Saccharomyces cerevisiae Mitoproteome Plasticity in Response to

Saccharomyces cerevisiae Mitoproteome Plasticity in Response to Recombinant Alternative Ubiquinol Oxidase Gre´ gory Mathy,†,§ Rachel Navet,†,§ Pascal Gerkens,‡ Pierre Leprince,# Edwin De Pauw,‡ Claudine M. Sluse-Goffart,† Francis E. Sluse,*,† and Pierre Douette† Laboratory of Bioenergetics and Laboratory of Mass Spectrometry, Baˆt. B6c, Alleé de la Chimie 3, 4000, and Centre de Recherche en Neurobiologie Cellulaire et Molećulaire, Baˆt. B36, Avenue de l'Hoˆpital 1, 4000 Lie`ge, Belgium. Received October 13, 2005

The energy-dissipating alternative oxidase (AOX) from Hansenula anomala was expressed in Saccharomyces cerevisiae. The recombinant AOX was functional. A comparative analysis by two-dimensional differential in-gel electrophoresis (2D-DIGE) of mitochondrial protein patterns found in wild-type and recombinant AOX strains was performed. 60 proteins exhibiting a significant difference in their abundance were identified. Interestingly, proteins implicated in major metabolic pathways such as Krebs cycle and amino acid biosynthesis were up-regulated. Surprisingly, an up-regulation of the respiratory-chain complex III was associated with a down-regulation of the ATP synthase complex. Keywords: alternative oxidase • yeast • mitoproteome • proteomics • two-dimensional differential in-gel electrophoresis

Mitochondria from all plants, as well as from some Protists and Fungi, can possess two types of proteins specialized in energy-dissipation: uncoupling protein (UCP) and alternative oxidase (AOX). Simultaneous expression of these two energydissipating systems has been demonstrated in mitochondria from Acanthamoeba castellanii1,2 and Candida parapsilosis,3,4 and in plants, e.g., in tomato fruit during ripening and senescence.5,6 On one side, UCPs belong to the mitochondrial carrier family. During phosphorylating respiration, they compete with ATP synthase for the proton electrochemical gradient built up by the proton pumps of respiratory chain, leading to a decrease in the efficiency of oxidative phosphorylation. Although the regulation of UCP1 homologues activity is still under vigorous debate, UCP1 found in brown adipocytes sustains a proton conductance that is stimulated by free fatty acids and inhibited by purine nucleotides (PN).7 Recent data have shown that the inhibition of UCP1 homologues by PN is under the control of the redox state of coenzyme Q (Q), a redox intermediate of the respiratory chain, in ADP-phosphorylating mitochondria from plants, mammals, and protists.8-10 On the other side, AOX is a terminal oxidase branched upstream from the ubiquinol-cytochrome c oxidoreductase (complex III), where it sustains a cyanide-resistant respiration. AOX transfers electrons from the Q pool to oxygen without coupling the four-electron reduction of oxygen to proton translocation.11 Moreover, the engagement of the alternative * To whom correspondence should be addressed. Fax. +32-4-366-2878. E-mail: [email protected]. † Laboratory of Bioenergetics. ‡ Laboratory of Mass Spectrometry. § These authors contribute equally to the work. # Centre de Recherche en Neurobiologie Cellulaire et Mole ´ culaire. 10.1021/pr050346e CCC: $33.50

 2006 American Chemical Society

respiration has been found to occur at high Q redox state in mitochondria from Acanthamoeba castellanii1 and in higher plants.12 Two types of AOX can be phylogenetically distinguished according to their regulatory mechanism of activation. First, the plant-type AOX is present in the form of a nonfunctional covalent homodimer13 that is activated by post-translational modification in the presence of soluble reductants (e.g., thioredoxin or glutathione) and R-ketoacids (e.g., pyruvate) through the generation of a thiohemiacetal on a highly conserved cysteine.14 Second, the fungi-type AOX is strictly monomeric and is activated allosterically by 5′-monophosphate PN, such as GMP.15 Both types of AOXs are inhibited by Q analogues, such as benzohydroxamic acid (BHAM). Although UCP and AOX affect the oxidative phosphorylation process at two different level, i.e., the proton gradient for UCP (proton leak) and the redox potential for AOX (electron leak), they both lead to the same final effect, which is a decrease in ATP synthesis yield.16 A thermogenic purpose has been associated to these energy-dissipating systems in very specialized tissues, including the brown adipose tissue of hibernating mammals17 and the spadices of Araceae18 that use UCP1 and AOX, respectively, as free energy spendthrifts. Since UCP and AOX have been also found in nonthermogenic tissues as well as unicellular organisms, other physiological roles have been hypothesized. Energy-dissipation could have a role in the control of energy balance, i.e., the matching between the nutrient-derived supply of reducing substrates and the energy/ carbon demand for biosynthesis, by increasing the flexibility of mitochondrial respiratory activities.16,22,23 Accordingly, AOX has been though to work as safety valve for the Krebs cycle carbon flux to proceed even when overloads in redox potential and/or in phosphate potential occur (the energy-overflow Journal of Proteome Research 2006, 5, 339-348

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research articles hypothesis).16,22 Another physiological role for AOX and UCP has been suggested in which these energy-dissipating systems could prevent the generation of oxygen free radicals by the respiratory chain.23-26 There is a major difference between the mitochondrial Saccharomyces cerevisiae respiratory chain compared to plants, mammals, and the other unicellulars: Saccharomyces cerevisiae lacks complex I, which usually allows the electrons transfer from internal NADH to ubiquinone, associated with a proton pumping across the inner mitochondrial membrane. In Saccharomyces cerevisiae, the electron transfer is ensured by the rotenone-insensitive external19 and internal20 NADH dehydrogenases (NDE-1, NDE-2, and NDI-1) that allow oxidation of cytosolic and matricial NADH respectively, without proton pumping. Moreover, Saccharomyces cerevisiae mitochondrion does not possess the two energy dissipating systems described above. In a previous study, we used the baker’s yeast Saccharomyces cerevisiae as a recombinant host for UCP1.28 This gainof-function study allowed us to map the proteomic adaptation occurring in yeast mitochondria in response to the heterologous expression of active UCP1. We found that UCP1 is strongly connected to the energy-related enzymes at the protein level. Among others, we found that complex III as well as ATP synthase were up-regulated in response to UCP1 expression. These results suggested that the introduction of UCP1 dramatically affects the energy balance of the yeast, leading to an adaptative response. In the present work, we designed a recombinant Saccharomyces cerevsisae strain expressing the AOX cDNA isolated from other yeast, Hansenula anomala. We found that the recombinant product is properly adressed to the host mitochondria where it is functional. Moreover, the expression of AOX leads to a decrease in the growth rate of the yeast, suggesting that recombinant AOX exhibits its energy-dissipating activity in vivo. Using the proteomic technology, we studied the mitoproteome adaptation occurring in response to the presence of the alternative respiration. We found that AOX expression affects the energy-related enzymes in a very specific manner, which is different from the mitoproteome adaptation observed in mitochondria exhibiting UCP1.28 For example, whereas complex III, the direct competitor of AOX for electrons from reduced Q, was up-regulated, ATP synthase was downregulated in AOX-expressing yeast. Moreover, increase in enzymes related to the Krebs cycle demonstrates the functional linkage that exists between AOX and the Krebs cycle, strenghtening the energy-overflow hypothesis.23 With regard to our results, we hypothesize that the yeast can trigger a tailor-made mitoproteome adaptation that is specific to the recombinant energy-dissipating system. These observations underlie a tremendous mitochondrial plasticity in response to various mitochondrial endogenous stresses in yeast more likely through a complex mitochondria-to-nucleus crosstalk.

Material and Methods Materials. Cyanine dyes (CyDyes: Cy2, Cy3, and Cy5) and immobilized IPG strips were purchased from GE Healthcare. Sequencing-grade modified trypsin was from Roche. All other chemical were of the highest purity grade and were purchased from Sigma. CyDye, DeCyder, Ettan are trademarks of G. E. Healthcare. Strain, Plasmid, and Cloning. The HaAOX cDNA from Hansenula anomala29 (kind gift of Dr. Sakajo) was amplified 340

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by PCR using a forward primer (5′-ATGATTAAAACATATCAATATCGTTCAATTTT-3′) and a reverse primer (5′-CTAATGATGATGATGATGATGTAAACGCATC-3′) containing a sixhistidine tag for Western Blotting analysis. The PCR product was purified by electrophoresis on 1% agarose gel and inserted into the pYES2.1/Topo TA cloning vector (Invitrogen). After plasmid propagation in Top10F′ E. coli strain, the purified plasmid pYES-AOX that contains the AOX gene under the control of the GAL1 promoter was subcloned into the competent S. cerevisiae diploid INVSc1 (a/R) strain (Invitrogen) using a lithium chloride method (S.c. Easy comp Transformation Kit, Invitrogen). To assess the efficiency of heterologous expression in S. cerevisiae, a Kozak sequence (underlined) was also introduced at the translation initiation site by PCR using a forward primer (5′-AACAAAATGATTAAAACATATCAATATCGTT3′).30 The PCR product was cloned into the pYES2.1/Topo TA cloning vector (pYES-kAOX). Growth Condition. Freshly transformed wild-type (WT, empty pEMBL) and AOX-expressing yeasts were grown 36hours to a final optical density at 650 nm of ∼2 in 1.2 liters flasks in SC minimal medium containing 0.67% (w/v) yeast nitrogen base, 0.25% (w/v) (NH4)2SO4, 0.01% or 0.05% (w/v) amino acids supplemented with 5% glycerol and 2% galactose. Galactose induced the expression of the GAL1 promoted AOX cDNA cloned in the pYES-AOX vector. Mitochondrial Preparation and Oxygen Consumption Measurements. Crude mitochondria for functional assays were obtained by enzymatic spheroplasting using zymolyase 20T (ICN Biomedicals) followed by osmotic lysis and differential centrifugations procedure described by Daum and co-workers.31 Mitochondria devoid of microsomal and cytoplasmic contaminations for proteomic analyses were purified in a threestep sucrose gradient (2.5 mL 23%, 4 mL 32%, 2.5 mL 60% as described by Meisinger and co-workers.32 Mitochondria and cell respirations were assayed at 25 °C using a Clark-type electrode (Hansatech, U.K.) as described elsewhere.33 Cellular respirations were performed on the fresh SC minimal medium with 1 mM KCN or 2 mM of BHAM. Mitochondrial respiration were performed an incubation medium (Mannitol 0.65 M, TrisHCl 20 mM, EGTA 0.1 mM, BSA 0.1%, pH 7.5) supplemented with NADH 3 mM, succinate 1 mM, KCN 1 mM, BHAM 2 mM and GMP 1 mM. CyDye Labeling and Two-Dimensional Differential In-Gel Electrophoresis (2D-DIGE). The CyDye labeling of yeast mitochondrial proteins was performed as described in 26. Briefly, the mitochondrial proteins were extracted into the lysis buffer (7 M urea, 2 M thiourea, 2% (w/v) ASB-14 (zwitterionic detergent)) at 10 mg of proteins per ml and vortexed for 10 min. After removing the insoluble material by centrifugation, the pH was adjusted to 8.5 with 100 mM NaOH suitable for efficient CyDye labeling. Protein concentration was evaluated with the RC/DC Protein Assay (Bio-Rad Laboratories). Mitochondrial protein samples (25 µg) were set to a final protein concentration of 5 mg/mL with lysis buffer and labeled separately with 0.2 nmol of CyDye (Cy3, Cy5) (Amersham Bioscience), vortexed, and incubated 30 min in the dark. At the same time, a pooled sample composed of the equal amount of mitochondrial proteins from WT and AOX-containing samples was labeled with Cy2 that constituted an internal standard used for matching and normalization between gels.34,35 After 30 min in the dark, the reaction was stopped with 10 mM lysine. To control the labeling efficiency, labeled proteins (0.5 µg) were subjected to SDS-PAGE separation and the gels were scanned

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with the Typhoon 9400 scanner (G.E Healthcare) at the wavelengths corresponding to each CyDye, namely 480 nm (Cy2), 532 nm (Cy3), and 633 nm (Cy5). Differentially labeled mitochondrial samples (25 µg of each Cy2-, Cy3-, and Cy5-labeled sample) were pooled and resolved isoelectrically on 24-cm IPG strips, pH 3-10 NL on an IPGphor isoelectric focusing unit (G. E. Healthcare). Isoelectric focusing (IEF) was successively conducted at 200 V for 200 Vh, 500 V for 500 Vh, 1 kV for 1 kVh, and 8 kV for 60 kVh at 20 °C and a maximum current setting of 50 µA per strip. IPG strips were then equilibrated according to Go¨rg and co-workers36 and sealed on top of 12% w/v acrylamide gels. The seconddimension electrophoresis was performed overnight at 20 °C in an Ettan Dalt II system (G. E. Healthcare) at 1 W per gel. Each gel was finally scanned with the Typhoon 9400 scanner (G. E. Healthcare) at the wavelengths corresponding to each CyDye. To account for experimental bias due to in-gel protein aggregation, 2D-gels of mitochondrial samples were run in triplicate. Image Analysis. Images were analyzed with the DeCyder software (G. E. Healthcare) according to the manufacturer. In brief, co-detection of the three CyDye-labeled forms of each spot was performed using the DIA (Differential In-gel Analysis) module. The DIA software allowed the calculation of ratios between samples and internal standard abundances for each spot. Inter-gels variability was corrected by normalization of the Cy2 internal standard spot maps present in each gel by the BVA (Biological Variance Analysis) module. Protein spots that showed a statistically significant Student’s t-test (p < 0.05) for an increased or decreased in intensity were accepted as being differentially expressed in WT and AOX-containing samples. Protein Identification. 2D-gels were silver-stained using a MS compatible protocol.37 Spots that showed a significant variation in their abundance were manually excised from the gel and submitted to tryptic digestion following protein reduction (135 mM DTT) and alkylation (55 mM iodoacetamide). The resulting digested peptides were extracted, lyophilized and solubilized in 0.1% formic acid prior to MS analysis. Peptide sequencing was performed as described elsewhere28,38 using an Esquire HCT ion trap mass spectrometer (Bruker Daltonics) coupled to Ultimate Liquid Chromatography system (LC Packings) completed with Famos auto-sampler and a Switchos II microcolumn switching device for sample cleanup and preconcentration. Raw data were analyzed and formatted (Data Analysis software, Bruker) for subsequent protein identification against the NCBI nonredundant protein database through MS/ MS ions search algorithm on the Mascot search engine. The mass tolerance of precursors and sequence ions were set at 0.5 and 0.3 Da respectively and carbamido-methylation of cysteines and methionine oxidation were set as fixed and variable modifications, respectively. 2D-HPLC Proteomic Analysis. Purified mitochondrial proteins (100 µg) were dissolved in 100 µL of 50 mM NH4HCO3. The protein sample was then reduced by addition of 5 µL of 200 mM DTT in 100 mM NH4HCO3 followed by boiling during 10 min. The sample was alkylated for 90 min in the dark by adding 4 µL of 100 mM iodoacetamide in 100 mM NH4HCO3. The remaining iodoacetamide was then neutralized by adding 20 µL of 200 mM DTT in 100 mM NH4HCO3 for 45 min. Digestion was performed overnight at 37 °C using 2 µL of modified trypsin (Roche) (trypsin:sample ratio 1:50). Digestion was stopped with 5 µL of 500 mM HCl during 45 min at 37 °C

Figure 1. (A) Western blotting performed against WT and AOX+ isolated mitochondria with the anti-histidine-tag antibody coupled to peroxidase. Mitochondrial proteins were loaded at different amounts (10 µg, 25 µg, and 50 µg respectively) on SDS-PAGE. Immunoblotting reveals that only mitochondria isolated from yeast strain expressing the alternative oxidase exhibit crossreactivity with the anti-histidine-tag antibody. (B) Western blotting performed against WT, kAOX+ (with Kozak sequence) and AOX+ (without Kozak sequence). Twenty-five and 50 µg of mitochondrial proteins were loaded per lane. Immunodetection shows no difference in AOX expression level between the kAOX+ and AOX+ strain mitochondria.

(pH ) 2). Finally the digested sample was centrifuged at 16 000 × g for 15 min and aliquoted. A 30-µL portion of the peptide mixture was loaded into the 2D-HPLC system (LC Packings, Dionex) combining a first SCX column (LC Packings, µ-PrecolumnTM, Bio-X-SCX (500 µm × 15 mm)) followed by a reverse-phase liquid chromatography composed of a microprecolumn cartridge (300 µm × 5 mm, packed with C18PepMap, 5 µm, 100 Å) and a separating nano-column (75 µm × 15 cm, packed with C18 PepMap100, 3 µm, 100 Å. Peptides were eluted from the SCX column by different steps of salt concentration (20, 30, 40, 60, 80, 100, 200, 300, 400, and 800 mM ammonium acetate) and separated on the reverse phase system as described for in-gel digestion and protein identification. SDS-PAGE and Western Blotting. Proteins were subjected to electrophoresis in 8% SDS-polyacrylamide gel. Proteins were then transferred to a PVDF membrane (Immobilon-P, Millipore) using a Trans-Blot semi-dry transfer cell (Bio-Rad Laboratories) and analyzed by Western Blotting with an anti-His6 antibody coupled to peroxidase (Roche) using a BM Chemiluminescence Western Blotting Kit (Roche).

Results and Discussion Functional Expression of Recombinant HaAOX in Saccharomyces cerevisiae. To investigate the effect of AOX on the yeast metabolism at the mitoproteome level, we designed a recombinant yeast strain expressing AOX from Hansenula anomala. We cloned the HaAOX cDNA in the pYES2.1/Topo TA cloning vector that allows the recombinant expression of a protein of interest under the control of the GAL1 promoter inducible on galactose addition. We included a histidine-tag at the Cterminal end of HaAOX in order to ease Western Blotting analysis of the recombinant product using an antibody directed against the histidine epitope. After induction with 2% galactose in the SC minimal medium (see Material and Methods), Western Blotting analysis revealed the presence of a ∼32-kDa protein in AOX-containing mitochondria (Figure 1A). To assess the efficiency of expression of the recombinant AOX, we also cloned and expressed HaAOX carrying a Kozak sequence upstream from its AUG initiation codon (kAOX). In Eukaryotes, Kozak sequences modulate the translation of mRNA, while Journal of Proteome Research • Vol. 5, No. 2, 2006 341

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Figure 2. Growth curve of WT, kAOX+, and AOX+ yeast strains. Cells were transferred in culture media at an initial optical density of 0.02 at 650 nm and the first measurement was taken 12 h after innoculation, then every 2 h. Results show a marked growth defect in both kAOX+ and AOX+ strains. Generation times are respectively 240 min for WT and 320 min for both AOX+ strains. The latter result is in agreement with the AOX protein expression level detected by immunoblotting (Figure 1B).

addition of a Kozak sequence might dramatically affect the efficiency of expression of recombinant proteins in yeast (e.g., 10-fold for mammalian UCP1 expressed in yeast using a pYES2.1 vector).28 We found no difference at the protein level between the two genetic constructions, i.e., pYES-AOX and pYES-kAOX (Figure 1B). We hypothesize that the close phylogenetic relationship existing between H. anomala and S. cerevisiae allows a suitable ribosomal recognition of HaAOX

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mRNA in S. cerevisiae leading to a high expression efficiency of the recombinant product even in the absence of a Kozak sequence. Interestingly, expression of HaAOX in S. cerevisiae induced a ∼30% growth defect on a nonfermentable carbon source as shown in Figure 2, with a doubling time of ∼240 min for the WT strain and of ∼320 min for the pYES-AOX-containing strain (AOX+ and kAOX+) during the exponential growing phase. Oxymetric assays on intact yeast cells showed the presence of a cyanide-resistant BHAM-sensitive respiration in AOX+ strain, typical of a functional AOX (Figure 3A). Moreover, GMP, a PN that regulates allosterically fungi-type AOXs, further activated (21%) the AOX-sustained respiration as evidenced on isolated mitochondria during nonphosphorylating (state 4) respiration (Figure 3B). These results suggested that the recombinant HaAOX is properly addressed to the mitochondria of S. cerevisiae where it is able to catalyze its energy-dissipating activity in vivo leading to a macroscopic phenotype, i.e. a growth defect. Two-Dimensional Differential In-Gel Electrophoresis (2DDIGE) and Comparative Analysis of WT and AOX-Containing Mitoproteomes. Since we found a specific adaptation of the yeast mitoproteome in response to the introduction of UCP1,28 which is another type of energy-dissipating system absent from the yeast, we then wondered whether HaAOX could also trigger an adaptative response in the yeast. Moreover, two-dimensional differential in-gel electrophoresis (2D-DIGE) has been successfully used for a quantitative comparative proteomic analysis of WT vs UCP1-carrying yeast mitochondria.28 Consequently, we used the 2D-DIGE technology to compare the mitopro-

Figure 3. (A) Cell respiratory measurements performed on AOX+ strain. Total cell respiration was inhibited by 79% with 1 mM KCN (left). The resulting cyanide-resistant respiration was inhibited 92% following addition of 2 mM BHAM. Total cell respiration was inhibited 72% by 2 mM BHAM (right). The resulting BHAM-insensitive respiration was inhibited 93% by 1 mM KCN. These results are in agreement with a functional activity of the recombinant alternative oxidase. (B) Respiratory measurements performed with nonphosphorylating mitochondria (state 4, AOX+ strain). Addition of 1 mM KCN induced a 61% decrease in the mitochondrial state 4 respiration. The remaining cyanide-insensitive respiration was then increased by 21% 2 mM GMP (an allosteric activator of fungi-type AOX). Finally, KCN-insensitive GMP-stimulated respiration was inhibited by 68% 2 mM BHAM, suggesting that the recombinant alternative oxidase is functional and suitably regulated in isolated mitochondria. 342


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S. cerevisiae Mitoproteome Plasticity Table 1. List of Proteins Exhibiting Different Abundance in AOX+ versus WT Yeasta spot

nom

Q

t-test

1 22 6 11 7 3 9 21 3

0.0017 0.00017 0.0095 0.0027 0.0022 0.0037 0.022 0.0027 0.00036

V

Mw

pI

2.09 1.72 1.52 2.32 1.72 1.62 1.31 1.31 1.68

39 886 39 300 85 836 85 836 85 836 53 151 53 574 47 156 70 229

8.84 9.81 8.15 8.15 8.15 8.26 8.48 7.09 7.47

1.4 -1.82 -1.69 -1.7 -1.73 -2.24 1.33 1.32 1.27 1.57 1.68

34 174 58 855 54 924 54 924 54 924 54 924 50 254 50 254 23 635 50 228 70 229

8.65 8.95 5.71 5.71 5.71 5.71 6.77 6.77 8.24 7.24 7.47

1.42 1.38 2.31 3.71 3.68 1.52 1.39 1.31 1.6 1.61 1.72 1.63

34 326 75 061 68 408 68 408 68 408 67 900 40 329 40 329 44 585 44 585 44 585 43 797

6.07 8.59 5.89 5.89 5.89 6.75 8.15 8.15 9.83 9.83 9.83 9.6

1.31 1.41 4.04 1.25

64 680 64 076 95 267 63 221

6.54 8.67 8.42 9.15

1.73

44 739

9.09

1.47 1.76

33 204 40 353

5.18 6.33

1.21 1.25 1.23 1.42 1.41 1.51 1.38 1.65 1.38 1.29 1.32 1.37

70 585 70 585 70 585 60 999 60 999 72 377 48 113 93 228 51 109 48 881 48 881 42 206

5.48 5.48 5.48 5.23 5.24 6.09 6.19 6.25 6.54 9.44 9.44 5.48

Krebs Cycle 1052 1051 144 150 152 1335 761 899 327 1463 1374 767 1260 1758 1761 837 838 1840 1067 327 1289 313 287 312 320 318 1028 1031 1078 1045 1043 1073 485 549 708 684 1061 1439 1441 717 289 297 409 411 238 861 836 796 895 939 904

isocitrate dehydrogenase IDH2 precursor isocitrate dehydrogenase IDH1 precursor Aconitate hydratase Aconitate hydratase Putative aconitase Fumarate hydratase, mitochondrial Fumarate hydratase probable succinate-CoA ligase (GDP-forming) succinate dehydrogenase (Flavoprotein)

Respiratory Chain NADH-cytochrome-b5 reductase 9 0.0021 F1F0-ATPase alpha subunit 13 1.80E-05 ATP synthase beta chain 2 0.002 ATP synthase beta chain 32 0.011 ATP synthase beta chain 4 0.00054 ATP synthase beta chain 9 0.001 ubiquinol-cytochrome-c reductase 44K core protein 13 0.011 ubiquinol-cytochrome-c reductase 44K core protein 33 0.0029 ubiquinol-cytochrome-c reductase Rieske iron-sulfur protein 9 0.024 Ubiquinol-cytochrome-c reductase complex core protein I 1 0.0022 succinate dehydrogenase 3 0.00036 Branched-Chain-Amino Acid Synthesis acetolactate synthase (EC 4.1.3.18) regulatory chain 3 0.024 acetolactate synthase (EC 4.1.3.18) 6 0.0027 2-isopropylmalate synthase I LEU4 6 0.00053 2-isopropylmalate synthase I LEU4 16 8.30E-05 2-isopropylmalate synthase I LEU4 1 0.00014 2-isopropylmalate synthase II LEU 9 2 0.0018 3-isopropylmalate dehydrogenase 16 0.003 3-isopropylmalate dehydrogenase 6 0.0047 Acetohydroxy-acid isomeroreductase 37 0.0022 Acetohydroxy-acid isomeroreductase 4 0.00094 Acetohydroxy-acid isomeroreductase 10 0.00051 Branched-chain-amino acid aminotransferase 4 0.00065 Amino Acid Biosynthesis and Degradation 1-pyrroline-5-carboxylate dehydrogenase 18 0.035 threonine ammonia-lyase 8 0.013 acetylglutamate kinase 2 1.80E-05 glycine hydroxymethyltransferase 18 0.0024 Haem biosynthesis Ferrochelatase, mitochondrial 11 0.0019 Oxidative Stress Cytochrome c peroxidase 13 0.00068 Cytochrome c peroxidase 5 0.00059 Protein Fate (folding, import) dnaK-type molecular chaperone SSC1 3 0.0024 dnaK-type molecular chaperone SSC1 15 0.0084 dnaK-type molecular chaperone SSC1 30 0.00013 heat shock protein HSP60 precursor, mitochondrial 21 0.013 heat shock protein HSP60 precursor, mitochondrial 21 0.00026 SSC 2 DNA K type molecular Chaperonne (mtHSP70) 4 0.038 translation elongation factor EF-Tu 5 0.00046 Elongation factor Tu 4 0.0072 mitochondrial processing peptidase 6 0.037 mitochondrial import protein MPI1 precursor 3 0.048 mitochondrial import protein MPI1 precursor 3 0.035 mitochondrial import receptor chain TOM40 7 5.40E-05

a Mitochondrial proteins that significantly varied in AOX+ versus WT yeast (p < 0.05, student’s t test) are organized according to their general function. Legend abbreviations: Q, queries, number of peptide identified by mass spectroscopy; t-test, value of the student’s t-test; V, amplitude of variation where a positive value means that the amount of protein is increased in AOX+ yeast mitochondria; MW, molecular weight; pI, isoelectric point.

teomes of WT and AOX+ mitochondria. 2D-DIGE enables the visualization of multiple protein samples on one 2D-gel by means of a differential minimal labeling of the protein samples with spectrally resolvable fluorescent dyes (N-hydroxy succinimide derivatives CyDyes, Cy2, Cy3, and Cy5). This fluorescencebased technology allows the detection of more subtle changes in protein expression profile supported by statistical confidence with a detection limit of ∼150-500 pg for a single protein and a very high dynamic range (∼105).39,40 Multiplexing of the Cy3-

and Cy5-labeled independent protein samples in one gel can be completed by the addition of a Cy2-labeled internal standard composed of equal amounts of the protein samples under comparison. The use of this pooled internal standard that is run among all gel replicates allows inter-gel normalization, which further decreases the gel-to-gel variation and enables the detection of 95% statistical confidence.35 Accordingly, purified mitochondria from WT and AOX+ strains were differentially labeled and Journal of Proteome Research • Vol. 5, No. 2, 2006 343

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Figure 4. Mitochondrial protein pattern by 2D-DIGE. Yeast mitochondria were isolated and mitochondrial proteins were extracted by using lysis buffer (7 M urea, 2 M thiourea, 2% ASB-14). WT and AOX+ yeast mitochondria were differentially labeled with Cy3 and Cy5. An internal standard composed of equal amount of each mitochondrial sample and labeled with Cy2 was added to improve comparative analysis. Labeled samples (25 µg of each Cy2, Cy3, and Cy5) were loaded on 24-cm 3-10 NL IPG-strips and subjected to isoelectrofocusing. Second dimension was performed in 12% acrylamide gels. Gels were then scanned in a wavelength-selective way and subsequent image analyses were performed with the DeCyder software including the DIA and BVA modules (G. E. Healthcare). The proteins, which were significantly up- or down-regulated ((1.2 f (∞, p < 0.05, Student’s t-test in three independent gels) in AOX+ mitochondria compared to WT and successfully identified by LC-MS/MS, are marked with the number allocated by the DeCyder software.

analyzed by 2D-DIGE. Each gel was scanned using the three wavelength specific of the CyDyes. About 2614 spots were detected on each gel. According to our statistical treshold (p < 0.05, Student’s t test), 143 protein spots of the overall 2614 spots exhibited differences in normalized spot volume ratios exceeding 20%. Among them, 113 spots were increased while 30 spots were decreased in the AOX+ mitoproteome. Protein spots that varied between WT and AOX+ mitoproteome were subjected to in-gel trypsin digestion and peptide sequences were determined by ion trap mass spectrometry coupled to liquid chromatography (Table 1). 57 proteins (Figure 4) were identified successfully with significant score. To assess the purification protocol we performed a 2D HPLC to check mitochondrial purity on WT mitochondria. On 224 proteins detected by HPLC, we observed that 97% were of mitochondrial origin with only small contaminations from the nucleus, the cytoplasm and the plasma membrane (Figure 5) Adaptation of Enzymes Related to the Energy Metabolism. In our comparative 2D-DIGE analysis, we found that 6 enzymes of the Krebs cycle (i.e., aconitase, isocitrate dehydrogenase-1 (Figure 6A) and isocitrate dehydrogenase-2, succinyl CoA lyase, succinate dehydrogenase (Figure 6B) and fumarase) were strongly up-regulated in response to the introduction of AOX. Since the Krebs cycle is the main source of reducing substrates (NADH, FADH2) for the respiratory chain, up-regulation of 344


Figure 5. Subcellular identity of proteins contained in the purified WT mitochondrial fraction isolated from the 32%/60% interface on the three-step sucrose gradient (as described under the “Material and Methods” section). Proteins were digested by trypsin in solution and the peptide mixture was separated by 2DHPLC prior to MS sequencing. 224 proteins were identified from WT mitochondrial fraction.

enzymes of the Krebs cycle would accelerate the substrate flux in order to increase the reducing substrate supply to the respiratory chain. The increase in the substrate flux is further confirmed by the increase in subunits of the pyruvate dehydrogenase that generates acetyl-CoA for the Krebs cycle from pyruvate.


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Figure 6. Comparative analysis of four protein spot intensities using the BVA module of the DeCyder software. The selected spots are displayed as partial view of the 2D-gel and as three-dimensional images. Spot boundary of selected proteins is displayed in pink and are pointed out by an arrow.

At the level of the respiratory chain, we found a ∼1.4-fold increase in the proton pumping complex III (ubiquinolcytochrome c oxidoreductase) (Figure 6C), which competes with AOX for the reduced Q, as well as a ∼1.7-fold increase in the succinate dehydrogenase that delivers electrons produced by the reduction of succinate into fumarate to the Q pool. In addition, the NADH-cytochrome c reductase (MCR1) that transfers electrons from the externally produced NADH directly to cytochrome c was ∼1.4-fold greater in AOX+ mitochondria. This increase in MCR1 could force electrons to pass through the cytochrome c oxidase, the other proton pump of the yeast respiratory chain. Up-regulation of the complex III as well as of MCR1 would influence the electron partitioning at the level of the Q pool in favor of the cytochrome pathway in order to diminish the impact of recombinant AOX on the oxidative phosphorylation and the energy conservation. One of the most striking observations takes place at the level of the ATP synthase. Indeed, we found a 2-fold decrease in three soluble subunits of the ATP synthase (Figure 6D). The ATP synthase consumes the proton electrochemical gradient (∆µH+) built up by the respiratory chain in order to synthesize ATP. Moreover, it is well-known that ATP synthase is very sensitive to minute changes in the membrane potential while it is turned off below ∼150-160 mV. On the other hand, it is noteworthy that, by diverting electrons from the cytochrome pathway, AOX decreases the efficiency of proton pumping and consequently affects the establishment of the proton electro-

chemical gradient (∆µH+). Therefore, a decrease in the ATP synthase content would allow the establishment of a novel steady state between the rate of ∆µH+ building and the rate of ∆µH+ consumption, favorable to the ATP synthase to perform ATP synthesis. Adaptation of Enzymes Related to the Biosynthesis of Branched-Chain Amino Acids. Among the pathways that were affected by the recombinant expression of AOX, we found increases in proteins that catalyze the biosynthesis of branchedchain amino acids, i.e., valine, leucine, and isoleucine. Acetohydroxyacid isomeroreductase (ILV5), which generates Rβdihydroxyisovalerate from R-acetolactate, was greater ∼1.6-fold in AOX+ mitochondria. Then R-ketoisovalerate can be converted either into R-isopropylmalate by the R-isopropylmalate synthase (LEU4/LEU9) at the expense of acetyl-CoA or into valine by the branched chain amino acid aminotransferase (BAT1). Finally R-isopropylmalate is exported to the cytosol where it is converted into leucine. LEU4, LEU9 and BAT1 were up-regulated ∼3-fold, ∼1.5-fold, and ∼1.7-fold respectively in AOX+ mitochondria, suggesting a strong increase in the biosynthesis of leucine as well as valine. Interestingly exported R-isopropylmalate is very important in the nitrogen metabolism in yeast.41 It is an activator of the Leu3p transcription factor that controls the expression of a number of target genes including ILV5, LEU4, and potentially BAT1 coding for the mitochondrial proteins mentioned above as well as GDH1 that is essential in the early step of nitrogen Journal of Proteome Research • Vol. 5, No. 2, 2006 345

research articles metabolism, i.e., assimilation of ammonia.42 Therefore, upregulations of ILV5, LEU4, and BAT1 could be a direct consequence of the accumulation in the cytosol of R-isopropylmalate that acts as a transactivator for these genes. Finally, since biosynthesis of branched-chain amino acids requires pyruvate and acetyl-CoA, an increase of this biosynthetic pathway is more likely related to the availability in acetyl-CoA. We also found 4 other enzymes related to other amino acid biosynthesis and degradation pathways. Glycine hydroxymethyltransferase that catalyzes the conversion of glycine to serine was ∼1.25fold increased in AOX+ mitochondria. We also observed increases in the acetylglutamate kinase (4-fold) that catalyzes the second step in arginine biosynthesis, the 1-pyrroline-5carboxylate dehydrogenase (∼1.3-fold) involved in proline catabolism and the threonine ammonia-lyase (∼1.4-fold) that catalyzes the degradation of both L-serine and L-threonine Impact of AOX on the Mitochondrial DNA Maintenance and on Mitochondrial Protein Biogenesis. Mitochondrial DNA maintenance is essential to maintain the respiratory activity of mitochondria and is regulated through protein-DNA complexes called mtDNA nucleoids. mtDNA nucleoids are composed of proteins related to four functional categories:43 (I) mtDNA activities; (II) mitochondrial protein biogenesis; (III) the Krebs cycle; and (IV) amino acid biosynthesis. Recent data established the direct relationship between the metabolic regulation and the mDNA maintenance through a switch of function between the Krebs cycle and the protein-DNA nucleoid complex.43 We found that several major nucleoid components were increased in response to the introduction of AOX, which belong to the category II (mitochondrial chaperonin Hsp60 and protein import chaperone SSC1), III (aconitase, isocitrate dehydrogenase) and IV (acetohydroxyacid isomeroreductase and threonine ammonia-lyase). Aconitase is particularly essential for the mitochondrial DNA maintenance. According to our results, expression of AOX might lead to an increased capacity of mtDNA maintenance. Moreover, increases of heat shock proteins (Hsp60, SSC1) as well as proteins of the TIM/TOM import machinery (TOM40, MPI1 and the mitochondrial processing peptidase MPP) strongly support the idea of a higher activity of mitochondrial protein biogenesis in order to sustain the mitochondrial adaptation occurring in AOX+ mitochondria. Accordingly, the mitochondrial elongation factor EF-Tu (EFT1) involved in the translation of mitochondrial mRNA for subunits of the respiratory complexes was up-regulated in response to AOX expression. We also found that ferrochelatase, the enzyme that catalyzes the last step of haem biosynthesis, was 1.7-fold greater in AOX+ mitochondria. It is noteworthy that haem is essential for catalytic activity of many proteins involved in electron transfer, such as complex III that was also found in greater amount when AOX was heterolougsly expressed.

Conclusion The heterologous expression of a plant-type AOX from Sauromattum guttatum has already been achieved in the yeast Schizosaccharomyces pombe leading to a 18% growth defect on nonfermentable carbon source (glycerol).44 The effect was related to the nonprotonmotive AOX activity that competes with the cytochrome pathway for the reducing substrates. In the present work, we were able to obtain a 30% growth defect by expressing AOX from the yeast Hansunela anomala in another yeast Saccharomyces cerevisiae. We found that HaAOX was highly expressed while the expression level did not seem 346


Mathy et al.

Figure 7. Scheme summarizing the effect of the recombinant alternative oxidase on the yeast mitoproteome. In red (+) and in blue (-) are pathways and proteins that were found to be up regulated and down regulated, respectively, in mitochondria expressing the alternative oxidase. Several important pathways such as Krebs cycle, amino acid biosynthesis, protein fate, haem biosynthesis, and respiratory chain are strongly affected by the expression of the AOX. Abbreviations: VDAC-1: Voltage dependent anionic channel (mitochondrial porin), NDE: external NADH dehydrogenase, NDI: internal NADH dehydrogenase, SDH: succinate dehydrogenase, MCR-1: NADH/cytochrome c oxidoreductase, PDH: pyruvate dehydrogenase, ILV-5: acetohydroxy-acid isomeroreductase.

to be influenced by the addition of a Kozak sequence upstream from the HaAOX cDNA. This high level of expression is more likely related to the close phylogenetic resemblance between the two types of yeast. Moreover, recombinant HaAOX induced a cyanide-resistant BHAM-sensitive respiration, which was sensitive to GMP, an allosteric activator of fungi-type AOX. By 2D-DIGE, we found that the introduction of HaAOX leads to a specific adaptative response from the yeast that mainly affects the energy metabolism, the biosynthesis of branched-chain amino acids as well as the biogenesis of mitochondrial proteins (Figure 7). These results strongly suggest that HaAOX is functional during the cell growth in vivo and affects energy conservation leading to growth impairment, depsite adaptations of the yeast mitoproteome. We previously observed that, despite the growth defect, the yeast tried to regulate its energy metabolism by adaptating specifically its mitoproteome in response to the introduction of UCP1, another type of energy-dissipating system.28 Expression of recombinant UCP1 leads to an increase of ATP synthase, the direct competitor of UCP1 for the proton electrochemical gradient, as well as of key-enzymes of the energy metabolism, including complex III and proteins of the Krebs cycle. This mitochondrial plasticity reflects the ability of the yeast to react to an endogenous metabolic stress of mitochondrial origin. In this context, we wondered whether the mitoproteome adaptation is specific to a general dysfunction of the oxidative phosphorylation process or is tailor-made depending on the affected respiratory function. Indeed, although AOX and UCP both lead to a decrease in ATP synthesis yield, they work on the respiratory chain at two different levels: the redox potential of the respiratory chain for AOX and the proton electrochemical gradient for UCP. As a general adaptation to the introduction of an energydissipating system, the Krebs cycle and the complex III are


increased. It is noteworthy that complex III is more likely the rate-limiting step of the yeast respiratory chain since it is the only respiratory enzyme affected in both conditions, i.e., UCP+ 28 and AOX (this work). These adaptations occur in order to increase the electron flux in the respiratory chain in order to counter the energy-dissipating action of UCP1 or AOX. Moreover, an increased content of complex III that is the direct competitor of AOX for the reduced Q would promote the electron partitionning in favor of the cytochrome pathway and consequently in favour of energy conservation. However, a striking result was the ∼2-fold decrease in ATP synthase when AOX was ectopically expressed (Table 1) while UCP1 induced a ∼2-fold increase in ATP synthase.28 This reflects the formidable plasticity of the yeast that can distinguish between mitochondrial endogenous stress of different origins and adequatly adapt its mitoproteome in return. Moreover, this phenomenon would be the result of a complex crosstalk between mitochondria and the nucleus, which seems to require several signaling elements sensing different bioenergetics parameters such as the redox state of the respiratory chain, the ATP/ADP status or the free radical generation. Finally, we provide strong evidence of a functional linkage existing between AOX and the Krebs cycle. Indeed, five enzymes of the Krebs cycle were up-regulated in response to the introduction of AOX. This observation sustains the hypothesis of the energy-overflow that connects the AOX activity to the control of the energy balance. Indeed, in our gain-of-function study, introducing AOX results in a metabolic imbalance that is unefficiently compensated through proteomic adaptations notably at the level of the Krebs cycle. In this context, AOX has a direct effect on the reducing substrate availability and may have a regulatory role in energy balance. Abbreviations: 2D-DIGE, two-dimensional differential ingel electrophoresis; AOX, alternative oxidase; BHAM, benzohydroxamic acid; GMP, guanosine 5′-monophosphate; PN, purine nucleotide; Q, coenzyme Q; UCP, uncoupling protein; WT, wild-type.

Acknowledgment. We first thank Dr. Sakajo for the kind gift of the HaAOX cDNA from Hansenula anomala. This work was supported by grants from the Fonds National de la Recherche Scientifique (FRFC 2.4532.03, FRSM 9.4573.04) and from the Fonds Spećiaux de Recherche dans les universite´s. It was also co-financed by the Centre d′Analyse des Re´sidus en Traces (CART), the Re´gion Wallonne (i-Maldi 114713), Fonds Social Europeén (FSE) and the Centre of Biomedical Integrative Genoproteomics (CBIG). P.D. is recipient of a Fonds pour la Recherche Industrielle et Agronomique fellowship. P.L. is a Research Associate of the FNRS. References (1) Jarmuszkiewicz, W.; Sluse-Goffart, C. M.; Hryniewiecka, L.; Michejda, J.; Sluse, F. E. Electron partitioning between the two branching quinol-oxidizing pathways in Acanthamoeba castellanii mitochondria during steady-state 3 respiration. J. Biol. Chem. 1998, 273, 10174-10180. (2) Jarmuszkiewicz, W.; Sluse-Goffart, C. M.; Hryniewiecka, L.; Sluse, F. E. Identification and characterization of a protozoan uncoupling protein in Acanthamoeba castellanii. J. Biol. Chem. 1999, 274, 23198-23202. (3) Jarmuszkiewicz, W.; Milani, G.; Fortes, F.; Schreiber, A. Z.; Sluse, F. E.; Vercesi, A. E. First evidence and characterization of an uncoupling protein in fungi kingdom: CpUCP of Candida parapsilosis. FEBS lett. 2000, 467, 145-149.

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