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Keywords:

  • chloroplasts;
  • evolution;
  • mitochondria;
  • pentatricopeptide repeat;
  • RNA editing

Abstract

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Expansion of the PPR family
  5. III. RNA editing and the evolution of PPR genes
  6. IV. Sequence drift in the organelle genomes of early land plants
  7. V. Loss of RNA editing in angiosperms
  8. VI. Other functions besides editing
  9. VII. Conclusion
  10. References

Contents

 Summary37
I.Introduction37
II.Expansion of the PPR family38
III.RNA editing and the evolution of PPR genes40
IV.Sequence drift in the organelle genomes of early land plants41
V.Loss of RNA editing in angiosperms43
VI.Other functions besides editing44
VII.Conclusion44
 References44

Summary

The pentatricopeptide repeat (PPR) is a degenerate 35-amino-acid structural motif identified from analysis of the sequenced genome of the model plant Arabidopsis thaliana. From the wealth of sequence information now available from plant genomes, the PPR protein family is now known to be one of the largest families in angiosperm species, as most genomes encode 400–600 members. As the number of PPR genes is generally only c. 10–20 in other eukaryotic organisms, including green algae, the family has obviously greatly expanded during land plant evolution. This provides a rare opportunity to study selection pressures driving a 50-fold expansion of a single gene family. PPR proteins are sequence-specific RNA-binding proteins involved in many aspects of RNA processing in organelles. In this review, we will summarize our current knowledge about the evolution of PPR genes, and will discuss the relevance of the dramatic expansion in the family to the functional diversification of plant organelles, focusing primarily on RNA editing.


I. Introduction

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Expansion of the PPR family
  5. III. RNA editing and the evolution of PPR genes
  6. IV. Sequence drift in the organelle genomes of early land plants
  7. V. Loss of RNA editing in angiosperms
  8. VI. Other functions besides editing
  9. VII. Conclusion
  10. References

Organellar function is essential for eukaryotic life, and depends upon activities that are maintained by proteins either internally synthesized within the organelle (organelle-encoded) or imported from cytosol (nucleus-encoded). Many of these functions are carried out by enzyme complexes with both organelle-encoded and nucleus-encoded subunits, and therefore coordination of the expression of organelle and nuclear genes is a critical matter. For example, the plant mitochondrial proteome can be estimated to consist of c. 2000 gene products (Millar et al., 2005). Of these, only c. 40 proteins are encoded within the mitochondrial genome, most of which encode essential subunits of oxidative phophorylation enzymatic complexes and ribosomal proteins (Kubo et al., 2000; Notsu et al., 2002; Handa, 2003; Ogihara et al., 2005; Sugiyama et al., 2005; Tian et al., 2006; Allen et al., 2007; Kubo & Newton, 2008; Fujii et al., 2010). Although these represent only a small proportion of all mitochondrial proteins, their importance to mitochondrial function means that incorrect regulation of these mitochondrial genes would severely affect the whole system. As plant mitochondria encode no machinery to manage their own RNA expression and post-transcriptional RNA modification processes, these essential steps are totally reliant on nuclear-encoded gene products (Binder & Brennicke, 2003).

RNA metabolism plays a particularly important role in organelle gene expression (Stern et al., 2010) and a wide array of different RNA binding proteins are found in organelles. Pentatricopeptide repeat (PPR) proteins are the most numerous of these. The first PPR protein to be described was the Saccharomyces cerevisiae mitochondrial protein Pet309, found to participate in translation of cox1 (Manthey & McEwen, 1995; Manthey et al., 1998; Tavares-Carreon et al., 2008). Subsequently, Pet309, as well as the protein P67 implicated in transcription in Triticum aestivum mitochondria (Ikeda & Gray, 1999) and CRP1 in Zea mays involved in translation of photosynthesis genes (Fisk et al., 1999; Schmitz-Linneweber et al., 2005), were recognized to be members of a large family of related proteins following the systematic analysis of the Arabidopsis thaliana (thale cress) genome (Aubourg et al., 2000; Small & Peeters, 2000). Since the discovery and definition of the PPR consensus sequence (Small & Peeters, 2000), predicted to form an antiparallel double alpha-helical motif, many studies have been conducted on PPR proteins covering biochemistry, molecular functions, cellular functions and roles in development (Schmitz-Linneweber & Small, 2008). The general picture that results from these studies is that PPR proteins form sequence-specific associations with RNA, and that these associations affect folding, processing and/or translation of the RNA, thus manipulating expression of the transcript. For example, CRP1 in maize is shown to associate with the 5′UTR region of photosynthetic genes psaC and petA, and mutants disrupted in CRP1 gene lack translation of these genes, leading to the defects in photosynthesis (Fisk et al., 1999; Schmitz-Linneweber et al., 2005). Arabidopsis CRR4 was the first gene found in plants to be directly involved in cytosine to uridine RNA editing (Kotera et al., 2005), since followed by many other PPR editing factors (Table 1). These are only a couple of examples, and for more discussion of the molecular functions of PPR proteins readers are redirected to reviews elsewhere that treat this topic comprehensively (Andres et al., 2007; Delannoy et al., 2007; Saha et al., 2007; Schmitz-Linneweber & Small, 2008; Chateigner-Boutin & Small, 2010). The PPR family is subdivided into two major classes, P and PLS. Whereas P-class PPR proteins consist of an orthodox tandem alignment of 35-amino-acid PPR (P) motifs, PLS-class proteins contain, in addition, slightly longer (L) or shorter (S) variant PPR motifs in tandem arrays of characteristic triplets, P-L-S (Lurin et al., 2004; Rivals et al., 2006; O’Toole et al., 2008). PLS-class proteins can be divided into two further groups, the E subclass and the DYW subclass, based on their C-terminal domains (Lurin et al., 2004; Rivals et al., 2006; O’Toole et al., 2008).

Table 1.   Known editing factors from plant organelles
OrganelleAGISubclassNameEdited geneReference
ChloroplastAt1g08070DYWOTP82ndhG, ndhBHammani et al. (2009)
ChloroplastAt1g11290DYWCRR22ndhB, ndhD, rpoBOkuda et al. (2009)
ChloroplastAt1g15510DYWAtECB2/VAC1accDYu et al. (2009), Tseng et al. (2010)
ChloroplastAt1g59720DYWCRR28ndhB, ndhDOkuda et al. (2009)
ChloroplastAt2g02980DYWOTP85ndhDHammani et al. (2009)
ChloroplastAt2g29760DYWOTP81rps12 intronHammani et al. (2009)
ChloroplastAt3g22690DYWYS1rpoBZhou et al. (2008)
ChloroplastAt3g57430DYWOTP84ndhF, psbZ, ndhBHammani et al. (2009)
ChloroplastAt3g63370DYWOTP86rps14Hammani et al. (2009)
ChloroplastAt5g13270DYWRARE1accDRobbins et al. (2009)
ChloroplastAt5g48910DYWLPA66psbFCai et al. (2009)
ChloroplastAt2g45350ECRR4ndhDKotera et al. (2005)
ChloroplastAt5g59200EOTP80rpl23Hammani et al. (2009)
ChloroplastAt1g05750ECLB19rpoA, clpPChateigner-Boutin et al. (2008)
ChloroplastAt5g55740ECRR21ndhDOkuda et al. (2007)
MitochondriaAt2g03880DYWREME1nad2, orfXBentolila et al. (2010)
MitochondriaAt2g25580DYWMEF8nad5Takenaka et al. (2010)
MitochondriaAt3g12770DYWMEF22nad3Takenaka et al. (2010)
MitochondriaAt4g14850DYWLOI1/MEF11cox3, nad4, ccb203Tang et al. (2010), Verbitskiy et al. (2010)
MitochondriaAt5g52630DYWMEF1rps4, nad7, nad2Zehrmann et al. (2009)
MitochondriaOryza sativaDYWOGR1cox2, cox3, ccmC, nad2, nad4Kim et al. (2009)
MitochondriaPhyscomitrella patensDYWPpPPR_56nad3, nad4Ohtani et al. (2010)
MitochondriaPhyscomitrella patensDYWPpPPR_71ccmFTasaki et al. (2010)
MitochondriaPhyscomitrella patensDYWPpPPR_77cox2, cox3Ohtani et al. (2010)
MitochondriaPhyscomitrella patensDYWPpPPR_91nad5Ohtani et al. (2010)
MitochondriaAt1g62260EMEF9nad7Takenaka (2010)
MitochondriaAt2g22410ESLO1nad4, nad9Sung et al. (2010)
MitochondriaAt3g05240EMEF19ccb206Takenaka et al. (2010)
MitochondriaAt3g18970EMEF20rps4Takenaka et al. (2010)
MitochondriaAt5g19020EMEF18nad4Takenaka et al. (2010)
MitochondriaAt2g20540EMEF21cox3Takenaka et al. (2010)

The aim of this review is to integrate our current knowledge about the evolution of the PPR gene family and connect this with the changes thought to have taken place in the sequence and expression of the organelle genomes during the history of land plants. Given the rapid expansion of PPR proteins in land plants, there is an interest in understanding what selective forces might have operated on them to increase their numbers. Recent progress in genome sequencing and the consequent enrichment of comparative genomic databases are starting to allow us to understand how and why the plant-specific expansion of the PPR family occurred.

II. Expansion of the PPR family

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Expansion of the PPR family
  5. III. RNA editing and the evolution of PPR genes
  6. IV. Sequence drift in the organelle genomes of early land plants
  7. V. Loss of RNA editing in angiosperms
  8. VI. Other functions besides editing
  9. VII. Conclusion
  10. References

Outside land plants, the organism with largest set of known PPR genes is the parasitic protozoan Trypanosoma brucei, commonly known as the causative agent for sleeping sickness, with 28 PPR members (Pusnik et al., 2007). This compares with the 450 and 477 PPR genes in Arabidopsis thaliana and Oryza sativa (rice), model species for eudicots and monocots, respectively (Lurin et al., 2004; O’Toole et al., 2008). The green alga Chlamydomonas reinhardtii (Merchant et al., 2007) contains only 12 PPR genes, whilst the moss Physcomitrella patens (Rensing et al., 2008) contains 103 (O’Toole et al., 2008; Schmitz-Linneweber & Small, 2008). These data suggest that the initial expansion of PPR genes took place after the separation of the land plant lineage from green algae such as Chlamydomonas but before the divergence of seed plants from bryophytes.

At present, sufficient information is available to conduct whole-genome comparative genomic studies for 18 plant species (from websites such as Phytozome: http://www.phytozome.net/). The distribution of PPR genes in these species is shown in Fig. 1. It is now certain that a wide range of angiosperms belonging to the core-eudicots or monocots contain a very large number of PPR genes, and that considering variations in genome size and total gene content between these species, the number of PPR genes is relatively constant. The least number of PPR genes is found in Medicago truncatula (365) and the most in Glycine max (629). The partially assembled Selaginella moellendorffii (spikemoss, lycophyte) genome contains > 1000 PPR genes (Fig. 1), although the numbers given here should be treated as preliminary and are likely to change with improved coverage and gene annotation.

image

Figure 1. Numbers of pentatricopeptide repeat (PPR) genes and plastid RNA editing sites in various plant species. Left histogram: light bars, P-class PPR genes; dark bars, PLS-class PPR genes. Right histogram: *, number of RNA editing sites detected from expressed sequence tags (Smith, 2009); **, number of RNA editing sites predicted by PREP (Mower, 2009).

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The mechanism of expansion is likely to have involved retrotransposition. As many as 80% of the PPR genes in Arabidopsis and Oryza are intron-less (Lurin et al., 2004; Rivals et al., 2006; O’Toole et al., 2008), whereas > 75% of P. patens PPR genes contain introns. Evolutionarily older PPR genes in rice and A. thaliana contain more introns (O’Toole et al., 2008) and some of the putative orthologues to the P. patens intron-containing genes lack introns in angiosperms (O’Toole et al., 2008). These data are consistent with amplification by retrotransposition.

Gene families can also increase in size by genome duplication events. Angiosperm genomes have undergone multiple large genome duplication events (Bowers et al., 2003; De Bodt et al., 2005). These duplication events have large effects on gene number in gene families that are already present in large numbers. However, as shown in Fig. 2, abundant genes containing short repeat motifs such as ankyrin, tetratricopeptide repeat (TPR) and WD40 motifs have not increased in number during the evolution of land plants, implying that, for these families, gene loss has balanced the creation of new genes by duplication. This contrasts with the large increase in number of leucine-rich repeat (LRR) genes and PPR genes (Fig. 2). This suggests there may have been some form of selection for retention of duplicated LRR and PPR genes early in land plant evolution. On the other hand, it was shown that PPR gene content is well conserved between A. thaliana and Oryza (O’Toole et al., 2008) and that the PPR genes duplicated in independent whole-genome duplication events occurring in these angiosperm lineages were rarely retained. This suggests that whatever the selective forces acting to increase PPR gene number in early land plant evolution, they appear no longer to act in this way in extant angiosperms.

image

Figure 2. Numbers of short tandem repeat motif genes present in plant taxa. The pentatricopeptide repeat (PPR) family has expanded more than other families of structurally similar genes that were probably present in greater numbers in the ancestors of land plants. TPR, tetratricopeptide repeat; LRR, leucine-rich repeat; PPR, pentatricopeptide repeat.

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In the case of LRR genes, their mode of action in the defence against pathogens can explain selection for increasing diversity. What might have caused selective pressures acting to retain large numbers of PPR genes following retrotransposition or genome duplications? In the next two sections, we will explore the growth of PPR family, focusing on the connection to one of its main molecular functions, RNA editing.

III. RNA editing and the evolution of PPR genes

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Expansion of the PPR family
  5. III. RNA editing and the evolution of PPR genes
  6. IV. Sequence drift in the organelle genomes of early land plants
  7. V. Loss of RNA editing in angiosperms
  8. VI. Other functions besides editing
  9. VII. Conclusion
  10. References

RNA editing in plant mitochondria or chloroplasts commonly involves post-transcriptional substitution of cytidine by uridine (C-to-U) or, less commonly, uridine by cytidine (Gray, 1996, 2009; Maier et al., 1996; Mulligan et al., 1999; Shikanai, 2006; Takenaka et al., 2008). Many RNA editing events play important roles in gene expression by generating start codons at ACG sites, correcting codons to encode conserved amino acids and generating required stop codons.

Beginning with the identification of CRR4 (Kotera et al., 2005), many PPR proteins have been identified as factors required for C-to-U RNA editing in chloroplasts (summarized in Table 1). All of these PPR proteins are members of the plant-specific PLS class, one of two major subgroups within the PPR family. A possible exception is the Arabidopsis P-class protein PPR596, which acts to suppress RNA editing in mitochondrial rps3 transcripts (Doniwa et al., 2010). It remains to be seen whether PPR596 acts in a mechanistically similar way to other editing factors or whether the effect on editing is an indirect result of alterations in RNA processing or turnover.

The majority of PLS-class members may be dedicated to RNA editing, given the large numbers of editing mutants affected in expression of these genes (Table 1). So far, the only exceptions are the DYW-subclass protein CRR2 involved in RNA cleavage of a chloroplast transcript (Hashimoto et al., 2003; Okuda et al., 2009) and the E-subclass protein OTP70 involved in splicing of chloroplast rpoC1 transcripts (Chateigner-Boutin et al., 2011). The presence of DYW domains is strictly correlated in plant evolution with the presence of RNA editing (Salone et al., 2007; Rudinger et al., 2008). Marchantia polymorpha, one of the earliest branching land plants, lacks both DYW-subclass PPR genes and RNA editing, although the molecular phylogeny suggests it is likely that Marchantia has lost these properties rather than never had them (Groth-Malonek et al., 2007; Salone et al., 2007; Rudinger et al., 2008). The origin of the DYW domain is unclear but there is a possibility that horizontal gene transfer has taken place between a plant and a heterolobosean protist (Knoop & Rudinger, 2010). PLS-class PPR proteins had not previously been found outside the plant kingdom, and even within it, they are not present in algae (O’Toole et al., 2008; Knoop & Rudinger, 2010). However, the recently sequenced genome of a protist species, Naegleria gruberi (Fritz-Laylin et al., 2010), contains 12 PLS-class PPR genes, with 10 of them carrying a DYW domain (Knoop & Rudinger, 2010). Additional investigation is required to map the phylogenetic distribution of these proteins in protists and settle the direction of any gene transfer.

Both RNA editing (of the type found in plant organelles) and PLS-class PPR genes first appeared in the earliest land plants. The number of C-to-U RNA editing sites in organelles is variable between plants, ranging from only one in Physcomitrella chloroplasts to > 500 in the chloroplasts of the bryophyte Anthoceros, and even more in most land plant mitochondria (Fig. 1) (Kugita et al., 2003; Tillich et al., 2006; Jobson & Qiu, 2008). Unfortunately, comprehensive RNA editing information and complete nuclear genome sequences (i.e. information on PPR gene numbers) only rarely coexist. Species with both sets of data available are limited to Physcomitrella, Selaginella, Oryza, Zea (maize) and Arabidopsis (Fig. 1). Only 10 organellar RNA editing sites are known in P. patens (Miyata & Sugita, 2004; Tasaki & Sugita, 2010) and 103 PPR genes (including 10 DYW-subclass PPR genes; Fig. 1). There are at least 104 RNA editing sites in S. moellendorffii chloroplasts (Smith, 2009), many more than in angiosperms (20–40 sites). Mitochondrial RNA editing information is not available for Selaginella; however, a high number of RNA editing sites is expected from the extremely high GC content of the mitochondrial genome in this species because of the positive correlation between GC content and RNA editing (Malek et al., 1996; Smith, 2009). This species is estimated to possess over 1000 PPR genes, whereas angiosperms, Oryza, Zea and Arabidopsis possess c. 450 (Fig. 1). Furthermore, it is the number of PLS-class PPR genes that correlates best (Pearson’s correlation = 0.96) with the number of editing sites (Fig. 1) rather than the P subclass (Pearson’s correlation = 0.5), in accordance with the mechanistic links between PLS-class proteins and RNA editing. The correlation between numbers of RNA editing sites and number of PPR genes implies that any selective pressures acting to increase RNA editing in early land plants would also have selected for increasing numbers of PPR genes.

IV. Sequence drift in the organelle genomes of early land plants

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Expansion of the PPR family
  5. III. RNA editing and the evolution of PPR genes
  6. IV. Sequence drift in the organelle genomes of early land plants
  7. V. Loss of RNA editing in angiosperms
  8. VI. Other functions besides editing
  9. VII. Conclusion
  10. References

Algal organelle genomes are generally AT-rich, including those of the nearest relatives to the land plant lineage (Smith, 2009). By contrast, many land plant plastid genomes are relatively GC-rich (see Fig. 3), implying a sustained sequence drift over hundreds of millions of years. Phylogenetic reconstructions show that the rate of genomic T[RIGHTWARDS ARROW]C transitions must have far exceeded the rate of reverse mutations during this long period (Fig. 4). The potential link with RNA editing is evident, given that it reverses these DNA mutations at the RNA level. Indeed, several studies have conclusively shown that the number of RNA editing sites is strongly positively correlated with genome GC content (Malek et al., 1996; Smith, 2009; Yura et al., 2009). The reason for this sustained mutation bias is not clear. One explanation that has been suggested is that increased exposure to DNA-damaging UV light in land plants might have led to a relative suppression of adjacent pyrimidines (especially adjacent thymines) as a result of the formation of cyclobutane pyrimidine dimers and other photoproducts that can block transcription or replication, and which can be mutagenic (Singer & Ames, 1970; Maier et al., 1996; Rozema et al., 2002; Yura et al., 2009). The support for this in extant sequences is weak, as judged by strand-specific dinucleotide frequencies (Fig. 4), but it remains a plausible hypothesis for changes in early land plants when UV exposure was probably considerably higher, because of both lower stratospheric ozone concentrations and the fact that land plants had not yet evolved the full set of UV-protecting traits possessed by extant plants (Takahashi & Ohnishi, 2004; Hader & Sinha, 2005).

image

Figure 3. Proportion of GC nucleotides at the third codon position in chloroplast genomes. Coding sequences from green algal and land plant chloroplast genomes were aligned. The nucleotide at the third position of each aligned codon was scored; the proportion of G + C is shown for each group. The standard deviation of the values for each group is indicated by a horizontal line. The number of genomes in each group is as follows: monocots (27), eudicots (79), basal dicots (10), conifers (10), gnetophytes (three), ferns (seven), lycophytes (three), streptophyte algae (six), chlorophyte algae (20).

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image

Figure 4. Genome sequence trends during land plant evolution. The topology of the dendrogram was reconstructed from the concatenated multiple alignment of 51 chloroplast genes deduced from 104 plant species, and by using the maximum-likelihood method (Stamatakis, 2006). Numbers of synonymous substitutions (dS) were estimated by PAML (Yang, 2007), and absolute branch length was calculated using the molecular divergence estimation program r8s (Sanderson, 2003). Divergence time of the root was fixed as 1200 million yr ago (MYA), as described in Lang et al. (2008), and the minimum divergence time of angiosperms was set as 132 MYA from the oldest fossil record (Crane et al., 1995). Magenta branches indicate when the T-to-C substitution rate was more than twice that of the C-to-T substitution rate, and green edges indicate when the C-to-T substitution rate was more than twice that of T-to-C substitutions. The histogram on the left indicates the number of RNA editing sites predicted by PREP (Mower, 2009) for each extant species. The histograms in the middle and to the right show the proportion of thymines in TT pairs within the sense strand or antisense (transcribed strand) of 51 chloroplast genes.

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An important point when considering the evolution of RNA editing is that phylogenetic reconstructions show very little long-term conservation of editing sites. There is little overlap between those present in extant ferns, mosses, spikeworts or angiosperms (Freyer et al., 1995, 1997; Malek et al., 1996; Tillich et al., 2009). This has two ramifications. First, it means we are unable to state with confidence how much RNA editing was present in the common ancestors of these various lineages, as almost all the extant sites arose after the lineages split. Second, it shows that many land plant lineages continue to show a high rate of appearance of new editing sites.

V. Loss of RNA editing in angiosperms

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Expansion of the PPR family
  5. III. RNA editing and the evolution of PPR genes
  6. IV. Sequence drift in the organelle genomes of early land plants
  7. V. Loss of RNA editing in angiosperms
  8. VI. Other functions besides editing
  9. VII. Conclusion
  10. References

Over 100 chloroplast editing sites are known in early-branching land plants such as Anthoceros, Alsophila, Adiantum or Selaginella (Freyer et al., 1997; Kugita et al., 2003; Wolf et al., 2004; Tillich et al., 2006; Smith, 2009), but fewer sites are edited in angiosperm chloroplast RNAs. From this, several authors have proposed that angiosperm chloroplasts are losing editing sites faster than they gain new ones (Freyer et al., 1997; Tillich et al., 2006; Jobson & Qiu, 2008). To demonstrate this, we have plotted the number of predicted RNA editing sites in each species against its evolutionary distance from the common ancestor of all angiosperms (Fig. 5). The two parameters are clearly anticorrelated, implying a steady overall loss of RNA editing sites in angiosperm chloroplasts. By using known divergence times to estimate phylogenetic branch length, an RNA editing site is estimated to be lost in 2.43 million yr, on average (Fig. 5). This loss of RNA editing sites is again linked to a concomitant change in DNA mutation rates; in angiosperm plastids, the rate of C[RIGHTWARDS ARROW]T transitions exceeds that of T[RIGHTWARDS ARROW]C transitions, a reversal of the situation in early land plants (Fig. 4).

image

Figure 5. Loss of RNA editing during angiosperm evolution. The vertical axis plots the number of plastid RNA editing sites predicted in each angiosperm species. The horizontal axis shows the cumulative edge length (number of synonymous substitutions per site, dS) from the common ancestor of angiosperms to each extant angiosperm species.

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An interesting question is whether the RNA editing process is advantageous to the plant, or whether, on the contrary, it carries an associated cost. Advantages might include permitting a finer control over gene expression or providing a form of ‘heterosis’ by permitting the production of proteins with slightly different sequence from the same gene. Costs include the cost of producing the editing machinery (i.e. large numbers of PPR proteins) and the cost of producing and disposing of incorrect proteins translated from incompletely edited transcripts. We compared the conservation of C sites in angiosperm chloroplast coding sequences by summing cumulative branch length from when they appeared to when they reverted to a T (Fig. 6). We then compared the conservation of those C sites that are known or predicted to be edited at the RNA level with those that are not (Fig. 6). The evidence is that sites subject to RNA editing revert to a T more quickly than sites that are not edited.

image

Figure 6. Conservation of RNA editing sites (ES), in comparison to other unedited C sites (Other C). (a) Values are an indication of the duration that these sites persisted as a C during angiosperm evolution. Boxes include 50% of distributions. Horizontal bars indicate means. *, < 0.05; **, < 0.01, Student’s t-test. (b) Schematic diagram of the values in the upper panel were calculated. In this example, solid edges indicate the period that a particular C was conserved. Thus the cumulative edge length for this site is 1.9 (values are relative to the average number of synonymous substitutions per site, dS).

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Several explanations are possible for why edited cytidines might revert more frequently than unedited cytidines. Sequence context affects nucleotide substitution frequency, and the pronounced bias in sequence context around editing sites might lead to more frequent reversion. Alternatively, retrotranscription of edited organelle transcripts is a likely explanation for reversion of multiple closely linked sites and may explain a general high rate of reversion for editing sites (Cuenca et al., 2010; Sloan et al., 2010). Finally, it is possible that there is a sufficient cost associated with editing (the cost of synthesizing and importing editing factors, plus the costs of dealing with the consequences of incompletely edited transcripts and off-target effects of editing factors) to drive selection of reversion events and the loss of superfluous editing factors. It has been demonstrated recently that reversion of an editing site can lead to loss of the relevant editing factor (Hayes & Mulligan, 2011). It seems likely that angiosperm chloroplasts are on their way to ridding themselves of RNA editing. In this context it is worth noting that the number of RNA editing sites reduces quite rapidly in organelle genomes with accelerated substitution rates (Parkinson et al., 2005; Sloan et al., 2010).

VI. Other functions besides editing

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Expansion of the PPR family
  5. III. RNA editing and the evolution of PPR genes
  6. IV. Sequence drift in the organelle genomes of early land plants
  7. V. Loss of RNA editing in angiosperms
  8. VI. Other functions besides editing
  9. VII. Conclusion
  10. References

Not all PPR proteins are implicated in RNA editing. Assuming as a first approximation that all PLS-class proteins are editing factors and all P-class proteins are not, then roughly half of the PPR proteins in angiosperms (c. 200–250) have other functions. This is still a huge increase in numbers over the 10–20 homologous proteins in algae and other eukaryotes, and this expansion probably occurred before, and independently of, the expansion in PLS-class PPR genes (O’Toole et al., 2008). These P-class PPR proteins have roles in many other RNA-processing activities, including splicing, RNA turnover and translation, with positive or negative effects depending on the target and the context (Schmitz-Linneweber & Small, 2008). Many of these activities probably passively result from tight and sequence-specific binding, and, unlike RNA editing, do not require additional catalytic activities (Pfalz et al., 2009; Prikryl et al., 2011). The selection pressures acting to drive expansion of these PPR genes are less evident than those encoding RNA editing factors, but are likely to be similar, involving correction at the RNA level of genome-level mutations and rearrangements (Maier et al., 2008). This is most evident in a particular clade of P-class proteins related to fertility restorer genes associated with cytoplasmic male sterility (CMS). These restorer-to-fertility-like (RFL) genes are numerous in angiosperms and show particularly rapid rates of evolutionary change (Geddy & Brown, 2007; Fujii et al., 2011); at least some of them are required to block expression of the recombinant mitochondrial genes that induce CMS (Bentolila et al., 2002; Brown et al., 2003; Desloire et al., 2003; Kazama & Toriyama, 2003; Koizuka et al., 2003; Komori et al., 2004; Wang et al., 2006; Kato et al., 2007; Kazama et al., 2008; Uyttewaal et al., 2008; Barr & Fishman, 2010).

VII. Conclusion

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Expansion of the PPR family
  5. III. RNA editing and the evolution of PPR genes
  6. IV. Sequence drift in the organelle genomes of early land plants
  7. V. Loss of RNA editing in angiosperms
  8. VI. Other functions besides editing
  9. VII. Conclusion
  10. References

The current understanding of PPR proteins is that they are modular sequence-specific binding factors (Schmitz-Linneweber & Small, 2008). This modularity, coupled with highly specific target recognition, is perhaps what has made this protein family such an attractive resource for evolution to make use of in various cellular activities for plants. Amongst these various activities, RNA editing is one of the most enigmatic processes in terms of understanding how it came about, how it could have expanded to the extent it has in some land plant lineages, and to what extent it is advantageous. The discoveries described in this review are helping to make sense of these questions and have shown us that the evolution of land plant organelle genomes is inextricably linked to the evolution of the nuclear PPR gene family, and vice versa. It now seems probable that a rapid expansion in numbers of PPR genes permitted RNA editing on a large scale, and that this in turn allowed genome sequence drift. If (and it is a big ‘if’) this sequence drift was advantageous to early terrestrial plants (by conferring increased tolerance to temperature extremes, UV or oxidative stress induced by conditions on land) then PPR proteins and RNA editing may have had an important role in accelerating the colonization of the land.

References

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Expansion of the PPR family
  5. III. RNA editing and the evolution of PPR genes
  6. IV. Sequence drift in the organelle genomes of early land plants
  7. V. Loss of RNA editing in angiosperms
  8. VI. Other functions besides editing
  9. VII. Conclusion
  10. References
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