• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of narLink to Publisher's site
Nucleic Acids Res. 2004; 32(12): 3615–3622.
Published online Jul 7, 2004. doi:  10.1093/nar/gkh695
PMCID: PMC484182

Rapid evolution of RNA editing sites in a small non-essential plastid gene

Abstract

Chloroplast RNA editing proceeds by C-to-U transitions at highly specific sites. Here, we provide a phylogenetic analysis of RNA editing in a small plastid gene, petL, encoding subunit VI of the cytochrome b6f complex. Analyzing representatives from most major groups of seed plants, we find an unexpectedly high frequency and dynamics of RNA editing. High-frequency editing has previously been observed in plastid ndh genes, which are remarkable in that their mutational inactivation does not produce an obvious mutant phenotype. In order to test the idea that reduced functional constraints allow for more flexible evolution of RNA editing sites, we have created petL knockout plants by tobacco chloroplast transformation. We find that, in the higher plant tobacco, targeted inactivation of petL does not impair plant growth under a variety of conditions markedly contrasting the important role of petL in photosynthesis in the green alga Chlamydomonas reinhardtii. Together with a low number of editing sites in plastid genes that are essential to gene expression and photosynthetic activity, these data suggest that RNA editing sites may evolve more readily in those genes whose transitory loss of function can be tolerated. Accumulated evidence for this ‘relative neutrality hypothesis for the evolution of plastid editing sites’ is discussed.

INTRODUCTION

Primary transcripts in plant cell organelles undergo a complex series of RNA processing events, including maturation of their 5′ and 3′ ends, intron splicing, cleavage of polycistronic into oligocistronic or monocistronic units and RNA editing. Editing is a post-transcriptional process changing the identity of individual pyrimidine nucleotides. In higher plant plastids, these changes appear to be restricted to cytidine-to-uridine conversions [for review see e.g. (1,2)] whereas in hornwort chloroplasts, ‘reverse’ editing by U-to-C transitions has also been reported (3,4). RNA editing by pyrimidine transitions also occurs in higher plant mitochondria (57) and, interestingly, the editing processes in the two DNA-containing cell organelles of higher plants share numerous features [reviewed e.g. in (8,2)] suggesting that they originate from common evolutionary roots.

With very few exceptions (9,10), the vast majority of known plastid editing events alter the coding properties of the affected mRNAs, usually resulting in the restoration of phylogenetically conserved amino acid residues (1113). Transgenic experiments creating plants with a non-editable version of a chloroplast gene have provided direct evidence for the functional importance of plastid RNA editing (14).

The seemingly unnecessary existence of RNA editing as an additional RNA processing step in plastids provides many interesting evolutionary puzzles, most of which remain unsolved at present. In terms of the evolutionary origin of RNA editing, plastid as well as mitochondrial editing have been shown to occur in all major lineages of land plant evolution (15,16) but appear to be absent from some bryophyte lineages as well as from algae and from the prokaryotic ancestors of present-day organelles (cyanobacteria and α-proteobacteria). Interestingly, many of the known plastid editing sites are interspecifically only poorly conserved pointing to a remarkable evolutionary flexibility with respect to loss and/or acquisition of RNA editing sites (17,18,15).

To gain more information about the evolution of plastid editing sites we have investigated here the phylogeny of RNA editing in a small plastid gene, petL, encoding subunit VI of the cytochrome b6f complex. We report the detection of several new RNA editing sites, an unusually high frequency of editing in this small gene and a remarkably dynamic pattern of editing site evolution. Taken together with reverse genetics data on petL function and with published data on the functions of other highly edited genes, our data support the hypothesis that RNA editing sites evolve more readily in genes whose transitory loss of function can be tolerated. This hypothesis can explain the extremely uneven genomic distribution of plastid RNA editing sites.

MATERIALS AND METHODS

Plant material

Green leaf tissue from the plant species examined was obtained from the Botanical Gardens of Münster, Freiburg and Bonn. For transgenic experiments, tobacco plants (Nicotiana tabacum cv. Petit Havana) were grown under aseptic conditions on agar-solidified MS medium containing 30 g/l sucrose (19). Transplastomic lines were rooted and propagated on the same medium in the presence of spectinomycin (500 mg/l).

Isolation of nucleic acids

Total plant nucleic acids were isolated from leaf tissue samples using a cetyltrimethylammoniumbromide (CTAB)-based method (20). RNAs were alternatively extracted with the TriFast reagent (Peqlab GmbH, Erlangen, Germany). If necessary, additional purification steps (phenol/chloroform extractions followed by ethanol precipitation or purification by the NucleoSpin® RNA Plant kit, Macherey-Nagel, Düren, Germany) were introduced to obtain nucleic acids of sufficient purity for cDNA synthesis and/or PCR amplification.

cDNA synthesis and PCR

RNA samples treated with RNase-free DNase I were reverse transcribed with M-MLV reverse transcriptase (Promega) following the instructions of the supplier. cDNA synthesis was primed with random hexanucleotide primers or specific primers derived from the petL 3′-untranslated region (3′-UTR) (temperature for DNA polymerization 45–55°C). DNA and cDNA samples were amplified with petL-specific primers in an Eppendorf thermocycler using Taq DNA polymerase (Promega) and standard protocols. A list of species-specific primers and reaction conditions is available upon request. DNA and cDNA amplification products were purified for sequencing by electrophoresis on 1.5% agarose gels and subsequent extraction from gel slices using the QIAEX II kit (Qiagen) or the GFX kit (Amersham). Direct sequencing of amplification products was performed by cycle sequencing followed by automated analysis in a MegaBACE capillary sequencer (Amersham).

Vector construction and cloning procedures

Plastid transformation vector pIpetL was derived from a cloned tobacco ptDNA fragment. A 2.4 kb SacI/BglII restriction fragment was subcloned into pUC19 cut with SacI and BamHI. A chimeric aadA gene (21) was subsequently inserted as a blunt-end Ecl136II/DraI fragment into the SpeI site within the petL coding region blunted by a fill-in reaction with the Klenow fragment of DNA polymerase I from Escherichia coli (New England BioLabs). A clone was selected that contained the plastid selectable marker gene aadA in the same transcriptional orientation as the petL gene. This clone was designated pIpetL and used as chloroplast transformation vector.

Plastid transformation and selection of transplastomic tobacco lines

Young leaves harvested from sterile tobacco plants were bombarded with plasmid DNA-coated gold particles using the BioRad PDS1000He biolistic gun. Spectinomycin resistant cell lines were selected on RMOP regeneration medium containing 500 mg/l spectinomycin dihydrochloride (21). Plastid transformation and successful inactivation of the petL were verified by PCR employing petL-specific primers flanking the aadA insertion site. Four independently generated transplastomic lines (designated Nt-pIpetL-3, Nt-pIpetL-14, Nt-pIpetL-18 and Nt-pIpetL-31) were subjected to three additional rounds of regeneration on RMOP/spectinomycin to eliminate residual wild-type genome copies. Shoots resulting from the final round of regeneration were rooted on phytohormone-free spectinomycin-containing medium, transferred to the soil and further cultivated in the greenhouse.

RFLP analyses

Total cellular DNA was digested with restriction enzymes, separated in 0.7% agarose gels and transferred onto Hybond nylon membranes (Amersham, Buckinghamshire, UK) by capillary blotting. A PCR product covering the entire petL sequence was radiolabeled with 32P-dCTP and used as probe. Hybridization was performed at 65°C in Rapid-Hyb buffer (Amersham) following the manufacturer's protocol.

Crosses and inheritance tests

Wild type and Nt-pIpetL plants were transferred to soil and grown to maturity under greenhouse conditions. Seed pods were collected from selfed plants and from reciprocal crosses of the Nt-pIpetL transplastomic lines with wild-type plants. Surface-sterilized seeds were germinated on MS medium with spectinomycin (500 mg/l) and analyzed for uniparental inheritance of the petL-disrupting resistance gene. Selfed transformants and crosses with a plastid transformant as the maternal parent give rise to resistant (green) progeny, whereas seeds collected from wild-type plants yield sensitive (white) seedlings.

RESULTS AND DISCUSSION

Phylogeny of mRNA editing in petL

Most genes encoded in the tobacco plastid genome specify photosynthetic proteins and constituents of the plastid gene expression machinery. petL is one of the smallest plastid genes (with only 31 codons for amino acids in tobacco), whose protein product was discovered to be a subunit of the cytochrome b6f complex by reverse genetics in Chlamydomonas reinhardtii (22). Insertional inactivation of petL produced algal strains whose photoautotrophic growth was severely impaired due to drastically reduced amounts of the cytochrome b6f complex, the redox-coupling multiprotein complex that connects the two photosystems. Expectedly, reduced abundance of the cytochrome b6f complex in the petL knockout strains was associated with a much slower photosynthetic electron transfer rate (22).

Upon analyzing the cytochrome b6f complex from spinach and tobacco (23) we discovered an RNA editing site in the spinach petL transcript (site 3 in Figure Figure1).1). This editing event changes a genomically encoded TCT serine codon into a UUU phenylalanine codon at the mRNA level. The phenylalanine residue in this position of the PetL protein appeared not to be very well conserved insofar as the closely related Chenopodiaceae species Beta vulgaris as well as the gymnosperm species Pinus thunbergii (24) have a CCT codon in this position. Interestingly, analysis of petL editing in P.thunbergii (24) revealed that this CCT proline codon does not undergo RNA editing. In order to trace the origin of the RNA editing activity at this site, we performed a phylogenetic analysis of RNA editing in petL. We included representatives from most major clades of seed plants and compared the petL DNA sequences with the corresponding cDNA sequences (Figure (Figure1)1) to detect sites of C-to-U editing. As the intergenic spacer downstream of the petL gene turned out not to be very well conserved, in most cases, a 3′ primer derived from the downstream petG gene was used for DNA amplification followed by design of species-specific primers binding in the 3′-UTR of the petL transcript for cDNA amplification.

Figure 1
Alignment of petL nucleotide sequences from the higher plant species analyzed with the petL gene from the liverwort Marchantia polymorpha (as a reference species that lacks RNA editing). Start and stop codons are indicated in bold. Sites of C-to-U RNA ...

Altogether this systematic analysis identified nine RNA editing sites, with most species harboring two or three sites (Figure (Figure1).1). Considering the small size of petL, this is the highest editing frequency detected to date in a plastid transcript from higher plants. Our phylogenetic data also revealed numerous other interesting features, including an editing site immediately upstream of the petL start codon [which is notable because editing outside coding regions is very rare; (10)] and a dynamic pattern of gains and losses of editing sites. Only a few selected aspects are discussed here in some detail. The evolution of site 2, creating the start codon for petL translation in gymnosperms, can most easily be explained with a single acquisition event followed by loss of the site due to a genomic C-to-T mutation in the lineage giving rise to the angiosperms (Figure (Figure1).1). A similarly simple scenario can be envisaged for site 8, creating the stop codon in Pinus species but not in the other gymnosperms analyzed here, indicating that, in this case, the site was either acquired in the genus Pinus or lost from the Abies/Larix lineage. In contrast, site 3 shows a much more dynamic evolutionary pattern involving at least four distinct loss events and several changes of the codon identity with altogether five different amino acids residing in this position in the species analyzed here. Interestingly, the CCT codon is not edited in pine and larch, indicating that editing may have evolved only in the angiosperm lineage (Figure (Figure1).1). This is remarkable in that the editable site appeared long before the evolution of the editing activity, suggesting that low functional constraints allowed for the long persistence of the site in the absence of correction by RNA editing.

When we aligned the amino acid sequences of the edited coding regions (data not shown), PetL turned out to be a well-conserved protein, as this has been seen previously for other small subunits of the photosynthetic apparatus [e.g. (23,10)].

The high frequency and dynamics of editing site evolution within petL was surprising to us because it appeared to contradict a seemingly straightforward hypothesis about the evolution of RNA editing sites which is discussed below.

A hypothesis for editing site evolution

One of the evolutionary enigmas surrounding RNA editing is the extremely uneven distribution of editing sites in the plastid genome. For example, in maize plastids [for which a comprehensive genome-wide survey of editing sites has been conducted; (25)], out of the 25 identified editing sites, 14 are located in just 3 genes: rpoB (4 sites), ndhA (4 sites) and ndhB (6 sites). In contrast, the almost 70 remaining protein-coding genes harbor altogether only 11 sites (25). It has been noted earlier that editing is most frequent in mRNAs of non-essential plastid genes (26). This is obviously the case for editing in the ndhA and ndhB genes that encode subunits of a plastid NAD(P)H dehydrogenase. Deletion of ndh genes from the tobacco plastid genome has no significant phenotypic consequences under standard growth conditions, but results in some measurable physiological effects when the knockout plants are exposed to various stress conditions (2731). The physiological data suggest that the Ndh protein complex participates in cyclic electron flow around photosystem I, a function that is compatible with the absence of significant phenotypic effects of ndh gene inactivation under normal growth conditions.

The situation is less obvious for RNA editing in rpoB, the gene encoding the β-subunit of the E.coli-like plastid-encoded RNA polymerase (PEP). Targeted inactivation of rpoB in tobacco plastid produces a severe non-photosynthetic phenotype (32), clearly indicating that loss of rpoB function is incompatible with photoautotrophic growth. However, closer inspection of the RNA editing pattern in the rpoB transcript reveals that the four editing sites in the maize rpoB transcript cluster in a domain that has been identified as so-called ‘dispensable region’: it is much less conserved than the rest of the RpoB amino acid sequence and, most importantly, can be deleted from the E.coli enzyme without any significant effect on the fidelity of the RNA polymerase (33,34,26).

Thus these three highly edited plastid sequences share a remarkable feature in that they are only of limited importance to plastid function and hence plant fitness. It is unlikely, though, that these editing events are entirely dispensable, because nearly all of them restore highly conserved amino acid residues (13,35,18). In the case of the ndh genes, this means that, although the Ndh gene products are non-essential under normal growth conditions, editing is most probably important for proper function of the Ndh protein complex (13,35), and hence of the cyclic electron transport, under stress conditions.

The evidence from the three cases of highly edited mRNAs discussed above may suggest that editing sites evolve more readily in genes (or domains of genes) whose transient loss of function can be tolerated. We will thereafter refer to this hypothesis as the ‘relative neutrality hypothesis for the evolution of plastid editing sites’ (Figure (Figure2).2). Fixation of a T-to-C mutation in a non-essential gene (or domain) is facilitated by the mutation's only slightly deleterious phenotypic effect (or neutrality under normal growth conditions). This in turn provides flexibility and sufficient time to evolve the mutation-compensating editing activity under only mild selective pressure.

Figure 2
The relative neutrality hypothesis for the evolution of editing sites. Owing to the polyploidy of the plastid genome, appearance of a mutation initially results in the simultaneous presence of wild-type and mutant genomes (heteroplasmy). Heteroplasmic ...

While the hypothesis provides a straightforward explanation for the extremely uneven distribution of plastid RNA editing sites, it self-evidently does not account for the evolution of every single editing site. Notably, it does not entirely exclude the evolution of sites in essential genes or domains. Instead, it just explains why such editing sites are much rarer. The hypothesis also does not make predictions about silent editing sites in third codon position or outside coding regions. One might expect, however, that such silent editing events are rare because of the absence of selection pressure to maintain the editing activity [(14); Figure Figure22].

The hypothesis can satisfactorily explain high-frequency editing in ndh genes and in rpoB, the three most highly edited plastid genes in higher plants (25). In addition, it makes testable predictions about the functions of plastid genes: one would expect genes (or domains of genes) with a high frequency of editing sites to be of limited functional importance, whereas genes encoding no or very few sites should be essential or at least make a significant contribution to plant fitness. In this respect, it was surprising to find that the small petL gene is highly edited and, in many species, even more highly edited than rpoB and the ndh genes. This seemed to contradict the relative neutrality hypothesis for the evolution of plastid editing sites since, extrapolating from the petL knockout experiments in Chlamydomonas (22), one would expect that a transitory loss of petL function (by mutations requiring correction through RNA editing) produces photosynthesis-deficient plants which would be rapidly eliminated by natural selection. Consequently, evolution would not be given enough time to acquire the editing activity necessary to counteract the mutation (Figure (Figure2).2). To experimentally test whether or not high-frequency petL editing disproves the relative neutrality hypothesis for the evolution of editing sites, we set out to determine the contribution of petL function to fitness in higher plants by reverse genetics.

Functional analysis of petL in tobacco

In order to test whether petL serves a similarly important function in higher plants as to the function it performs in C.reinhardtii, we produced an analogous petL knockout construct for targeted gene disruption (Figure (Figure3A)3A) as had been used in Chlamydomonas (22). Biolistic chloroplast transformation in tobacco yielded several transplastomic lines four of which were further characterized (Nt-pIpetL-3, Nt-pIpetL-14, Nt-pIpetL-18 and Nt-pIpetL-31). Homoplasmic petL disruption lines were isolated after subjecting the primary transformants to three additional rounds of regeneration under antibiotic selection. Correct integration of the selectable marker gene aadA into the petL reading frame was confirmed by PCR (data not shown) and Southern blot analyses (Figure (Figure3B).3B). Homoplasmy and maternal transgene transmission were ultimately verified by reciprocal crosses (Figure (Figure3C).3C). We were next interested in analyzing the phenotypic effects of petL disruption. As inactivation of a photosynthesis gene can produce a mutant phenotype in a light-dependent manner, we assayed wild-type and mutant plants under widely different light intensities (ranging from 4 to 400 μmol quanta per square meter per second). In no case did the mutant plants show any visible phenotype or growth retardation compared with wild-type plants (Figure (Figure3D3D and data not shown), suggesting that, at least under the conditions tested, the PetL protein makes no significant contribution to photosynthesis and plant fitness. It is noteworthy in this respect that petL is not present in the fully sequenced genome of the cyanobacterium Synechocystis (36), further supporting the idea that, at least in some photosynthetic organisms, the PetL protein is not critical for proper cytochrome b6f function. Its importance in Chlamydomonas may be due to structural differences among the cytochrome b6f complexes in algae, cyanobacteria and higher plants.

Figure 3
Targeted inactivation of petL in tobacco. (A) Map of the plastid transformation vector pIpetL used for disruption of petL. Genes above the line are transcribed from the right to the left; genes below the line are transcribed in the opposite direction. ...

In summary, we have shown that high-frequency editing in petL, one of the smallest plastid genes, coincides with a non-essential function of petL in higher plants, in contrast to the importance of petL to photosynthesis in green algae. Together with low-frequency editing or complete lack of editing in functionally important genes from higher plant plastids [e.g., (25,15,26,37)] and high-frequency editing in the non-essential ndh genes as well as dispensable domains of rpoB, these data lend support to the relative neutrality hypothesis for the evolution of RNA editing sites in higher plant plastids. With reverse genetics becoming a more routine technology for tobacco plastids, additional non-essential genes and/or domains are likely to be identified in the near future and to become available for analysis of RNA editing site evolution.

ACKNOWLEDGEMENTS

We thank H. Voigt, J. Hausfeld, M. Voss (Botanical Garden Münster) and R. Oberle (Botanical Garden Freiburg) for providing material from various plant species, Dr. W. Lobin (Botanical Garden Bonn) for providing plant material from Amborella trichopoda, Dr Stefanie Hartmann for help with vector construction and Professor Dr Focke Albers for stimulating discussion. This research was supported by grants from the Deutsche Forschungsgemeinschaft to R.B.

REFERENCES

1. Bock R. (2000) Sense from nonsense: how the genetic information of chloroplasts is altered by RNA editing. Biochimie, 82, 549–557. [PubMed]
2. Bock R. (2001) RNA editing in plant mitochondria and chloroplasts. In Bass,B. (ed.), Frontiers in Molecular Biology: RNA Editing. Oxford University Press, New York, NY, pp. 38–60.
3. Yoshinaga K., Iinuma,H., Masuzawa,T. and Ueda,K. (1996) Extensive RNA editing of U to C in addition to C to U substitution in the rbcL transcripts of hornwort chloroplasts and the origin of RNA editing in green plants. Nucleic Acids Res., 24, 1008–1014. [PMC free article] [PubMed]
4. Kugita M., Yamamoto,Y., Fujikawa,T., Matsumoto,T. and Yoshinaga,K. (2003) RNA editing in hornwort chloroplasts makes more than half the genes functional. Nucleic Acids Res., 31, 2417–2423. [PMC free article] [PubMed]
5. Gualberto J.M., Lamattina,L., Bonnard,G., Weil,J.-H. and Grienenberger,J.-M. (1989) RNA editing in wheat mitochondria results in the conservation of protein sequences. Nature, 341, 660–662. [PubMed]
6. Covello P.S. and Gray,M.W. (1989) RNA editing in plant mitochondria. Nature, 341, 662–666. [PubMed]
7. Hiesel R., Wissinger,B., Schuster,W. and Brennicke,A. (1989) RNA editing in plant mitochondria. Science, 246, 1632–1634. [PubMed]
8. Hanson M.R., Sutton,C.A. and Lu,B. (1996) Plant organelle gene expression: altered by RNA editing. Trends Plant Sci., 1, 57–64.
9. Hirose T., Fan,H., Suzuki,J.Y., Wakasugi,T., Tsudzuki,T., Kössel,H. and Sugiura,M. (1996) Occurrence of silent RNA editing in chloroplasts: its species specificity and the influence of environmental and developmental conditions. Plant Mol. Biol., 30, 667–672. [PubMed]
10. Kudla J. and Bock,R. (1999) RNA editing in an untranslated region of the Ginkgo chloroplast genome. Gene, 234, 81–86. [PubMed]
11. Hoch B., Maier,R.M., Appel,K., Igloi,G.L. and Kössel,H. (1991) Editing of a chloroplast mRNA by creation of an initiation codon. Nature, 353, 178–180. [PubMed]
12. Kudla J., Igloi,G.L., Metzlaff,M., Hagemann,R. and Kössel,H. (1992) RNA editing in tobacco chloroplasts leads to the formation of a translatable psbL mRNA by a C to U substitution within the initiation codon. EMBO J., 11, 1099–1103. [PMC free article] [PubMed]
13. Maier R.M., Hoch,B., Zeltz,P. and Kössel,H. (1992) Internal editing of the maize chloroplast ndhA transcript restores codons for conserved amino acids. Plant Cell, 4, 609–616. [PMC free article] [PubMed]
14. Bock R., Kössel,H. and Maliga,P. (1994) Introduction of a heterologous editing site into the tobacco plastid genome: the lack of RNA editing leads to a mutant phenotype. EMBO J., 13, 4623–4628. [PMC free article] [PubMed]
15. Freyer R., Kiefer-Meyer,M.-C. and Kössel,H. (1997) Occurrence of plastid RNA editing in all major lineages of land plants. Proc. Natl Acad. Sci. USA, 94, 6285–6290. [PMC free article] [PubMed]
16. Steinhauser S., Beckert,S., Capesius,I., Malek,O. and Knoop,V. (1999) Plant mitochondrial RNA editing—extreme in hornworts and dividing the liverworts? J. Mol. Evol., 48, 303–312. [PubMed]
17. Freyer R., Lopez,C., Maier,R.M., Martin,M., Sabater,B. and Kössel,H. (1995) Editing of the chloroplast ndhB encoded transcript shows divergence between closely related members of the grass family (Poaceae). Plant Mol. Biol., 29, 679–684. [PubMed]
18. Zeltz P., Hess,W.R., Neckermann,K., Börner,T. and Kössel,H. (1993) Editing of the chloroplast rpoB transcript is independent of chloroplast translation and shows different patterns in barley and maize. EMBO J., 12, 4291–4296. [PMC free article] [PubMed]
19. Murashige T. and Skoog,F. (1962) A revised medium for rapid growth and bio assays with tobacco tissue culture. Physiol. Plant., 15, 473–497.
20. Doyle J.J. and Doyle,J.L. (1990) Isolation of plant DNA from fresh tissue. Focus, 12, 13–15.
21. Svab Z. and Maliga,P. (1993) High-frequency plastid transformation in tobacco by selection for a chimeric aadA gene. Proc. Natl Acad. Sci. USA, 90, 913–917. [PMC free article] [PubMed]
22. Takahashi Y., Rahire,M., Breyton,C., Popot,J.-L., Joliot,P. and Rochaix,J.-D. (1996) The chloroplast ycf7 (petL) open reading frame of Chlamydomonas reinhardtii encodes a small functionally important subunit of the cytochrome b6f complex. EMBO J., 15, 3498–3506. [PMC free article] [PubMed]
23. Hager M., Biehler,K., Illerhaus,J., Ruf,S. and Bock,R. (1999) Targeted inactivation of the smallest plastid genome-encoded open reading frame reveals a novel and essential subunit of the cytochrome b6f complex. EMBO J., 18, 5834–5842. [PMC free article] [PubMed]
24. Wakasugi T., Hirose,T., Horihata,M., Tsudzuki,T., Kössel,H. and Sugiura,M. (1996) Creation of a novel protein-coding region at the RNA level in black pine chloroplasts: The pattern of RNA editing in the gymnosperm chloroplast is different from that in angiosperms. Proc. Natl Acad. Sci. USA, 93, 8766–8770. [PMC free article] [PubMed]
25. Maier R.M., Neckermann,K., Igloi,G.L. and Kössel,H. (1995) Complete sequence of the maize chloroplast genome: gene content, hotspots of divergence and fine tuning of genetic information by transcript editing. J. Mol. Biol., 251, 614–628. [PubMed]
26. Corneille S., Lutz,K. and Maliga,P. (2000) Conservation of RNA editing between rice and maize plastids: are most editing events dispensable? Mol. Gen. Genet., 264, 419–424. [PubMed]
27. Burrows P.A., Sazanov,L.A., Svab,Z., Maliga,P. and Nixon,P.J. (1998) Identification of a functional respiratory complex in chloroplasts through analysis of tobacco mutants containing disrupted plastid ndh genes. EMBO J., 17, 868–876. [PMC free article] [PubMed]
28. Sazanov L.A., Burrows,P.A. and Nixon,P.J. (1998) The chloroplast Ndh complex mediates the dark reduction of the plastoquinone pool in response to heat stress in tobacco leaves. FEBS Lett., 429, 115–118. [PubMed]
29. Endo T., Shikanai,T., Takabayashi,A., Asada,K. and Sato,F. (1999) The role of chloroplastic NAD(P)H dehydrogenase in photoprotection. FEBS Lett., 457, 5–8. [PubMed]
30. Horváth E.M., Peter,S.O., Joet,T., Rumeau,D., Cournac,L., Horváth,G., Kavanagh,T.A., Schäfer,C. and Medgyesy,P. (2000) Targeted inactivation of the plastid ndhB gene in tobacco results in an enhanced sensitivity of photosynthesis to moderate stomatal closure. Plant Physiol., 123, 1337–1349. [PMC free article] [PubMed]
31. Li X.-G., Duan,W., Meng,Q.-W., Zou,Q. and Zhao,S.-J. (2004) The function of chloroplastic NAD(P)H dehydrogenase in tobacco during chilling stress under low irradiance. Plant Cell. Physiol., 45, 103–108. [PubMed]
32. Allison L.A., Simon,L.D. and Maliga,P. (1996) Deletion of rpoB reveals a second distinct transcription system in plastids of higher plants. EMBO J., 15, 2802–2809. [PMC free article] [PubMed]
33. Severinov K., Kashlev,M., Severinova,E., Bass,I., McWilliams,K., Kutter,E., Nikiforov,V., Snyder,L. and Goldfarb,A. (1994) A non-essential domain of Escherichia coli RNA polymerase required for the action of the termination factor Alc. J. Biol. Chem., 269, 14254–14259. [PubMed]
34. Borukhov S., Severinov,K., Kashlev,M., Lebedev,A., Bass,I., Rowland,G.C., Lim,P.P., Glass,R.E., Nikiforov,V. and Goldfarb,A. (1991) Mapping of trypsin cleavage and antibody-binding sites and delineation of a dispensable domain in the beta subunit of Escherichia coli RNA polymerase. J. Biol. Chem., 266, 23921–23926. [PubMed]
35. Maier R.M., Neckermann,K., Hoch,B., Akhmedov,N.B. and Kössel,H. (1992) Identification of editing positions in the ndhB transcript from maize chloroplasts reveals sequence similarities between editing sites of chloroplasts and plant mitochondria. Nucleic Acids Res., 20, 6189–6194. [PMC free article] [PubMed]
36. de las Rivas J., Lozano,J.J. and Ortiz,A.R. (2002) Comparative analysis of chloroplast genomes: Functional annotation, genome-based phylogeny, and deduced evolutionary patterns. Genome Res., 12, 567–583. [PMC free article] [PubMed]
37. Tsudzuki T., Wakasugi,T. and Sugiura,M. (2001) Comparative analysis of RNA editing sites in higher plant chloroplasts. J. Mol. Evol., 53, 327–332. [PubMed]
38. Judd W.S., Campbell,C.S., Kellogg,E.A., Stevens,P.F. and Donoghue,M.J. (2002) Plant Systematics: A Phylogenetic Approach, 2nd edn. Sinauer Associates, Inc., Sunderland, MA.
39. Davies T.J., Barraclough,T.G., Chase,M.W., Soltis,P.S. and Soltis,D.E. (2004) Darwin's abominable mystery: insights from a supertree of the angiosperms. Proc. Natl Acad. Sci. USA, 101, 1904–1909. [PMC free article] [PubMed]
40. Bock R. (2001) Transgenic chloroplasts in basic research and plant biotechnology. J. Mol. Biol., 312, 425–438. [PubMed]

Articles from Nucleic Acids Research are provided here courtesy of Oxford University Press
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...