* Genomic Disorders Research Centre, University of Melbourne Department of Medicine, St. Vincent's Hospital, Victoria, Australia
Department of Pathology and Laboratory Medicine, Center for Aging and Developmental Biology, University of Rochester Medical Center
Centre for Neuroscience, University of Melbourne, Victoria, Australia
Correspondence: E-mail: i.trounce{at}unimelb.edu.au.
![]() |
Abstract |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key Words: cybrid xenomitochondrial cytochrome b mtDNA respiratory chain coevolution murids
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Coevolution of mtDNA-encoded and nuclear-encoded RC subunits had been suggested since the finding of heterogeneous mutation rates of different mtDNA genes in cross-species comparisons, in particular for cytochrome oxidase subunit II (COII) and cytochrome oxidase subunit IV (COIV)(Cann, Brown, and Wilson 1984; Lomax et al. 1992). Considerable evidence has been gained for a coevolutionary interaction of primate COII and cytochrome c (Osheroff et al. 1983; Adkins, Honeycutt, and Disotell 1996; see Blier, Dufresne, and Burton 2001 for review).
Kenyon and Moraes (1997) found that mtDNAs from the primate species most closely related to humans, gorilla and chimpanzee, could repopulate human cells without endogenous mtDNA (0 cells), whereas more divergent primate mtDNAs could not. These primate "xenomitochondrial cybrids" showed reduced complex I activity with preserved function of the other respiratory chain complexes (Kenyon and Moraes 1997). Several groups subsequently showed that mouse
0 cells could be repopulated with mtDNA from both the closely related Mus spretus and the divergent Rattus norvegicus (Dey, Barrientos, and Moraes 2000; McKenzie and Trounce 2000; Yamaoka et al. 2000). We found that in the Rattus xenocybrids, the catalytic activity of RC complexes I, III, and IV were all affected, with complex III showing the greatest defect (McKenzie and Trounce 2000). This contrasted with the primate xenocybrid findings of specific complex I defects (Kenyon and Moraes 1997), and also with human-orangutan hybrid cell studies where a complex IV defect was most pronounced (Barrientos et al. 2000).
Unlike the small number of primate species closely related to humans, there are dozens of species of murids closely related to the laboratory mouse, Mus musculus domesticus. The genus Mus alone has around 38 species, the murinae subfamily over 500 species, and the Muridae family over 1,300 species (Michaux and Catzeflis 2000). This richness of closely related species therefore provides a unique opportunity to model in more detail the functional constraints of nuclear/mitochondrial genome coevolution using transmitochondrial cell technology.
Here we describe the creation of a panel of xenocybrids using as mitochondrial donors, cells from species diverged from M. m. domesticus by around 2, 4, 6, and 12 Myr before present. Functional studies of the resulting xenocybrid respiratory chain complexes revealed progressive impacts on all the complexes with dual genetic origins, as expected. Interestingly, complex III showed the most severe defects in the most divergent xenocybrids, with a partial loss of electron transfer function in Rattus xenocybrids and a complete loss of activity in the Otomys xenocybrid. Complex I and complex IV were not as severely defective in these cybrids. Since cytochrome b is the only mtDNA-encoded subunit of complex III, this suggests that the nuclear subunits of the holocomplex that interact with cytochrome b face the highest coevolutionary constraint in murids.
![]() |
Materials and Methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Primary fibroblast lines were created from a 2-day-old laboratory mouse (M. m. domesticus, B6, CBA background) and a 5-week-old Mus spretus (SPRET/Ei) mouse, as described previously (McKenzie and Trounce 2000).
Mus caroli primary fibroblasts were a gift from Rachel Waugh O'Neill (University of Connecticut, Storrs); Mus pahari primary fibroblasts were prepared from animals obtained from the Jackson Laboratory (Bar Harbor, Me), and Otomys irroratus fibroblasts were a gift from Terry Robinson (Stellenbosch University, South Africa). The RN1T Rattus norvegicus mammary tumor cell line and the III8C Mus dunni primary fibroblast cell lines were obtained from the American Type Culture Collection (Manassas, Va). The mouse 0 (mtDNA-less) cell clone LMEB3 was derived from the parental line LMTK by exposure to ethidium bromide (Trounce et al. 2000).
Production of Xenomitochondrial L-Cell Cybrids
Mouse L-cell cybrids were produced by enucleation of mitochondrial donor cells and fusion of the cytoplasts with mouse 0 cells, followed by selection for respiratory competent transformants, as previously described (McKenzie and Trounce 2000). Cells used as mitochondrial donors included the M. m. domesticus, Mus spretus, Mus caroli, Mus dunni, Mus pahari, and Otomys irroratus primary fibroblast lines and the Rattus norvegicus mammary tumor line described above.
Seven to 10 days postfusion, cybrid clones were isolated using cloning cylinders, expanded and viably frozen. Three independent clones were frozen from each fusion experiment, and cybrids were used in experiments from passage 5 through passage 10.
Extraction of Total DNA from Cultured Cells and Mouse Tissue
Approximately 1 x 106 cells or 0.5 g of mouse tissue were digested with 100 µg/ml proteinase K (Invitrogen) in STE buffer (100 mM NaCl, 25 mM EDTA, and 10 mM Tris-HCl pH 8.0) with 0.5% SDS for 4 h at 55°C, followed by digestion with 40 µg/ml RNase A (Invitrogen) for 1 h at 37°C. DNA was extracted using phenol/chloroform/isoamyl alcohol (25:24:1) followed by ethanol precipitation. DNA pellets were resuspended to 200 µg/ml in TE pH 8.0 (1 mM EDTA and 10 mM Tris-HCl pH 8.0).
mtDNA PCR Amplification
A 461-bp mtDNA fragment containing part of the D-loop and the tRNAPhe gene was generated using the forward primer 5'-CTC AAC ATA GCC GTC AAG GC-3' (representing nucleotides 15934 to 15953 [Bibb et al. 1981]) and the reverse primer 5'-ACC AAA CCT TTG TGT TTA TGG G-3' (representing nucleotides 80 to 59) using 1.0 U Taq polymerase (Invitrogen) and 35 cycles at 94°C for 30 s, 55°C for 1 min, and 72°C for 1 min (McKenzie and Trounce 2000).
The complete cytochrome b gene was amplified using the forward primer 5'-CGA AGC TTG ATA TGA AAA ACC ATC GTT G-3' and the reverse primer 5'-TCT TCA TTT YWG GTT TAC AAG AC-3' using 1.0 U Taq polymerase (Invitrogen) and 35 cycles at 94°C for 30 sec, 55°C for 1 min, and 72°C for 1 min (Jansa, Goodman, and Tucker 1999).
mtDNA Sequencing and Creation of Phylogenetic Trees
Sequencing of mouse mtDNA was performed using the primers described for PCR amplification and the primers UMMZ12 5'-RTA DGG GTG RAA TGG RAT TTT WTC-3' and UMMZ13 5'-CAY GAA WCA GGV TCA AAY AAY CC-3' for the cytochrome b gene (Jansa, Goodman, and Tucker 1999). PCR templates were purified using "Wizard" purification columns (Promega, Madison, Wis.), and sequencing reactions were performed using the BigDye terminator cycle sequencing kit (Applied Biosystems, Foster City, Calif.) as per the manufacturer's instructions. Reactions were analyzed using an ABI 377 automated sequencer and Sequencing Analysis software (Applied Biosystems).
D-loop and cytochrome b gene sequences were aligned using ClustalX software (Thompson et al. 1997). Maximum-parsimony trees were created using ClustalX and NJPlot software.
Lactate Measurement
Lactate was measured in medium using a commercial kit (Sigma). Approximately 1 x 106 cells were harvested into 15 ml tubes, resuspended in 1 ml of RPMI/GUP medium, and incubated at 37°C/5% CO2 for up to 66 h. Ten milliliters of medium were removed at various time intervals and combined with 500 ml of reaction mixture (0.2 M glycine and hydrazine pH 9.2, 33 units/ml lactate dehydrogenase, and 3.3 mg/ml NAD). Reactions were incubated at 37°C for 15 min before spectrophotometric determination of NAD reduction by measuring absorbance at 340 nm. Measurements were compared with a lactate standard and standardized per cell.
Mitochondrial Isolation
Cultures were expanded to approximately 109 cells by seeding 2 x 107 cells into roller bottles (Corning) in 300 ml RPMI/GUP medium, expanded after 4 or 5 days to 1,000 ml, and then harvested after another 4 days. Mitochondrial isolation was performed as previously described (McKenzie and Trounce 2000), with mitochondrial isolates frozen in aliquots at -70°C.
OXPHOS Enzymology
Frozen mitochondrial aliquots were thawed and diluted to 2 µg/µl in mitochondrial isolation buffer and used to determine mitochondrial respiratory chain activity. Complex I (NADH:ubiquinone oxidoreductase, EC 1.6.5.3), complex II (succinate:ubiquinone oxidoreductase, EC 1.3.5.1), complex II+III (succinate:cytochrome c oxidoreductase), and complex IV (ferrocytochrome c:oxygen oxidoreductase, or cytochrome oxidase, EC 1.9.3.1) activities were measured spectrophotometrically as previously described (Trounce et al. 1996). Five micrograms of rotenone were also added to the complex I reaction mixture of duplicate reactions to measure complex I rotenone-insensitive activity.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Fusions of enucleated hamster cells with LMEB3 failed to produce any cybrids in two experiments, one using Chinese hamster (Cricetulus grigaeus) cells and the other using Syrian golden hamster (Mesocricetus aureus) cells, as previously reported (McKenzie and Trounce 2000).
Genetic Comparison of Murid Species
The mitochondrial genomes of the rodent species M. m. domesticus, Mus spretus, Mus caroli, Mus dunni, Mus pahari, Otomys irroratus, and Rattus norvegicus were sequenced at two positions. The first region contained the D-loop and the tRNAPhe gene, and the second region contained the cytochrome b gene.
The D-loop sequence of the M. m. domesticus primary culture was identical to the published sequence (Bibb et al. 1981). The Mus spretus D-loop showed 89% homology with the published M. m. domesticus sequence, the Mus caroli D-loop showed 77% homology, Mus dunni showed 80% homology, Mus pahari showed 76% homology, Rattus norvegicus showed 65% homology, and Otomys irroratus showed 65% homology. The D-loop sequence from the Rattus norvegicus RN1T cells was also identical to the published R. norvegicus sequence (Gadaleta et al. 1989).
The cytochrome b sequence of the M. m. domesticus primary culture was identical to the published sequence (Bibb et al. 1981); the Mus spretus sequence showed 91% homology with the published M. m. domesticus sequence, Mus caroli showed 86% homology, Mus dunni showed 87% homology, Mus pahari showed 86% homology, Rattus norvegicus showed 83% homology, and Otomys irroratus showed 82% homology (table 1).
|
Translation of these sequences revealed a high degree of conservation, with amino acid sequence homology higher than genomic sequence homology as expected (fig. 1 and table 1). Comparisons were also made between the amino acid sequences determined here and the published sequences. M. m. domesticus, Mus caroli, Mus pahari, and Rattus norvegicus amino acid sequences were identical to their respective published sequences, whereas Mus spretus showed one amino acid difference (an alanine to threonine change at position 23) and Otomys irroratus showed 16 differences (table 1). Alignment of the cytochrome b amino acid sequences is shown in figure 1.
|
Both D-loop and cytochrome b gene sequences were combined and used to create a phylogenetic tree (fig. 2). The mtDNA tree predicts the same evolutionary relationship between the murid species examined as the consensus nuclear gene tree, except that Otomys irroratus did not nest between Rattus and Mus as suggested by a recent study using nuclear genes (Michaux, Reyes, and Catzeflis 2001). Mus dunni also appeared to be more closely related to M. m. domesticus than Mus caroli. Lepus granatensis (Granada hare) D-loop (GenBank accession number AF157432) and cytochrome b gene (GenBank accession number AF157465) sequence data were used to root the tree.
|
|
OXPHOS Enzymology
The mean activity of OXPHOS complexes I, II, II+III, and IV was measured in triplicate using three independent mitochondrial isolates from each of the control cybrid and each xenocybrid and normalized to the control cybrid (100% activity). No significant differences were found for the Mus spretus, Mus caroli, or Mus dunni xenocybrids when compared with the control cybrid (P>0.2). The Mus pahari xenocybrid had normal complex I, II, and III activities with a significant defect in complex IV, with activity reduced to 59% (P < 0.05; fig. 4).
|
The Otomys xenocybrid exhibited the most severe respiratory defect overall, with complex I activity reduced to 72% (P < 0.05) and complex IV reduced to 44% (P < 0.05). Again, the most striking defect was measured in the complex II+III assay, with complex III activity reduced to only 2% of the control activity (P < 0.005; fig. 4). Together with the relatively preserved complex II activity, these results lead us to suggest that a severe complex III defect exists as this linked assay is normally rate-limited by complex II (Taylor et al. 1993). For this reason we will not see partial defects of complex III with this assay, and it remains possible that lesser defects exist in the other xenocybrids.
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
When the more distantly related Rattus norvegicus and Otomys irroratus mtDNAs were transferred to the mouse nuclear background, more severe respiratory chain dysfunction was evident. Lactate production in the Rattus xenocybrid was four times greater than the control cybrid after 12 h, indicating a large increase in glycolytic ATP generation to compensate for the impaired oxidative phosphorylation. The Otomys xenocybrid exhibited the greatest increase in lactate production, with a 10-fold increase observed over the control cybrid after 12 h. This lactate production was similar to the LMEB3 0 cell line, indicating little or no respiratory chain function in the Otomys xenocybrid.
Enzymological analysis was used to define the specific RC defects. In the spretus, caroli, and dunni cybrids, no significant differences were found. In the pahari cybrids complexes I, II, and II+III activities were normal, but complex IV activity was significantly lower at 59% of the control activity. Severe respiratory defects were confirmed by enzymological analysis in the two most divergent xenocybrids. The Rattus xenocybrid complex III was reduced to 37% of the control activity, complex I was reduced to 44%, and complex IV was reduced to 78%. In the Otomys xenocybrids, complex III also showed the greatest defect (2% of control cybrid) followed by complex IV (44% of control cybrid) and complex I (72% of control cybrid).
As mtDNA replication, transcription, and translation are maintained in the xenocybrids (Dey, Barrientos, and Moraes 2000; McKenzie and Trounce 2000), the respiratory defects observed are consequent to the mismatches between the mouse nuclear gene encoded OXPHOS subunits and the xenotypic mtDNA-encoded subunits. The cybrid system used here depends on some respiratory chain function to overcome the auxotrophy for uridine and pyruvate exhibited by mtDNA-less cells (Morais et al. 1988; King and Attardi 1989). In the Otomys cybrid, there is virtually no respiratory chain function due to the severe complex III defect, as indicated by the lactate production being indistinguishable from the 0 cells. It is likely that cybrids with more divergent mtDNAs would not support any respiratory chain function, and we have therefore found the functional limits of murid xenotypic nuclear/mitochondrial respiratory chain subunit interaction.
Kenyon and Moraes (1997) showed in human-gorilla and human-chimpanzee xenocybrids that only complex I had measurable defects. Barrientos et al. (2000) used human-orangutan hybrids to further explore interactions of the different primate species respiratory chain subunits. In hybrids formed by fusion of human 0 cells with orangutan whole cells or microcells plus mitochondria, they found that only RC complex IV had measurable defects, probably consequent to defective assembly of the holocomplex (Barrientos et al. 2000). This "xenohybrid" system differs by having both human and orangutan nuclear subunits expressed, allowing demonstration of dominant negative effects by competing, homologous nuclear subunits. In our pure cybrid system, such interference effects do not complicate the defects seen. We were therefore more likely to see effects directly related to decreased electron transfer efficiency in the mismatched holocomplexes.
Our results show that complex I and IV defects are also present in the murid xenocybrid system, but the ability to further extend the system in murids due to the numerous intermediate species has allowed us to further demonstrate that complex III appears to suffer most from xenotypic nuclear subunits. The severe complex III defect is particularly interesting since cytochrome b is the only mtDNA-encoded subunit of this complex. Cytochrome b shows 26 amino acid differences between M. m. domesticus and both Rattus norvegicus and Otomys irroratus, with many of these changes differing between Rattus and Otomys.
The tertiary structure of bovine holocomplex III has been solved (Iwata et al. 1998). Cytochrome b interacts closely with both the cytochrome c1 subunit and the "Rieske" iron-sulphur protein, the latter appearing to swivel to accomplish electron transfer from ubiquinol to cytochrome c1 (Iwata et al. 1998). Our results suggest that neighboring nuclear subunits do not correctly support the electron flow via cytochrome b. That is, the mouse cytochrome c1 and/or Rieske subunits are not compatible with efficient electron transfer from the Rattus or Otomys cytochrome b. Possible residues involved in this incompatibility include Met 249 (replaced by Leu in Rattus and Ala in Otomys) and Met 257 (to Thr in both Rattus and Otomys [see fig.1]). These changes occur in the loop ef of cytochrome b, which is thought to restrict the catalytic movement of the iron-sulfur protein between cytochromes b and c1 (Darrouzet and Daldal 2002). Residue 60, Met in all species except in Otomys where it is replaced with Thr, may also be important since this is in the helix ab, which also interacts with cytochrome c1 (Iwata et al. 1998).
In complex IV, all the redox centers (CuA, CuB, and hemes a and a3) are housed within the mtDNA-encoded core subunits COI and COII, with cytochrome c binding COII at the CuA site (Tsukihara et al. 1996; Abramson et al. 2001). Thus, we speculate that even in the most divergent murid xenocybrids, complex IV activity is not as severely affected since these vital electron transfer centers retain their native interactions. It would be of interest to use the various mouse species cytochromes c in this analysis, instead of the horse heart cytochrome c that was used. This may provide further evidence of coevolutionary interactions between cytochrome c and COII, as suggested by intercrossing allopatric subpopulations of a marine copepod (Edmunds and Burton 1999), and further evidenced by measuring complex IV activity using cloned cytochrome c molecules from these populations (Rawson and Burton 2002).
Complex I, having around 45 subunits at last count (Carroll et al. 2002) and a molecular mass approaching 1,000 KDa, is by far the most complex of the RC electron carriers, and the atomic structure is yet to be resolved. The L-shaped complex comprises an integral membrane arm and a globular arm projecting from the membrane, the latter housing up to nine redox iron-sulfur clusters and FMN. This projecting arm component of the complex acts as an NADH dehydrogenase. All seven mtDNA-encoded subunits are hydrophobic, integral membrane arm components with homologs in the simpler, 14-subunit bacterial enzyme (Friedrich et al. 1998). This integral membrane arm component is believed to catalyze ubiquinone reduction and proton translocation (Walker 1992). It is possible that since all the redox centers in our xenocybrids are contained in subunits derived from the M. m. domesticus nuclear genome, they are able to assemble and retain some electron transfer activity. We used the soluble ubiquinone analog, Q1, to measure complex I activity, and it is possible that this more mobile substrate can be reduced more efficiently by the chimeric complex than the native substrate.
Further investigation of putative coevolution in murid complex III genes requires sequence information from the key nuclear subunits (cytochrome c1 and the iron-sulfur protein) in these species, which are not yet available. Complete mtDNA sequences are also not yet available except for M. m. domesticus and Rattus norvegicus. It is interesting that cytochrome b, along with COI and COII in cytochrome oxidase, shows the greatest deviation from "clock-like" mutation rates in the mammalian mitochondrial genomes analyzed to date (Ma et al. 1993; Janke et al. 1994; Andrews, Jermiin, and Easteal 1998; Wu et al. 2000). We would predict that in the case of the species investigated in the present work, coadaptive amino acid changes may be seen in the Rattus and Otomys cytochrome c1 and/or iron-sulfur protein nuclear genes. We also speculate that the separation of genetic encoding of the apoproteins housing the redox centers of complex III (into both nuclear and mtDNA genes) results in the highest coevolutionary constraints among nuclear and mtDNA-encoded RC subunits.
![]() |
Supplementary Material |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
Acknowledgements |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
Footnotes |
---|
![]() |
Literature Cited |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Abramson, J., M. Svensson-Ek, B. Byrne, and S. Iwata. 2001. Structure of cytochrome c oxidase: a comparison of the bacterial and mitochondrial enzymes. Biochim. Biophys. Acta 1544:1-9.[ISI][Medline]
Adkins, R. M., R. L. Honeycutt, and T. R. Disotell. 1996. Evolution of eutherian cytochrome c oxidase subunit II: heterogeneous rates of protein evolution and altered interaction with cytochrome c. Mol. Biol. Evol. 13:1393-1404.
Andrews, T. D., L. S. Jermiin, and S. Easteal. 1998. Accelerated evolution of cytochrome b in simian primates: adaptive evolution in concert with other mitochondrial proteins? J. Mol. Evol. 47:249-257.[ISI][Medline]
Barrientos, A., S. Muller, R. Dey, J. Wienberg, and C. T. Moraes. 2000. Cytochrome c oxidase assembly in primates is sensitive to small evolutionary variations in amino acid sequence. Mol. Biol. Evol. 17:1508-1519.
Bibb, M. J., R. A. Van Etten, C. T. Wright, M. W. Walberg, and D. A. Clayton. 1981. Sequence and gene organization of mouse mitochondrial DNA. Cell 26:167-180.[ISI][Medline]
Blier, P. U., F. Dufresne, and R. S. Burton. 2001. Natural selection and the evolution of mtDNA-encoded peptides: evidence for intergenomic co-adaptation. Trends Genet. 17:400-406.[CrossRef][ISI][Medline]
Bonhomme, F., and J.-L. Guenet. 1996. The laboratory mouse and its wild relatives.Pp.15771596 in M. F. Lyon, S. Raston, and S. D. M. Brown, eds. Genetic variants and strains of the laboratory mouse. Oxford University Press, Oxford.
Cann, R. L., W. M. Brown, and A. C. Wilson. 1984. Polymorphic sites and the mechanism of evolution in human mitochondrial DNA. Genetics 106:479-499.
Carroll, J., R. J. Shannon, I. M. Fearnley, J. E. Walker, and J. Hirst. 2002. Definition of the nuclear encoded protein composition of bovine heart mitochondrial complex I: identification of two new subunits. J. Biol. Chem. 277:50311-50317.
Catzeflis, F. M., A. W. Dickerman, J. Michaux, and J. A. W. Kirsch. 1993. DNA hybridization and rodent phylogeny.Pp. 159172 in F. S. Szalay, M. J. Novacek, and M. McKenna, eds. Mammal phylogeny. Vol. 2. Placentals. Springer-Verlag, New York.
Darrouzet, E., and F. Daldal. 2002. Movement of the iron-sulfur subunit beyond the ef loop of cytochrome b is required for multiple turnovers of the bc1 complex but not for single turnover Qo site catalysis. J. Biol. Chem. 277:3471-3476.
Dey, R., A. Barrientos, and C. T. Moraes. 2000. Functional constraints of nuclear-mitochondrial DNA interactions in xenomitochondrial rodent cell lines. J. Biol. Chem. 275:31520-31527.
Ducroz, J. F., V. Volobouev, and L. Granjon. 2001. An assessment of the systematics of arvicanthine rodents using mitochondrial DNA sequences: evolutionary and biogeographical implications. J. Mammal. Evol. 8:173-206.[CrossRef]
Edmunds, S., and R. S. Burton. 1999. Cytochrome c oxidase activity in interpopulation hybrids of a marine copepod: a test for nuclear-cytoplasmic coadaptation. Evolution 53:1972-1978.[ISI]
Friedrich, T., A. Abelmann, and B. Brors, et al. (11 co-authors). 1998. Redox components and structure of the respiratory NADH:ubiquinone oxidoreductase (complex I). Biochim. Biophys. Acta 1365:215-219.[ISI][Medline]
Gadaleta, G., G. Pepe, G. De Candia, C. Quagliariello, E. Sbisa, and C. Saccone. 1989. The complete nucleotide sequence of the Rattus norvegicus mitochondrial genome: cryptic signals revealed by comparative analysis between vertebrates. J. Mol. Evol. 28:497-516.[ISI][Medline]
Iwata, S., J. W. Lee, K. Okada, J. K. Lee, M. Iwata, B. Rasmussen, T. A. Link, S. Ramaswamy, and B. K. Jap. 1998. Complete structure of the 11-subunit bovine mitochondrial cytochrome bc1 complex. Science 281:64-71.
Janke, A., G. Feldmaier-Fuchs, W. K. Thomas, A. Von Haessler, and S. Pääbo. 1994. The marsupial mitochondrial genome and the evolution of placental mammals. Genetics 137:243-256.
Jansa, S. A., S. M. Goodman, and P. K. Tucker. 1999. Molecular phylogeny and biogeography of the native rodents of Madagascar (Muridae: Nesomyinae): a test of the single-origin hypothesis. Cladistics 15:253-270.[CrossRef][ISI]
Kenyon, L., and C. T. Moraes. 1997. Expanding the functional human mitochondrial DNA database by the establishment of primate xenomitochondrial cybrids. Proc. Natl. Acad. Sci. USA 94:9131-9135.
King, M. P., and G. Attardi. 1989. Human cells lacking mtDNA: repopulation with exogenous mitochondria by complementation. Science 246:500-503.[ISI][Medline]
Lomax, M. I., D. Hewett-Emmett, T. L. Yang, and L. I. Grossman. 1992. Rapid evolution of the human gene for cytochrome c oxidase subunit IV. Proc. Natl. Acad. Sci. USA 89:5266-5270.[Abstract]
Lundrigan, B. L., S. A. Jansa, and P. K. Tucker. 2002. Phylogenetic relationships in the genus Mus, based on paternally, maternally, and biparentally inherited characters. Syst. Biol. 51:410-431.[CrossRef][ISI][Medline]
Lundrigan, B. L., and P. K. Tucker. 1994. Tracing paternal ancestry in mice, using the Y-linked, sex-determining locus, Sry. Mol. Biol. Evol. 11:483-492.[Abstract]
Ma, D-P., A. Zharkikh, D. Graur, J. L. VendeBerg, and W-H. Li. 1993. Structure and evolution of opossum, guinea pig and porcupine cytochrome b genes. J. Mol. Evol. 36:327-334.[ISI][Medline]
McKenzie, M., and I. Trounce. 2000. Expression of Rattus norvegicus mtDNA in Mus musculus cells results in multiple respiratory chain defects. J. Biol. Chem. 275:31514-31519.
Michaux, J., and F. Catzeflis. 2000. The bushlike radiation of muroid rodents is exemplified by the molecular phylogeny of the LCAT nuclear gene. Mol. Phylog. Evol. 17:280-293.[CrossRef][ISI][Medline]
Michaux, J., A. Reyes, and F. Catzeflis. 2001. Evolutionary history of the most speciose mammals: molecular phylogeny of muroid rodents. Mol. Biol. Evol. 18:2017-2031.
Morais, R., P. Desjardins, C. Turmel, and K. Zinkewich-Peotti. 1988. Development and characterization of continuous avian cell lines depleted of mitochondrial DNA. In Vitro Cell. Dev. Biol. 24:649-658.[ISI][Medline]
Osheroff, N., S. H. Speck, E. Margoliash, E. C. I. Veerman, J. Wilms, B. W. Konig, and A. O. Muijsers. 1983. The reaction of primate cytochromes c with cytochrome oxidase. J. Biol. Chem. 258:5731-5738.
Rawson, P. D., and R. S. Burton. 2002. Functional coadaptation between cytochrome c and cytochrome c oxidase within allopatric populations of a marine copepod. Proc. Natl. Acad. Sci. USA 99:12955-12958.
Saraste, M. 1999. Oxidative phosphorylation at the fin de siecle. Science 283:1488-1493.
Silver, L. M. 1995. Mouse genetics. Oxford University Press, Oxford.
Taylor, R. W., M. A. Birch-Machin, K. Bartlett, and D. M. Turnbull. 1993. Succinate-cytochrome c reductase: assessment of its value in the investigation of defects of the respiratory chain. Biochim. Biophys. Acta 1181:261-265.[ISI][Medline]
Thompson, J. D., T. J. Gibson, F. Plewniak, F. Jeanmougin, and D. G. Higgins. 1997. The Clustal_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 25:4876-4882.
Trounce I., Y. L. Kim, A. S. Jun, and D. C. Wallace. 1996. Assessment of mitochondrial oxidative phosphorylation in patient muscle biopsies, lymphoblasts and transmitochondrial cell lines. Methods Enzymol. 264:484-509.[Medline]
Trounce, I., J. Schmiedel, H. C. Yen, S. Hosseini, M. D. Brown, J. J. Olson, and D. C. Wallace. 2000. Cloning of neuronal mtDNA variants in cultured cells by synaptosome fusion with mtDNA-less cells. Nucleic Acids Res. 28:2164-2170.
Tsukihara, T., H. Aoyama, E. Yamashita, T. Tomizaki, H. Yamaguchi, K. Shinzawa-Itoh, R. Nakashima, R. Yaono, and S. Yoshikawa. 1996. The whole structure of the 13-subunit oxidised cytochrome c oxidase at 2.8 Å. Science 272:1136-1144.[Abstract]
Walker, J. E. 1992. The NADH: ubiquinone oxidoreductase (complex I) of respiratory chains. Q. Rev. Biophys. 25:253-324.[ISI][Medline]
Wu, W., T. R. Schmidt, M. Goodman, and L. I. Grossman. 2000. Molecular evolution of cytochrome c oxidase subunit 1 in primates: Is there coevolution between mitochondrial and nuclear genomes? Mol. Phylogenet. Evol. 17:294-304.[CrossRef][ISI][Medline]
Yamaoka, M., K. Isobe, H. Shitara, H. Yonekawa, S. Miyabayashi, and J. I. Hayashi. 2000. Complete repopulation of mouse mitochondrial DNA-less cells with rat mitochondrial DNA restores mitochondrial translation but not mitochondrial respiratory function. Genetics 155:301-307.