Abstract

In yeast, mammals, and land plants, mitochondrial F1FO-ATP synthase (complex V) is a remarkable enzymatic machinery that comprises about 15 conserved subunits. Peculiar among eukaryotes, complex V from Chlamydomonadales algae (order of chlorophycean class) has an atypical subunit composition of its peripheral stator and dimerization module, with nine subunits of unknown evolutionary origin (Asa subunits). In vitro, this enzyme exhibits an increased stability of its dimeric form, and in vivo, Chlamydomonas reinhardtii cells are insensitive to oligomycins, which are potent inhibitors of proton translocation through the FO moiety.

In this work, we showed that the atypical features of the Chlamydomonadales complex V enzyme are shared by the other chlorophycean orders. By biochemical and in silico analyses, we detected several atypical Asa subunits in Scenedesmus obliquus (Sphaeropleales) and Chlorococcum ellipsoideum (Chlorococcales). In contrast, complex V has a canonical subunit composition in other classes of Chlorophytes (Trebouxiophyceae, Prasinophyceae, and Ulvophyceae) as well as in Streptophytes (land plants), and in Rhodophytes (red algae). Growth, respiration, and ATP levels in Chlorophyceae were also barely affected by oligomycin concentrations that affect representatives of the other classes of Chlorophytes. We finally studied the function of the Asa7 atypical subunit by using RNA interference in C. reinhardtii. Although the loss of Asa7 subunit has no impact on cell bioenergetics or mitochondrial structures, it destabilizes in vitro the enzyme dimeric form and renders growth, respiration, and ATP level sensitive to oligomycins.

Altogether, our results suggest that the loss of canonical components of the complex V stator happened at the root of chlorophycean lineage and was accompanied by the recruitment of novel polypeptides. Such a massive modification of complex V stator features might have conferred novel properties, including the stabilization of the enzyme dimeric form and the shielding of the proton channel. In these respects, we discuss an evolutionary scenario for F1FO-ATP synthase in the whole green lineage (i.e., Chlorophyta and Streptophyta).

Introduction

The F1FO-ATP synthase is a ubiquitous rotary motor enzyme that couples proton flow through its membrane-embedded FO channel to ATP synthesis that occurs on its F1 moiety (Boyer 2000). In fungi, mammals, and flowering plants, mitochondrial ATP synthase is composed of at least 14–15 conserved subunits of dual genetic origin: Up to five subunits are usually encoded by the mitochondrial genomes, whereas the remainder are nuclear gene products. Altogether, they build the F1 catalytic domain, the FO proton pore, and two stalks that link and hold F1 to FO. One of these stalks is thought to act as a rotor and the other as a peripheral stator (Weber and Senior 2003; Cardol et al. 2005; Vázquez-Acevedo et al. 2006; Wittig and Schagger 2008).

In contrast, biochemical and computational analyses revealed that the enzyme from three chlorophycean algae belonging to the Chlamydomonadales order (Chlamydomonas reinhardtii, Polytomella sp. Pringsheim 198.80, and Volvox carteri) lacks eight subunits (b, d, e, f, g, h, F6, and IF1) that are conserved in mammals and fungi and participate in the building of the peripheral stalk and in the dimerization of the enzyme. Instead, the algal enzyme contains nine nucleus-encoded subunits of unknown evolutionary origin, which were named Asa1 to 9 for “ATP Synthase-Associated” proteins (Cardol et al. 2005; Vázquez-Acevedo et al. 2006; Van Lis et al. 2007). It was thus hypothesized that Asa subunits build a novel peripheral stator and dimerization module architecture. Indeed, electron microscopy studies revealed that the structures of the ATP synthase dimeric forms of beef heart (Minauro-Sanmiguel et al. 2005) and Polytomella (Dudkina et al. 2005) differ, the latter exhibiting two large, robust, protruding arms that extend from the membrane to the upper region of the F1 moieties. Moreover, in contrast to other known F1FO-ATP synthases, the dimeric complex V of chlorophycean algae is highly stable in vitro (Van Lis et al. 2003, 2007; Vázquez-Acevedo et al. 2006; Villavicencio-Queijeiro et al. 2009). However, because of the lack of information on ATP synthase subunit composition in other green organisms (i.e., between flowering plants and Chlamydomonadales), the question arose to know whether these Asa subunits were genuinely atypical components rather than highly divergent homologs of classical complex V proteins. In this work, we first investigated the subunit composition of mitochondrial ATP synthase in the green photosynthetic organisms, with emphasis on the green algal phylum (i.e., Chlorophyta). Our data allowed us to propose an evolutionary scenario for F1FO-ATP synthase diversification. To uncover novel specific properties conferred by the atypical subunits in Chlorophyceae, we investigated the role of the 19.5-kDa subunit (Asa7) by inactivating its gene expression in C. reinhardtii. We found that the silencing of ASA7 destabilizes the dimeric enzyme complex in vitro and renders cell growth, respiration, and ATP level sensitive to oligomycins.

Materials and Methods

Strain and Growth Conditions

The C. reinhardtii strain used in this study is the cw15 arg7-8 mt+ mutant. This strain lacks a cell wall and is auxotroph for arginine because of a mutation in the ARG7 gene coding for argininosuccinate lyase (Debuchy et al. 1989). The other Chlorophyta used in this work originated from axenic cultures available at the University of Göttingen (Sammlung von Algenkulturen [SAG], Germany): Chlorococcum ellipsoideum (63.80), Uronema acuminata (33.86), Chlamydomonas moewusii (21.90), Chlorogonium elongatum (12–2b), Scenedesmus obliquus (276–3b), Chlorella sorokinia (211-31), Chlorella vulgaris (2II-11b), Nannochloris sp. (251-2), Coccomyxa pringsheimii (216.7), Leptospira obovata (445-1), Pseudendoclonium basiliense (466-1), Gloeotilopsis paucicellulare (463.1), Ulothrix Fimbriata (36.86), Micromonas pusilla (39.85), and Tetraselmis chuii (8-6).

Cells were routinely grown in liquid or on solid agar medium under moderate light (50-μmol photon m−2 s−1) at 25 °C. Tris-minimal-phosphate medium (TMP) supplemented or not with acetate (TAP, 5 or 17 mM) was used for cultivating the algae (Harris 1989), except for T. chuii and M. pusilla that were grown on solid Marine medium (Difco Marine Broth 2216, BD, United States). Biomass calculated as the product of the cell density by the mean volume of cells is proportional to the turbidity (A750nm). Both parameters were determined using a Coulter counter (Coulter electronics, Harpenden Herts, United Kingdom).

Construction of Plasmid pASA7-RNAi (4.08 kbp)

Escherichia coli DH5α was used for cloning, and E. coli transformants were grown in LB medium in the presence of ampicillin (50 μg ml−1) at 37 °C. The pNB1 plasmid (2,895 bp) was used to express double-stranded RNA (dsRNA). A NIA1/TUB1 promoter was inserted in the XbaI and HindIII sites of the pUC19 vector (Cardol et al. 2006). An ASA7 cDNA (287 bp) and the corresponding genomic (934-bp) fragment were amplified by polymerase chain reaction (PCR) using as forward primer ASA7-RNAi-1F (5′-AGCTTAGCACCCTAGTCGAA-3′) and as reverse primers ASA7-RNAi-2R (5′-CGTCAGTGTCAGCAGGTAGT-3′) and ASA7-RNAi-3R (5′-GCGCGGTAGTAGTAATCCTT-3′), respectively. The oligonucleotides contained ClaI/HindIII (forward) or HindIII/NcoI (reverse) restriction sites at their 5′ ends for further constructions. These PCR fragments were cloned into pGEM-T Easy Vector (Promega) to obtain pASA7-13 (ASA7-RNAi-1F/ASA7-RNAi-2R cDNA) and pASA7-3 (ASA7-RNAi-1F/ASA7-RNAi-3R genomic), respectively. The excised HindIII fragment of pASA7-13 was inserted into the pNB1 plasmid, and the construct with inverse orientation of ASA7-RNAi-1F/ASA7-RNAi-2R fragment was selected by a PCR analysis to obtain pASA7-AS. The ClaI–NcoI fragment of pASA7-3 was then inserted into the ClaI–NcoI site of pASA7-AS, giving the plasmid pASA7-RNAi (where RNAi is RNA interference), used for RNA inactivation of ASA7.

Transformation of C. reinhardtii

Transformation of the C. reinhardtii cw15 arg7-8 mt+ strain was carried out using the glass-bead method (Kindle 1990) with 4 μg of plasmid pRNAi (linearized with SacI) and 1 μg of pASL, linearized with BamHI. This pASL plasmid bears the Chlamydomonas ARG7 gene encoding for the argininosuccinate lyase (Debuchy et al. 1989) and is used as a selectable marker. Prototroph transformants were selected on TAP agar plates. The presence of sequences belonging to the right and to the left part of the RNAi plasmids in the transformants was checked by PCR with primers hybridizing with the ASA7 sequences and the vector (universal primers 5′-GTAAAACGACGGCCAG-3′ and 5′-CAGGAAACAGCTATGAC-3′) on a total nucleic acid extract prepared according to standard procedures (Newman et al. 1990) directly on algal colonies as described in Remacle et al. (2006). The stability of the phenotype observed for the transformants mentioned in this study was confirmed 2 years after their original isolation.

RNA Analyses

Total RNA (15 μg) prepared according to Newman et al. (1990) was separated on 0.8% agarose–formaldehyde gels and transferred onto Hybond-N membrane (Amersham Pharmacia Biotech). Digoxigenin-labeled PCR products of cDNA fragments were used as gene probes and detected with antidigoxigenin-AP conjugates and CDP-Star as substrate (Roche, Basel, Switzerland). Hybridization and washing steps were performed according to standard protocols. ASA7-RNAi-1F/ASA7-RNAi-2R and ATP2-RNAi-1F (GTGGATGTGCGTTTCG)/ATP2-RNAi-2R (CCGGTCACCAGGATCT) primers were used to synthesize the probe for detection of ASA7 and ATP2 transcripts, respectively.

Protein Analyses

Chlamydomonasreinhardtii, S. obliquus, and C. ellipsoideum crude total membrane fractions were obtained according to Remacle et al. (2001). For S. obliquus, a nebulizer (BioNeb, Cell disruption System, Glas-Col) was used for cell disruption prior to sonication. Crude mitochondrial fraction was obtained according to Cardol et al. (2002) and loaded onto a discontinuous Percoll gradient (13%/21%/45% in mannitol-EDTA-Tris (MET) Buffer [280 mM mannitol, 10 mM Tris–HCl pH 7, 0.5 mM ethylenediaminetetraacetic acid (EDTA), and 0.1% BSA]). Purified mitochondria were recovered at the 21%/45% interface and washed twice in MET buffer by a 10-min centrifugation at 11,000 × g. The final pellet was resuspended in 20-mM 4-(2-Hydroxyethyl)-1-piperazineethane sulfonic acid (HEPES-KOH) (pH 7.2), 150-mM mannitol, and 4-mM MgCl2. The protein content was determined by the Bradford method (Bradford 1976). To conduct Blue native polyacrylamide gel electrophoresis (BN-PAGE) analyses (Schägger and Von Jagow 1991), protein complexes were first solubilized in the presence of either N-dodecyl-β-D-maltoside or Triton X-100, 375 mM 6-aminohexanoic acid, 250 mM EDTA, and 25 mM Bis–Tris, pH 7.0, and centrifuged for 20 min at 15,000 × g at 4 °C to remove insoluble matter. 0.4% (w/v) sodium taurodeoxycholate was then added to the supernatant prior to separation by electrophoresis on a 4–12% polyacrylamide gradient BN gel. ATP synthase activity was detected by incubating the gel in 50 mM HEPES, pH 8.0, containing 10 mM ATP and 30 mM CaCl2. Coomassie blue staining and the second dimensional Tricine–sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) procedure were performed as described in Cardol et al. (2004). The molecular size of the proteins was calculated by comparison with known markers (PageRulerPrestained Protein Ladder Plus, Fermentas, Ontario, Canada). For Western blot analysis, protein extracts were loaded onto 10% SDS gels and electroblotted according to standard protocols onto polyvinylidene fluoride membranes (Amersham GE Healthcare). Detection was performed using a BM Chemiluminescence Western blotting kit (Roche, Basel, Switzerland) with antirabbit peroxidase–conjugated antibodies. We used rabbit sera obtained against Polytomella sp. Pringsheim 198.80 Atp2 (1:200,000) or C. reinhardtii Asa7 (1:50,000) (Genscript, Piscataway, NJ).

ATP Determination

ATP was extracted according to Gans and Rebeille (1990). ATP cellular level was determined using the Enliten luciferase/luciferin kit (Promega, Madison, WI).

Oxygen Evolution

Cells grown mixotrophically in TAP liquid medium were sampled during the exponential phase. Dark respiration rates were measured using a Clark Electrode (Hansatech Instruments, King′s Lynn, United Kingdom) as previously described (Duby and Matagne 1999). The cytochrome pathway and the alternative pathway of respiration were inhibited by addition of 1-mM potassium cyanide in aqueous solution and 1-mM salicylhydroxamic acid (SHAM) in ethanol (final concentration 1%), respectively. The possible inhibitory effect of ethanol alone was subtracted from the measurements. The apparent capacity of each pathway corresponds to the following respiratory rates: For the cytochrome pathway, the oxygen consumption inhibited by KCN after addition of SHAM; for the alternative pathway, the oxygen consumption inhibited by SHAM after addition of KCN.

In Silico Analyses

Multiple-sequence alignments of polypeptides were performed with MUSCLE program available at http://www.ebi.ac.uk/Tools/muscle/index.html (Edgar 2004). The tree shown in figure 2 was conservatively assembled from recent phylogenetic studies of the green lineage (Lewis and McCourt 2004; Müller et al. 2004; Pombert et al. 2004, 2005; Rodriguez-Ezpeleta et al. 2007). Gene gain and losses for mitochondrial ATP synthase subunits were then mapped onto the tree from presence–absence data derived from table 1 using unweighted Dollo parsimony as implemented in DOLMOVE (PHYLIP package; Felsenstein J, 2005. PHYLIP [phylogeny inference package] version 3.6. Distributed by the author. Department of Genome Sciences, University of Washington, Seattle, WA). As DOLMOVE cannot handle polytomies, nine variants of the tree were successively considered to account for uncertain relationships within Chlorophyceae and among classes of Chlorophytes, though yielding highly similar mapping. Basic alignment search tool (Blast) searches (Altschul et al. 1997) were carried out on the NCBI portal (Johnson et al. 2008) using sensitive parameters (e.g., BLOSUM45 matrix, smaller word size, and masking of low complexity regions for look-up only). Both PSI-BlastP/BlastX searches against the nonredudant protein (nr) database and TBlastN/TBlastX searches against nonhuman/nonmouse expressed sequence tags (ESTs) (est_others) were conducted.

Table 1.

Subunit Composition of Mitochondrial F1FO-ATP Synthase in Eukaryotes.

    Streptophyta
 
Chlorophyta
 
Name Subunit Metazoa Fungi Rodophyta Tracheophyta Marchantiophyta Bryophyta Charophyceae Prasinophyceae
 
Trebouxiophyceae
 
Ulvophyceae
 
Chlorophyceae
 
B.t. S.c. C.m. A.t. Ma.p. P.p. C.v. O.t. Mi.p P.w. C.v. P.a. U.l. S.o. C.e. C.r. V.c. 
α (ATP1) nu nu nu mt mt mt mt mt nu mt  mt   nu nu 
β (ATP2) nu nu nu nu nu nu  nu nu nu nu  nu nu nu nu 
γ (ATP3) nu nu nu nu nu nu  nu nu nu nu  nu nu  nu nu 
δ (ATP16) nu nu nu nu nu nu  nu nu nu nu     nu nu 
ε (ATP15) nu nu nu nu nu nu  nu nu nu nu     nu nu 
OSCP (ATP5) nu nu nu nu nu nu  nu nu  nu    nu nu 
A6L (ATP8) mt mt mt mt mt mt mt mt mt mt  mt    — — 
F6 nu nu — —  —  — —  —     — — 
IF1 nu nu — nu nu nu  — —  —     — — 
a (ATP6) mt mt mt mt mt mt mt mt mt mt  mt  mt nu nu 
b (ATP4) nu nu nu mt mt mt mt mt mt mt  mt    — — 
c (ATP9) nu nu nu mt mt mt mt nu mt mt  mt  mt  nu nu 
d nu nu nu nu nu nu  nu nu nu nu     — — 
e nu nu — — — —  — —  —     — — 
f nu nu nu nu nu nu  nu nu nu nu     — — 
g nu nu nu nu nu nu  nu nu  nu     — — 
Asa4/FAd (ATP7) — — — nu nu nu  nu nu  nu   nu  nu nu 
Asa1 — — — — — —  — —  —    nu nu 
Asa2 — — — — — —  — —  —    nu nu 
Asa3 — — — — — —  — —  —     nu nu 
Asa5 — — — — — —  — —  —   nu  nu nu 
Asa6 — — — — — —  — —  —     nu nu 
Asa7 — — — — — —  — —  —   nu  nu nu 
Asa8 — — — — — —  — —  —     nu nu 
Asa9 — — — — — —  — —  —   nu  nu nu 
stf1 — nu — — — —  — —  —     — — 
Stf2 — nu — — — —  — —  —     — — 
— nu — — — —  — —  —     — — 
— nu — — — —  — —  —     — — 
nu — — — — —  — —  —     — — 
    Streptophyta
 
Chlorophyta
 
Name Subunit Metazoa Fungi Rodophyta Tracheophyta Marchantiophyta Bryophyta Charophyceae Prasinophyceae
 
Trebouxiophyceae
 
Ulvophyceae
 
Chlorophyceae
 
B.t. S.c. C.m. A.t. Ma.p. P.p. C.v. O.t. Mi.p P.w. C.v. P.a. U.l. S.o. C.e. C.r. V.c. 
α (ATP1) nu nu nu mt mt mt mt mt nu mt  mt   nu nu 
β (ATP2) nu nu nu nu nu nu  nu nu nu nu  nu nu nu nu 
γ (ATP3) nu nu nu nu nu nu  nu nu nu nu  nu nu  nu nu 
δ (ATP16) nu nu nu nu nu nu  nu nu nu nu     nu nu 
ε (ATP15) nu nu nu nu nu nu  nu nu nu nu     nu nu 
OSCP (ATP5) nu nu nu nu nu nu  nu nu  nu    nu nu 
A6L (ATP8) mt mt mt mt mt mt mt mt mt mt  mt    — — 
F6 nu nu — —  —  — —  —     — — 
IF1 nu nu — nu nu nu  — —  —     — — 
a (ATP6) mt mt mt mt mt mt mt mt mt mt  mt  mt nu nu 
b (ATP4) nu nu nu mt mt mt mt mt mt mt  mt    — — 
c (ATP9) nu nu nu mt mt mt mt nu mt mt  mt  mt  nu nu 
d nu nu nu nu nu nu  nu nu nu nu     — — 
e nu nu — — — —  — —  —     — — 
f nu nu nu nu nu nu  nu nu nu nu     — — 
g nu nu nu nu nu nu  nu nu  nu     — — 
Asa4/FAd (ATP7) — — — nu nu nu  nu nu  nu   nu  nu nu 
Asa1 — — — — — —  — —  —    nu nu 
Asa2 — — — — — —  — —  —    nu nu 
Asa3 — — — — — —  — —  —     nu nu 
Asa5 — — — — — —  — —  —   nu  nu nu 
Asa6 — — — — — —  — —  —     nu nu 
Asa7 — — — — — —  — —  —   nu  nu nu 
Asa8 — — — — — —  — —  —     nu nu 
Asa9 — — — — — —  — —  —   nu  nu nu 
stf1 — nu — — — —  — —  —     — — 
Stf2 — nu — — — —  — —  —     — — 
— nu — — — —  — —  —     — — 
— nu — — — —  — —  —     — — 
nu — — — — —  — —  —     — — 

NOTE.—Subunits present in the indicated organisms are marked as (mt) if mitochondria encoded or as (nu) if nucleus encoded. (—), no homolog could be identified in the nuclear or mitochondrial genomes. Blank spaces indicate that we cannot yet decide for the presence or for the absence of a particular subunit due to the lack of biochemical or molecular data. Subunits marked in italics are conserved in all mitochondrial F1FO-ATP synthases. Subunits marked in bold are conserved in all eukaryotes except in Chlorophyceae. Abbreviations: B.t., Bos taurus; S.c., Saccharomyces cerevisiae; C.m. Cyanidioschyzon merolae; A.t., Arabidopsis thaliana; Ma.p., Marchantia polymorpha; P.p., Physcomitrella patens; C.v. Chara vulgaris; O.t., Ostreococcus tauri; Mi.p., Micromonas pusilla; P.w. Prototheca wickerhamii; C.v., Chlorella vulgaris; P.a., Pseudendoclonium akinetum; U.l. Ulva linza; S.o., Scenedesmus obliquus; C.e. Chlorococcum ellipsoideum; C.r., Chlamydomonas reinhardtii; V.c. Volvox carteri; OSCP, Oligomycin Sensitivity-Conferring Protein. See supplementary table 1, Supplementary Material online, for accession numbers.

Transmission Electron Microscopy

Chlamydomonasreinhardtii cells were fixed with 2.5% glutaraldehyde in phosphate buffered saline (PBS) (pH 7.2) for 2 h at 4 °C and washed three times with PBS by centrifugation in a table-top centrifuge. The algal cells were postfixed with 1% osmium tetroxide. Dehydration was carried out at room temperature in a graded series of ethanol at a concentration from 40% to 100% (v/v) in 10% increments. Then, samples were placed two times for 15 min each in propylene oxide. Pre-embedding in 1:1 propylene oxide–epoxy resin was conducted overnight. Thin sections (50–60 nm thick) were cut with an ultramicrotome (Leica Ultracut R) and placed onto formvar-coated copper grids. Grids were contrasted with uranyl acetate and lead citrate and examined under a JEOL 1200 EX II transmission electron microscope operating at 60 or 70 kV.

Mass Spectrometry Analyses

Coomassie blue–stained proteins associated with spots or bands of interest were manually excised. Gels plugs were transferred together with 200-μl high performance liquid chromatography water into 0.5-ml polypropylene Protein LoBind Eppendorf tubes. Water was replaced by 200 μl 50 mM ammonium carbonate (pH 8.0) in 50% acetonitrile. After incubation for 5 min at 20 °C under shaking, the solution was replaced by 200 μl 10% acetonitrile. After incubation for 5 min, acetonitrile was removed, and gels plugs were dried under vacuum (Savant Speed Vac Concentrator). Gel plugs were rehydrated in 20 μl of a digestion buffer containing 50 mM ammonium carbonate (pH 8.0) and 0.5 μg of trypsin. Proteolysis was performed for 16 h at 37 °C and stopped by adding 10 ml 1% (v/v) trifluoroacetic acid (TFA). Supernatants of each tube were transferred into new tubes. Fifty microliters of peptide extraction solution containing 50% (v/v) acetonitrile and 0.1% (v/v) TFA were added to the gel plugs. After incubation for 5 min, extracts were combined with the first ones. A second extraction with 50 μl of 100% acetonitrile was performed, and after 5 min, the extract was combined with the previous two ones and dried under vacuum (Savant Speed Vac Concentrator). Peptides were solubilized in 20 μl 0.1% TFA, desalted, and concentrated using a C18 ZipTip (Millipore, Billerica, MA) according to the manufacturer protocol. Two microliters of 10 mg ml−1 of alpha-cyano-4-hydroxycinnamic acid (alpha-cyano matrix-assisted laser desorption/ionization [MALDI] matrix) in 50% (v/v) acetonitrile and 0.1% (v/v) TFA were mixed with 2 μl of each ZipTip concentrated peptide solution. From this, 0.5 μl was layered on an Opti-TOF 384 Well Insert MALDI plate (Applied Biosystems). MS and MS/MS spectra were acquired using an Applied Bisosystems 4800 MALDI time of flight (TOF/TOF) Analyzer spectrometer with a 200-Hz solid-state laser operating at a wavelength of 355 nm. MS spectra were obtained using 3,200 and 2,000 laser shots per spot in a range of m/z between 800 and 4,000. MS/MS spectra were obtained by selecting the 15 most intense precursor ions per spot and using 3,800 and 2,100 laser shots per precursor. The automatically selected precursors were submitted to a collision energy of 1 kV with collision gas (air) at a pressure of about 1 × 106 Torr. Data were collected with the Applied Biosystems 4000 Series Explorer software.

MS and MS/MS queries were performed using the Applied Biosystems GPS Explorer 3.6 software together with the Matrix Science Ltd MASCOT Database search engine v2.1 using nonredudant protein (nr) database from NCBI. Precursor tolerance of 150 ppm for MS spectra and 0.1-Da fragment tolerance for MS/MS spectra were allowed. A charge state of +1 was selected. A single trypsin miscleavage and variable modifications consisting of methionine oxidation and acrylamide-modified cystein were allowed. For protein direct identification with MASCOT, protein scores greater than 68 were considered as significant (P < 0.05). The Applied Biosystems GPS Explorer— DeNovo Explorer Version 3.6 software was used for identifying proteins that had not been previously characterized or are not contained in protein databases. MS/MS data were submitted to the DeNovo Explorer software to generate amino acid sequences, using mass differences between peaks. The obtained sequences were manually verified and submitted to UniProt database using the FASTS program (EMBL-EBI). Proteins matching two or more peptides were taken into consideration.

Results

Within Plantae, the Presence of Asa Subunits Is Limited to Chlorophycean Algae

To obtain evidence of the presence of Asa subunits in chlorophycean species that do not belong to the order of Chlamydomonadales, we first analyzed a crude membrane preparation of C. ellipsoideum cells (order of Chlorococcales). Membrane proteins solubilized by addition of n-dodecyl-maltoside or Triton X-100 were separated by BN-PAGE. This technique allows the separation of mitochondrial complexes in their native form (Schägger and Von Jagow 1991). A subsequent in-gel detection of ATPase activity (leading to a white calcium phosphate precipitate) indicated that complex V from C. ellipsoideum migrates at approximately 1,700 kDa (fig. 1a). This observation is consistent with previous works in which it was shown that complex V from C. ellipsoideum, C. reinhardtii and other chlorophycean species exhibits the same electrophoretic mobility (∼1,600–1,700 kDa) and migrates under dimeric form (V2) (Vázquez-Acevedo et al. 2006; Villavicencio-Queijeiro et al. 2009). An additional band bearing ATPase activity could be visualized at about 300 kDa. It might correspond to either a mitochondrial or a chloroplastic F1 moiety. The band corresponding to the dimeric complex V was excised from the BN gel and its constitutive subunits were then separated by denaturating SDS-PAGE. A minimum of 10 bands of molecular mass ranging from 65 to 8 kDa were observed after Coomassie blue staining (fig. 1a, lane 4). Mass spectrometry analysis (MS combined with MS/MS MALDI TOF/TOF) allowed the identification of bands 2, 3, 5, and 6 as mitochondrial subunits β (ATP2), α (ATP1), a (ATP6), and oligomycin sensitivity-conferring protein (oligomycin sensitivity-conferring protein [OSCP], ATP5), respectively. Although peptide MS/MS spectra were obtained for the other bands, they did not match any known ATP synthase subunits. De novo sequencing from the MS/MS spectra was then undertaken, and this allowed us to identify bands 1 and 4 as homologs to atypical subunits Asa1 and Asa2 of C. reinhardtii (fig. 1b).

FIG. 1.

Identification of Asa proteins in chlorophycean algae Chlorococcum ellipsoideum. (A) Hundred-microgram proteins of a crude membrane fraction from C. ellipsoideum cells were solubilized in the presence of 10% (w/w) Triton X-100 (lanes 1 and 3) or 20% (w/w) n-dodecyl-β-maltoside (lane 2) and analyzed by BN-PAGE. After electrophoresis, the gel was submitted to Coomassie blue staining (lanes 1 and 2) or to ATPase activity staining (lane 3; negative picture). The subunits of dimeric complex V (V2) (obtained as in lane 2 but from 500 μg proteins of crude membrane fractions) were resolved in a 10% acrylamide 2D-Tricine–SDS-PAGE and stained with Coomassie blue (lane 4). n.i., not identified. (B) Alignment between the C. ellipsoideum (Chlel) sequences obtained by MS de novo sequencing (see Materials and Methods for details) and the homologous sequences of Chlamydomonas reinhardtii (Chlre) Asa1 and Asa2. Conserved residues are shaded in black; similar residues are shaded in gray. Positions of C. reinhardtii residues are indicated along the alignment.

FIG. 1.

Identification of Asa proteins in chlorophycean algae Chlorococcum ellipsoideum. (A) Hundred-microgram proteins of a crude membrane fraction from C. ellipsoideum cells were solubilized in the presence of 10% (w/w) Triton X-100 (lanes 1 and 3) or 20% (w/w) n-dodecyl-β-maltoside (lane 2) and analyzed by BN-PAGE. After electrophoresis, the gel was submitted to Coomassie blue staining (lanes 1 and 2) or to ATPase activity staining (lane 3; negative picture). The subunits of dimeric complex V (V2) (obtained as in lane 2 but from 500 μg proteins of crude membrane fractions) were resolved in a 10% acrylamide 2D-Tricine–SDS-PAGE and stained with Coomassie blue (lane 4). n.i., not identified. (B) Alignment between the C. ellipsoideum (Chlel) sequences obtained by MS de novo sequencing (see Materials and Methods for details) and the homologous sequences of Chlamydomonas reinhardtii (Chlre) Asa1 and Asa2. Conserved residues are shaded in black; similar residues are shaded in gray. Positions of C. reinhardtii residues are indicated along the alignment.

We undertook the same approach with S.obliquus (Sphaeropleales order) and found that complex V behaves as a 1,600-kDa dimer in BN-PAGE (data not shown), as previously observed (Vázquez-Acevedo et al. 2006). We however failed to obtain enough material to perform SDS-PAGE and subsequent MS analysis. As an alternative approach, we investigated the nucleic sequences available at NCBI for S. obliquus. In addition to subunits a and c encoded in the mitochondrial genome of S. obliquus, we identified putative homologous sequences for subunits β,γ, Asa4, Asa5, Asa7, and Asa9 in the EST data set (∼6,600 ESTs) (table 1 and supplementary table S1, Supplementary Material online). Altogether, the detection of ASA genes in species belonging to three different chlorophycean orders (ASA1–9 in Chlamydomonadales; ASA1–2 in Chlorococcales; and ASA4–5, ASA7, and ASA9 in Sphaeropleales) led us to explore other species.

The green algae are divided into the phyla Streptophyta and Chlorophyta. Streptophyta contains land plants and green algae belonging to the class Charophyceae, whereas Chlorophyta contains the members of the classes Chlorophyceae, Trebouxiophyceae, Prasinophyceae, and Ulvophyceae (e.g., Lewis and McCourt 2004). To investigate complex V subunit composition in these organisms, we took advantage of the availability of nuclear genome sequences of Chlorella vulgaris C-169 (Trebouxiophyceae, available at http://www.jgi.doe/gov/), Ostreococcus tauri (Derelle et al. 2006), M. pusilla (Worden et al. 2009) (Prasinophyceae), and Physcomitrella patens (Bryophyte). We also explored the EST data sets available at NCBI for Prothoteca wickerhamii (Trebouxyophyceae, ∼6,000 ESTs), Ulva linza (Ulvophyceae, ∼2,000 ESTs), and Marchantia polymorpha (Marchantiophyte, ∼34,000 ESTs). We finally took benefit of the availability of mitochondrial genome sequences of Pseudendoclonium akinetum (Ulvophyceae), P. wickerhamii, M. pusilla, O. tauri, Chara vulgaris (Charophyceae), P. patens and M. polymorpha. Looking for genes encoding classical or atypical subunits of mitochondrial ATP synthase, we identified coding sequences of α, β, γ, a, b, c, and A6L subunits in Ulvophyceae. A larger gene set coding for classical subunits α, β, γ, δ, ϵ, a, c, OSCP, b, d, f, g, and A6L was found in mosses and liverworts, Prasinophyceae, Trebouxiophyceae (table 1 and see also supplementary table 1, Supplementary Material online, for accession numbers). In these two latter algal classes, we also identified a protein that appears to be a distant homolog of both subunit FAd (ATP7) from Arabidopsis thaliana complex V (Heazlewood et al. 2003) and subunit Asa4 of Chlamydomonadales (see supplementary fig. 1, Supplementary Material online, for details). In contrast, we did not identify any sequence sharing similarities with Chlamydomonadales Asa1-3 or Asa5-9 polypeptides (see also below), with classical subunits e, IF1 and F6, or with subunits present in mammals and fungi. This deduced subunit composition is rather similar to that found in A. thaliana (Heazlewood et al. 2003; Cardol et al. 2005) and to that that we could deduce from mining nucleic sequence database (http://merolae.biol.s.u-tokyo.ac.jp/) of the red alga Cyanidioschyzon merolae.

As to date, Asa subunits are still of unknown evolutionary origin (Vázquez-Acevedo et al. 2006; Van Lis et al. 2007), we performed sensitive database searches aiming at identifying potential homologs in other organisms (i.e., all eukaryotes and prokaryotes). For Asa1, Asa5–7, and Asa9 subunits, we did not observe any significant hit (E ≤ 10−3) beyond chlorophycean algae. In contrast, a very significant hit (GD169774; E = 10−32) was obtained for Asa8 with an EST from the ciliate Sterkiella (Oxytricha) histriomuscorum. When using this EST as the query of a reciprocal TBlastX search, the best match was with an EST (DV203963; E = 10−36) from Haematococcus pluvialis (Chlamydomonadales), while an additional Sterkiella EST (GD170269; E = 10−25) was also identified. Because these searches did not yield any nonchlorophycean organism other than this ciliate, which is known to preys on algae, we interpret these two ESTs as probable Chlamydomonadales contaminations of Sterkiella libraries. In the cases of Asa2 and Asa3, a few weak hits (E > 10−11) were observed with a flavoprotein monooxygenase found in Prasinophyceae (e.g., CAL55851) as well as with a series of oxygenase-related ESTs from Dinophyceae (e.g., Karenia brevis, Karlodinium micrum, and Alexandrium catenella). Further examination of these results suggests that these matches are likely spurious as 1) both ASA2 and ASA3 proteins lack an oxygenase domain and 2) most of these hits disappear when enabling the low complexity filter of the Blast engine.

In a next step, we inferred gene gains and losses for mitochondrial ATP synthase subunits using Dollo parsimony. To this end, we conservatively assembled an evolutionary tree featuring relationships drawn from previous phylogenetic studies (Müller et al. 2004; Pombert et al. 2004; Keeling et al. 2005; Rodriguez-Ezpeleta et al. 2007). In Chlorophyta, aside from the well-accepted basal position of Prasinophyceae, the relative branching order of Chlorophyceae, Trebouxiophyceae, and Ulvophyceae is still a matter of debate, which led us to consider three different subtrees (Pombert et al. 2004, 2005; Rodriguez-Ezpeleta et al. 2007). Similarly, unresolved relationships within Chlorophyceae (e.g., Lewis and McCourt 2004) entail three variants of each subtree, thus amounting to nine possible evolutionary scenarios. Whatever the scenario examined, however, highly similar gene gains and losses were obtained (data not shown); hence, we present only the most consensual scenario in figure 2. Using E. coli as the bacterial representative, 8 subunits (α,β,γ,δ, a, b, c, and OSCP) are shared between Bacteria and eukaryotes, whereas 6 subunits (ϵ, d, f, g, IF1, A6L) were acquired deeply in the eukaryotic tree. Opisthokonts later acquired subunits e and h, and a few additional subunits were further acquired either by Fungi or along the lineage leading to mammals. In Plantae (represented here by the green lineage and the rhodophyte Cyanidioschyzon merolae), all organisms appear to share a similar set of subunits, to the notable exceptions of IF1, which has been lost independently twice, and of chlorophycean subunits. In the latter class, indeed, five classical subunits disappeared (b, d, f, g, and A6L), whereas eight new subunits (Asa) were recruited.

FIG. 2.

Phylogenetic distribution of mitochondrial ATP synthase gene gains and losses in the green lineage. Clades and species for which the subunit composition of the mitochondrial ATP synthase has been (at least partially) determined are indicated. Gene gains and losses were mapped onto the tree from presence–absence data derived from table 1 using unweighted Dollo parsimony Phylogenetic relationships were drawn from other studies (see text). Incoming and outgoing arrows from the tree represent gains and losses, respectively, of the subunits shown in the box. Numbers accompanying species names indicate the presumed number of complex V subunits. *This class may comprise several distinct lineages that warrant class-level ranking (e.g., Lewis and McCourt 2004).

FIG. 2.

Phylogenetic distribution of mitochondrial ATP synthase gene gains and losses in the green lineage. Clades and species for which the subunit composition of the mitochondrial ATP synthase has been (at least partially) determined are indicated. Gene gains and losses were mapped onto the tree from presence–absence data derived from table 1 using unweighted Dollo parsimony Phylogenetic relationships were drawn from other studies (see text). Incoming and outgoing arrows from the tree represent gains and losses, respectively, of the subunits shown in the box. Numbers accompanying species names indicate the presumed number of complex V subunits. *This class may comprise several distinct lineages that warrant class-level ranking (e.g., Lewis and McCourt 2004).

Altogether, our observations thus strengthen the proposal that atypical Asa subunits might be component characteristic of the mitochondrial ATP synthase of Chlorophyceae.

Loss of Asa7 Atypical Subunit in C. reinhardtii Leads to an Unstable Complex V

In a next step, we decided to initiate the study of Asa subunit function in Chlorophyceae. In C. reinhardtii, efficient targeted gene disruption by homologous recombination is not yet available for nuclear genes. However, interference with the expression of specific genes by dsRNA (RNAi) is a powerful method to investigate protein function in this organism (Schroda 2006; Molnar et al. 2009). To suppress the expression of ASA7 gene, a gene sequence also identified in S. obliquus, a strain of C. reinhardtii lacking the cell wall and auxotrophic for arginine was cotransformed with the plasmid pASL (bearing the ARG7 gene as a selectable marker) and the plasmid designed for RNAi (pASA7-RNAi) (see Materials and Methods section). Among 100 Arg+ prototrophic transformants selected on TAP agar plates in the light, 40 had integrated the RNAi construction, as found by PCR experiments (data not shown). Levels of ASA7 transcript and ATP2 (encoding ATP synthase β subunit) transcripts taken as a control were estimated by RNA blotting. ASA7:ATP2 transcript ratio was strongly diminished in transformants T17 (fig. 3a) and T30 (data not shown). To check the amount of Asa7 protein in T17 mutant comparatively to wild-type, crude membrane extracts were analyzed by immunoblotting using polyclonal antibodies raised against C. reinhardtii Asa7 and Polytomella sp. Pringsheim 198.80β/Atp2 subunits. Although a strong signal was obtained with Asa7 antibodies for the wild type, no signal was detected in transformant T17 (hereafter referred as ASA7-silenced strain) (fig. 3b). Conversely, no significant difference between the two strains could be detected when using the antibodies against β subunit.

FIG. 3.

Analysis of ASA7 and ATP2 gene transcripts and corresponding protein amounts and study of ATP synthase dimer stability in wild-type (WT) and ASA7-silenced transformant T17. (A) RNA blot analysis. Hybridization patterns were obtained with ATP2 and ASA7 probes on RNA (15-μg) blots from WT and ASA7-silenced transformant T17. (B) Western blot analysis. Proteins from WT and ASA7-silenced transformant T17 crude membrane fractions (15 μg per lane) were resolved by SDS-PAGE, transferred onto PVDF membranes and immunoblotted with the indicated antibodies (from top to bottom: anti-Atp2 and anti-Asa7 from Chlamydomonas reinhardtii and Polytomella sp. Pringsheim 198.80, respectively). In the case of Atp2, only the upper band corresponds to mitochondrial ATP synthase β subunit from purified complex V (Lapaille M, Remacle C, Cardol P, unpublished results). (C,D) BN-PAGE analysis. Purified mitochondria (50 μg of protein) of C. reinhardtii WT and ASA7-silenced transformant 17 (T17) strains were loaded onto a BN gel after solubilization with 10% or 40% (w/w) of n-dodecyl-β-D-maltoside as indicated on top of the lanes. After electrophoresis, the gel was stained for ATPase activity (C) or with Coomassie blue (D). V2, dimeric complex V; I, Complex I; III2, dimeric complex III; and F1, F1 moiety of complex V.

FIG. 3.

Analysis of ASA7 and ATP2 gene transcripts and corresponding protein amounts and study of ATP synthase dimer stability in wild-type (WT) and ASA7-silenced transformant T17. (A) RNA blot analysis. Hybridization patterns were obtained with ATP2 and ASA7 probes on RNA (15-μg) blots from WT and ASA7-silenced transformant T17. (B) Western blot analysis. Proteins from WT and ASA7-silenced transformant T17 crude membrane fractions (15 μg per lane) were resolved by SDS-PAGE, transferred onto PVDF membranes and immunoblotted with the indicated antibodies (from top to bottom: anti-Atp2 and anti-Asa7 from Chlamydomonas reinhardtii and Polytomella sp. Pringsheim 198.80, respectively). In the case of Atp2, only the upper band corresponds to mitochondrial ATP synthase β subunit from purified complex V (Lapaille M, Remacle C, Cardol P, unpublished results). (C,D) BN-PAGE analysis. Purified mitochondria (50 μg of protein) of C. reinhardtii WT and ASA7-silenced transformant 17 (T17) strains were loaded onto a BN gel after solubilization with 10% or 40% (w/w) of n-dodecyl-β-D-maltoside as indicated on top of the lanes. After electrophoresis, the gel was stained for ATPase activity (C) or with Coomassie blue (D). V2, dimeric complex V; I, Complex I; III2, dimeric complex III; and F1, F1 moiety of complex V.

To determine the assembly state and the ATPase activity of complex V in ASA7-silenced strain, respiratory-chain complexes from purified mitochondria were solubilized with n-dodecyl-maltoside and further separated by BN-PAGE. Both in-gel ATPase activity assay (fig. 3c) and Coomassie blue staining (fig. 3d) revealed that in wild-type control extracts, F1FO-ATP synthase is present at 1,700 kDa (dimeric form) and exhibits ATPase activity. In contrast, ATPase activity is almost exclusively observed as one or two bands of ∼400 kDa in ASA7-silenced when mitochondria are solubilized by 10% n-dodecyl-maltoside. Surprisingly, solubilization with 40% n-dodecyl-maltoside led to a moderate signal for the dimer and to the occurrence of an additional ATPase band of intermediary molecular mass (about 520 kDa). Similar observations could be made with other nonionic detergents (digitonin and Triton X-100, data not shown).

The subunit composition of wild-type dimeric complex V and mutant 400-kDa ATPase was determined by further analyzing the corresponding bands by SDS-PAGE experiments (fig. 4). It is to be noted that the 520-kDa ATPase band (fig. 3c) is almost undetectable in Coomassie blue–stained gel (data not shown). For this reason, it was not analyzed in more detail. From the wild-type dimeric enzyme, we resolved 11 bands. Their subsequent analysis by MS identified 15 of the 17 subunits belonging to the C. reinhardtii enzyme (Vázquez-Acevedo et al. 2006; Van Lis et al. 2007) (fig. 4a and supplementary table S2, Supplementary Material online). Only the small ϵ subunit and the 60-kDa Asa1 protein were missing in our analysis. From the 400-kDa doublet investigated as a unique band, three spots were resolved and unambiguously identified by MS as α, β, and γ subunits of the mitochondrial ATP synthase F1 moiety (fig. 3b and supplementary table S2, Supplementary Material online). Based on the predicted molecular mass of individual proteins (Vázquez-Acevedo et al. 2006), the theoretical molecular mass of the F1 fraction (α3β3γδϵ) was deduced to be 406.6 kDa, a value fitting very well with the estimated mass of the ATPase subcomplex found in the ASA7-silenced mutant (fig. 3) and also with the 400 kDa apparent molecular mass of the F1 fraction obtained after heat-treatment dissociation of Polytomella sp. Pringsheim 198.80 complex V (Vázquez-Acevedo et al. 2006; Van Lis et al. 2007). In this respect, the 520-kDa ATPase band shown in figure 3 should correspond to the F1 moiety associated with few other proteins. Taking into account that a small amount of the fully assembled enzyme could be observed in ASA7-silenced strain (figure 3c), one can hypothesize that Asa7 is involved in the stability of the dimeric F1FO mitochondrial ATP synthase.

FIG. 4.

Two-dimensional BN/SDS-PAGE analysis of Chlamydomonas wild-type (A) and ASA7-silenced mutant (B) mitochondria. Whole lanes from BN-PAGE (see fig. 2D) were loaded on a Tricine–SDS gel (10% acrylamide) for subsequent resolution of the protein complexes into their respective components. The main respiratory-chain complexes are indicated in the first dimension (BN-PAGE, see legend of figure 3 for details). Gels were stained with Coomassie blue. Numbers refer to supplementary table 2, Supplementary Material online.

FIG. 4.

Two-dimensional BN/SDS-PAGE analysis of Chlamydomonas wild-type (A) and ASA7-silenced mutant (B) mitochondria. Whole lanes from BN-PAGE (see fig. 2D) were loaded on a Tricine–SDS gel (10% acrylamide) for subsequent resolution of the protein complexes into their respective components. The main respiratory-chain complexes are indicated in the first dimension (BN-PAGE, see legend of figure 3 for details). Gels were stained with Coomassie blue. Numbers refer to supplementary table 2, Supplementary Material online.

To analyze the possible phenotypical consequences of Asa7 loss, we compared the growth of the ASA7-silenced mutant and wild-type strains cultivated in the light (50 μmol photons m−2 s−1) or in the dark on agar medium containing acetate (5 mM) as an exogenous carbon source. Consequences of Asa7 loss on cell fitness and bioenergetics were then investigated. The photoauto, auxo, or mixotrophic growth of wild-type control and asa7 mutant cells was then compared on agar plates. A marked phenotype was expected, especially in auxotrophic conditions where energy production, mainly relying on the sole mitochondria of Chlamydomonas, should be deeply affected by the ATPase activity of free F1 moieties. As shown in fig. 5a, the ASA7-silenced strain behaved exactly as the control strain under both conditions. Additional combinations of light intensities (0, 5, 50, or 400 μmol of photons m−2 s−1), acetate concentrations (0, 5, or 17 mM) and temperatures (16, 25, or 34 °C) were explored, but no abnormal phenotype could be observed for the mutant (data not shown). In the yeast S. cerevisiae, subunits e, g, and k are those involved in complex V dimerization (Arnold et al. 1998). Although exhibiting a normal growth phenotype, yeast mutants altered in subunits e or g exhibit abnormal mitochondrial structures, probably related to the absence of oligomeric ATP synthases (Arselin et al. 2004). To evaluate the impact of Asa7 loss on mitochondrial structure in C. reinhardtii, we investigated mitochondria morphology by electron microscopy. Mitochondria size, shape, and cristae were quite similar in both genotypes (Fig. 5B). These observations suggest that in vivo, the dimeric state of complex V in C. reinhardtii is not affected by the loss of Asa7 subunit.

FIG. 5.

Growth phenotype and mitochondrial structure of wild-type and ASA7-silenced mutant cells. Cells were cultivated in mixotrophic (light, 5 mM acetate) or heterotrophic (dark, 5 mM acetate) conditions. (A) Cell suspension were spotted at two different cell densities (upper line, A750 = 0.05; bottom line, A750 = 0.005) on solid agar plates, and growth was evaluated after 3 days in the light and 10 days in the dark. (B) Electron microscopy analysis showing the mitochondrial ultrastructure from mixotrophically grown cells. Bar, 100 nm.

FIG. 5.

Growth phenotype and mitochondrial structure of wild-type and ASA7-silenced mutant cells. Cells were cultivated in mixotrophic (light, 5 mM acetate) or heterotrophic (dark, 5 mM acetate) conditions. (A) Cell suspension were spotted at two different cell densities (upper line, A750 = 0.05; bottom line, A750 = 0.005) on solid agar plates, and growth was evaluated after 3 days in the light and 10 days in the dark. (B) Electron microscopy analysis showing the mitochondrial ultrastructure from mixotrophically grown cells. Bar, 100 nm.

Higher Sensitivity to Oligomycin in C. reinhardtii Lacking Subunit Asa7

As a next step, we investigated the effect of two respiratory inhibitors: myxothiazol, which inhibits the cytochrome bc1 complex activity, and reduces the H+ gradient formation (Von Jagow and Link 1986), and oligomycin, which prevents H+ channeling through the FO moiety of mitochondrial ATP synthase, with no or only a weak effect on chloroplast photophosphorylation (reviewed in Hong and Pedersen 2008). During the exponential growth phase under standard conditions (50 μmol photon m−2 s−1, 5 mM acetate, 25 °C), the cell doubling time was similar for the wild-type control (11.9 ± 0.9 h) and the ASA7-silenced mutant (12.5 ± 0.8 h). In the presence of myxothiazol, inhibition of cell growth was similar in control and mutant strains (fig. 6a), and its magnitude was in good agreement with previous reports in C. reinhardtii (e.g., Cardol et al. 2002). Surprisingly, wild-type cells were rather insensitive to the presence of oligomycin. In contrast, a significant sensitivity to oligomycin was observed for ASA7-silenced mutant cells. The same effect could be observed for cells grown heterotrophically on solid medium supplemented with 1, 5, or 20 μM oligomycin (fig. 6B). Wild-type cells were almost insensitive to oligomycin, whereas ASA7-silenced mutant growth was gradually inhibited as the oligomycin concentration increased.

FIG. 6.

Impact of inhibitors of the mitochondrial respiratory chain on growth of WT and ASA7-silenced cells. (A) Biomass accumulated after 24 h (starting concentration: 0.2 mm3 ml−1) under mixotrophic conditions in the absence or presence of the mitochondrial respiratory inhibitor oligomycin (5 μM) or myxothiazol (5 μM). Biomass was calculated as the product of the cell density (cell/ml) by the mean cell volume (mm3). Error bars indicate standard deviation of the mean from three independent measurements. (B) Cells were cultivated in heterotrophic conditions (dark, 5 mM acetate) in the absence or presence of oligomycin (1, 5, or 20 μM). Cell suspensions (A750 = 0.1) were spotted on solid agar plates, and growth was evaluated after 10 days in the dark.

FIG. 6.

Impact of inhibitors of the mitochondrial respiratory chain on growth of WT and ASA7-silenced cells. (A) Biomass accumulated after 24 h (starting concentration: 0.2 mm3 ml−1) under mixotrophic conditions in the absence or presence of the mitochondrial respiratory inhibitor oligomycin (5 μM) or myxothiazol (5 μM). Biomass was calculated as the product of the cell density (cell/ml) by the mean cell volume (mm3). Error bars indicate standard deviation of the mean from three independent measurements. (B) Cells were cultivated in heterotrophic conditions (dark, 5 mM acetate) in the absence or presence of oligomycin (1, 5, or 20 μM). Cell suspensions (A750 = 0.1) were spotted on solid agar plates, and growth was evaluated after 10 days in the dark.

To evaluate the possible impact of Asa7 loss on mitochondrial respiratory activity, oxygen uptake of ASA7-silenced cells in the dark was compared with control cells. The results presented in table 2 show that the total respiration rate was slightly weaker in ASA7-silenced cells (80% of WT). Myxothiazol and SHAM were used to inhibit the cytochrome pathway and the alternative oxidase pathway of respiration, respectively. From the data of table 2, the apparent cytochrome pathway capacities of wild-type and ASA7-silenced cells (expressed in μmol O2 h−1 mg chlorophyll−1 and calculated as described in the Materials and Methods section) were 43 ± 3 and 33 ± 4, respectively, whereas the apparent alternative pathway capacities were 13 ± 3 and 15 ± 3, respectively. Addition of an uncoupler (10-μM CCCP) had no effect on wild-type respiration but accelerated the respiration rate of ASA7-silenced cells up to values similar of those of the control. These results suggest that the decrease in cytochrome pathway activity in ASA7-silenced mitochondria could be due to a slight limitation in ATP synthase activity. Addition of 10 μM oligomycin did not significantly reduce the respiratory rate of wild-type cells, even after 4 h of incubation in the dark. In this case, SHAM (1 mM) was added at the same time as oligomycin to inhibit the alternative oxidase pathway. This one is not under the control of the electrochemical proton gradient, and its activity could mask a decreased cytochrome pathway activity when complex V is inhibited (Peltier and Thibault 1985). In contrast to our observations for wild-type cells, the respiration of ASA7-silenced cells dropped by about 40% in the presence of oligomycin. The steady-state levels of intracellular ATP were also measured. No difference was observed between the two strains. As expected, the addition of myxothiazol dramatically reduced the ATP levels in both strains after 4-h incubation in the dark. In the same conditions, the presence of oligomycin did not affect the ATP level in wild-type cells. In contrast, the steady-state ATP level was reduced by ∼30% in ASA7-silenced cells.

Table 2.

Dark Respiratory Rate and Steady-State ATP Levels in Wild-Type and ASA7-Silenced Mutant Cells.

  WT ASA7-Silenced 
Respiration No addition 45 ± 3 37 ± 4 
μmoles O2 · h−1 · mg chl−1 + Myxothiazol (5 μM) 15 ± 4 18 ± 3 
 + SHAM (1 mM) 45 ± 1 36 ± 6 
 + Myxothiazol (5 μM) 2 ± 2 3 ± 2 
 + SHAM (1 mM) 
 + CCCP (10 μM) 46 ± 2 48 ± 4 
 + Oligomycin (10 μM) 46 ± 4 23 ± 3 
 + SHAM (1 mM) 
ATP level No addition 73 ± 2 77 ± 5 
nmol ATP · mg chl−1 + Myxothiazol (5 μM) 28 ± 4 23 ± 5 
 + Oligomycin (10 μM) 78 ± 3 57 ± 5 
  WT ASA7-Silenced 
Respiration No addition 45 ± 3 37 ± 4 
μmoles O2 · h−1 · mg chl−1 + Myxothiazol (5 μM) 15 ± 4 18 ± 3 
 + SHAM (1 mM) 45 ± 1 36 ± 6 
 + Myxothiazol (5 μM) 2 ± 2 3 ± 2 
 + SHAM (1 mM) 
 + CCCP (10 μM) 46 ± 2 48 ± 4 
 + Oligomycin (10 μM) 46 ± 4 23 ± 3 
 + SHAM (1 mM) 
ATP level No addition 73 ± 2 77 ± 5 
nmol ATP · mg chl−1 + Myxothiazol (5 μM) 28 ± 4 23 ± 5 
 + Oligomycin (10 μM) 78 ± 3 57 ± 5 

From these analyses, we conclude that ATP synthase activity is still efficient in the absence of Asa7 subunit but becomes partially sensitive to oligomycin.

Resistance to Oligomycin Is a Property Shared by Chlorophycean Algae

From the functional data presented above, it appears that the lack of Asa7 atypical subunit leads to a higher sensitivity to oligomycin. To determine whether the presence of an atypical or classical ATP synthase in Chlorophytes is correlated to an in vivo resistance or sensitivity to oligomycin, species belonging to various orders of the four classes were selected (see table 3) and tested for their ability to grow under phototrophic and/or heterotrophic conditions on agar plates supplemented or not with 0.2, 1, or 10 μM oligomycin. As exemplified in figure 7, phototrophic growth of C. ellipsoideum cells was insensitive to 10 μM oligomycin. Conversely, C. vulgaris, G. paucicellulare, and T. chuii grew much slower in the presence of 10 μM oligomycin. Table 3 summarizes our observations for each species. Without any exception, chlorophycean algae were all insensitive to oligomycin at the concentrations tested, whereas the algae belonging to the other three classes showed a growth reduction already detected for concentrations of 0.2 or 1 μM oligomycin.

Table 3.

Oligomycin Sensitivity in Chlorophytes.

  Phototrophic Growth
 
Heterotrophic Growth
 
Dark Respiratory Rate ATP Level 
 Oligomycin (μM) 0.2 10 0.2 10 10 10 
Chlorophyceae 
    Chlorococcales Chlorococcum ellipsoideum         98% 98% 
    Chaetophorales Uronema acuminata         nd nd 
    Chlamydomonadales Chlamydomonas reinhardtii         nd nd 
 Chlamydomonas moewusii     † † † † nd nd 
 Chlorogonium elongatum         nd nd 
    Sphaeropleales Scenedesmus obliquus         98% 100% 
Trebouxiophyceae 
    Chlorellales Chlorella sorokinia    † † 43% 37% 
 Chlorella vulgaris    † † 40% 65% 
 Nannochloris sp.  † † † † † nd nd 
    Coccomyxaceae Coccomyxa pringsheimii  † † † † † nd nd 
    Ctenocladales Leptospira obovata   † † † † nd nd 
Ulvophyceae 
    Ulvales Pseudendoclonium basiliense   † † † † nd nd 
    Ulotrichales Gloeotilopsis paucicellulare   † † † † 68% 46% 
 Ulothrix Fimbriata   † † † † nd nd 
Prasinophyceae 
    Mamiellales Micromonas pusilla  nd nd nd nd nd nd 
    Chlorodendrales Tetraselmis chuii  nd nd nd nd 39% 44% 
  Phototrophic Growth
 
Heterotrophic Growth
 
Dark Respiratory Rate ATP Level 
 Oligomycin (μM) 0.2 10 0.2 10 10 10 
Chlorophyceae 
    Chlorococcales Chlorococcum ellipsoideum         98% 98% 
    Chaetophorales Uronema acuminata         nd nd 
    Chlamydomonadales Chlamydomonas reinhardtii         nd nd 
 Chlamydomonas moewusii     † † † † nd nd 
 Chlorogonium elongatum         nd nd 
    Sphaeropleales Scenedesmus obliquus         98% 100% 
Trebouxiophyceae 
    Chlorellales Chlorella sorokinia    † † 43% 37% 
 Chlorella vulgaris    † † 40% 65% 
 Nannochloris sp.  † † † † † nd nd 
    Coccomyxaceae Coccomyxa pringsheimii  † † † † † nd nd 
    Ctenocladales Leptospira obovata   † † † † nd nd 
Ulvophyceae 
    Ulvales Pseudendoclonium basiliense   † † † † nd nd 
    Ulotrichales Gloeotilopsis paucicellulare   † † † † 68% 46% 
 Ulothrix Fimbriata   † † † † nd nd 
Prasinophyceae 
    Mamiellales Micromonas pusilla  nd nd nd nd nd nd 
    Chlorodendrales Tetraselmis chuii  nd nd nd nd 39% 44% 

NOTE.—Phototrophic and heterotrophic growths were tested on agar plates supplemented (see fig. 7 for details) with the indicated oligomycin concentration. White box, no growth delay compared with control plate; #, growth inhibition was observed; and †, no growth. Percentages of residual dark respiratory rate and steady-state ATP levels after 1 h in the dark in the presence of 10 μM oligomycin and 1 mM SHAM. nd, not determined.

FIG. 7.

Oligomycin sensitivity in Chlorophytes. The sensitivity of four chlorophycean algae representative of the four classes of Chlorophytes (see table 3) was tested after 5 days of growth in the light (50-μmol photons m−2 s−1) on agar plates supplemented or not with 10 μM oligomycin as indicated on the picture. Cells were plated at three different cell densities (from top to bottom, A750 = 0.2, 0.05, and 0.01).

FIG. 7.

Oligomycin sensitivity in Chlorophytes. The sensitivity of four chlorophycean algae representative of the four classes of Chlorophytes (see table 3) was tested after 5 days of growth in the light (50-μmol photons m−2 s−1) on agar plates supplemented or not with 10 μM oligomycin as indicated on the picture. Cells were plated at three different cell densities (from top to bottom, A750 = 0.2, 0.05, and 0.01).

Finally, we evaluated the effect of oligomycin (10-μM) and SHAM (1-mM) addition on the dark respiratory rate of light-adapted cultures for some species representative of the four algal classes. We also determined the steady-state ATP levels after incubation for 1 h in the dark in the presence or in the absence of oligomycin. Values presented in table 3 indicate that, as previously observed for C. reinhardtii wild-type cells (table 2), both parameters were unaffected by oligomycin addition in the two chlorophycean algae S. obliquus and C. ellipsoideum. In contrast, oligomycin markedly reduced dark respiratory rate and ATP steady-state level in Trebouxiophyceae Chlorella species, in the Ulvophyceae G. paucicellulare, and in the Prasinophyceae T. chuii. Altogether these results indicate a strong in vivo resistance of Chlorophyceae to oligomycin.

Discussion

In this work, we gathered and obtained sequence data confirming that core subunits of the bacterial ATP synthase are highly conserved in most eukaryotes. These core proteins are subunits α,β,γ, and δ, of the catalytic sector F1, subunits a and c of the H+ translocation membrane sector FO, as well as subunits b and OSCP of the peripheral stator stalk (e.g., Velours and Arselin 2000). In Plantae and Opisthokonts (i.e., Fungi and Metazoa), the mitochondrial F1FO-ATP synthase (complex V) comprises at least 14 conserved subunits, among which six subunits (ϵ, d, f, g, IF1, and A6L) must have been recruited before the divergence of these two eukaryotic super-groups. Opisthokonts acquired the subunits e and h and a few additional subunits were further acquired either by Fungi or along the lineage leading to mammals. In Plantae, apart from Chlorophyceae (see below), all organisms investigated at the genomic level share a rather similar set of subunits. However, due to the lack of biochemical characterization of the enzyme complex in most of these species, we cannot exclude that other species-specific subunits may exist. In contrast, five classical subunits (b, d, f, g, and A6L) were not observed in representative of three orders of Chlorophyceae (Chlamydomonadales [Cardol et al. 2005; Vázquez-Acevedo et al. 2006; Van Lis et al. 2007], Chlorococcales, and Sphaeropleales), whereas up to eight Asa subunits of unknown origin were identified. Although molecular data are not yet available for other orders of Chlorophyceae (e.g., Chaetophorales and Oedogoniales), these findings suggest that Asa subunits could be components characteristic of the mitochondrial ATP synthase in the whole chlorophycean lineage.

This conclusion is in line of our previous hypothesis (Vázquez-Acevedo et al. 2006) according to which the absence of atp4 (subunit b) and atp8 (subunit A6L) genes in the mitochondrial genome is a clue pointing to the presence of an atypical enzyme. Both genes are indeed absent from chlorophycean mitochondrial genomes but are found in the mitochondrial genome of nonchlorophycean Chlorophytes and Streptophytes (Vázquez-Acevedo et al. 2006) (see also table 1). Moreover, chlorophycean mitochondrial genomes exhibit a distinct pattern of evolution (Turmel et al. 1999), with an extremely compact and sometimes linear organization, and contain scrambled rRNA genes along to only a few short noncoding regions. Gene transfer from mitochondria to the nucleus also occurred massively, thus leading to relocalization of several genes (e.g., cox2, cox3, nad3, nad4L, nad7, and nad9) coding for core subunits of respiratory-chain complexes that are still mitochondria encoded in other green plants and in the vast majority of eukaryotes (e.g., Bullerwell and Gray 2004; Vázquez-Acevedo et al. 2006). In the cases of atp4 and atp8, no homolog could be found in the nuclear genome of C. reinhardtii or V. carteri (Cardol et al. 2005; Vázquez-Acevedo et al. 2006). Marx and collaborators suggested that the migration of mitochondrial genes to the nucleus may actually be the underlying cause for the recruitment of additional subunits (Marx et al. 2003). In addition, these authors proposed that the number of genes that have migrated to the nucleus is correlated with the number of secondarily acquired subunits. Two scenarios could be invoked for the origin of Asa subunits: 1) the retargeting of previously plastidic/cytosolic proteins to the mitochondria (possibly after gene duplication), and their assembly into a new scaffold in the peripheral stalk of the ATP synthase, or 2) the acquisition of novel genes by lateral gene transfer (either from bacterial or eukaryotic origin), and the utilization of the corresponding proteins as novel subunits of the ATP synthase. From our database searches, it not possible to conclude as we did not find any homolog (neither ortholog or paralog) for ASA genes (except ASA4) beyond Chlorophyceae. Thus, whatever their origin, Asa subunits are likely to have undergone extensive evolutionary divergence. In Chlorophyceae, the recruitment of Asa subunits might be concomitant to the loss of mitochondrial genes for ATP synthase proteins (including subunits b and A6L) along with the loss of secondarily acquired subunits d, f, and g. Subunits b and d are the main building blocks of the peripheral stator architecture in eukaryotes (Velours and Arselin 2000; Walker and Dickson 2006), whereas subunit g is critical for dimer stability (Arnold et al. 1998; Arselin et al. 2004). In C. reinhardtii, lack of Asa7 leads to a less stable dimeric enzyme that partially dissociates after detergent exposure and BN-PAGE, hence releasing the F1 moiety. This observation provides a consistent proof of the involvement of this atypical subunit in the peripheral stator and/or in the dimerization architecture of the chlorophycean ATP synthase. This is not the first example of a modification of the stator and dimerization module architecture during species evolution. Indeed, contrarily to F1 and FO subunit composition, the composition of the peripheral stator differs between bacterial and classical mitochondrial enzymes (reviewed in Walker and Dickson 2006): formed by OSCP (named δ in prokaryotes) and by an homodimer or heterodimer of subunit b/b′ in prokaryotes, the peripheral stator contains four main subunits in Metazoa and Fungi (subunits b, d, h, and OSCP) (Walker and Dickson 2006).

In the yeast S. cerevisiae, mutations in the stator also have no effect on F1 assembly but do affect its ability to interact with the FO part in vivo (reviewed in Rak et al. 2009). This in vivo effect is not observed in the ASA7-silenced mutant, because ATP synthase activity (and presumably complex V assembly) in the mutant is barely affected. The number of subunits known to be involved in the dimerization process also differs from one group to another: three in yeast (e, g, and k), two in mammals (e and g), and one in plants (g). In contrast to the stator, the dimerization module is dispensable for cell survival (Arnold et al. 1998). Functional roles of ATP synthase oligomers have been, however, suggested and were recently reviewed (Wittig and Schagger 2009). Briefly, they include dynamic regulation of ATP synthase activity, stabilization of the holoenzyme, bending of membranes to alter the local pH gradient, and participation in supramolecular arrangement of respiratory complexes. In this respect, unlike other known ATP synthases, dimeric mitochondrial complex V of chlorophycean algae hardly dissociates upon solubilization with nonionic detergents, whereas a dimeric enzyme in nonchlorophycean Chlorophytes, if present, still remains to be identified (Van Lis et al. 2003, 2007; Vázquez-Acevedo et al. 2006). This strongly indicates that the stability of dimeric complex V as judged from BN-PAGE experiments could be taken as an indicator of the enzyme subunit composition. In green algae at least, a highly stable dimeric enzyme would be correlated with an atypical subunit composition, whereas an unstable dimeric enzyme would reflect a classical subunit composition. In C. reinhardtii and Polytomella sp. Pringsheim 198.80, monomeric F1FO or free F1 moieties were observed only after heat treatment or addition of biliary salts, the monomeric form exhibiting a very short half-life time (∼1 min) in these conditions (Van Lis et al. 2007; Villavicencio-Queijeiro et al. 2009). The dimeric organization of the mitochondrial ATP synthase from C. reinhardtii is also unaffected by the cellular metabolic state, in contrast with the chloroplastic dimeric complex V (Rexroth et al. 2003; Schwassmann et al. 2007). Lack of Asa7 does not seem to alter mitochondrial structure, cell growth, or ATP level. It is thus tempting to conclude that in vivo, the dimeric state prevails in the absence of Asa7 and that other Asa subunits might be involved in dimer stability. Asa6 could be instrumental in maintaining the dimeric synthase conformation as suggested by the presence of a possible coiled coil (Van Lis et al. 2007) and evidenced by cross-linking experiments in Polytomella sp. Pringsheim 198.80 (Villavicencio-Queijeiro et al. 2009). However, it is important to remember that Asa6 and Asa7 subunits do not share any similarity with classical subunits e, g, and k (Vázquez-Acevedo et al. 2006; Van Lis et al. 2007), which mediate dimerization of the yeast ATP synthase (Arnold et al. 1998; Brunner et al. 2002). In Polytomella sp. Pringsheim 198.80, dimeric complex V is also more resistant than the monomeric form to heat treatment, high hydrostatic pressure, and protease degradation (Villavicencio-Queijeiro et al. 2009). These results point out possible evolutionary advantages in terms of structural integrity brought by the Asa subunits. Due to this highly stable nature, the dimeric ATP synthase from Chlorophyceae is a good candidate to work with in order to further elucidate the functional role of oligomeric state of ATP synthase in eukaryotes.

As highlighted in this work, atypical Asa7 protein is not essential for mitochondrial F1FO-ATPase complex activity and assembly. Surprisingly, an ASA7-silenced mutant is more sensitive to oligomycin than the wild type. Perhaps even more remarkably, species belonging to four orders of Chlorophyceae (see table 3) are insensitive to oligomycin concentrations that reduce cell growth, respiration, and ATP levels of algae belonging to other Chlorophyte classes (i.e., Trebouxiophyceae, Prasinophyceae, and Ulvophyceae).

Produced in various strains of Streptomyces, oligomycins are potent and classical inhibitors of mitochondrial ATP synthase H+ channel in vitro and also in vivo in various organisms (reviewed in Hong and Pedersen 2008). Two distinct mechanisms for oligomycin resistance, a direct and an indirect one, have been so far described in the yeast S. cerevisiae. The direct mechanism involves mutated residues in subunits a and c (John and Nagley 1986; Nagley et al. 1986). These amino acids are conserved in Chlorophyceae (C. reinhardtii and S. obliquus), as they are in other green algae (supplementary fig. 2, Supplementary Material online). In addition, expression of C. reinhardtii subunit a in an ATP6-deficient human cell line has been claimed to restore oligomycin-sensitive ATP synthesis in permeabilized cells (Ojaimi et al. 2002). At last, the activity of purified complex V from Polytomella sp. Pringsheim 198.80 is fully inhibited by oligomycin once it is detergent-activated (Villavicencio-Queijeiro et al. 2009). Altogether, these data strongly indicate that the oligomycin binding site is well conserved in the chlorophycean complex V. The indirect mechanism described in yeast requires overexpression of YOR1 protein, an ABC transporter located in the plasma membrane and responsible for multidrug resistance (e.g., Katzmann et al. 1995; Grigoras et al. 2008). Although a hypothetical YOR1 homolog was identified in Chlamydomonas (XP_001693422), overexpression of such a plasma membrane transporter would not explain the previous report by Eriksson and coworkers in 1995, who showed that ADP-stimulated respiration of Chlamydomonas purified mitochondria is almost insensitive to oligomycin concentrations (up to 1 μM) that fully inhibit the same parameter in isolated pea mitochondria (Eriksson et al. 1995). These authors already suggested that low sensitivity to oligomycin might be related to different properties of the mitochondrial ATP synthase from Chlamydomonas as compared with higher plants. The sensitivity to oligomycin of Asa7-deprived cells demonstrates the link between oligomycin resistance in vivo and the presence of atypical complex V subunit composition in Chlorophyceae. Asa7 protein would be positioned in a manner that limits or prevents the access of oligomycin to its action site when the dimeric enzyme is embedded within the inner mitochondrial membrane. This proposal is in good agreement with a working model proposed for the Polytomella enzyme (Van Lis et al. 2007), in which Asa7 would be located in the dimerization region in the vicinity of the subunit a, which is involved in H+ channeling and oligomycin sensitivity (see above).

In conclusion, the drastic modification in subunit composition of the peripheral stator and dimerization module that occurred through the recruitment of Asa7 along with other Asa subunits probably conferred on the chlorophycean ancestor new properties in terms of oligomer stability and shielding the H+ channel against inhibitory molecules. Future work will determine whether other roles might be ascribed to the other Asa subunits. In other respects, we propose that four criteria could be independently taken into consideration to detect an atypical ATP synthase in Chlorophyceae 1) an in vivo resistance to oligomycin, 2) the in vitro stability of the dimeric form of the enzyme, 3) the presence of atypical subunits, or 4) the absence of ATP4 mitochondrial gene coding for subunit b.

Supplementary Material

Supplementary figures 1 and 2 and supplementary tables S1 and S2 are available at Molecular Biology and Evolution online (http://www.mbe.oxfordjournals.org/).

This work was supported by grants from the Belgian Fonds pour la Recherche Scientifique (F.R.S.-FNRS 1.C057.09 and F.4735.06 to P.C., 2.4638.05 and 1.5.255.08 to C.R. as well as 2.4601.08 to M.B. and C.R.), from Action de la Recherche Concertee ARC07/12-04 to C.R., and from the Interuniversity Attraction Poles Program, Belgian Science Policy to M.B. We also acknowledge funding from grants 56619 from CONACyT and IN217108 from PAPIIT, DGAPA, UNAM (Mexico). The authors warmly thank René Matagne for careful reading of the manuscript and also thank Michèle Radoux and Miriam Vázquez-Acevedo for technical assistance. M.L. is supported by the F.R.S.-FNRS. P.C. is F.R.S.-FNRS research associate. E.R.-S. receives a fellowship from CONACyT (203465).

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Author notes

Associate editor: Richard Thomas