Mitochondrial biogenesis during fungal spore germination

125

Biochimica et Biophysica Acta, 6 0 6 ( 1 9 8 0 ) 1 2 5 - - 1 3 7 © Elsevier/North-Holland Biomedical Press

BBA 99582

MITOCHONDRIAL BIOGENESIS DURING FUNGAL SPORE GERMINATION PURIFICATION, PROPERTIES AND BIOSYNTHESIS OF CYTOCHROME v OXIDASE FROM BOTR YODIPLODIA THEOBROMAE

MARK JOSEPHSON

* and ROBERT

BRAMBL **

Graduate Program in Plant Physiology, The University of Minnesota, Saint Paul, MN 55108 (U.S.A.) (Received April 5th, 1979)

Key words: Mitochondrial biogenesis; Spore germination; Cytochrome c oxidase synthesis; Cytochrome c oxidase structure; (Botryodiplodia theobromae)

Summary 1. Cytochrome c oxidase (ferrocytochrome c:oxygen oxidoreductase, EC 1.9.3.1} was purified from the mycelial fungus Botryodiplodia theobromae by sodium cholate and ammonium sulfate solubilization, ammonium sulfate fractionation, and DEAE-cellulose chromatography. The purified enzyme contained 6--7 nmol heme a/mg of protein. The specific activity of the purified enzyme was 2.1--2.3 • 103 k (min -1) per mg of protein with 15 ~mol ferrocytochrome c and at pH 5.9 and optimal phosphate and Tween 80 concentrations (65 mM and 0.1%, respectively). The Km for ferrocytochrome c was determined to be 1.2--1.3 • 10 -s M, while at infinite substrate concentration the enzyme catalyzed the oxidation of about 60 pmol of ferrocytochrome c/min per mg of protein. Sedimentation behavior and kinetic evidence suggest that the purified enzyme exists as aggregates of the single molecule. The purified B. theobromae cytochrome c oxidase with its 428 nm Soret absorption maximum may be similar if not identical to oxygenated forms of the enzyme from other fungal species. 2. The purified enzyme was shown by sodium dodecyl sulfate polyacrylamide gel electrophoresis to consist of seven polypeptides with the following respective molecular weights: I, 41 000; II, 28 000; III, 19 000; IV, 14 800; * Present address: Department o f L a b o r a t o r y Medicine and P a t h o l o g y , The University o f Minnesota Medical School, Minneapolis, MN 55455, U.S.A. * * T o w h o m c o r r e s p o n d e n c e s h o u l d be addressed. Abbreviation: SDS, s o d i u m d o d e c y l sulfate; k, rate c o n s t a n t has units o f m i n - I .

126 V, 12 800; VI, 11 500; and VII, 9300. Biosynthesis studies showed that the three highest molecular weight polypeptides of the enzyme were synthesized on mitochondrial ribosomes, and the four smaller polypeptides were products of cytoplasmic ribosomes.

Introduction C y t o c h r o m e c oxidase, the terminal protein of the mitochondrial electron transport system, has been the object of extensive catalytic and spectral studies. In recent years studies of the structural properties of this enzyme and its biosynthesis have shown that c y t o c h r o m e c oxidase is a complex of at least seven subunits [1--3], the three largest of which are synthesized on mitochondrial ribosomes (presumably as products of mitochondrial genes); the remaining four subunits are synthesized on cytoplasmic ribosomes and are imported into the mitochondria for assembly into the enzyme complex (see Ref. 4 for citations of recent research). In our studies of germinating spores of the fungus Botryodiplodia theobromae, we are particularly interested in the structural and enzymic properties of cytochrome c oxidase and in describing the biosynthesis of this enzyme during germination. The dormant spores of Botryodiplodia contain an incomplete mitochondrial c y t o c h r o m e electron transport system which becomes functional during the first 120--150 min of the spore germination sequence through steps requiring cytoplasmic protein synthesis [5--7] and fatty acid biosynthesis [8]. The mitochondria of the dormant spores contain cytochrome c and one of the two b-type cytochromes of the germinated spores, but they do n o t contain c y t o c h r o m e a or heme a [6]. Importantly, c y t o c h r o m e c oxidase activity cannot be detected in the spore mitochondria until a b o u t 150 min of germination, after which time the extractable enzymic activity increases dramatically [7]. The development of cyanide-sensitive oxygen uptake [5] and elaboration of c y t o c h r o m e c oxidase [7] are both sensitive to cycloheximide, b u t n o t to inhibitors of the mitochondrial genetic system. It is possible that translation of the latent messenger R N A stored in the dormant spores [5,9] yields products which, early in germination, activate or supplement the incomplete, dormant spore respiratory system. The absence of c y t o c h r o m e c oxidase in the dormant spores of Botryodiplodia and its rapid elaboration during spore germination provide a novel opportunity to study regulation of respiratory membrane biogenesis in cells which are rapidly assembling a functional respiratory system. With this objective, we have devised a purification scheme for c y t o c h r o m e c oxidase, and we have described the properties of the purified enzyme from this organism.

Experimental procedures Techniques for production and germination of the Botryodiplodia spores have been described in detail previously [ 5,6]. The 16 h mycelium grown from germinated spores was collected by filtration, washed three times with mitochondrial extraction buffer solution, and stored at --70°C. The mitochondrial

127 extraction solution contained 0.25 M sucrose, 50 mM Tris-HCl (pH 8.0), and 1.0 mM EDTA. For large-scale isolation of mitochondria, frozen mycelium (1 kg) was shattered into small pieces, submerged in liquid nitrogen, and converted to a powder in a Waring Blendor by four 15-s bursts at high speed. Extraction solution (1.6--2.0 1) was added and the brei was thoroughly mixed for 3--4 min at high speed. The homogenate was centrifuged at 1750 X g (g values are gay) for 15 min, and the resulting pellets of cell debris were re-extracted by the above liquid nitrogen step. The pooled 1750 X g supernatant fractions were centrifuged at 25 000 Xg for 20 min or, alternatively, the mitochondria were pelleted by centrifugation of the 1750 X g supernatant fraction through a Beckman JCF-Z continuous-flow rotor at 18 000 rev./min with a flow rate of 90 ml/min. The mitochondrial fraction was resuspended in extraction solution, collected by centrifugation at 25 000 X g for 20 min, washed in a phosphate buffer (below) and re-centrifuged. The mitochondrial fraction was suspended to 20-25 mg protein/ml (in 50 mM potassium phosphate, pH 7.5, 0.9% (w/v) KC1, 1.0 mM EDTA) and stored at --70°C. Procedures for isolation of mitochondria from small quantities of isotopically labeled spores have been described [7]. All operations of the cytochrome c oxidase purification were carried out between 0 and 6°C and percentage saturation values for the ammonium sulfate fractionations were for 0°C. Suspensions of crude mitochondrial protein at 20--25 mg/ml (about 5 g of protein) were subjected to four 15-s bursts of sonic irradiation (with 30-s pauses) from a 0.75 inch horn of a Braunsonic 1510 sonic generator operated at 50 W. The suspension was then centrifuged at 10 000 Xg for 5 min and the supernatant fluid containing the submitochondrial particles was centrifuged at 65 000 X g for 180 min; the resulting supernatant fluid was discarded and the pellet resuspended (to 25 mg of original crude mitochondrial protein/ml) in 0.1 M sodium phosphate (pH 7.5), 0.5 mM EDTA. The submitochondrial particle suspension was made 4% (w/v) in cholic acid (decolorized and recrystallized) with the addition of a 20% stock solution (titrated to about pH 8.0) and stirred 15 min, after which time solid ammonium sulfate (164 mg/ml of submitochondrial particle/cholic acid suspension) was slowly added (taking about 30 min). This suspension (pH of about 7.5) was gently stirred for another 60 min and centrifuged at 65 000 Xg for 60 min. The pellet was discarded; solid ammonium sulfate was added (117 mg/ml of solution, 20--25 min for addition) to the yellow-green supernatant fluid and the mixture was stirred for 30 min and then centrifuged at 25 000 Xg for 30 min. The supernatant fluid was discarded; the red-brown pellet was resuspended in sucrose/ cholate buffer solution (0.25 M sucrose, 10 mM Tris-HC1 (pH 7.4), 0.5% cholate) at 15 ml/g crude mitochondrial protein and clarified by centrifugation at 25 000 Xg for 20 min. The resulting supernatant fluid was taken to 20% saturation (106 mg/ml) with saturated neutralized (NH4)2SO4. (All subsequent fractionations were made with this saturated neutralized (NH4)2SO4 solution, while centrifugations refer to 25 000 × g for 20 min.) The suspension was immediately centrifuged and the supernatant fluid was adjusted to 45% saturation (143 mg/ml) with (NH4)2SO4. After 15 min of stirring, the precipitated enzyme was collected by centrifugation, resuspended in 20 mM sodium phos-

128 phate (pH 7.0) and 0.25% (v/v) Tween 80 (5 ml/g crude mitochondrial protein), clarified by centrifugation, and mixed with 2 vols. of the sucrose/ cholate buffer solution. This solution was taken to 25% saturation (134 mg/ml) with (NH4)2SO4, stirred for 15 min and centrifuged. The 25% saturation supernatant fluid was adjusted to 45% saturation (115 mg/ml) with (NH4)2SO4, stirred 15 min and centrifuged. The 45% saturation pellet was resuspendcd in 50 mM Tris/acetate (pH 8.0) and 0.1% (v/v) Tween 80 at 2.5 ml/g of crude mitochondrial protein and clarified by centrifugation. The supernatant fluid was mixed with 2 vols. of sucrose/cholate buffer solution and taken up to 30% saturation (164 mg/ml) with (NH4)2SO4, stirred for 15 min and centrifuged. The supernatant fluid was taken to 40% saturation (56 mg/ml) with (NH4)~SO4, stirred 15 min and centrifuged. The 40% saturation pellet was resuspended in 50 mM Tris/acetate (pH 8.0), 0.1% Tween 80 (1 ml/g of crude mitochondrial protein); this solution was desalted by passage over a 2 cm × 30 cm Sephadex G-25 column equilibrated with the same buffer solution and was immediately applied to a precycled, equilibrated, DEAE-cellulose column (1.5 cm ~ 15 cm; DE-32, Whatman). The column was washed sequentially with two column volumes of application buffer solution containing 50 mM KC1. The flow rates of the application and washings were 50 ml/h. The enzyme was eluted at 10 ml/h with application buffer solution containing 300 mM KC1; the high 428nm absorbing fractions were pooled, mixed with 1 vol. of sucrose/cholate buffer and adjusted to 35% saturation (194 mg/ml) with (NH4)2SO4. The redgreen pellet obtained after centrifugation was dissolved in 50 mM Tris/acetate (pH 8.0), 0.1% Tween 80 {0.5 ml/g crude mitochondrial protein), mixed with 1 vol. of sucrose/cholate buffer solution and adjusted to 30% saturation (164 mg/ml) with (NH4)~SO4. This solution was stirred for 15 min and centrifuged; the resulting red-green pellet was dissolved in a minimal volume of 50 mM Tris/ acetate (pH 8.0), 0.1% Tween 80. Heme a to protein rations and enzymic activity determinations were performed immediately with this purified fraction. For storage at --70°C, the purified enzyme was diluted one to one with 50% glycerol (w/v). In some instances, the purified enzyme was fractionated further with ammonium sulfate, centrifuged through glycerol gradients (5--20%, w/v, at 300 000 × g), or applied a second time to DEAE-cellulose. Absorption spectra were recorded with a Cary 118C spectrophotometer equipped with a scattered transmission accessory, high-intensity light source and a 1 cm pathlength microcell attachment for work with 50-pl volumes. Lowtemperature absorption spectra were recorded with an Aminco-Chance DW-2 s p e c t r o p h o t o m e t e r at the temperature of liquid nitrogen. The heme a content of purified c y t o c h r o m e c oxidase was determined by preparing a direct pyridine hemochrome of the isolated enzyme [12]. In intermediate, turbid samples of the purification procedure, the differential coefficient of extinction 12.0 mM -1 • cm -1 [13] at 604 nm was used instead. Enzymic activity was measured according to Smith and Conrad [14] at 24--25°C in a buffer solution consisting of 65 mM potassium phosphate (pH 5.9), 0.1% Tween 80 and 15 ~mol ferrocytochrome c/ml. The first order velocity constant, k (min-1), is defined by and calculated from the equation of Smith and Conrad [14]. Specific activity is expressed as k (min -1) per mg protein, while the V is expressed as t~mol of ferrocytochrome c oxidized/min

129 per mg protein. Ferrocytochrome c was prepared and stored as previously described [7]. SDS-polyacrylamide gel electrophoresis of radioactively labeled and nonlabeled samples, and subsequent molecular weight determinations of the separated polypeptides were performed as previously described [15,16], with 10 cm X 0.6 cm 15% resolving gels and 1 cm X 0.6 cm 4% stacking gels. Gels with non-labeled samples were fixed overnight in 50% methanol/7% acetic acid, stained with Coomassie blue and destained in 5% methanol/7% acetic acid at 50°C. Previously published procedures were used for the determination of radioactivity is sliced gels [16], and the channels ratio method was employed for analysis of the radioactivity in each gel slice contributed by the isotopes in dual-labeled samples. Immunoprecipitations were performed with an antiserum raised in rabbits against holo-cytochrome c oxidase of Neurospora crassa (74A) as previously described by Werner [17]; our preliminary experiments were performed with an antiserum donated to us by Dr. Sigurd Werner. Cytochrome c oxidase was purified from Neurospora by Werner's procedure [ 18], with minor adjustments (as required) of the deoxycholate and ammonium sulfate fractionations. Neurospora was grown as described [18], and the mitochondria were prepared as outlined above. Radioisotopic labeling of the cells was performed as previously described [16]. Protein was determined by the method of Lowry et al. [19] for mitochondrial fractions and by the microbiuret method of Goa [20] for the purified enzyme and assorted fractions of the purification scheme. Hydrogen peroxide (0.3%, v/v) was included in the microbiuret procedure to bleach the hemoproteins in the various samples [21]. Bovine serum albumin was the protein standard for both methods of determination. Results

Purification of the enzyme Cytochrome c oxidase of Botryodiplodia was solubilized from submitochondrial particles with cholic acid and ammonium sulfate. It was then precipitated, resuspended, and fractionated with ammonium sulfate. The enzyme was further purified by DEAE-cellulose chromatography and subsequent ammonium sulfate fractionation. In some instances, purified enzyme preparations were recycled through DEAE-cellulose or were centrifuged through glycerol density gradients. The data in Table I show that the Botryodiplodia enzyme was purified about 35-fold with respect to heme a and approximately 25-fold with respect to catalytic activity. The purified enzyme, which in the reduced state had light absorption maxima at 604 nm and 443 nm, was judged to be free of contaminating cytochromes by room temperature absolute spectrophotometry (Fig. 1) and by examination of the direct pyridine hemochrome of the purified enzyme (data not shown), which yielded absorption peaks only at 587 nm and 430 nm, characteristic heme a absorption maxima [12]. Furthermore, contaminating cytochromes were not detectable by room temperature or low temperature --196°C) reduced minus oxidized difference spectrophotometry of the purified enzyme (data not shown).

130

TABLE I PURIFICATION

O F C Y T O C H R O M E c O X I D A S E F R O M B. ' Y ' I f E O B R O M A E Protein

Submitochrondrial particles 50% ( N H 4 ) 2 S O 4 1st 45% (NH4)2SO 4 2rid 4 5 % ( N H 4 ) 2 S O 4 40% (NH4)hSO 4 G-25 filtrate Pooled D E A E f~actions P o s t - D E A E 35% ( N H 4 ) 2 S O 4 P o s t - D E A E 30% ( N H 4 ) 2 S O 4

mg/ml

Total mg

Specific activity ~ (rain -I ) per rag)

15.4 10.3 15.1 14.1 15.7 6.4 5.4 9.3 9.1

3080.0' 659.0 408.0 169.0 67.5 59.5 38.9 23.3 20.0

8] 1105 1039 1300 787 937 1020 2133 2062

H e i n e a/ protein (nmol/mg)

P u r i f i c a t i o n (-fold) Activity

Heme a

0.18 0.48 0.59 1.39 3.13 -3.40 5.60 6.52

-13.6 12.8 16.0 9.7 11.6 12.6 26.3 25.5

-2.7 3.3 7.7 17.4 -18.9 31.1 36.2

We attempted to purify Botryodiplodia cytochrome c oxidase by other published procedures. Methods which were successful for yeast [10,11,22] gave unsatisfactory results with Botryodiplodia. In attempts to adapt the procedure of Mason et al. [10], we discovered that the Botryodiplodia enzyme was unstable in buffered Triton X-100 solutions. The instability of Botryodiplodia cytochrome c oxidase was characterized by degradation of the oxidized absorption spectrum at room temperature while recording a trace, although an undegraded spectrum could be recorded at low temperature (--196°C). Replacement of the Triton X-100 with Tween 80 permitted the recording of a stable spectrum at room temperature. This stability of the

I 28O

443

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T

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604

42B

C I 250

-----

300 -

3 350

--

450

Wovelength Fig. 1. A b s o l u t e a b s o r p t i o n oxidase. The oxidized trace was then treated with a few (B) w a s r e c o r d e d . T r a c e C c h a n g e in a b s o r b a n c c .

500

550

600

650

(nm)

s p e c t r a (at r o o m t e m p e r a t u r e ) o f t h e p u r i f i e d Botryodiplodia c y t o c h r o m e c (A) of the purified e n z y m e (1.16 m g protein/ml) was recorded. The sample g r a i n s o f s o d i u m d i t h i o n i t e , a n d a f t e r 25 r a i n o f i n c u b a t i o n t h e r e d u c e d t r a c e is t h e b a s e l i n e . N o t e t h e d i f f e r e n t v a l u e s o f o r d i n a t e scale e x p a n s i o n . AA,

131

enzyme in Tween 80 solutions was used in the subsequent development of a procedure for the purification of catalytically active and spectrally stable enzyme. Attempts also were made to employ the procedure of Rubin and Tzagoloff [22] and the adaptation of this method by Werner [18] for our purification of B. theobromae cytochrome c oxidase. However, we could n o t achieve a distinct separation of b and c-type cytochromes from the cytochrome c oxidase by differential deoxycholate-KC1 extraction of submitochondrial particles. Also, a 30°C or 37°C incubation step to precipitate contaminating proteins, when applied either to the successful purification procedure described here or to previously attempted methods [18,22], instead yielded a form of the Botryodiplodia enzyme which was insoluble in buffer solutions containing Tween 80.

Kinetic and spectral properties With the Tween 80 and ferrocytochrome c concentrations in the assay buffer solution at 0.1% (v/v) and 15 ~mol/ml, respectively, the optimum pH for the purified enzyme was found to be 5.90, and the optimum phosphate concentration (at pH 5.90) was 65 mM. These reaction conditions (at 24--25°C) gave purified enzyme specific activity values of 2.1--2.3 • 103 k (min -1) per mg of protein. Tween 80 enhanced the enzyme activity when it was present in the assay buffer solution at 0.05 or 0.1%, while higher concentrations were n o t as effective. Triton X-100 (0.05--1.0%) in the assay buffer solution inhibited enzymic activity (up to 70%} while Tween 20 and Tween 80 largely relieved the deleterious effect of Triton X-100. The K m for ferrocytochrome c, determined from Lineweaver-Burk plots, was 1.2--1.3" 10 -s M, while the V was about 60 ~mol ferrocytochome c oxidized/min per mg of protein. Cyanide (1 mM) inhibited (greater than 98%) the oxidation of ferrocytochrome c by the Botryodiplodia enzyme, and the oxidation of ferrocytochrome c was dependent upon the presence of the enzyme. SDS (0.01%, w/v) in the assay buffer solution eliminated the lag phase in the oxidation of ferrocytochrome c, suggesting that the isolated enzyme may exist in an aggregated form [23,24]. A room temperature absolute absorption spectrum of purified cytochrome c oxidase is shown in Fig. 1; the enzyme as isolated had absorption maxima at 604 nm, 428 nm, and 280 nm, and when the enzyme was reduced with dithionite, the absorption maxima appeared at 604 nm and 443 nm with a slight fl band at about 517 nm. In the reduced spectral trace, a slight shoulder appeared at about 420 nm which most likely represents a modified form of the enzyme [25] since no other cytochromes were detected in the preparation by difference spectrophotometry and only heme a was found in the direct pyridine hemochrome of the purified enzyme. The spectral properties of the purified Botryodiplodia enzyme show that the ratios of absorbance at 280 nm and 443 nm, and 443 nm and 428 nm (2.15 and 1.32, respectively} fit well within the limits of purity established by Yonetani [21]. The h e m e a content of the purified enzyme was 6--7 n m o l / m g of protein; this value was determined from the pyridine hemochrome extinction coefficient of 21.6 mM -1. cm -I at 604 nm derived for the native enzyme [12], or 27.4 mM -~ • cm -1 at 587 nm for the direct pyridine hemochrome [26]. The 604 nm minus 630 nm differential extinction coefficient of 16.5 mM -1. cm -1 [27] gave a heme a/

132 protein ratio of about 7--8 n m o l / m g of protein, but this value may be erroneous because the extinction coefficient 16.5 mM -I • cm -1 has been reported to be incorrect [ 28], yielding higher than actual heme a values. The oxidized spectrum of Botryodiplodia cytochrome c oxidase, with its Sorer absorption m a x i m u m at 428 nm, appears to be identical to oxygenated forms of the enzyme prepared from some other species by peroxide incubation or vigorous aeration of a previously reduced enzyme [29,30]. It is known that Tween 80 may contain a peroxide contaminant [31], and we therefore purified c y t o c h r o m e c oxidase in the absence of this detergent to learn whether the 428 nm absorption peak was generated by an artifactual oxygenation of the enzyme. With cholic acid as the exclusive solubilizing agent, the enzyme nevertheless yielded an absorption spectrum with a Soret m a x i m u m at 428 nm. This result showed that this enzyme spectrum was not generated by oxygenation of the cytochrome c oxidase in the presence of Tween 80. While cytochrome c oxidase from some organisms possesses an oxidized Soret peak at 418--422 nm [10,29,30,32--34], certain preparations of the enzyme from yeast [22] and those from Neurospora [18,35,36] yield a spectrum with the Soret peak at or near 428 nm, and it seems likely that the enzyme from Botryodiplodia is spectrally identical to these latter fungal enzymes. It is notable that when Botryodiplodia cytochrome c oxidase was treated with the chemical oxidant potassium ferricyanide, there was a shift of the Soret m a x i m u m from 428 nm to 423--425 nm. This observation and the fact that the 428 nm form of the Botryodiplodia enzyme does not revert to the 418--422 nm form upon standing for prolonged periods are in contrast to the findings of others [30]. The a peak absorbance of the ferricyanide-treated enzyme also was diminished and shifted to a shorter wavelength (about 600 nm); the ratio of the reduced a peak absorbance to the oxidized a peak absorbance increased from about 1.5 (untreated) to about 2 (treated). Indeed, oxygenated forms of c y t o c h r o m e c oxidase are characterized as having higher a peak absorbances than their oxidized (or ferric) counterparts [30]. These findings indicate that the Botryodiplodia enzyme (which exhibits a 428 nm Soret m a x i m u m and significantly reduced a peak as isolated) may represent a cytochrome c oxidase which somehow became altered in its spectral properties during its purification.

Structural properties Purified cytochrome c oxidase from Botryodiplodia was examined by SDSpolyacrylamide gel electrophoresis to determine the subunit composition of the enzyme. A typical Coomassie blue-stained gel electrophoresis pattern (Fig. 2) shows that this enzyme consists of at least seven polypeptide subunits with molecular weights of 41 000 (I); 28 000 (II); 19 000 (III); 14 800 {IV); 12 800 (V); 11 500 (VI), and 9300 (VII), as determined by co-electrophoresis with standard proteins of known molecular weights. All these polypeptides are likely to be authentic subunits of the Botryodiplodia cytochrome c oxidase because they co-purified through extensive a m m o n i u m sulfate fractionation, DEAE-cellulose chromatography, further a m m o n i u m sulfate fractionation, and a second DEAE-cellulose cycle or glycerol density gradient centrifugation (5--20% gradients). The subunit polypeptides (Fig. 2) do n o t appear to exist in stoichiometric quantities in these stained gel electrophoresis patterns, and this

133 IV V

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2

Relative Mobility Fig. 2. S D S - p o l y a c r y l a m i d e gel e l e c t r o p h o r e s i s of p u r i f i e d Botryodiplodia c y t o c h r o m e c oxidase. E l e c t r o phoresis, staining, a n d d e s t a i n i n g w e r e p e r f o r m e d a c c o r d i n g to t h e p r o c e d u r e d e s c r i b e d in t h e t e x t . A b o u t 50 ttg of protein w e r e a p p l i e d t o t h e gel. T h e C o o m a s s i e b l u e - s t a i n e d gels w e r e s c a n n e d a t 560 rim. T h e m o l e c u l a r w e i g h t s of t h e p o l y p e p t i d e b a n d s l a b e l e d are given in t h e t e x t . T h e a r r o w d e n o t e s t h e p o s i t i o n o f t h e t r a c k i n g d y e ( P y r o n i n Y) f r o n t .

effect is probably due to different affinities of the subunits for the Coomassie blue protein stain [35]. We learned that subunit I of the B. theobromae enzyme was uniquely sensitive to the brief boiling step (in the presence of SDS) which is routinely employed to ensure complete dissociation of SDS-denatured polypeptides. Exposure of the enzyme to heat usually causes an aggregation of the subunit I polypeptides so that they do not penetrate the gel. Throughout our earlier research we seldom saw more than trace amounts of subunit I in the electrophoresis patterns after boiling of the samples. Incubation of enzyme samples at 50°C for prolonged time periods (12 h or more) in the presence of SDS also was effective at promoting this aggregation of subunit I. Denaturation of the enzyme in SDS solutions was best achieved by incubation at room temperature for a b o u t 4 h or by overnight incubation at 4°C. Samples treated in this manner exhibited an intense subunit I band as shown by Coomassie blue staining. The unique sensitivity of subunit I to the heat treatment is unexplained, but the effect also exists in Neurospora, although not in yeast (Werner, S., personal communication). A possibly related observation with the rat liver c y t o c h r o m e c oxidase has been reported [37]. When the Botryodiplodia cytochrome c oxidase was centrifuged through glycerol gradients, it sedimented at a rate greater than that of catalase, which has a molecular weight of 240 000. In contrast, some reports [10,34] give a value of a b o u t 200 000 for the isolated c y t o c h r o m e c oxidase from other sources. This discrepancy may be due to the formation of aggregates by the purified Botryodiplodia enzyme, a condition which has been observed by others with enzyme from other sources [23,24] ; this explanation is supported by the observation that dilute concentrations of SDS (0.01%, w/v) in the assay buffer solution can eliminate a kinetic lag phase in the oxidation of ferrocytochrome c.

134

Biosynthesis of the enzyme subunit polypeptides To examine the ribosomal origin of the subunit polypeptides and to study the biosynthesis of the subunits during spore germination, we treated Triton X-100-solubilized membranes of labeled mitochondria from Botryodiplodia with an antiserum raised against holo-cytochrome c oxidase of Neurospora. The antiserum against the Neurospora enzyme accurately recognized and precipitated the counterpart enzyme from Botryodiplodia, as shown by the results in Fig. 3 in which the [14C]leucine-labeled mitochondrial extracts from Neurospora were mixed with the same fraction from [3H]leucine-labeled Botryodiplodia, treated with antiserum, and the resulting precipitate was electrophoresed. The Botryodiplodia polypeptides precipitated with this antiserum were identical in number and molecular weights to those of the purified enzyme described in the previous section. This experiment also provides a direct comparison of the molecular weights of the subunits of cytochrome c oxidase from the two organisms; the two largest subunits from each are identical in molecular weight, while the remaining five subunits are all of slightly lower molecular weight in Botryodiplodia. We confirmed the immunoprecipitation results by electrophoresis of a purified, [3H]leucine-labeled preparation of c y t o c h r o m e c oxidase from Botryodiplodia. We labeled the Botryodiplodia cells in the presence of cycloheximide or

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F i g . 3 . S D S - p o l y a c r y l a m i d e gel c o - e l e c t r o p h o r e t i c analysis o f i m m u n o p r e c i p i t a t e s o b t a i n e d f r o m l a b e l e d m i t o e h o n d r i a o f 16-h c u l t u r e s of Botryodiplodia ($) a n d Neurospora (o). T h e cells w e r e l a b e l e d f o r 1 8 0 r a i n w i t h [ 3 H ] l e u c i n e (Botryodiplodia) or [ 1 4 C ] l e u c i n e (Neurospora), a n d t h e e x t r a c t e d m i t o c h o n dria ( f o l l o w i n g T r i t o n X - 1 0 0 s o l u b i l i z a t i o n ) w e r e t r e a t e d w i t h a n t i s e r u m t o Neurospora c Y t o c h r o m e c o x i d a s e . M i g r a t i o n is f r o m l e f t t o right, a n d t h e c l o s e d b l o c k i n d i c a t e s t h e p o s i t i o n o f c y t o c h r o m e c at t h e termination of t h e e l e c t r o p h o r e s i s . B a c k g r o u n d c o u n t s h a v e b e e n s u b t r a c t e d f r o m t h e d a t a in this a n d t h e s u b s e q u e n t figure.

Fig. 4. S D S - p o l y a c r y l a m i d e gel c o - e l e c t r o p h o r e t i c analysis of i m m u n o p r e c i P i t a t e s o b t a i n e d f r o m l a b e l e d m i t o c h o n d r i a o f 16-h Botryodiplodia c u l t u r e s l a b e l e d for 1 8 0 rain in t h e a b s e n c e o f i n h i b i t o r w i t h [ 1 4 C ] l e u c i n e (o) o r l a b e l e d f o r 6 0 rain in t h e p r e s e n c e o f c y c l o h e x i m i d e w i t h [ 3 H ] l e u c i n e (e). T h e closed b l o c k indicates p o s i t i o n o f c y t o c h r o m e c.

135 chloramphenicol and determined the radioactivity in the electrophoretically separated subunit polypeptides precipitated by the antiserum against the c y t o c h r o m e c oxidase. This use of ribosome-specific inhibitors allows a demonstration of the ribosomal origin of the subunit polypeptides. We found, however, that a direct cycloheximide inhibition-labeling experiment would not ensure incorporation of [3H]leucine into the subunit polypeptides I--III. We observed, as have others with Neurospora [3,35], that a transitory incubation (30 min) of the cells in the presence of chloramphenicol is essential to cause an accumulation of cytoplasmic precursors, so that when the cells are washed free of the chloramphenicol and labeled in the presence of cycloheximide, there is a stimulation of mitochondrial protein synthesis [3,35]. In Fig. 4 is shown the results of an experiment in which an immunoprecipitate from the uninhibited [14C]leucine-labeled cells (16 h) was subjected to co-electrophoresis with an immunoprecipitate from cells labeled with [3H]leucine in the presence of cycloheximide (0.1 mg/ml) after a preliminary chloramphenicol (3.5 mg/ml) incubation. The results indicate that the synthesis of the four smaller cytochrome c oxidase subunits are sensitive to cycloheximide and that these four polypeptides are synthesized on cytoplasmic ribosomes. Conversely, when the Botryodiplodia cells were labeled with [3H]leucine in the presence of chloramphenicol (3.5 mg/ml) and the residual label and inhibitor were washed away to allow the cells to recover from the inhibition [3,35], we observed large, discrete peaks of radioactivity in the gel which corresponded exactly to subunits IV--VII (data not shown), as would be expected if these subunits were products of cytoplasmic ribosomes. Labeling of subunits I--III was sharply reduced by the chloramphenicol treatment, but it was n o t completely abolished. Nevertheless, this experiment demonstrates that subunits I--III likely are products of mitochondrial ribosomes. Discussion The catalytically active cytochrome c oxidase which we have purified from the ascomycetous fungus B. theobromae possesses spectral characteristics (such as the 280 nm/443 nm and the 443 nm/428 nm ratios) which fit well within the criteria of purity proposed by Yonetani [21] for c y t o c h r o m e c oxidase from beef heart. The enzyme purified from Botryodiplodia consists of at least seven polypeptide subunits which range in molecular weight from 41 000 to 9300. These polypeptides co-purified with the catalytic activity when the enzyme was subjected to extensive ammonium sulfate fractionation, DEAEcellulose chromatography, and glycerol density gradient centrifugation. All seven subunits were precipitated from radioactively labeled Botryodiplodia mitochondria after detergent lysis and treatment with an antiserum raised against the holoenzyme of Neurospora. The biosynthesis of the polypeptide subunits of the Botryodiplodia cytochrome c oxidase is dependent upon both the nucleocytoplasmic and mitochondrial genetic systems. The three high molecular weight subunits (I, 41 000; II, 28 000, and III, 19 000) were synthesized in the presence of cycloheximide, and we presume them to be products of the mitochondrial genetic system. Subunits IV--VII were labeled in the presence of chloramphenicol, indicating a

136

cytoplasmic ribosome synthesis of these polypeptides. The sites of synthesis of the Botryodiplodia enzyme subunits are identical to those of yeast [39] and Neurospora [ 35,38 ]. Cytochrome c oxidase activity is not detectable during the first 120 min of the 240 min spore germination sequence of B. theobromae, but after 120 rain of incubation there is a dramatic increase in the a m o u n t of enzyme activity [7]. Cytoplasmic (but not mitochondrial) ribosome function is essential for elaboration of cytochrome c oxidase activity up to about 100 min of germination; if cycloheximide is present during this period, development of the enzyme activity is completely inhibited [7]. Whether this required cytoplasmic translation product is' a regulatory protein or one or more of the cytoplasmic subunits of cytochrome c oxidase is n o t yet known. In other experiments (unpublished data) we found that only subunit polypeptides II and V were present in labeled enzyme immunoprecipitates from germinating spores, but this result could be attributed to the possible inability of our antiserum to the holo-cytochrome c oxidase to recognize all of the seven (unassociated) subunits of the enzyme or to differences in pool sizes of the unassociated subunit polypeptides. Experiments are now in progress with an antiserum against all enzyme subunit polypeptides to examine the timing of synthesis during spore germination o f each of the subunits of cytochrome c oxidase and to determine at which point they become associated with heme a to form a functional enzyme in the respiratory membrane. Because spore germination, respiration, and developm e n t of cytochrome c oxidase activity are dependent upon cytoplasmic protein synthesis, we also are testing whether the latent messenger RNA of the dorm a n t spores may code for one or more of the cytoplasmic subunits of the cytochrome c oxidase and whether translation of this RNA early in germination may contribute to the rapid subsequent development of the enzyme activity. Acknowledgments The Deutscher Akademischer Austauschdienst provided R.B. with a fellowship for a short-term study at the Institute for Physiological Chemistry and Physical Biochemistry of the University of Munich where Dr. Sigurd Werner generously contributed valuable help and advice for this research. This research was supported by Public Health Service research grant GM-19398 from the National Institute of General Medical Sciences and by a Faculty Grant-in-Aid of Research from the University of Minnesota Graduate School. References 1 T z a g o l o f f , A., R u b i n , M.S. a n d Sierra, M.F. ( 1 9 7 3 ) B i o c h i m . B i o p h y s . A c t a 3 0 1 , 7 1 - - 1 0 4 2 S c h a t z , G. a n d M a s o n , T . L . ( 1 9 7 4 ) A n n u . R e v . B i o c h e m . 4 3 , 5 1 - - 8 7 3 Weiss, H., S c h w a b , A . J . a n d W e r n e r , S. ( 1 9 7 5 ) i n M e m b r a n e B i o g e n e s i s ( T z a g o l o f f , A., e d . ) , p p . 1 2 5 - 1 5 3 , P l e n u m Press, N e w Y o r k a n d L o n d o n 4 T z a g o l o f f , A., M a c i n o , G. a n d Sebald, W. ( 1 9 7 9 ) A n n u . R e v . B i o c h c m . 4 8 , 4 1 9 - - 4 4 1 5 B r a m b l , R. ( 1 9 7 5 ) B i o c h i m . B i o p h y s . A c t a 3 9 6 , 1 7 5 - - 1 8 6 6 B r a m b l , R. a n d J o s e p h s o n , M. ( 1 9 7 7 ) J. B a c t e r i o l . 1 2 9 , 2 9 1 - - 2 9 7 7 B r a m b l , R. ( 1 9 7 7 ) A r c h . B i o c h e m . B i o p h y s . 1 8 2 , 2 7 3 - - 2 8 1 8 B r a m b l , R., W e n z l e r , H. a n d J o s e p h s o n , M. ( 1 9 7 8 ) J. B a c t e r i o l . 1 3 5 , 3 1 1 - - 3 1 7 9 Brambl, R.M. and Van Etten, J.L. (1970) Arch. Biochem. Biophys. 137, 442--452

137 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39

Mason, T.L., Poyton, R.O., Wharton, D.C. and Schatz, G. (1973) J. Biol. Chem. 248, 1346--1354 Eytan, G.D. and Schatz, G. (1975) J. Biol. Chem. 250, 767--774 Morrison, M. and Horie, S. (1965) Anal. Biochem. 12, 77--82 Van Gelder, B.F. (1966) Biochim. Biophys. Acta 118, 36--46 Smith, L. and Conrad, H. (1956) Arch. Biochem. Biophys. 63, 403--413 Maizel, J.V. (1971) in Methods in "virology (Maramorasch, K. and Koprowsld, H., eds.), Vol. 5, pp. 179--246, Academic Press, New York Brambl, R. and Handschin, B. (1976) Arch. Biochem. Biophys. 175, 606--617 Werner, S. (1974) Eur. J. Biochem. 43, 39--48 Werner, S. (1977) Eur. J. Bioehem. 79, 103--110 Lowry, O.H., Rosebrough, N.J., Farr, A.L. and Randall, R.J. (1951) J. Biol. Chem. 193, 265--275 Goa, J. (1953) Scand. J. Clin. Lab. Invest. 5, 218--222 Yonetani, T. (1961) J. Biol. Chem. 236, 1680--1688 Rubin, M.S. and Tzagoloff, A. (1973) J. Biol. Chem. 248, 4269--4274 Shakespeare, P.G. and Mahler, H.R. (1971) J. Biol. Chem. 246, 7649--7655 Vanneste, W.H., Ysebaert-Vanneste, M. and Mason, H.S. (1974) J. Biol. Chem. 249, 7390--7401 Elllott, W.B., Hulsmann, W.C. and Slater, E.C. (1959) Biochim. Biophys. Acta 33, 509--517 Warburg, O., Gewitz, H.S. and V61ker, W. (1955) Z. Naturforsch. 10b, 541--545 Griffiths, D.E. and Wharton, D.C. (1961) J. Biol. Chem. 236, 1850--1856 Vanneste, W.H. (1965) Biochem. Biophys. Res. Commun. 18, 563--568 Lemberg, M.R. (1969) Physiol. Rev. 49, 48--121 Lemberg, M.R. and Barrett, J. (1973) Cytochromes, Academic Press, New York Orii, Y. and Okunuki, K. (1965) J. Biochem. (Tokyo) 58, 561--568 Wainio, W.W. (1970) The Mammalian Mitochondrial Respiratory Chain, pp. 282--330, Academic Press, New York Sekuzu, I. and Takemori, S. (1972) in Electron and Coupled Energy Transfer in Biological Systems (King, T.E. and Klingenberg, M., eds.), pp. 325--378, Marcel Dekker, New Y ork Keyhani, J. and KeyhanL E. (1975) Arch. Biochem. Biophys. 167, 588--595 Sebald, W., Machleidt, W. and Otto, J. (1973) Eur. J. Biochem. 38, 311--324 Weiss, H., Sebald, W. and Bficher, Th. (1971) Eur. J. Biochem. 22, 19--26 Hundt, E. and Kadenbach, B, (1977) Hoppe-Seyler's Z. Physiol. Chem. 358, 1309--1314 Schwab, A.J., Sebald, W. and Weiss, H. (1972) Eur. J. Biochem. 30, 511--516 Mason, T.L. and Schatz, G. (1973) J. Biol. Chem. 248, 1355--1360