (1977) Arch. Biochem. Bioghyg. 178, 195- Lin, c. Y., and Kumar, S. (1972) 1, Biol. Chem. 247, 604. Linder, M. C., Moor, J. R., and Munro, H. N. (1974) 1. Biol. Chem. 249, 7707. Linn, T.1§éé Stark, M. J., and Srere, P. A. (1980) 1. Biol. Chem. 255, Lornitzo, F. A., Qureshi, A. A., and Porter, J. W. (1974) 1. Biol. Chan. 249, 1654. Lornitzo, F. A'., Qureshi, A. A., and Porter, J. W. (1975) 1. Biol. Chem. 250, 4520. Lust, G., and Lynen, F. (1968) 3111;. _.1. Biochem. I, 68. Lynen, F. (1959) 1, Cell. Comm. thsiol. 51 Suppl. 1, 55. Lynen, F. (1961) F351. Proc. _2_0, 941. Lynen, F. (1962) Proc. Robert 11. Welsh Found. Conf. Chem. Res. 112. 5, Houston, TX, p. 293. Lynen, F. (1967) Biochem. 1. $2, 381. Lynen, F. (1969) Meth. Enzymol. &, 17. Lynen, F. (1980) Eu_r. ,1. Biochem. 112, 431. Lynen, F., Hopper-Kessel, 1., and Eggerer, H. (1964) Biochem. g. 515), 95. Macara, I. G., Hoy, T. G., and Harrison, P. M. (1972) Biochem. 1. 126, 151. Maitra, S. K., and Kumar, S. (1974) 1, Biol. Chem. 249, 117. Majerus, P. w. (1967) 1. Biol. Chem. 242, 2525. Majerus, P. W., Alberts, A. W., and Vagelos, P. R. (1965) :1. Biol. Chem. 240, 618. Majerus, P. W., Alberts, A. W., and Vagelos, P. R. (1969) Meth. Enzymol. 1_4, 64. Marsh, J. B., and James, A. T. (1962) Biochim. BioPhys. Acta _6_9_, 320. Martin, D. B., Horning, M. G., and Vagelos, P. R. (1961) 1. Biol. Chem. 256, 663. Martin, R. G., and Ames, B. N. (1961) ,1. Biol. Chem. a6, 1372. Massey, T. H., and Deal, W. C. (1973) ,1. Biol. Chem. 248, 56. Massover, W. H. (1978) Biochim. BiOphys. Acta 552, 202. 143 Mattick, J. 3., Zehner, z. E., Calabro, M. A., and Wakil, S. J. (1981) Eur. .1. Biochem. 114, 643. Mazur, A., and Carleton, A. (1963) ,1. Biol. Chem. 58, 1817. McKay, R., Druyan, R., Getz, G. 8., and Rabinowitz, M. (1969) Biochem. J. 114, 455. . Michaelis, L. (1947) Adv. Prot. Chem. 5, 55. Michaelis, L., Coryell, C. D., and Granick, S. (1943) _1. Biol. Chem. 8 6 1__: 3- Muesing, R. A., and Porter, J. w. (1975) Meth. Enzymol. 55, 45. Municio, A. M., Lizarbe, M. A., Relano, E., and Ramos, J. A. (1977) Biochim. Biophys. Acts 487, 175. Munro, H. N., and Linder, M. c. (1978) Physiol. Rev. 55, 517. Muraoka, N., Nakajima, s., Kato, T., and Shimada, T. (1966) _J_. Biochem. Q. 1#89- Nepokroeff, C. M., Lakshmsnan, M. R., and Porter, J. W. (1975) Meth. Enzml. 55, 37. Nepokroeff, C. M., Lau, H.-P., and Porter, J. W. (1979) Int. 1. Biochem. 19, 791. Nepokroeff, C. M., and Porter, J. w. (1978) _J_. _111_o1. 9112. 255, 2279. Niederer, W. (1970) Experientia 36, 218. Nugteren, D. H. (1965) Biochim. Biophys. M £6, 280. Nunez, M. T., Cole, E. s., and Class, J. (1980) M55, 1051. O'Brien, J. S. (1965) Science 141, 1099. Otsuka, 8., Maruyama, H., and Listowsky, I. (1981) _Biochemistry 90, 5226. Paskin, N., and Mayer, R. J. (1976) Biochem. ,1. 159, 181. Popjak, C., and Tietz, A. (1955) Biochem. :1. 6_0, 147. Porter, J. W., Kumar, 8., and Dugan, R. E. (1971) Progr. Biochem. Pharmacol. 6, 1. Qureshi, A. A., Jenik, R. A., Kim, M., Lornitzo, F. A., and Porter, J. w. (1975) Biochem. Biophys. Res. Commun. 66, 544. Qureshi, A. A., Solomon, J. K., and Eichelman, B. (1978) Biochem. Bioghxs. Res. Commun. 65, 1191. Robinson); J. D., Brady, R. 0., and Bradley, R. M. (1965) J_. Lipid 1193. 144 J O Romsos, D. R., and Leveille, G. A. (1974) 5511. 13:13 .1335. 12, 97. Roncari, D. A. K. (1975) 9433. _J_. Biochem. 55, 135. Rous, S. (1974) mgr, £111, 55. Russell, S. M., and Harrison, P. M. (1978) Biochem. 1. 175, 91. 144 Rutkoski, A. and Jaworkski, J. G. (1978) Biochem. BiophL. Res. Commun. 1311. 428. Schaefer, F. V., and Theil, E. c. (1981) 1. Biol. Chem. 256, 1711. Schmiedeberg, o. (1894) Arch. Exptl. Path. Pharmakol. 55, 101. Schweizer, E., and Bolling, H. (1970) Proc. Natl. Acad. Sci. USA 67, 660. Schweizer, E., Dietlein, G., Gimmler, G., Knobling, A., Tahedl, H. W., Schwietz, H., and Schweizer, M. (1975) Proc. FEBS Meet. 99, 85. Schweizer, E., Kniep, B., Castorph, H., and Holzner, U. (1975) Eur. 1. Biochan. 59, 353. Seckbach, J. (1972) 1. Ultrastruct. Res. 59, 65. Segel, I. H. (1975) 1.11 Enzyme Kinetics, p. 926, Wiley-Interscience, New York. Seubert, w., Gruell, G., and Lynen, F. (1957) 16363. 2119:.- 69, 559. Seubert, W., and Podack, E. R. (1973) Mol. Cell. Biochem. _l, 29. Simon, E. J., and Shemin, D. (1953) 1. Amer. Chem. Soc. 15, 2520. Sirivech, 8., Driskell, J., and Frieden, E. (1977) 1. 111151. 191, 739 Sirivech, 8., Frieden, E., and Osaki, 8. (1974) Biochem. 1. 145, 311. Smith, 8., and Abraham, 8. (1970) Arch. Biochem. Bioths. 116, 112. Smith, 8., and Abraham, 8. (1971s) 1. Biol. Chem. 246, 2537. Smith, 8., and Abraham, S. (1971b) 1. Biol. Chem. 246, 6428. Smith, 3., and Abraham, S. (1975a) 14;. £121.12 59;. 15, 195. Smith, 8., and Abraham, 8. (1975b) M_e_t_:_l_1. Enzthol. 55, 65. Sobhy, C. (1977) E1. gr_og. 56, 2659. Sobhy, c. (1979) 1. §i_o_1_. £1192- 25$ 8561. Speake, B. K., Dils, R., and Mayer, R. J. (1976) Biochem. 1. _1511, 559. Stoops, J. K., Arslanian, M. J., Aune, K. C., and Wakil, s. J. (1978) Arch. Biochem. BioghL. 188, 348. Stoops, J. K., and Wakil, 8. J. (1978) Biochem. Biophys. Res. Conlnun. 651, 225. 8t00ps, J. K., and Wakil, S. J. (1981a) 1. Biol. Chem. 256, 8364. Stoops, J. K., and Wakil, S. J. (1981b) _J_. Biol. Chem. 256, 5128. Strawser, L. D., and Larrabee, A. R. (1976) 1. Biol. Chem. 2L1, 270. Strom, K. A., Galeos, W. L., Davidson, L. A., and Kumar, 8. (1979) 1. Biol. Chem. 254, 8155. Strom, K. A., and Kumar, 8. (1979) 1. Biol. Chem. 254, 8159. Taketa, K., and Pogell, B. M. (1966) 1. Biol. Chem. 241, 720. Taylor, J. M., and Schimke, R. T. (1974) ,1. Biol. Chem. 249, 5597. 145 Thiel, E. C. (1973) 1. Biol. Chem. 248, 622. Towe, K. M. (1981) 1. Biol. Chem. 256, 9377. Trams, E. G., and Brady, R. O. (1960) 1, Amer. Chem. Soc. 69, 2972. Treffry, A., and Harrison, P. M. (1978) Biochem. 1. 1L1 515. Treffry, A., and Harrison, P. M. (1980a) Biochem. Soc. Trans. 6, 655. Treffry, A., and Harrison, P. M. (1980b) Biochem. Soc. Trans. 6, 656. Treffry, A. ., and Harrison, P. M. (1980c) Biochim. Biophys. Acta 610, 421 Tubbs, P. K., and Garland, P. B. (1965) Biochem. 1. 69, 25?. Tweto, J., and Larrabee, A. R.. (1972) 1. Biol. Chem. 247, 4900. Tweto, Liberati, ‘M., and Larrabee, A. R. (1971) 1. Biol. Chem. 246, 2468. Ulvik, R. J. (1981) _F_B_B_S_Le_t_t_. _159, 281. Ulvik, R. J., and Romslo, I. (1978) Biochim. Biophys. 1665 5_4_l, 251. Ulvik, R. J., and Romslo, I. (1979) Biochim. Biophys. Acts 568, 256. Ulvik, R. J., and Romslo, I. (1981) Biochim. Bioghzs. ALta 655, 457. Ulvik, R. J., Romslo, 1., Roland, F., and Crichton, R. R. (1981) Biochim. BiOEhXS. Acta6 6], SO. Urushizski, 1., Niitsu, Y., Ishitani, K., Mstsuds, M, and Fukuda, M. (1971) Biochim. Biophzs. ALta 24 5, 187. Vagelos, P. R. (1974) Biochem. Li id 4 99. Vagelos, P. R., Alberts, A. W., and Majerus, P. W. (1969) Meth. Enzymol. l_,4 59. Valimiri, L., Linder, M. C., Munro, H. N., and Catsimpoolss, N. (1977) Biochim.B Biophzs. ALta 491, 67. Vandsmme, E. (1980) Doctoral thesis, Uhiversite Cstholique de Louvain. Volpe, J. J., and Kishomoto, Y. (1972) 1. Neurochem. 19, 757. Volpe, J. J., and Vagelos, P. R. (1973) Annu. Rev. Biochem. fig, 21. Wakil, s. J. (1958) _J_. m. _C_hLem. 65. 69, 6465. Wakil, s. J. (1961) 1. E32529; g, l. Wagstsff, M., Worwood, M., and Jacobs, A. (1978) Biochem. 1, 115, 969. Wstanabe, N., and Drysdale, J. (1981a) Biochem. Bioghzs. Res. Commun. 10 05, 207. Watansbe, 8N., and Drysdale, J. (1981b) Biochem. Bioghzs. Res. Commun. L 507- Weeks, 0., and Wakil, S. J. (1968)'1. Biol. Chem. 245, 1180. Wetz, K., and Crichton, R. R. (1976) Eur. 1. Biochem. 61, 545. Wherest, A. F. (1971) 11!. LiEid Res. 9, 119. 146 Wielsnd, F., Renner, L., Verfurth, C., and Lynen, F. (1979) Eur. 1. Biochem. 94, 189. Williamson, I. P., and Wakil, S. J. (1966) 1. Biol. Chem. 241, 2526. Yang, P. C., Butterworth, P. H. W., Bock, R. M., and Porter, J. W, (1967) 1, Biol. Chem. 242, 5501. Yariv, J., Kalb, A. J., Sperling, R., Bauminger, E. R., Cohen, S. G., and Ofer, S. (1981) Biochem. 1. 191, 171. Yu, H. L., and Burton, D. N. (1974a) Arch. Biochem. Biophys. 161, 297. Yu, H. L., and Burton, D. N. (1974b) Biochem. Biophy1. Res. Commun. 61, 485. Yun, s. L., and Hsu, R. Y. (1972) 1. Biol. Chem. 247, 2689. Zahringer, R. J., Baliga, B. S., and Munro, H. N. (1976) Proc. Natl. Acad. Sci. USA 15, 857. Zsman, 2., and Verwilghen, R. L. (1977) Biochem. Soc. Trans. 9, 506. Zehner, 2. E., Joshi, V. C., and Wakil, S. J. (1977) 1. Biol. Chem. 292, 7015. M71111111111111fififlijfifilflifliflfliflfiflfil'“