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Aconitase from the oyster Crassostrea virginica
Comp. Biochem. Phy.~iol.Vol. 72B, No. 2, pp. 321 to 324. 1982
0305-0491/82/020321-04503.00/0 Pergamon Press Ltd
Printed in Great Britain.
ACONITASE FROM THE OYSTER CRASSOSTREA VIRGINICA MALAK I. SHOUKRY* Department of Chemistry, University of Maryland, College Park, MD 20742, U.S.A.
(Received 14 September 1981) 1. The presence of aconitase activity in the oyster, Crassostrea riryinica, has been demonstrated. 2. Low levels of activity were found in the different tissues with highest level in digestive diverticular and lowest level in muscle. 3. The conversion of both citrate and iso-citrate to cis-aconitate suggests the presence of an enzyme system capable of utilizing these compounds at a slow but demonstrable rate to give classically expected results. 4. Comparison of the oyster enzyme with aconitase from mammalian tissue indicated great similarity between the two enzyme systems. Abstract
INTRODUCTION Isolated reports by investigators using molluscs have suggested that in this phylum, some organisms may have substantially modified pathways for carbohydrate metabolism. Jodrey & Wilbur (1955) have reported that the activities of the citric acid cycle enzymes in the mantle tissues of the American oyster, Crassostrea cirginica, were 1 3.8% of rat liver activities, aconitase was not detected. They also observed an increase in respiratory rate when iso-citrate was added to the mantle tissue but no increase when citrate was added. They correlated this with the absence of aconitase and suggested that citric acid was missing from the cycle. Black (1962) found aconitase in eggs a n d larvae of the same species of oyster. N o further investigation was made, and it has been tacitly assumed following the concept of biochemical unity of life, that aconitase is p r o b a b l y present in the oyster. The present study was undertaken to verify this suggestion in order to establish clearly the presence or absence of aconitase in the oyster and to study its properties.
MATERIALS AND METHODS All reagents were obtained commercially from Sigma Chemical Company (St. Louis, Missouri). Oysters were purchased at the Lexington Market, Baltimore, Maryland.
Preparation of the enzyme 100 g of freshly shelled oysters were homogenized with 300 ml of 4 m M Na-citrate buffer containing 10 mM fl-mercaptoethanol, 0.05 mM EDTA, adjusted to pH 5.0 with NaOH and 40 ml chloroform (Villafranca and Mildvan, 1971). Homogenization was carried out in Sorval "omni mixer" for 3 min at maximum speed. The crude extract was centrifuged at 20,000 ff for 20 min and the precipitate was discarded. The supernatant was brought to * Present address: Department of Biochemistry, Faculty of Medicine, Kuwait University, P.O. Box 24923, Kuwait. 321
50% saturation with solid ammonium sulfate (0.291 g/ml). The mixture was allowed to stand for 30 min then centrifuged at 18,000 g for 15 rain. The precipitate was discarded and ammonium sulfate (0.091 g/ml) was added to the supernatant to bring it to 65% saturation. The mixture w a s centrifuged as described above and the resulting precipitate was dissolved in a minimal volume of 20 mM Tris, 1 mM citrate 0.1 mM EDTA, and 0.1 mM fl-mercaptoethanol, at pH 7.4. 20 ml of this solution was applied to the top of a column (2.5 x 50 cm) of AcA 34 polyacrylamide agarose gel (LKB-Ultrogel), which had been equilibrated with 15 mM tricarballylate-Tris buffer containing 0.5 mM EDTA, l0 mM fl-mercaptoethanol, adjusted to pH 7.4, tricarballylate is not inhibiting the enzyme at the concentration used. The enzyme was eluted with the same buffer. This resulted in seven fold increase in specific activity, Table I.
Enzyme assay Aconitase was routinely assayed by the method of Racker (1950) by following the disappearance of cis-aconitate at 240nm. At 25°C, the assay solution contained 20 mM Tris HCI, 2 mM cis-aconitate, 100 mM NaCI and enzyme in a final volume of 0.5 ml at pH 7.4. Absorbance changes were measured in a Gilford 2400 recording spectrophotometer. A quartz insert was used to reduce the light path in the cuvette. When citrate or iso-citrate were used as substrates, the assay solutions contained 20 mM citrate or 5 mM O(+)iso-citrate, 25mM Tris HC1, and enzyme in a final volume of 1 ml at pH 7.4. One unit of enzyme activity is defined as the production of l llmol of product per min. A millimolar absorption coefficient of 4.88 m M - 1 cm- ~ for cis-aconitate at 240 nm was used. RESULTS
Aconitase activity Preliminary experiments indicated no aconitase activity in acetone powder preparations of whole oyster or from mantle and gill tissues. However, low levels of activity were found in fresh oyster tissues; the order of activity in the different anatomical parts was: digestive diverticular > mantle a n d gill > muscle (Table 2). The enzyme solution prepared was extremely un-
322
MALAK I. SHOUKRY Table 1. Preparation of aconitase from whole oyster tissues Purification Step Crude extract 50-65~o Ammonium sulfate Polyacrylamide agarose chromatography
U/ml
Protein mg/ml
Specific activity U/mg protein
109
5.03
21.7
1
404
5.96
67.8
3.12
41
140
0.88
7.33
17"
Purification
159
Y °'o 100
* Approximately one half of the 50-65~ ammonium sulfate fraction was processed on the polyacrylamide agarose column. The yield (Y), however, refers to the total volume. Table 2. Aconitase content of oyster anatomical parts Tissue Muscle Mantle and Gill Digestive diverticula
Units/ ml
mg protein/ ml
Specific activity units/mg protein
36.9 12.3 41.0
19.7 3.98 8.3
1.87 3.08 4.94
The reaction mixture contained Tris HC1 buffer, pH 7.4. 100mM NaC1. 0.1 mM cis-aconitate and enzyme in a final volume of 3 ml.
stable, losing 40-50~o of its activity in one day. However the a m m o n i u m sulfate fraction was less unstable, and after 10 days at 0°C, this fraction was found to have 60% of its activity remaining.
Michaelis constants Apparent Km's were determined for the three substrates at 25°C in Tris-HC1 buffer, at pH 7.4. Substrate concentrations were over the range of: 2 - 2 0 m M for citrate, 0 . 3 - 2 m M for D ( + ) iso-citrate and 0.2-2 m M for cis-aconitate. The following values were obtained: K,, citrate = 4.19mM, K,~ isocitrate = 0.57 m M and Km cis-aconitate = 0.21 mM. Although K,, values reported in the literature show a wide range even for aconitase from the same source (Table 3) the order is the same as found here, namely, K,~ citrate > K m iso°citrate > K m cis-aconitate. This wide variations is probably due to the different assay methods,
Optimum pH and temperature The pH activity relationship of aconitase over the range pH4.1-10.6 was also studied. The reactions Table 3. Comparison of Michaelis constants of aconitase* Source of enzyme Pig heart Pig heart Rabbit liver Beef heart Beef liver Mustard Oyster'["
K~, citrate (mM)
K,, cis-aconitate (mM)
Km isocitrate (raM)
3.6 0.62 0.9 0.95 0.20 4.0-4.4 4.19 (n = 4 )
0.12 0.015 0.09 0.099 0.007 0.1 0.21 (n = 3)
0.48 0.2 0.32 0.139 0.034 0.15 0.57 (n = 4 )
* From The Enzymes, Vol. 5, p. 426. f This study.
[citrate ---*cis-aconitate], [isocitrate---, cis-aconitate] and cis-aconitate hydration all exhibited the same optima, namely pH 7.0 in phosphate buffer and pH 7.5 in 3, 3 dimethylglutarate buffer. The above three reactions, examined in the 8-60°C range, exhibited the same optimum temperature of 42°C. Aconitase is heat labile, being completely denatured in 5 min at 60°C. The enzyme exhibited half optimal activity at 8°C.
lnhibitors of aconitase (a) Tricarboxylic acids. Inhibition patterns for trans-aconitate with the three substrates of aconitase are summarized in Table 4. It can be seen that the nature of inhibition depends on the substrate used. Thus trans-aconitate is a competitive inhibitor when cis-aconitate is the substrate but has a non-competitive component when citrate or iso-citrate are the substrates. Tricarballylate which is a substrate analogue of citrate and iso-citrate, exhibited similar behaviour, in that it is a competitive inhibitor of aconitase when citrate is the substrate but is a non-competitive inhibitor when cis-aconitate is used as substrate. An essential feature of a Kinetic scheme which shows both competitive and non-competitive behaviour is that Kt value derived from data, where I is a competitive inhibitor, must equal the K~ (slope) value where I is a non-competitive inhibitor (Villafranta, 1974). This was confirmed by the reasonably close values for Kt and Kis obtained in the present study (see Table 4). (b) SulJhydryl reagents. Dickman & Cloutier (1951), demonstrated that aconitase from pig heart could be rapidly and completely inhibited by I mM p-chloromercuribenzoate. In this study it was found that 1.5 m M p-chloromercuribenzoate completely inhibited the oyster enzyme suggesting that sulfhydryl groups might be necessary for enzymatic activity. The inhibition is non-competitive. N-Ethylmaleimide and
Aconitase from the oyster Crassostrea viryinica
323
Table 4. Inhibition constants for aconitase and substrate analogues Inhibitor
Substrate
Pattern
Kt*
K~st KHt
Trans-aconitate Trans-aconitate Trans-aconitate Tricarballylate Tricarballylate
cis-aconitate Citrate Isocitrate Citrate cis-aconitate
Competitive Noncompetitive Noncompetitive Competitive Noncompetitive
3.6 --
-3.9 2.11 -11
9
-0.6 0.9 0.14
* Calculated from the equation: 1__
K,,
v
v[s]
17+ E1 + [ 1KtJ l v"
"~Kts and Ktt are inhibition constants calculated from the slopes and intercepts respectively using the equation: ; = v[s]
l + K~s_l + ] + K ~ J v "
iodoacetamide were also tested (see Table 5), but were found to be far less effective. (c) Metal chelators. Compounds that bind iron are known to inhibit aconitase from other sources. In this study two such iron chelators, O-phenanthroline and ~-picolinic acid were tested at 2 and 10 m M concentrations respectively. As shown in Table 5, they had little effect on enzyme activity. A pre-incubation of the inhibitor with the enzyme is necessary for the inhibition to be demonstrable.
Activation by ferrous iron and cysteine It is well known that the rate of inactivation of an aging aconitase preparation can be decreased, by the addition of Fe 2 + and cysteine. The addition of these same compounds can also reactivate dialyzed samples. Aconitase prepared from oyster tissue was also found to lose activity upon storage or dialysis, so activation with iron and cysteine was tested. The effects of these compounds are shown in Table 6.
Table 5. Effect of metal chelators and sulfhydryl reagents on aconitase hydration (n = 3)
Reagent
Final conc. (mM)
Incubation period (min)
2 2 10 10
0 5 0 5
Metal chelators: O-phenanthroline ~-picolinate Sulfhydryl reagents: P-Chloromercuir benzoate N-Ethylmaleimide Iodoacetamide
0.5 1.5
5 0
1
5
2 2 2 3 3
5 5 15 5 15
~ Inhibition I1 33 4.8 17 17 100 32 46 11 21 11 22
Table 6. Restoration of activity of aconitase by Fe(lI) and cysteine Sample
9/0 Remaining activity*
(A) Enzyme solution dialyzed
85 against buffer1" n = 2 (B) Enzyme solution dialyzed against 65 buffer containing 1.0 mM O-phenanthroline n = 2 Addition of 2.0 mM Fe z+ + 10 mM cysteine hydrochloride Sample A 147 Sample B 135 * Relative to undialyzed sample. t 20 mM Tris buffer, containing 1.0 mM citrate, 0.1 mM EDTA, 0.1 mM fl-mercaptoethanol, pH 7.4.
324
MAI.AK 1. SHOUKRY tO0
o o o
o
o
Zg+, o
6C
Ni + ÷
•~ U"~4-
~' 40
•
2O
0 -4
-3
-2
log m o l a r i t y of metal ion
Fig. 1. Inhibition of [isocitrate---+ cis-aconitate] reaction by the metal ions, Cu 2+, Zn 2+ and Ni 2+. In addition to the metal ion, the reaction mixture contained 5.0ram D(+) isocitrate, 20raM Tris HCI buffer pH7.4 and enzyme in 1.0 ml. It must be noted here that b o t h iron and cysteine are required for the activation, since neither of them alone had any effect. Effects o f M e t a l Ions
Z n 2+, Cu 2+, a n d Ni 2. were tested for their effects on the enzyme activity. Cu 2+ at 1 m M completely inhibited the enzyme. However Z n 2+ up to 10mM caused no significant decrease in activity. With Ni 2+ there was a decrease in activity but even at 1 0 m M Ni 2+, some 50 60{~, of the activity remained. {See Fig. 1.) DISCUSSION The data presented d e m o n s t r a t e unequivocally that aconitase activity is present in oyster tissues, although at an extremely low level. In view of this low activity, it is u n d e r s t a n d a b l e why other investigators (Jodrey & Wilbur, 19551, could not detect any activity in this species. Additional reasons might be (1) the use of dilute homogenates (1:20W/V), (2) the use of phosphate buffer which is k n o w n to inhibit the enzyme (Morrison, 1954), and (3) the choice of mantle tissue as the sole source of enzyme. Localization of the enzyme to a particular subcellular fraction was difficult because of the low yield of enzyme as well as the lack of established methods for the fractionation and characterisation of subcellular organelles in the oyster which is k n o w n to be rich in glycogen and fibrous tissue. Aconitase from the oyster seems not to differ kinetically from aconitase from other sources. The enzyme loses activity upon storage or dialysis, partly due to the removal of iron, as indicated by the restoration of activity to dialyzed enzyme with iron (It) and cysteine, indicating the oyster aconitase is an Fe 2 + sulphur enzyme similar to the nonh a e m iron-sulphur proteins of the electron transport chain. This is supported by the observation that other
divalent metals, such as Z n 2 ~ and C u 2 ' , did not activate the dialysed enzyme in connection with aconitases from other sources. Inhibition studies with st, bstrate analogues suggest that two forms of aconitase exists, one preferentially binds cis-aconitate and its structu,al analogues, while the other preferentially binds citrate and its structural analogue. More detailed study with other tricarboxylic acids are needed to confirm this conclusion. These results however are consistent with the finding of Villafranca with pig heart aconitase. The interconvcrsion of the three acids: citric, cisaconitic and iso-citric by the isolated oyster aconitase, indicates that this portion of the citric acid cycle is operative in the oyster. The relative rate of the reaction [iso-cilrate--,cis-aconitate] was found to be twice that of [citrate---. cis-aconitate] in support of the k n o w n fact that the mitochondrial carbon flux is in the direction of citrate to iso-citrate. In the cytoplasm the flux is in the direction of iso-citrate to citrate, making citrate available for fatty acid biosynthesis and regulation of glycolysis. This may lead one to speculate that it is possible that the role of aconitase in the oyster may be different from the role of aconitase in the mitachondrial citric acid cycle in m a m m a l i a n systems. The oyster, Crassostrea rir,qinica has a high concentration of citrate, but the origin of this acid in this system has not been established. It is possible that citrate synthetase is present in the oyster. A low level of citric acid cycle enzymes are also present. These finding would suggest that the oyster m i t o c h o n d r i o n may have different control systems from the m a m m a l i a n mitochondrion. Perhaps the organelle has a different permeability 1o citrate. However such a theory could only be tested by further study. REFERENCES
BLACK R. E. (19621 Respiration, electron transport enzymes, and Krebs-cycle enzymes in early developmental stages of the oystcr Crassostrea virqinica. Biol. Bull. 123, 58 70. BLACK R. E. (1962) The concentrations of some enzymes of the citric acid cycle and electron transport system in the large granule fraction of eggs and trochophores of the oyster, Crassostrea vir~linica. Biol. Bull. 123, 71 79. DlC:KMa,N S. R. & CLOI T,iR A. A. (1951} Factors affecting the activit~ of aconitasc .l. hiol. Chem. 188, 379 388. HAMMIY C. S. (1969) Metabolism of the oyster. Crassostrea rir~tinica. Zoologist 9, 309 318. H()('HA('HKA P. W. & MtlSIAI:A T. (1972) Invertebrate facultative anaerobiosis. Science, N.E 178, 1056 1060. JODRr!'{ L. H. & WI~BtJR K. M. [1955l Studies on shell formation. IV, The respiratory metabolism of the oyster mantle. Biol. Bull. 108, 346 358. MORRISON J. F. (1954) The purification of aconitase. BiochenL d. 56, 99 105. RA('KER E. (1950) Spectrophotometric measurements of the enzymatic formation of fumaric and cis-aconitic acids. Biochem. Biophys. Acta 4, 211 214. Vn.LAFRANCA J. J. (19741 The mechanism of aconitase action, lI. Evidence for an enzyme isomerization by studies of inhibition by tricarboxylic acids. J. biol. Chem. 249, 6149 6155. V I L L A F R A N C A J . J . ~ . M I L D V A N A. S. (1971) The mechanism of aconitase action. I. Preparation, physical properties of the enzyme, and activation by iron(Ill. J. biol. Chem. 246, 772 779.
0305-0491/82/020321-04503.00/0 Pergamon Press Ltd
Printed in Great Britain.
ACONITASE FROM THE OYSTER CRASSOSTREA VIRGINICA MALAK I. SHOUKRY* Department of Chemistry, University of Maryland, College Park, MD 20742, U.S.A.
(Received 14 September 1981) 1. The presence of aconitase activity in the oyster, Crassostrea riryinica, has been demonstrated. 2. Low levels of activity were found in the different tissues with highest level in digestive diverticular and lowest level in muscle. 3. The conversion of both citrate and iso-citrate to cis-aconitate suggests the presence of an enzyme system capable of utilizing these compounds at a slow but demonstrable rate to give classically expected results. 4. Comparison of the oyster enzyme with aconitase from mammalian tissue indicated great similarity between the two enzyme systems. Abstract
INTRODUCTION Isolated reports by investigators using molluscs have suggested that in this phylum, some organisms may have substantially modified pathways for carbohydrate metabolism. Jodrey & Wilbur (1955) have reported that the activities of the citric acid cycle enzymes in the mantle tissues of the American oyster, Crassostrea cirginica, were 1 3.8% of rat liver activities, aconitase was not detected. They also observed an increase in respiratory rate when iso-citrate was added to the mantle tissue but no increase when citrate was added. They correlated this with the absence of aconitase and suggested that citric acid was missing from the cycle. Black (1962) found aconitase in eggs a n d larvae of the same species of oyster. N o further investigation was made, and it has been tacitly assumed following the concept of biochemical unity of life, that aconitase is p r o b a b l y present in the oyster. The present study was undertaken to verify this suggestion in order to establish clearly the presence or absence of aconitase in the oyster and to study its properties.
MATERIALS AND METHODS All reagents were obtained commercially from Sigma Chemical Company (St. Louis, Missouri). Oysters were purchased at the Lexington Market, Baltimore, Maryland.
Preparation of the enzyme 100 g of freshly shelled oysters were homogenized with 300 ml of 4 m M Na-citrate buffer containing 10 mM fl-mercaptoethanol, 0.05 mM EDTA, adjusted to pH 5.0 with NaOH and 40 ml chloroform (Villafranca and Mildvan, 1971). Homogenization was carried out in Sorval "omni mixer" for 3 min at maximum speed. The crude extract was centrifuged at 20,000 ff for 20 min and the precipitate was discarded. The supernatant was brought to * Present address: Department of Biochemistry, Faculty of Medicine, Kuwait University, P.O. Box 24923, Kuwait. 321
50% saturation with solid ammonium sulfate (0.291 g/ml). The mixture was allowed to stand for 30 min then centrifuged at 18,000 g for 15 rain. The precipitate was discarded and ammonium sulfate (0.091 g/ml) was added to the supernatant to bring it to 65% saturation. The mixture w a s centrifuged as described above and the resulting precipitate was dissolved in a minimal volume of 20 mM Tris, 1 mM citrate 0.1 mM EDTA, and 0.1 mM fl-mercaptoethanol, at pH 7.4. 20 ml of this solution was applied to the top of a column (2.5 x 50 cm) of AcA 34 polyacrylamide agarose gel (LKB-Ultrogel), which had been equilibrated with 15 mM tricarballylate-Tris buffer containing 0.5 mM EDTA, l0 mM fl-mercaptoethanol, adjusted to pH 7.4, tricarballylate is not inhibiting the enzyme at the concentration used. The enzyme was eluted with the same buffer. This resulted in seven fold increase in specific activity, Table I.
Enzyme assay Aconitase was routinely assayed by the method of Racker (1950) by following the disappearance of cis-aconitate at 240nm. At 25°C, the assay solution contained 20 mM Tris HCI, 2 mM cis-aconitate, 100 mM NaCI and enzyme in a final volume of 0.5 ml at pH 7.4. Absorbance changes were measured in a Gilford 2400 recording spectrophotometer. A quartz insert was used to reduce the light path in the cuvette. When citrate or iso-citrate were used as substrates, the assay solutions contained 20 mM citrate or 5 mM O(+)iso-citrate, 25mM Tris HC1, and enzyme in a final volume of 1 ml at pH 7.4. One unit of enzyme activity is defined as the production of l llmol of product per min. A millimolar absorption coefficient of 4.88 m M - 1 cm- ~ for cis-aconitate at 240 nm was used. RESULTS
Aconitase activity Preliminary experiments indicated no aconitase activity in acetone powder preparations of whole oyster or from mantle and gill tissues. However, low levels of activity were found in fresh oyster tissues; the order of activity in the different anatomical parts was: digestive diverticular > mantle a n d gill > muscle (Table 2). The enzyme solution prepared was extremely un-
322
MALAK I. SHOUKRY Table 1. Preparation of aconitase from whole oyster tissues Purification Step Crude extract 50-65~o Ammonium sulfate Polyacrylamide agarose chromatography
U/ml
Protein mg/ml
Specific activity U/mg protein
109
5.03
21.7
1
404
5.96
67.8
3.12
41
140
0.88
7.33
17"
Purification
159
Y °'o 100
* Approximately one half of the 50-65~ ammonium sulfate fraction was processed on the polyacrylamide agarose column. The yield (Y), however, refers to the total volume. Table 2. Aconitase content of oyster anatomical parts Tissue Muscle Mantle and Gill Digestive diverticula
Units/ ml
mg protein/ ml
Specific activity units/mg protein
36.9 12.3 41.0
19.7 3.98 8.3
1.87 3.08 4.94
The reaction mixture contained Tris HC1 buffer, pH 7.4. 100mM NaC1. 0.1 mM cis-aconitate and enzyme in a final volume of 3 ml.
stable, losing 40-50~o of its activity in one day. However the a m m o n i u m sulfate fraction was less unstable, and after 10 days at 0°C, this fraction was found to have 60% of its activity remaining.
Michaelis constants Apparent Km's were determined for the three substrates at 25°C in Tris-HC1 buffer, at pH 7.4. Substrate concentrations were over the range of: 2 - 2 0 m M for citrate, 0 . 3 - 2 m M for D ( + ) iso-citrate and 0.2-2 m M for cis-aconitate. The following values were obtained: K,, citrate = 4.19mM, K,~ isocitrate = 0.57 m M and Km cis-aconitate = 0.21 mM. Although K,, values reported in the literature show a wide range even for aconitase from the same source (Table 3) the order is the same as found here, namely, K,~ citrate > K m iso°citrate > K m cis-aconitate. This wide variations is probably due to the different assay methods,
Optimum pH and temperature The pH activity relationship of aconitase over the range pH4.1-10.6 was also studied. The reactions Table 3. Comparison of Michaelis constants of aconitase* Source of enzyme Pig heart Pig heart Rabbit liver Beef heart Beef liver Mustard Oyster'["
K~, citrate (mM)
K,, cis-aconitate (mM)
Km isocitrate (raM)
3.6 0.62 0.9 0.95 0.20 4.0-4.4 4.19 (n = 4 )
0.12 0.015 0.09 0.099 0.007 0.1 0.21 (n = 3)
0.48 0.2 0.32 0.139 0.034 0.15 0.57 (n = 4 )
* From The Enzymes, Vol. 5, p. 426. f This study.
[citrate ---*cis-aconitate], [isocitrate---, cis-aconitate] and cis-aconitate hydration all exhibited the same optima, namely pH 7.0 in phosphate buffer and pH 7.5 in 3, 3 dimethylglutarate buffer. The above three reactions, examined in the 8-60°C range, exhibited the same optimum temperature of 42°C. Aconitase is heat labile, being completely denatured in 5 min at 60°C. The enzyme exhibited half optimal activity at 8°C.
lnhibitors of aconitase (a) Tricarboxylic acids. Inhibition patterns for trans-aconitate with the three substrates of aconitase are summarized in Table 4. It can be seen that the nature of inhibition depends on the substrate used. Thus trans-aconitate is a competitive inhibitor when cis-aconitate is the substrate but has a non-competitive component when citrate or iso-citrate are the substrates. Tricarballylate which is a substrate analogue of citrate and iso-citrate, exhibited similar behaviour, in that it is a competitive inhibitor of aconitase when citrate is the substrate but is a non-competitive inhibitor when cis-aconitate is used as substrate. An essential feature of a Kinetic scheme which shows both competitive and non-competitive behaviour is that Kt value derived from data, where I is a competitive inhibitor, must equal the K~ (slope) value where I is a non-competitive inhibitor (Villafranta, 1974). This was confirmed by the reasonably close values for Kt and Kis obtained in the present study (see Table 4). (b) SulJhydryl reagents. Dickman & Cloutier (1951), demonstrated that aconitase from pig heart could be rapidly and completely inhibited by I mM p-chloromercuribenzoate. In this study it was found that 1.5 m M p-chloromercuribenzoate completely inhibited the oyster enzyme suggesting that sulfhydryl groups might be necessary for enzymatic activity. The inhibition is non-competitive. N-Ethylmaleimide and
Aconitase from the oyster Crassostrea viryinica
323
Table 4. Inhibition constants for aconitase and substrate analogues Inhibitor
Substrate
Pattern
Kt*
K~st KHt
Trans-aconitate Trans-aconitate Trans-aconitate Tricarballylate Tricarballylate
cis-aconitate Citrate Isocitrate Citrate cis-aconitate
Competitive Noncompetitive Noncompetitive Competitive Noncompetitive
3.6 --
-3.9 2.11 -11
9
-0.6 0.9 0.14
* Calculated from the equation: 1__
K,,
v
v[s]
17+ E1 + [ 1KtJ l v"
"~Kts and Ktt are inhibition constants calculated from the slopes and intercepts respectively using the equation: ; = v[s]
l + K~s_l + ] + K ~ J v "
iodoacetamide were also tested (see Table 5), but were found to be far less effective. (c) Metal chelators. Compounds that bind iron are known to inhibit aconitase from other sources. In this study two such iron chelators, O-phenanthroline and ~-picolinic acid were tested at 2 and 10 m M concentrations respectively. As shown in Table 5, they had little effect on enzyme activity. A pre-incubation of the inhibitor with the enzyme is necessary for the inhibition to be demonstrable.
Activation by ferrous iron and cysteine It is well known that the rate of inactivation of an aging aconitase preparation can be decreased, by the addition of Fe 2 + and cysteine. The addition of these same compounds can also reactivate dialyzed samples. Aconitase prepared from oyster tissue was also found to lose activity upon storage or dialysis, so activation with iron and cysteine was tested. The effects of these compounds are shown in Table 6.
Table 5. Effect of metal chelators and sulfhydryl reagents on aconitase hydration (n = 3)
Reagent
Final conc. (mM)
Incubation period (min)
2 2 10 10
0 5 0 5
Metal chelators: O-phenanthroline ~-picolinate Sulfhydryl reagents: P-Chloromercuir benzoate N-Ethylmaleimide Iodoacetamide
0.5 1.5
5 0
1
5
2 2 2 3 3
5 5 15 5 15
~ Inhibition I1 33 4.8 17 17 100 32 46 11 21 11 22
Table 6. Restoration of activity of aconitase by Fe(lI) and cysteine Sample
9/0 Remaining activity*
(A) Enzyme solution dialyzed
85 against buffer1" n = 2 (B) Enzyme solution dialyzed against 65 buffer containing 1.0 mM O-phenanthroline n = 2 Addition of 2.0 mM Fe z+ + 10 mM cysteine hydrochloride Sample A 147 Sample B 135 * Relative to undialyzed sample. t 20 mM Tris buffer, containing 1.0 mM citrate, 0.1 mM EDTA, 0.1 mM fl-mercaptoethanol, pH 7.4.
324
MAI.AK 1. SHOUKRY tO0
o o o
o
o
Zg+, o
6C
Ni + ÷
•~ U"~4-
~' 40
•
2O
0 -4
-3
-2
log m o l a r i t y of metal ion
Fig. 1. Inhibition of [isocitrate---+ cis-aconitate] reaction by the metal ions, Cu 2+, Zn 2+ and Ni 2+. In addition to the metal ion, the reaction mixture contained 5.0ram D(+) isocitrate, 20raM Tris HCI buffer pH7.4 and enzyme in 1.0 ml. It must be noted here that b o t h iron and cysteine are required for the activation, since neither of them alone had any effect. Effects o f M e t a l Ions
Z n 2+, Cu 2+, a n d Ni 2. were tested for their effects on the enzyme activity. Cu 2+ at 1 m M completely inhibited the enzyme. However Z n 2+ up to 10mM caused no significant decrease in activity. With Ni 2+ there was a decrease in activity but even at 1 0 m M Ni 2+, some 50 60{~, of the activity remained. {See Fig. 1.) DISCUSSION The data presented d e m o n s t r a t e unequivocally that aconitase activity is present in oyster tissues, although at an extremely low level. In view of this low activity, it is u n d e r s t a n d a b l e why other investigators (Jodrey & Wilbur, 19551, could not detect any activity in this species. Additional reasons might be (1) the use of dilute homogenates (1:20W/V), (2) the use of phosphate buffer which is k n o w n to inhibit the enzyme (Morrison, 1954), and (3) the choice of mantle tissue as the sole source of enzyme. Localization of the enzyme to a particular subcellular fraction was difficult because of the low yield of enzyme as well as the lack of established methods for the fractionation and characterisation of subcellular organelles in the oyster which is k n o w n to be rich in glycogen and fibrous tissue. Aconitase from the oyster seems not to differ kinetically from aconitase from other sources. The enzyme loses activity upon storage or dialysis, partly due to the removal of iron, as indicated by the restoration of activity to dialyzed enzyme with iron (It) and cysteine, indicating the oyster aconitase is an Fe 2 + sulphur enzyme similar to the nonh a e m iron-sulphur proteins of the electron transport chain. This is supported by the observation that other
divalent metals, such as Z n 2 ~ and C u 2 ' , did not activate the dialysed enzyme in connection with aconitases from other sources. Inhibition studies with st, bstrate analogues suggest that two forms of aconitase exists, one preferentially binds cis-aconitate and its structu,al analogues, while the other preferentially binds citrate and its structural analogue. More detailed study with other tricarboxylic acids are needed to confirm this conclusion. These results however are consistent with the finding of Villafranca with pig heart aconitase. The interconvcrsion of the three acids: citric, cisaconitic and iso-citric by the isolated oyster aconitase, indicates that this portion of the citric acid cycle is operative in the oyster. The relative rate of the reaction [iso-cilrate--,cis-aconitate] was found to be twice that of [citrate---. cis-aconitate] in support of the k n o w n fact that the mitochondrial carbon flux is in the direction of citrate to iso-citrate. In the cytoplasm the flux is in the direction of iso-citrate to citrate, making citrate available for fatty acid biosynthesis and regulation of glycolysis. This may lead one to speculate that it is possible that the role of aconitase in the oyster may be different from the role of aconitase in the mitachondrial citric acid cycle in m a m m a l i a n systems. The oyster, Crassostrea rir,qinica has a high concentration of citrate, but the origin of this acid in this system has not been established. It is possible that citrate synthetase is present in the oyster. A low level of citric acid cycle enzymes are also present. These finding would suggest that the oyster m i t o c h o n d r i o n may have different control systems from the m a m m a l i a n mitochondrion. Perhaps the organelle has a different permeability 1o citrate. However such a theory could only be tested by further study. REFERENCES
BLACK R. E. (19621 Respiration, electron transport enzymes, and Krebs-cycle enzymes in early developmental stages of the oystcr Crassostrea virqinica. Biol. Bull. 123, 58 70. BLACK R. E. (1962) The concentrations of some enzymes of the citric acid cycle and electron transport system in the large granule fraction of eggs and trochophores of the oyster, Crassostrea vir~linica. Biol. Bull. 123, 71 79. DlC:KMa,N S. R. & CLOI T,iR A. A. (1951} Factors affecting the activit~ of aconitasc .l. hiol. Chem. 188, 379 388. HAMMIY C. S. (1969) Metabolism of the oyster. Crassostrea rir~tinica. Zoologist 9, 309 318. H()('HA('HKA P. W. & MtlSIAI:A T. (1972) Invertebrate facultative anaerobiosis. Science, N.E 178, 1056 1060. JODRr!'{ L. H. & WI~BtJR K. M. [1955l Studies on shell formation. IV, The respiratory metabolism of the oyster mantle. Biol. Bull. 108, 346 358. MORRISON J. F. (1954) The purification of aconitase. BiochenL d. 56, 99 105. RA('KER E. (1950) Spectrophotometric measurements of the enzymatic formation of fumaric and cis-aconitic acids. Biochem. Biophys. Acta 4, 211 214. Vn.LAFRANCA J. J. (19741 The mechanism of aconitase action, lI. Evidence for an enzyme isomerization by studies of inhibition by tricarboxylic acids. J. biol. Chem. 249, 6149 6155. V I L L A F R A N C A J . J . ~ . M I L D V A N A. S. (1971) The mechanism of aconitase action. I. Preparation, physical properties of the enzyme, and activation by iron(Ill. J. biol. Chem. 246, 772 779.