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Journal of Experimental Botany, Vol. 66, No. 22 pp. 6957–6973, 2015 doi:10.1093/jxb/erv415 Advance Access publication 9 September 2015 DARWIN REVIEW Plastid RNA polymerases: orchestration of enzymes with different evolutionary origins controls chloroplast biogenesis during the plant life cycle Thomas Pfannschmidt1,2,3,4,*, Robert Blanvillain1,2,3,4, Livia Merendino1,2,3,4, Florence Courtois1,2,3,4, Fabien Chevalier1,2,3,4, Monique Liebers1,2,3,4, Björn Grübler1,2,3,4, Elisabeth Hommel1,2,3,4 and Silva Lerbs-Mache1,2,3,4 1 Université Grenoble-Alpes, F-38000 Grenoble, France CNRS, UMR5168, F-38054 Grenoble, France 3 CEA, iRTSV, Laboratoire de Physiologie Cellulaire & Végétale, F-38054 Grenoble, France 4 INRA, USC1359, F-38054 Grenoble, France 2 * To whom correspondence should be addressed. Email: [email protected] Received 9 July 2015; Revised 9 August 2015; Accepted 11 August 2015 Editor: Christine Foyer Abstract Chloroplasts are the sunlight-collecting organelles of photosynthetic eukaryotes that energetically drive the biosphere of our planet. They are the base for all major food webs by providing essential photosynthates to all heterotrophic organisms including humans. Recent research has focused largely on an understanding of the function of these organelles, but knowledge about the biogenesis of chloroplasts is rather limited. It is known that chloroplasts develop from undifferentiated precursor plastids, the proplastids, in meristematic cells. This review focuses on the activation and action of plastid RNA polymerases, which play a key role in the development of new chloroplasts from proplastids. Evolutionarily, plastids emerged from the endosymbiosis of a cyanobacterium-like ancestor into a heterotrophic eukaryote. As an evolutionary remnant of this process, they possess their own genome, which is expressed by two types of plastid RNA polymerase, phage-type and prokaryotic-type RNA polymerase. The protein subunits of these polymerases are encoded in both the nuclear and plastid genomes. Their activation and action therefore require a highly sophisticated regulation that controls and coordinates the expression of the components encoded in the plastid and nucleus. Stoichiometric expression and correct assembly of RNA polymerase complexes is achieved by a combination of developmental and environmentally induced programmes. This review highlights the current knowledge about the functional coordination between the different types of plastid RNA polymerases and provides working models of their sequential expression and function for future investigations. Key words: Chloroplast biogenesis, developmental regulation, gene expression, nucleo–plastid interaction, plants, plastid RNA polymerases. Introduction Today’s plastids are a group of morphologically and functionally diverse cellular organelles that are surrounded by a double membrane and are found specifically in plant and algae cells. The most prominent representative of these organelles is the chloroplast, which is found in green tissues of vascular plants, mosses, and green algae. This plastid type is the site of photosynthesis, but also of nitrate and sulphate fixation, and parts of many other important cellular biosynthetic pathways © The Author 2015. Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved. For permissions, please email: [email protected] 6958 | Pfannschmidt et al. including amino acid and lipid biosynthesis. Besides chloroplasts, other non-green plastid forms exist in plants such as amyloplasts and elaioplasts, which are important for starch and oil accumulation, respectively, in roots and storage organs, or chromoplasts, which generate carotenoids responsible for the colours in fruits or flowers. All these different plastid types develop from a common undifferentiated precursor form, the proplastid, which is also the form involved in cytoplasmic inheritance. The respective plastid type that is generated from this proplastid is determined by the tissue context of the cell in which it resides. Furthermore, the different plastid types are convertible into each other, even after differentiation, when respective environmental and/or cellular influences change (e.g. the chloroplast-to-chromoplast conversion upon fruit ripening or in the opposite direction following illumination). Thus, plant plastids display a remarkable morphological and functional diversity and flexibility (Lopez-Juez and Pyke, 2005; Jarvis and Lopez-Juez, 2013). Plant plastids evolved from a monophyletic evolutionary event called primary endosymbiosis whereby a heterotrophic mitochondriated protist engulfed a photosynthetically active, free-living cyanobacterium. In the course of evolution, this cyanobacterium was ‘tamed’ by the host cell and transformed into a stable and inheritable cell compartment, the plastid (Cavalier-Smith, 1999; Reyes-Prieto et al., 2007). This included a massive horizontal gene transfer from the endosymbiont towards the genome of the host, reshaping its biochemical properties (Martin et al., 2002; Timmis et al., 2004; Reyes-Prieto et al., 2006). Current genome analyses strongly suggest that the process of endosymbiosis was facilitated by an infection of the host cell with Chlamydiae (Ball et al., 2013). The result of this complex evolutionary event was the generation of the first photosynthetic eukaryote. This ancient lineage split into three branches, the ‘green’ lineage (chloroplastida), the ‘red’ lineage (rhodophyta), and the glaucophyta. From the green lineage around 450 million years ago, the green algae and plants developed. The benefit provided by plastids was such an advantage that several secondary and even tertiary endosymbiotic events occurred within the green and red algae resulting in the generation of further ‘plastidiated’ eukaryotes such as the dinoflagellates, the diatoms, the euglenoids, and the apicomplexa (Delwiche, 1999; Stoebe and Maier, 2002). This fascinating complexity is one of the prime examples of the power of evolution within the tree of life. As a remnant of their cyanobacterial origin, today’s plastids still contain their own genome, the plastome, and a fully functional machinery for its expression. The plastome is comprised of around 120 genes in plants and up to around 200 genes in some red algae; it encodes mainly components of the photosynthetic and gene expression machineries (Sugiura, 1992; Martin et al., 2002; Green, 2011). This is, however, only a small part of the original coding capacity of the genome of the ancient cyanobacterial precursor. Recent proteome analyses identified up to 1660 non-redundant proteins within Arabidopsis thaliana plastids [available in the online databases AT_CHLORO and the Plant Proteome Data Base (PPDB)], indicating that the vast majority of the chloroplast proteome is encoded in the nucleus, translated in the cytosol, and subsequently imported into the organelle (Sun et al., 2009; Ferro et al., 2010). The present protein catalogues are probably not complete, as bioinformatic predictions of transit peptides identify many more potential plastid-localized proteins in the genome. Estimates range from 2000 to 3500 proteins that may be present in plastids. Low abundance and spatiotemporal expression variations probably represent the major constraints in detection and identification of the lacking proteins (Abdallah et al., 2000; Van Wijk and Baginsky, 2011). The dependency of plastids on the import of proteins from the cytosol gained the host cell full control over development and differentiation of the organelle (i.e. the cell type can fully determine the plastid type). Therefore, usually only one type of plastid is found in a given cell. Rare exceptions are some cells at an early developmental stage in variegation mutants that possess both normal chloroplasts and aberrant plastids. Segregation of these plastids during further cell division subsequently leads to the formation of white and green patches or stripes in the leaves of the mutants (Wetzel et al., 1994; Sakamoto, 2003). On the other hand, all major plastid protein complexes involved in photosynthesis or gene expression contain several plastid-encoded subunits that are essential for the assembly of these complexes (Allen et al., 2011; Pfalz and Pfannschmidt, 2013). Thus, the plastids retained some influence on the build-up of their own protein machineries and are, therefore, regarded as genetically semi-autonomous. The plastid protein complexes, in consequence, represent an evolutionary patchwork of subunits encoded in two different genetic compartments. Stoichiometric production and correct assembly of these subunits thus require a faithful and tight communication between the two compartments (Rodermel, 2001; Pogson et al., 2008; Woodson and Chory, 2008). This communication must coordinate the expression of nuclear genes for plastid proteins (typically present in one or few gene copies in the nucleus) with that of plastid-encoded genes (present in up to around 100 plastome copies per plastid and up to ~100 plastids per cell). This huge difference in gene copy number needs to be adjusted by the gene expression machineries in the nucleus and plastids (Lopez-Juez and Pyke, 2005; Dietzel et al., 2008). The RNA polymerases found within the different genetic compartments are among the key regulators of this complex interplay. By selective transcription of genes essential for the corresponding developmental stage of the tissue, they initiate a coordinated adjustment of gene expression events according to the needs of the organelles. The RNA polymerases and their associated regulatory proteins comprise factors of eukaryotic, prokaryotic, and phage ancestry, and reflect the complex evolutionary integration of a prokaryotic gene expression system into a eukaryotic host cell. Most of our current knowledge about this interplay was obtained by studying chloroplast biogenesis, while much less is known about these processes in other plastid types (e.g. in chromoplasts or amyloplasts), although some studies exist (Kahlau and Bock, 2008; Valkov et al., 2009). For reasons of simplicity, this review focuses largely on processes occurring during the light-dependent transition from the proplastid/ eoplast to the chloroplast, accepting that some aspects of the described events might be different in other plastid types. Plastid RNA polymerases govern chloroplast biogenesis | 6959 The different types of plastid RNA polymerase Plastids possess two types of RNA polymerase: (i) one multisubunit, plastid-encoded prokaryotic-type RNA polymerase (PEP), and (ii) two different single-subunit, nuclear-encoded phage-type RNA polymerase(s) (NEP) (Shiina et al., 2005; Liere et al., 2011; Börner et al., 2015). The first requires interaction with sigma-like factors that mediate promoter recognition as well as with additional PEP-associated proteins (PAPs) of various functions, all encoded by the nuclear genome (Fig. 1) (Schweer et al., 2010; Lerbs-Mache, 2011; Pfalz and Pfannschmidt, 2013; Yu et al., 2014; Chi et al., 2015). A proper function of the plastid transcription machinery therefore requires first the action of the eukaryotic nuclear RNA polymerase II, which is responsible for the expression of the nuclear-encoded components. The development of functional plastids, then, depends on the coordinated action of the two different types of plastid RNA polymerases, which initiate a sequential and highly organized expression cascade of essential plastid genes followed by further maturation of transcripts and appropriate translation. The RNA metabolism and transcript maturation as well as the regulation of plastid translation are highly sophisticated processes, especially in mature chloroplasts, and involve a high number of regulatory proteins (Barkan and Goldschmidt-Clermont 2000; Monde et al., 2000; Eberhard et al., 2002), which cannot be covered by this review. It is, however, important to keep in mind that these levels of gene expression are tightly interlinked. While plastid RNA polymerases provide transcripts for proteins (including transcripts for components of the large and small ribosome subunits) and tRNAs as substrate for translation (Williams-Carrier et al., 2014), plastid translation is, in turn, essential for the generation of plastomeencoded Rpo subunits. In addition, both levels are highly affected by the number of plastid DNA copies available (Udy et al., 2012). Transcription, RNA maturation, and translation thus exhibit a mutual interdependency that needs to be highly coordinated. This coordination during differentiation and build-up phases of the organelle is mediated primarily by developmental programmes yielding the typical tissue-specific plastid types such as chromoplasts, amyloplasts, or chloroplasts. In fully matured chloroplasts, a functional regulation that depends on environmental cues such as light variations becomes more dominant. This functional regulation is mediated mainly by the chloroplast itself, for instance by redox signals from photosynthesis (Pfannschmidt et al., 2003). The orchestration of the different transcription machineries, thus, Fig. 1. Schematic overview summarizing gene location, protein trafficking, and transcriptional activity of all components of the plastid transcriptional machinery and their mutual interdependency. The three genetic compartments of plant cells are depicted. Coloured boxes within them represent genes, with ovals and circles corresponding protein components. The nomenclature of genes and proteins given in these symbols follows that given in the text. Black arrows indicate the traffic of information from a gene via transcripts and translation (indicated by white boxes) towards the place of action of the corresponding protein. Blue symbols: Nuclear genes of NEP enzymes are represented by boxes. Plastid-located NEPs (ovals) are depicted above their target genes in plastids (classes II and III, including rpoA, -B, -C1, and -C2). Rpo subunits are translated within the plastid and generate the PEP core complex. Green symbols: PAP genes in the nucleus are probably expressed as a regulon requiring an as-yet-unknown master regulator (orange oval) for coordinated expression. PAP subunits assemble around the PEP core. Relative positions of PAPs are based on literature data describing protein–protein interactions in a yeast two-hybrid system (see main text). PAPs drawn with a red line display either a true (full line) or potential (broken line) second location in the nucleus. The still-unknown pathways of nuclear trafficking via a cytosolic or a plastid route are indicated by broken red arrows. Yellow symbols: Nuclear-encoded plastid sigma factors are transported into the plastid and mediate specific promoter recognition for the PEP complex (blue Rpo subunits assembled with all green PAPs) for the target genes (yellow boxes: classes I and II as well as sigma factor-specific genes psbN, atpH, and ndhF). For definition of classes I, II and III see text. 6960 | Pfannschmidt et al. requires the integration of developmental as well as physiological regulation networks and integrates signals from the nucleus towards the plastid (anterograde signalling) and signals from the plastid towards the nucleus (retrograde or plastidial signalling) (Pogson and Albrecht, 2011). The bacteria-like DNA-dependent RNA polymerase of plastids (PEP) Core complex Transcriptional activity in plastids was already reported in the 1960s (Kirk, 1964), but the nature of this activity remained obscure until molecular biology techniques allowed the sequencing of plastid genes (Hudson et al., 1988; Igloi et al., 1990). Nowadays, it is common knowledge that plastids possess a RNA polymerase complex of prokaryotic origin that is composed of subunits homologous to the well-studied subunits of RNA polymerase from Escherichia coli or other bacteria. So far, all sequenced chloroplast genomes of vascular plants have revealed the presence of genes that exhibit significant sequence homologies to bacterial RNA polymerase (rpo) genes (Fig. 1) (Igloi and Kossel, 1992). A comparison was performed of structural Rpo homology between Arabidopsis and E. coli at the protein level using Phyre2 and found homologies that ranged from 37% (RpoC2) to 85% (RpoA) and 95% (RpoB, RpoC1) (with respect to the lengths of the proteins). The genes for these proteins are typically arranged as a large operon comprising the genes rpoB–rpoC1–rpoC2 and a single gene rpoA being part of another operon that contains several genes for ribosomal components (Green, 2011). These rpo genes encode the catalytic subunit RpoB (or β-subunit), the subunit RpoC1 (or β′-subunit), which has no clearly assigned function yet, and the subunit RpoC2 (or β″-subunit), which is believed to possess a DNA binding function. Most likely, the genes rpoC1 and rpoC2 emerged from a split of the original rpoC gene of the cyanobacterial ancestor. In addition, in dicotyledonous plants, an intron can be found within the rpoC1 gene that is not present in monocotyledons. This observation might suggest different evolutionary constraints for the evolving transcription machinery in these two plant clades (Igloi and Kossel, 1992).Whether this is connected to specific functional differences of the RNA polymerase subunits between the two clades and/or whether this reflects an adaptation to the different developmental programmes underlying chloroplast biogenesis in monocotyledons and dicotyledons still needs to be elucidated. Interestingly, the rpo genes in the green algae Chlamydomonas reinhardtii are monocistronic units dispersed over the plastome, indicating that the plastome organization within the green lineage was not thoroughly maintained (Maul et al., 2002). The subunit RpoA (or α-subunit) probably serves as a stabilizing subunit of the complex and, in analogy to its bacterial counterpart, is assumed to occur as a dimer, although direct supporting evidence does not exist. The basic RNA polymerase complex with the stoichiometry α2, β, β′, β″ is referred to as the core enzyme and is capable of transcriptional elongation in vitro and probably also in vivo. Plastid knockout mutants in tobacco or nuclear knockdown mutants in Arabidopsis, both with structural or functional deficiency for any of these rpo genes, display an albino or yellowish phenotype with arrested plastid development, indicating the absolute requirement of Rpo subunits for proper chloroplast development. All these mutant plants have been reported to be viable on sucrose-supplemented medium, indicating that the missing chloroplast function can largely be rescued by an external carbon source (Allison et al., 1996; Hajdukiewicz et al., 1997; De SantisMaciossek et al., 1999; Chateigner-Boutin et al., 2008, 2011; Zhou et al., 2009). Sigma factors The typical promoters of PEP-transcribed genes resemble prokaryotic promoters and correspondingly comprise –35 (TTGACA)- and –10 (TATAAT)-like-sequence motifs (Shiina et al., 2005; Liere et al., 2011; Chi et al., 2015). For specific promoter recognition and transcription initiation, the PEP core complex is thus dependent on the interaction with prokaryotic-like sigma factors (Allison, 2000), which were first identified by biochemical means (Bülow and Link, 1988; Lerbs et al., 1988). In contrast to the other Rpo subunits, the genes for these factors were found to be encoded in the nucleus (Liu and Troxler, 1996; Tanaka et al., 1996). Like many other genes that were originally located in the genome of the engulfed cyanobacterium, they went through a horizontal gene transfer towards the nucleus of the host cell and are therefore under the control of the nuclear transcription system. In Arabidopsis, six different sigma factors are known and are designated Sig1–Sig6 (Fig. 1) (Schweer et al., 2010; Lerbs-Mache, 2011; Chi et al., 2015). In contrast to mutants with defects in the core components, single sigma-factor T-DNA inactivation mutants in Arabidopsis do not display severe phenotypic defects, pointing to a certain level of functional redundancy between the factors. Exceptions to this are sig2 and sig6 knockout mutants, which exhibit lightgreen phenotypes indicating a more pronounced role of the encoded Sig2 and Sig6 factors in plastid transcription in the early stages of chloroplast differentiation. Apparently, their loss can only partly be replaced by the other sigma factors during the early steps of seedling development. Studies from several laboratories indeed have revealed partial redundancy for sigma factor functions but also identified gene-specific functions for these regulatory factors (Schweer et al., 2010; Lerbs-Mache, 2011; Malik Ghulam et al., 2012). The specific functions, gene expression profiling, regulation of sigma activity/specificity by post-transcriptional modifications, and some evolutionary aspects have already been described and discussed in detail in a number of previous reviews (Lysenko and Kusnetsov, 2005; Schweer et al., 2010; LerbsMache, 2011; Chi et al., 2015). This review will focus mainly on some evolutionary aspects of sigma diversification and functional integration into the eukaryotic host during land plant evolution. Evolutionary origin of plant sigma factors Previous phylogenetic analysis of sigma factor genes indicated that, in plants, they all derived from the principal cyanobacterial sigma factor SigA (Hakimi et al., 2000; Mache et al., 2002; Plastid RNA polymerases govern chloroplast biogenesis | 6961 Lysenko and Kusnetsov, 2005). This suggests that, out of the multiple sigma factors present in the ancient cyanobacterium, only the primary sigma factor SigA has been retained in the chloroplast ancestor. Comparison of intron positions of plant sigma factor genes further suggests that Sig1 and Sig2 originated from this SigA common ancestor by gene duplication and that all other sigma factors were later derived from Sig2 (Lysenko and Kusnetsov, 2005). The early evolutionary origin of Sig1 and Sig2 was recently confirmed by showing the presence of the corresponding sigma factor genes in the nuclear genomes of the liverwort Marchantia polymorpha (Kanazawa et al., 2013; Ueda et al., 2013), of the moss Physcomitrella patens (Hara et al., 2001; Ichikawa et al., 2004), and of the spikemoss Selaginella moellendorffii (Banks et al., 2011). In addition to these two genes, the nuclear genome of M. polymorpha encodes two other different sigma factors. One was named Mp_SIGX because it does not relate to anyone of the six higher plant sigma factors (Kanazawa et al., 2013). The other one found to be orthologous to Sig5 of higher plants. Thus, Sig5 has evolved over the same time scale as Sig1 and Sig2 but probably does not originate from Sig2, nor from the cyanobacterial ancestor, because none of the cyanobacterial sigma factors group together with Sig5 (Mache et al., 2002). Sig5 has the weakest similarity to other plant sigma factors. Noticeably, A. thaliana Sig5 is orthologous to E. coli sigma factor RpoH, which is known to be implicated in heat-shock and stress responses of the bacterium (Mache et al., 2002). It seems that this function has been evolutionary conserved in basal and higher land plants, as suggested by the induction of Sig5 gene expression under several stress conditions in liverwort as well as in Arabidopsis (Nagashima et al., 2004; Kanazawa et al., 2013). However, the evolutionary origin of Sig5 remains to be elucidated. Diversification of plant sigma factors during plant evolution, and co-evolution of sigma factors and promoters As mentioned above, results obtained so far from non-vascular land plants reveal the presence of sigma orthologues to higher plant factors Sig1, Sig2, and Sig5. The other sigma factors, namely Sig3, Sig4, and Sig6, emerged during theevolution of vascular plants after the separation of lycophytes and euphyllophytes. They probably originated by gene duplication from Sig2. Sig4 represents the most recent acquisition of a new sigma factor. It has been found so far only in Superrosidae (Rosales and Geraniales). Its absence in currently known nuclear genomes of monocotyledonous plants suggests that it evolved after the separation of dicots and monocots. The recent emergence of Sig4 is of special interest as it can be related to a specific functional diversification of sigma function (Schweer et al., 2010; Lerbs-Mache, 2011; Chi et al., 2015). While Sig2 and Sig6 seem to be more general factors required for transcription initiation of many various genes or operons, Sig3 and Sig4 exclusively recognize the psbN and atpH promoters (Sig3) (Zghidi et al., 2007) and the ndhF promoter (Sig4) in Arabidopsis (Fig. 1) (Favory et al., 2005). The first insights into functional specification of sigma factors in non-vascular land plants came from a recent analysis of the plastid transcriptome of a sig1 mutant of M. polymorpha. These results revealed changes that are similar to those observed in mutants of higher plants, suggesting conservation of principal sigma activity of Sig1. However, the ndhF gene is also transcribed by Sig1 in liverwort. A comparison of ndhF promoter regions from liverwort and higher plants reveals considerable alterations in promoter elements between liverwort, several monocotyledons species, and Arabidopsis, testifying to co-evolution of sigma factors and promoters (Ueda et al., 2013). Co-evolution of Sig4 and ndh gene expression is also testified by the concomitant lack of Sig4 gene expression (Zhang et al., 2015) and the loss of ndh genes from the plastid genome in some Geraniaceae (Blazier et al., 2011; Weng et al., 2014). Geraniaceae plastid genomes are highly rearranged (Weng et al., 2014) and are characterized by remarkably elevated evolutionary rates, especially in rpoB, rpoC1, and rpoC2 genes (Guisinger et al., 2008; Weng et al., 2012). In addition, numerous gene duplications of Sig5 and Sig6 have been observed in various species (Zhang et al., 2015). Therefore, Geraniaceae represent an attractive system to study plastid–nuclear genome co-evolution with respect to the PEP transcriptional apparatus. Variations in genes for RNA polymerase subunits have already been shown to be useful for taxonomy classification and phylogenetic analyses (Taillardat-Bisch et al., 2003; Iyer et al., 2004; Lane and Darst, 2010). Using Geraniaceae as a model system, coordinated evolution rates between interacting plastid and nuclear genes coding for PEP subunits and sigma factors, respectively, have recently been shown (Zhang et al., 2015). The authors suggested that this strong correlation of evolutionary rates might provide an explanation for the observed plastome–genome incompatibility between species within the Geraniaceae. Emergence of sigma factor phosphorylation as a mechanism for regulating activity/specificity of transcription Plant sigma factors are composed of evolutionarily conserved C-terminal parts and variable N-terminal parts. The N-terminal parts have probably been acquired during the process of integration of the ancestral genes after duplication into different regions of the host’s nuclear genome. The N-terminal parts have functionally diverged during evolution. They are incompatible with prokaryotic transcription systems as shown for E. coli (Hakimi et al., 2000; Mache et al., 2002), and they are not interchangeable between different plant sigma factors (Schweer et al., 2009). In contrast to their bacterial counterparts, plant sigma factors can be modified post-translationally by phosphorylation, and this modification seems to occur exclusively in the N-terminal variable region. The functional consequences of sigma factor phosphorylation have so far been investigated in vivo for Sig1 (Shimizu et al., 2010) and Sig6 (Schweer et al., 2010) in Arabidopsis; however, using bioinformatics, hypothetical phosphorylation sites were predicted in all plant sigma factors. Phosphorylation has been observed in Sig2 by an in vitro approach, and phosphorylation of Sig1 and Sig6 have been associated with changes in the binding efficiency of RNA polymerase–sigma complexes to selected promoters (Türkeri et al., 2012). Phosphorylation of Sig1 is regulated by 6962 | Pfannschmidt et al. redox signals and serves to adapt photosystem stoichiometry to daily changing light conditions via changes in psbA and psaA transcript levels (Shimizu et al., 2010). Direct phosphorylation of sigma factors represents a mechanism of transcriptional regulation that has been acquired after the endosymbiotic event. However, it is not clear yet whether this mechanism evolved before or after the separation of Viridiplantae and Rhodophyta. Western blot analyses of the four sigma factors of the red alga Cyanidioschyzon merolae (note that these sigma factors cluster together and are not related to Sig1–Sig4 of higher plants as shown for Cyanidium caldarium; Mache et al., 2002) indicate posttranslational modifications for Sig3 and Sig4 (Kanesaki et al., 2012). The nature of these modifications is not known, but phosphorylation might be considered. PEP-associated proteins (PAPs) While genetic data imply a prokaryotic structure of the plastid RNA polymerase complex, biochemical purifications of this enzyme from chloroplasts yielded protein complexes with a much higher subunit number (15–20 depending on the species). These multi-protein complexes thus resemble more the eukaryotic-type RNA polymerases from the nucleus (Lerbs et al., 1985; Rajasekhar et al., 1991). Biochemical studies on mustard cotyledons then revealed that the enzyme in its simple prokaryotic composition (i.e. α2, β, β′, β″) could be purified from etioplasts of dark-grown seedlings. This complex can be referred to as the PEP core complex and forms a holo-enzyme together with the sigma factors for transcription initiation. It is able to rapidly acquire additional subunits upon light exposure, which triggers chloroplast formation (Pfannschmidt and Link, 1994). These additional subunits were subsequently identified by mass spectrometry approaches in mustard, tobacco, and Arabidopsis (Pfannschmidt et al., 2000; Suzuki et al., 2004; Pfalz et al., 2006; Steiner et al., 2011). The corresponding genes were identified in the Arabidopsis nuclear genome by homology searches and all displayed a predicted chloroplastic transit peptide at the N terminus. The majority of the subunits of the plastid-encoded RNA polymerase are thus imported from the cytosol (see below). A major challenge in studying the additional subunits of the plastid RNA polymerase complex by biochemical means is to discriminate between bona fide subunits from contaminating, irrelevant, or non-essential subunits. Plastid DNA and the enzymes responsible for its replication and expression are organized into huge structures resembling bacterial nucleoids. These plastid nucleoids can be visualized in vivo by various optical methods, including DAPI staining, revealing that they are first located at the proplastid envelope and then attach to thylakoids in fully differentiated chloroplasts (Krupinska et al., 2014; Powikrowska et al., 2014). Although nucleoids appear as distinctly shaped bodies in microscopy, they cannot be isolated as such since they do not possess any enclosing membrane or other stabilizing outer structure, and therefore precise subunit catalogues are difficult to obtain. The best approach so far is the purification of the transcriptionally active chromosome (TAC), a huge DNA–protein complex that is capable of in vitro RNA synthesis upon addition of nucleotides. It consists of around 40–60 proteins and was analysed in detail by mass spectrometry in Arabidopsis, mustard, and spinach (Pfalz et al., 2006; Melonek et al., 2012). A total of 35 proteins were identified in Arabidopsis from which 18 were novel and were called pTACs (Pfalz et al., 2006). Many of them are not involved in transcription directly but cover other important functions including structural stabilization of the DNA, membrane anchoring, RNA processing, and ribosome formation. The plastid RNA polymerase complex is an intrinsic part of this TAC, but it can also be purified by alternative protocols as a soluble enzyme activity. This enzyme complex is much smaller than the TAC and requires exogenously added DNA as template for transcription (Pfalz and Pfannschmidt, 2013). The smallest complex reproducibly purified from mustard chloroplasts comprises ten subunits in addition to the Rpo subunits (Steiner et al., 2011) and was formerly assigned as peak A enzyme (Pfannschmidt and Link,1994). The core complex consisting only of Rpo subunits could be purified as peak B enzyme from etioplasts or young immature chloroplasts (Pfannschmidt and Link, 1994). Complexes with intermediate subunit composition have never been found, strongly suggesting that the large peak A enzyme is only stable when all ten additional subunits are present at the same time. It therefore can be defined as the minimum RNA polymerase complex of chloroplasts, which may be larger by transient addition of further subunits. The potential functions of these ten subunits were predicted by domain analysis of the corresponding genes in Arabidopsis and verified in some cases by functional characterization of the respective T-DNA inactivation mutants (see below for detailed description). Intriguingly, all these mutants display an albino/ivory or pale-green phenotype with arrested plastid development, no or minor PEP activity, and an enhanced NEP activity. The mutations are seedling lethal on soil, but the albino mutants can be maintained and grown on sucrosesupplemented medium (Schröter et al., 2010). These phenotypic defects are highly reminiscent of those of plants in which PEP activity was inactivated by genetic approaches (plastomic Δrpo mutants), indicating that the presence of each single subunit is as essential for the activity of PEP as the rpo-core subunits themselves. Therefore, it is postulated that proteins that (i) are present in this minimum RNA polymerase complex, and (ii) cause severe defects in chloroplast biogenesis when inactivated (albinos) define a set of true RNA polymerase subunits that have been named PEP-associated proteins (PAPs) (Pfalz and Pfannschmidt, 2013). Not surprisingly these subunits were also found within the group of pTAC proteins (Pfalz et al., 2006). This allows a distinction to be made between true RNA polymerase subunits (PAPs) and more peripheral components among the pTACs that are less directly involved in transcription and for which chloroplast biogenesis phenotypes in inactivation mutants are less pronounced or even wild-type like. In Arabidopsis, so far 12 different PAPs have been identified that meet the mentioned criteria (Fig. 1). In co-expression analyses, they display a high degree of correlation in their transcript accumulation patterns, suggesting that they probably generate a regulon (Steiner et al., 2011). For PAP1, Plastid RNA polymerases govern chloroplast biogenesis | 6963 it was demonstrated that both RNA and protein are rapidly induced by illumination (Yagi et al., 2012). More detailed in silico expression analyses were performed using available databases at the eFP browser to test this observation for the other pap genes. To this end, the average expression value for the pap gene regulon under all stages of the life cycle (as far as available) was calculated and compared it with the highest expression value found in mature rosette leaves (set to 100%) (Fig. 2, see bar diagram). In general, high pap transcript accumulation was found in green tissues containing chloroplasts (such as carpel, sepals, rosette leaves, and the shoot apical meristam. In contrast, only low expression was observed in non-green tissues such as flower petals and stamen, pollen, roots, and dry seeds (Fig. 2). Interestingly, imbibition of seeds alone led already to significant induction of pap genes. This suggests that pap genes might be induced not only by light but also by internal developmental programmes. In summary, the data indicated that pap genes are mainly expressed in chloroplast-containing tissues and in tissues that are prone to generate chloroplasts (Fig. 2). This coincides with the initial biochemical observations describing a light-induced restructuring of the PEP complex upon etioplast-to-chloroplast transition (Pfannschmidt and Link, 1994). It is thus conceivable that pap gene expression is enhanced by nuclear RNA polymerase II as part of the developmental programmes that serve seedling development and photomorphogenesis. The controlling factors/regulators for this distinct response remain to be determined but most likely are connected to Fig. 2. Expression analysis of the PAP ‘regulon’ during the life cycle of Arabidopsis. Using the eFP browser (http://bar.utoronto.ca), the developmental series of expression levels was obtained for an average of 11 PAPs (centre plot). Means and standard deviations were calculated and plotted as percentages of the highest score set to 100% (marked by arrow head). The stage of life cycle tested can be identified by the positions numbered on the x-axis. 1, Dry seed (ds); 2, imbibed seed, 24 h (imb); 3, first node (not shown on the life cycle); 4, flower stage 12, stamens (not shown); 5, cauline leaf (cl); 6, cotyledon (c); 7, root (r); 8, entire rosette after transition to flowering (not shown); 9, flower stage 9 (f9); 10, flower stage 10/11 (f10); 11, flower stage 12 (f12); 12, flower stage 15 (f15); 13, flower stage 12, carpels (not shown); 14, flower stage 12, petals (not shown); 15, flower stage 12, sepals (not shown); 16, flower stage 15, carpels (floral diagram); 17, flower stage 15, petals (floral diagram); 18, flower stage 15, sepals (floral diagram); 19, flower stage 15, stamen (floral diagram); 20, flowers stage 15, pedicels (pl); 21, leaf 1 + 2 (not shown); 22, leaf 7, petiole (p); 23, leaf 7, distal half (d); 24, leaf 7, proximal half (px); 25, hypocotyl (h); 26, root (r); 27, rosette leaf 2 (rl2); 28, rosette leaf 4 (rl4); 29, rosette leaf 6 (rl6); 30, rosette leaf 8 (rl8); 31, rosette leaf 10 (rl10); 32, rosette leaf 12 (rl12); 33, senescing leaf (sl); 34, shoot apex, inflorescence (ifm); 35, shoot apex, transition (ft); 36, shoot apex, vegetative (sam); 37, stem, second internode (not shown); 38, mature pollen (mp); 39, seeds stage 3 with siliques (s3); 40, seeds stage 4 with siliques (s4); 41, seeds stage 5 with siliques (s5); 42, seeds stage 6 without siliques (s6); 43, seeds stage 7 without siliques (s7); 44, seeds stage 8 without siliques (s8); 45, seeds stage 9 without siliques (s9); 46, seeds stage 10 without siliques (s10); 47, vegetative rosette (arrowhead, reference set at 100%). Each value was then color coded and attributed to the schematic features on the life cycle. F, fertilization; EG, embryogenesis (inset picture depicts embryos in their photosynthetic phase); G, germination; PMG, photomorphogenesis (green arrow, light-dependent etioplast-to-chloroplast transition); FT, flowering transition; FMG; floral morphogenesis; GG, gametogenesis; FG, female gametophyte. Colours range from yellow (no expression) to red (full expression as described above for the plot); white, not documented. Displayed features were inspired by the eFP browser developmental map. 6964 | Pfannschmidt et al. the processes in the early steps of germination and embryo development. Although all Arabidopsis pap knockout mutants display highly similar phenotypes, the predicted functions and structures of the respective PAPs are quite diverse and bring a multitude of functional as well as structural features to the complex. For an exhaustive summary, all 12 pap gene sequences including their predicted functional domains and their corresponding predicted three-dimensional structure around the E. coli RNA polymerase core complex have been complied in Fig. 3 (for more details see the figure legend). Our current working hypothesis is that all the PAPs are required for the formation and/or stability of the PEP complex and that the lack of any of these subunits is detrimental to the whole complex, hence leading to defects broader than the one corresponding solely to the functional loss of any particular subunit. According to structural prediction of functional domains, the proteins can be subdivided into different groups comprising gene expression- or redox-related functions and a third group with unknown or metabolism-related functions. In addition, our predictions identified numerous PAPs that possess a nuclear localization signal (NLS) in addition to the N-terminal chloroplastic transit peptide (PAP1, -5, -7, -8, -9, and -12) (Fig. 3). They therefore probably belong to a special group of proteins recently identified that are dually localized to plastids and the nucleus (Krause and Krupinska, 2009). However, only for PAP5/HMR/pTAC12 has a dual localization in the nucleus and plastids been shown so far, both in Arabidopsis and in maize (Fig. 1) (Chen et al., 2010; Pfalz et al., 2015). The functionality of the NLS in the other PAPs awaits experimental proof. For the nucleus-localized PAP5/ HMR/pTAC12 of Arabidopsis, a function in photomorphogenesis has been attributed (Chen et al., 2010). The mutant allele hmr-5 shows defects in photomorphogenesis corresponding to hypocotyl elongation under red light and an albino phenotype that is typical of pap mutants (Chen and Chory, 2011). It could be demonstrated that HMR physically interacts with phytochrome-interacting-factors (PIFs) and mediates the degradation of PIF1 and PIF3 (Galvao et al., 2012), known to be an important step in the transition from skotomorphogenesis to photomorphogenesis. A 9 aa transcriptional activation domain at the C terminus of PAP5/HMR/pTAC12 mediates the expression of PIF target genes (Qiu et al., 2015), indicating that PAP5/HMR/pTAC12 can directly affect gene expression in the nucleus. This coincides with the action of plastidial PAP5/HMR/pTAC12, as shown recently in maize. It could be demonstrated that the orthologous ZmpTAC12 possesses a binding activity for Fig. 3. Predicted protein structure of the PAP proteins in Arabidopsis and a structural model of the PEP complex. PEP core: Structural model borrowed from bacterial-type rpo core complex (Ebright lab at http://rutchem.rutgers.edu); PAP1–PAP12: yellow, chloroplastic transit peptide (cTP); red, bipartite or monopartite NLS (bNLS or mNLS). PAP1: magenta, PNLR domain, stabilization of rbcL RNA; dark green, PPR (pentatricopeptide repeat) motif, RNA binding involved in RNA maturation and translation; cyan, SAP domain, a putative DNA binding motif involved in chromosomal rearrangement. PAP2: dark green, PPR domain composed of nine motives; light green, SMR small MutS related involved in mismatch repair. PAP3: cyan, S1-like domain, involved in RNA binding. PAP4 and PAP9: cyan and sky blue, SOD Fe/Mn superoxide dismutase C-terminal and N-terminal domains, respectively. PAP5: gold, glutamine-rich region; green, PEST signature of short-lived molecules (adapted from Chen et al., 2010). PAP6: green, fructokinase with substrate and ATP binding site. PAP7: magenta, SET domain, involved in methyl transferase activity; cyan, Rubisco binding domain sharing similarity with histone binding motives. PAP8: no homology with known functional domain. PAP10: magenta, thioredoxin domain. PAP11: pink and magenta, MurE ligase domain, involved in the incorporation of amino acids into peptidoglycan. PAP12: no homology with known functional domains. Blue lines indicate protein regions used for structural models predicted by Phyre2 (http://www.sbg.bio.ic.ac.uk/phyre2). Plastid RNA polymerases govern chloroplast biogenesis | 6965 single-stranded nucleic acids, but without displaying a genespecific function (Pfalz et al., 2015). Its essentiality for PEP complex formation could also be confirmed in this monocotyledonous plant. Interestingly, ZmpTAC12 exists in two differentially migrating forms that are present in both the nucleus and plastids. This poses questions for the trafficking of these proteins within the cell (see below) and the impact of the chloroplastidial PAP5/HMR/pTAC12 function on its nuclear action; i.e. would a lack of the PEP complex interfere with photomorphogenesis because the greening has become impossible and how is the retrograde signalling impacted by PAP5/HMR/pTAC12 loss of function? Transcriptional expression of the PAPs has been observed not only in green tissues but also in tissues preceding their greening, e.g. during imbibition of seeds (Fig. 2), whereas the fully assembled PEP complex has only been observed in lightexposed cotyledons or fully developed leaves. It is possible that all potentially dually localized PAPs may hold different functions in plastid transcription and in photomorphogenesis as described for PAP5/HMR/pTAC12. It has been surmised that a subgroup of the PAPs may assemble into a complex in both the nucleus and the plastid and participate in gene regulation, by interacting with two different RNA polymerases. Extensive research will be necessary to elucidate these events in detail. Urgent questions that remain open are those for the targeting of newly expressed PAPs to nucleus and plastid, and determining which localization signal becomes dominant under which condition. Does nuclear localization require differential translational initiation for removal of the chloroplastic transit peptide or does trafficking towards the nucleus occur through a plastidial route in an unknown process, as proposed for Whirly1 (Fig. 1) (Isemer et al., 2012)? In the PEP complex, one major functional group is comprised of PAPs involved in DNA/RNA metabolism and gene expression regulation, while the second group is related to redox regulation and reactive oxygen species protection (Steiner et al., 2011). The first group includes PAP1, PAP2, PAP3, and PAP7 (but also PAP5 and PAP12, as described recently; Kindgren and Strand, 2015). PAP1 is a protein with a predicted size of 110 kDa and an apparent size of 99 kDa in SDS-PAGE (Yagi et al., 2012). The gene-derived amino acid sequence displays a predicted SAP domain that is known to be involved in DNA/RNA binding (Fig. 3). It also displays additional domains such as a pentatricopeptide repeat (PPR) motif and a domain associated with the stabilization of rbcL transcripts. PPR proteins are known to be involved in RNA metabolism such as processing, splicing, editing, and translation (Delannoy et al., 2007; Schmitz-Linneweber and Small, 2008; Pfalz et al., 2009). It is reported that pap1 mRNA and the corresponding protein accumulate within 1–3 h, respectively, after illumination of dark-grown Arabidopsis seedlings (Yagi et al., 2012). PAP2 is another PPR-containing protein of 89 kDa that displays many structural similarities with PAP1 (Fig. 3) or a small MutS-related mismatch repair domain protein, MRL1, mainly because of the characteristic fold of the PPR domain (Pfalz et al., 2006; Johnson et al., 2010; Steiner et al., 2011). A more detailed analysis of the specific functions of PAP1 or PAP2 regarding their RNA target(s) is, however, still required. PAP3, a 77 kDa protein with an S1-like domain, was found to interact with other subunits of the PEP complex and PAP12 in particular (see below) (Yu et al., 2013). It is predicted to interact with RNAs through its S1-like domain (Fig. 3) in Nicotiana benthamiana (Jeon et al., 2012). PAP7/ pTAC14 is a SET domain-containing protein of 49 kDa that may play a role in the transfer of methyl groups on target proteins (Gao et al., 2011). However, so far there is no evidence for this kind of function for PAP7/pTAC14. Other PAPs may fall into the category of subunits necessary for (redox-dependent or redox-related) gene regulation. PAP6/FLN1 is a protein of 49 kDa and, like its paralogous protein FLN2, displays a phosphofructokinase domain that spans the entire length of the protein (Steiner et al., 2011; Gilkerson et al., 2012). However, a fructokinase activity of the protein has not been confirmed (Arsova et al., 2010) and its true function is still elusive. FLN2, interestingly, was not identified as a PAP and its corresponding knockout mutant displays a less severe phenotype. It thus represents a more transient or peripheral component within the TAC (Gilkerson et al., 2012, Pfalz and Pfannschmidt, 2013). Other protein modifications are possible within the PEP complex by interaction with the thioredoxin domain of the 12 kDa protein PAP10/TrxZ, which has been proven to interact with PAP6 (Arsova et al., 2010; Schröter et al., 2010). It might confer redox regulation of the transcription activity; however, experimental evidence for this is still lacking. Interestingly PAP10 is the first protein for which it was shown that, despite the specific loss of its functional domain, it still rescues the pap10 mutant phenotype with respect to albinism and plastid gene expression (Wimmelbacher and Börnke, 2014). This coincides with the proposed model of PAPs as essential structural components of the PEP complex (see above). Two other paralogous proteins found in the complex are PAP4 and PAP9, both with iron superoxide dismutase domains (Myouga et al., 2008). These 26 and 29 kDa proteins may serve as protection against oxidative stresses generated during the first activities of the photosynthetic apparatus. A partially functional redundancy is observed in the single mutants, as they show palegreen phenotypes rather than the albino phenotype observed with the double mutant. Interestingly, PAP4 displays a highly aberrant apparent molecular weight in SDS-PAGE, suggesting that it might exist as a trimer (Steiner et al., 2011). Three proteins among the PAPs are less well understood. PAP11 is a MurE domain-containing protein with a theoretical mass of 85 kDa, as concluded from structure prediction and its degree of homology to bacterial proteins able to incorporate amino acids into peptidoglycan (Garcia et al., 2008), but such a function is difficult to reconcile with plastid transcription. PAP8, a 31 kDa protein, is the most enigmatic protein of the complex, as it displays no homologies to any known proteins so far and can only be linked to the formation of the PEP complex by its physical presence within it. No other data about it are available, but sequence analysis of the gene identified a potential NLS, suggesting the possibility of having a second nuclear function. The same is true also for PAP12, a 12.47 kDa protein that was shown to interact with PAP3, PAP5, PAP6, and PAP7 in a yeast-two-hybrid 6966 | Pfannschmidt et al. approach (Yu et al., 2013). Future studies will have to unravel the precise functions for each of these proteins in the PEP complex and whether or not the functions are related to transcription. Phage-type DNA-dependent RNA polymerases of plastids (NEPs) First indications for the existence of a nuclear-encoded plastid-localized RNA polymerase activity came from studies on plant systems that either do not possess or do not express the plastid rpo genes due to gene loss or ribosome deficiency (Morden et al., 1991; Hess et al., 1993). Since plastids in these plants cannot express the Rpo subunits, they were expected to be defective in plastid transcription. However, numerous plastid transcripts were readily detectable, pointing to a second, probably nuclear-encoded enzyme activity. A first candidate for this activity was purified from spinach as a potential single-subunit RNA polymerase of 110 kDa (Lerbs-Mache, 1993). Further studies using trans-plastomic tobacco lines in which the rpo genes were genetically inactivated confirmed the presence of a nuclear-encoded transcription system in plastids that uses a distinct set of promoters (see below). In order to distinguish between the two activities, the terms nuclearencoded polymerase (NEP) and plastid-encoded polymerase (PEP) were coined (Hajdukiewicz et al., 1997). Final proof then came from studies in Arabidopsis that identified two nuclear genes for a T3/T7 phage-like single-subunit RNA polymerase being targeted to mitochondria and plastids, respectively (Hedtke et al., 1997). These genes were called rpoT for RNA polymerase of phage T3/T7 type. In Arabidopsis, three different genes exist that encode RNA polymerases targeted to mitochondria (RpoTm) and plastids (RpoTp) and, in addition, according to transit peptide::reporter gene fusion experiments, an enzyme (RpoTmp) that is targeted to both organelles (Fig. 1) (Hedtke et al., 2000; Liere et al., 2011). The gene variants probably emerged from duplication events multiplying the gene for the mitochondrial RNA polymerase, redirecting this RNA polymerase activity also to plastids by acquisition of corresponding transit peptides. However, so far a gene for RpoTmp has not been detected in monocotyledons, suggesting that it is a late evolutionary acquisition in dicotyledons (Liere et al., 2011). All rpoT genes encode phage-type enzymes composed of only a single subunit of around 110 kDa. In vitro experiments with recombinant enzyme activities or overexpression of these proteins indicate faithful promoter recognition, transcription initiation, and elongation, as well as for a specific transcription initiation factor (Liere et al., 2004; Kühn et al., 2007). Specificity factors mediating promoter recognition and transcription initiation as known for yeast or human mitochondria have not been identified yet in plants (Richter et al., 2010). Nevertheless, the NEP enzymes specifically recognize three classes of distinct promoters in vivo designated class Ia, class Ib, and class II (also called type 1a, type 1b, and type 2). Class Ia contains a conserved YRT motif several nucleotides upstream of the transcription start site (Liere and Maliga, 1999). Class Ib contains an additional GAA box around 20 nt upstream of the YRT motif, and class II represents all promoters without the YRT motif (Weihe and Börner, 1999). These distinct types of promoters suggest the existence of protein factors that specifically direct the NEP enzymes to these sites, and at least one protein factor called CDF2 has been reported in spinach that is able to direct RpoTmp to the PC promoter of the rRNA operon (Bligny et al., 2000) (see also below). However, no specificity factor could be unambiguously identified by mass spectrometry or other techniques. A cross-interaction with specificity factors of the PEP enzyme as well as with the PEP complex itself also appears very unlikely, as NEP enzymes have never been identified in any mass spectrometry analysis of PEP complexes. Furthermore, neither NEP nor sigma factors (with the exception of Sig2), or eukaryotic transcription factors with predicted plastidial localization (Wagner and Pfannschmidt, 2006), have ever been identified in nucleoidenriched fractions or TAC preparations (Pfalz et al., 2006; Majeran et al., 2012; Melonek et al., 2012). This lack of evidence suggests that NEP enzymes as well as transcription regulators are probably present in very low abundance or at very restricted time points outside the harvesting time point of current mass spectrometry studies. The regulation of NEP activity is still an open question. NEP activity cannot be inferred from the protein level since it has been shown that during plastid maturation the NEP protein level decreases but its activity increases (Cahoon et al., 2004). In addition, plastidial RpoTmp activity differs in different plant species. While in Arabidopsis RpoTmp activity is limited to the seed imbibition stage and declines rapidly after cold release (Courtois et al., 2007), in spinach it can be detected during all developmental stages (Bligny et al., 2000; Azevedo et al., 2008). Some results suggest that the intraplastidial localization of the enzyme might play a role in its activity in spinach. For instance, immunodetection with specific antisera detected RpoTp and RpoTmp in both stroma and the membrane fraction of spinach chloroplasts (Azevedo et al., 2006). These studies demonstrate an increase in thylakoid membrane association of RpoTmp during leaf development that is mediated by a RING H2-protein named NIP for NEP interacting protein. A regulatory role of this membrane association was suggested because it correlates with a decrease in rDNA transcription (Azevedo et al., 2006, 2008). In contrast, regulation of RpoTmp activity in Arabidosis should be different. RpoTmp exclusively transcribes the rRNA gene operon at the PC promoter, and this activity is limited to the period of seed imbibition (Courtois et al., 2007). The plastid-localized NEP polymerases are very active at early steps of seed germination (Demarsy et al., 2006, 2012; see below), and it is at this specific time that they are intensively expressed. In particular, accumulation of RpoTmp transcripts is strongly increased during seed stratification and then declines after root protrusion (Demarsy et al., 2006). Mitochondrial RpoTmp activity was found to be essential for the expression of a subset of mitochondrial genes coding for the subunits of respiratory chain complexes I and IV (Kühn et al., 2009). These expression data were obtained at an early step of plant development (when the first pair of leaves comes out) and at the rosette stage. However, it was not investigated Plastid RNA polymerases govern chloroplast biogenesis | 6967 whether the mitochondrial RpoTmp enzyme is also active during seed germination, as shown for the plastid equivalent. Interestingly, these early steps of seed germination are characterized by a strong production of anti-sense RNAs (Demarsy et al., 2012). NEP polymerases are known to exhibit a high readthrough transcription activity (Legen et al., 2002). This might cause the generation of anti-sense RNAs that are complementary to coding regions that are placed on the opposite DNA strand. As suggested for the psbT gene, hybridization of anti-sense RNAs with the corresponding sense RNAs might inhibit translation (Zghidi-Abouzid et al., 2011). In addition, in plants where the expression of RNase J was silenced, anti-sense RNAs were accumulated at high levels (Sharwood et al., 2011). This increased presence of anti-sense RNAs was causing a strong blockage of translation in these silenced plants. Interestingly, in situations where the NEP activity is very high, as for seeds at early steps of germination (Demarsy et al., 2012) and tobacco rpo-knockout plants that are deficient in PEP activity (Legen et al., 2002), transcripts were also not translated. Whether the anti-sense RNAs are just an unspecific by-product of the NEP readthrough transcription activity or whether they possess a distinct role in regulation of translation is not yet understood. Investigation of chloroplast development in plants Light-dependent chloroplast development in vascular plants has been investigated extensively as part of photomorphogenesis and represents a prime example for the influence of light on developmental programmes in biological systems. The most common biological test system for chloroplast development used over the years is the light-induced de-etiolation of dark-grown seedlings when shifted from dark to the light (Pogson and Albrecht, 2011). Dark-grown angiosperms such as Arabidopsis and Sinapis typically perform a special developmental programme called skotomorphogenesis exhibiting hypocotyl elongation, formation of an apical hook, and yellow cotyledons that contain a dark-specific plastid type, the etioplast. These plastids lack a thylakoid membrane system but instead possess a so-called prolamellar body, which is comprised of paracrystalline structures of protochlorophyllide, NADPH, and the enzyme protochlorophyll-oxidoreductase. Upon illumination, chlorophyll biosynthesis and thylakoid membrane formation are initiated, and within a few hours a fully developed chloroplast is generated while in parallel the apical hook is opening and hypocotyl elongation is stopped (Solymosi and Schoefs, 2010). In Arabidopsis, numerous mutants with defects at different steps of these developmental programmes have been developed, and essential regulators such as constitutive photomorphogenesis in the dark (cop) (a repressor of photomorphogenesis) or hy5 (the major activator of the light-dependent response) were identified (Casal and Yanovsky, 2005). Extensive research led to an understanding and build-up of the complex regulatory networks behind these light-dependent developmental programmes (Jiao et al., 2007), and chloroplast biogenesis is mainly understood as a consequence of the action of these networks. The etioplast–chloroplast transition is, however, only one case of chloroplast biogenesis among others. Under natural conditions, chloroplast biogenesis in seedlings occurs more often as a concomitant process of germination without a prolonged dark phase, since the seeds often are not fully covered from the light. Furthermore, while studies on seedling development are well accepted as developmental models, these approaches neglect the chloroplast formation in growing or mature plants. Here, chloroplasts are permanently generated in all meristems of the aerial and illuminated parts of a plant starting from proplastids in the meristematic stem cells. Recent studies of chloroplast formation in the shoot apical meristam of Arabidopsis have revealed very dynamic and tissue-dependent revertible chloroplast development (Charuvi et al., 2012). These observations suggest that, in plants beyond the seedling stage, modified or different regulation pathways of chloroplast biogenesis might be active that cannot be fully represented by studies on seedlings. It must be noted that the primary steps of chloroplast biogenesis occur only in meristematic cells, while in mature green cells chloroplasts multiply by division without running through the early developmental programme. Finally, Arabidopsis (but also many other plants) displays a transient greening stage of the embryo during seed formation within their siliques (Allorent et al., 2013). This includes chloroplast formation within the early embryo, which is then reversed by de-differentiation into so-called eoplasts in the late embryo during the desiccation of the seed. It should be noted that, while in Arabidopsis the maturing seeds turn to a brown colour, other plants maintain their seeds in a green stage until full maturation of seeds, provoking a question about the function of chloroplasts during this stage (Borisjuk et al., 2013). The primary chloroplast biogenesis in Arabidopsis embryos is largely not understood and requires further detailed investigation (see below). Additional complexity in this context is introduced by the differences in plastid development between monocotyledonous and dicotyledonous plants. Leaf development in monocotyledonous plants depends on the activity of a basal meristem leading to an age-dependent gradient of cells and plastids from the bottom towards to the tip of the leaf (for a recent review, see Pogson et al., 2015). In conclusion, we need to learn much more about the decisive and limiting steps in chloroplast biogenesis of terrestrial plants. One major aspect (and the focus of this review) is the interaction of the various RNA polymerases within the genetic compartments of the plant cell, which are required for the faithful formation of chloroplasts. Regulation and interplay of plastid RNA polymerases during embryogenesis, germination, and seedling development The initial model for the interaction of NEP and PEP enzymes has been developed based on research data obtained in 6968 | Pfannschmidt et al. transplastomic tobacco lines in which plastid rpo genes were inactivated by the insertion of an aadA gene cassette (Allison et al., 1996; Hajdukiewicz et al., 1997). Homoplastomic tissues and lines of these Δ rpo-plants are deficient in plastid PEP activity but retain the NEP enzyme activity (see above), allowing a discrimination between PEP- and NEP-dependent transcription units. It was proposed that the NEP enzyme is responsible for the early expression of so-called ‘housekeeping’ genes (including rpo genes and other gene expression-related genes), while the PEP enzyme transcribes genes required for later chloroplast biogenesis such as the genes for the photosynthetic complexes (Hajdukiewicz et al., 1997). In simple terms, the model proposes that the NEP enzyme transcribes the rpo genes on the plastome of proplastids or non-green plastids in meristematic cells. The resulting PEP enzyme then becomes important for the expression of photosynthesis genes and the build-up of the plastid photosynthetic apparatus. This proposed sequence of action of the two types of RNA polymerases is mainly based on their differential use of different types and/or combinations of promoter elements (see above). According to this usage, plastome genes were subdivided into three classes that possess promoters only for PEP (class I) (photosystem genes), for PEP and NEP (class II) (most other genes), or only for NEP (class III) (the rpo operon, accD and ycf2) (see Fig. 1). This model was a major breakthrough as it could explain the complex process of chloroplast biogenesis as a simple sequential action of different RNA polymerases activating different gene groups depending on the developmental stage. Data and observations raised in the last 15 years expanded our knowledge about the action of PEP and NEP transcription systems. There is growing evidence that both polymerases are active at the same time in all green or non-green tissues but with different degrees of activity. Systematic comparison of plastid transcription in green and white tissues of ΔrpoA tobacco plants and in primary leaves in the barley mutant albostrians revealed that PEP is the predominant transcriptional activity in green tissues, generating most of the mRNAs, tRNAs, and rRNAs (Legen et al., 2002; Zhelyazkova et al., 2012). The NEP enzymes display a minor activity in green tissues but are enhanced in white tissues of the mutants where they transcribe even photosynthesis genes from remote promoters, although to a low degree (Courtois et al., 2007; Zhelyazkova et al., 2012). An enhanced NEP activity in white plastids of PEP-deficient mutants appears to be characteristic for this type of mutants, as it has also been reported in many other albino mutants and especially in mutants with pap gene expression defects (Pfalz and Pfannschmidt, 2013). Apparently, PEP deficiency causes a condition in plastids that leads to either a compensatory or accidental enhancement of NEP activity, although this cannot rescue the chloroplast biogenesis programme. A recent study performed in maize proposes a predominant role of PEP in the generation of tRNAs supporting active translation in chloroplasts (Williams-Carrier et al., 2014). This coincides with the low levels of photosynthesis proteins found in white plastids. At the same time, this negatively affects the chlorophyll biosynthesis pathway, since the glutamyl-tRNA trnE represents the precursor of aminolevulinic acid, the entrance substrate of the tetrapyrrole biosynthesis pathway (Rüdiger and Grimm, 2006; Tanaka and Tanaka, 2007). These and other new data (e.g. about the requirement of PAP subunits) allow refining of the initial sequential model of NEP and PEP activity and adapting it to the timely and spatial dynamics that occur during tissue development in the life cycle of a plant. Action of RNA polymerases during embryogenesis The plant life cycle of a diploid dicotyledonous angiosperm such as Arabidopsis starts with the fertilization of the egg cell, resulting in a zygote. This zygote develops via distinct gene expression programmes and developmental steps (octant, globular stage, heart stage, and torpedo stage) into a fully developed plant embryo comprising two cotyledons, a shoot axis and a primary root embedded into a seed shell (Le et al., 2010). In Arabidopsis but also in many other plants, this involves a transient photosynthetic stage in which the complete embryo turns green and generates fully developed chloroplasts. The function of these chloroplasts is not fully understood yet, but repression of this photosynthetic phase by electron transport inhibitors (e.g. DCMU) negatively affects the germination ability of the resulting mature seeds (Allorent et al., 2015). The development of the embryo, still connected to the mother plant, might be influenced by hormones from the mother plant at this stage, but may also include independent processes that are controlled by the embryo itself. A clearcut separation of these influences is, however, difficult at this stage. Based on the observations given above, it must be assumed that all steps of the RNA polymerase-driven expression cascade for the full development of a chloroplast become active in the embryo. This includes the activation of NEP and PEP enzymes, as well as the expression of interacting sigma factors and PAPs. Indeed, expression and action of both NEP and PEP activities were identified in this particular stage (Allorent et al., 2013). Likewise activation of pap gene expression could be identified in publicly available expression data sets during the embryonic stage of Arabidopsis (Kremnev and Strand, 2014) (Fig. 2). As already mentioned above, Arabidopsis seeds lose their chloroplasts during maturation and turn into brown seeds with chlorophyll-less embryos. This requires an active block of chloroplast function followed by degradation of chloroplast membrane structures, protein components, and pigments within the embryo. Seed maturation, desiccation, and dormancy are largely determined by hormonal regulation via abscisic acid from the mother plant. A recent study reports that abscisic acid induces the expression of RSH2 and RSH3, two plastid-localized enzymes capable of synthesizing guanosine tetraphosphate (ppGpp), an alarmone that induces the stringent response in bacteria (Yamburenko et al., 2015). This ancient signalling pathway of probable endosymbiotic origin has been retained in plants; however, no real physiological function has been attributed to plant ppGpp yet (van der Biezen et al., 2000; Takahashi et al., 2004). As for bacterial RNA polymerases, the molecule was shown to inhibit transcriptional activities of purified PEP preparations in vitro (Sato, 2009). It is thus conceivable Plastid RNA polymerases govern chloroplast biogenesis | 6969 that abscisic acid shuts down PEP activity in chloroplasts of embryos during seed maturation by enhancing ppGpp production. This would lead to a halt in gene expression and could initiate subsequent disassembly of the photosynthetic apparatus. It has been shown recently that the plastids in the desiccated seeds do not fully return to the proplastid stage but retain the transcriptional apparatus at both the protein and transcript level in order to allow an immediate regain of transcription during germination and subsequent seed development (Allorent et al., 2013). Action of RNA polymerases during germination and plant development Upon imbibition and germination of the seed, a second phase of chloroplast development can be observed within the now-growing embryo. This process is, however, restricted to the tissue of the cotyledons, which is in contrast to the embryo stage during seed formation where all tissues become green. Apparently during seedling development, chloroplast formation in the hypocotyl and root is either actively prevented or not promoted. This would require either expression of an inhibiting factor or the repression of an activating factor. A recent study suggests that chloroplast formation in the roots is actively repressed by hormone signals from the shoot (Kobayashi et al., 2012). This observation also demonstrates that chloroplast formation is a developmental subprogramme that can be switched on and off and does not simply represent a concomitant process of illumination. In the cotyledons, it has been shown that the same RNA polymerase expression cascade runs as before in the seed embryo, leading to a quick generation of chloroplasts in the leaf blade tissues (Demarsy et al., 2006). NEP transcripts both for RpoTp and RpoTmp are already present in the dry seed (Demarsy et al., 2006; Allorent et al., 2013). Upon imbibition and germination, NEP transcripts and activities increase and peak at 2–3 d after germination. They trigger a rapid expression of plastid rpo genes, increasing subsequently PEP activity. This coincides with a high pap gene expression detected after imbibition of the seed (Fig. 2). Whether NEP and PAP expression is controlled by the same regulators in the nucleus is not known yet. PEP activity stored in the eoplast after de-differentiation of embryo chloroplasts appears to be important for efficient germination, as inhibition of this activity by tagetin treatment delays significantly germination (Demarsy et al., 2006). However, these starter molecules are apparently not sufficient to induce full chloroplast development and require supplementation of activity by generation of novel PEP complexes. This process of complementation probably becomes interrupted when germination occurs in the dark. In mustard, this leads to accumulation of the PEP core enzyme, which is rapidly reconstructed by the addition of PAPs upon illumination (Pfannschmidt and Link, 1994). The precise sequence of events that occur during this light-triggered reconstruction of PEP is largely not understood and will require careful dissection in future studies. Existing data suggest that it may include regulation at the levels of transcription, translation,and even post-translational mechanisms (Steiner et al., 2011; Yagi et al., 2012). The strong albino phenotype of pap knockout mutants indicates that the process of PEP reconstruction is an essential bottleneck in the generation of chloroplasts. Interruption of any single step in it should lead to similar phenotypes. A detailed characterization of all pap mutants as well as additional genetic screens for regulatory factors will be necessary to fully understand this process. The high speed at which chloroplasts are generated in dark-grown seedlings upon illumination suggests that the time window for the course of events is probably in the hour to minute range. In the meristems of aerial parts of adult plants, such as the shoot apical meristem, it should be expected that the processes run even faster, since no artificial pausing due to darkening does occur. A study analysing the presence of thylakoid membranes in plastids of the different cell layers in the shoot apex of Arabidopsis could identify fairly developed thylakoids in the L1 and L3 layers, while in the L2 layer true proplastids were identified (Charuvi et al., 2012). This suggests that the differential action of NEP and PEP enzyme must be under a tight cellular control. The described secondary loss of chloroplasts in epidermal cells of leaves (Charuvi et al., 2012) could be explained by a targeted repressive effect on PEP activity e.g. by ppGpp, followed by a specific degradation of plastids similar to the processes described for the late embryonic phase. How these processes are performed and regulated is largely unexplored. The events described above are based mainly on observations in dicotyledonous plants. In principle, it should be expected that similar processes also occur in monocotyledonous plants, but adapted to the specific developmental gradient of the plastids caused by the action of the basal meristem (Mullet, 1993). Indeed, many similarities have been reported. In maize, lack of ZmpTAC2, ZmpTAC10, and ZmpTAC12 (the orthologues of Arabidopsis PAP2, PAP3, and PAP5) each leads to PEP deficiency exactly as described for Arabidopsis (Pfalz et al., 2015). Analysis of the barley mutant albostrians indicates enhanced NEP activities in plastids of tissues with PEP deficiency (Zhelyazkova et al., 2012). Nevertheless, there also exist differences from dicotyledonous plants, for instance the existence of two variants of ZmpTAC12 (Pfalz et al., 2015). Whether this represents a species-specific difference or a difference between the plant clades needs to be elucidated in the future. Concluding remarks The transcription of plastidial genes is essential for all plastid types regardless of whether they perform photosynthesis or not. Proper plastid development therefore involves in all cases the action of the plastid RNA polymerases. It should be noted that, in green tissues with mature chloroplasts, the organelles do multiply by fission rather than by biogenesis from proplastids. The role of plastid RNA polymerases here is different from in meristems, embryos, or cotyledons, and is mainly restricted to the maintenance of transcript levels to allow sufficient protein production in the mature chloroplast. Future studies focusing on plastid development will therefore 6970 | Pfannschmidt et al. need to address a complete understanding of functioning and regulation of plastid RNA polymerases in both a spatial and time-resolved manner at the cellular and tissue levels. This will also include the search for essential (master?) regulators controlling the expression of the nuclear-encoded components of the plastid transcription machineries (NEPs, Sig1–6, and PAPs) and putative signalling molecules for retrograde signals from the plastids. Identification of such regulators and signalling molecules may help us to understand the coupling (and its limitations) between plastid development and developmental programmes that control plant morphology and provides interesting targets for potential biotechnological applications. A detailed understanding of the action of plastid RNA polymerases will be essential for future ‘green’ biotechnology and bioenergy applications. This includes functional and structural (crystallographic) analyses of the complex. Necessary experimental tools are readily available, and corresponding working models await experimental proof promising fundamental new discoveries in this research field. Acknowledgements Work in Team 7 ‘Nucleo-plastidic interaction: Chloroplast biogenesis and redox control’ at the Laboratoire de Physiologie Cellulaire & Végétale (LPCV) was funded by the DFG to TP (Pf323-4 and Pf323-5), by Université Grenoble-Alpes (AGIR2015) to RB and the Labex GRAL (Grenoble Alliance for Integrated & Structural Biology) to LPCV. Copyright for colorized scanning electron microscopy images of the Arabidopsis flower used in Fig. 2: J. Berger and D. Weigel, Max-Planck-Institute for Developmental Biology, Tübingen, Germany. References Abdallah F, Salamini F, Leister D. 2000. A prediction of the size and evolutionary origin of the proteome of chloroplasts of Arabidopsis. Trends in Plant Science 5, 141–142. Allen JF, de Paula WB, Puthiyaveetil S, Nield J. 2011. A structural phylogenetic map for chloroplast photosynthesis. Trends in Plant Science 16, 645–655. Allison LA. 2000. The role of sigma factors in plastid transcription. Biochimie 82, 537–548. Allison LA, Simon LD, Maliga P. 1996. Deletion of rpoB reveals a second distinct transcription system in plastids of higher plants. EMBO Journal 15, 2802–2809. Allorent G, Courtois F, Chevalier F, Lerbs-Mache S. 2013. Plastid gene expression during chloroplast differentiation and dedifferentiation into non-photosynthetic plastids during seed formation. Plant Molecular Biology 82, 59–70. Allorent G, Osorio S, Vu JL, et al. 2015. Adjustments of embryonic photosynthetic activity modulate seed fitness in Arabidopsis thaliana. New Phytologist 205, 707–719. Arsova B, Hoja U, Wimmelbacher M, Greiner E, Ustun S, Melzer M, Petersen K, Lein W, Bornke F. 2010. Plastidial thioredoxin z interacts with two fructokinase-like proteins in a thiol-dependent manner: evidence for an essential role in chloroplast development in Arabidopsis and Nicotiana benthamiana. The Plant Cell 22, 1498–1515. Azevedo J, Courtois F, Hakimi MA, Demarsy E, Lagrange T, Alcaraz JP, Jaiswal P, Marechal-Drouard L, Lerbs-Mache S. 2008. Intraplastidial trafficking of a phage-type RNA polymerase is mediated by a thylakoid RING-H2 protein. Proceedings of the National Academy of Sciences, USA 105, 9123–9128. Azevedo J, Courtois F, Lerbs-Mache S. 2006. Sub-plastidial localization of two different phage-type RNA polymerases in spinach chloroplasts. Nucleic Acids Research 34, 436–444. Ball SG, Subtil A, Bhattacharya D, Moustafa A, Weber APM, Gehre L, Colleoni C, Arias MC, Cenci U, Dauvillee D. 2013. Metabolic effectors secreted by bacterial pathogens: essential facilitators of plastid endosymbiosis? The Plant Cell 25, 7–21. Banks JA, Nishiyama T, Hasebe M, Bowman JL, Gribskov M. 2011. The Selaginella genome identifies genetic changes associated with the evolution of vascular plants. Science 332, 960–963. Barkan A, Goldschmidt-Clermont M. 2000. Participation of nuclear genes in chloroplast gene expression. Biochimie 82, 559–572. Blazier JC, Guisinger MM, Jansen RK. 2011. Recent loss of plastidencoded ndh genes within Erodium (Geraniaceae). Plant Molecular Biology 76, 263–272. Bligny M, Courtois F, Thaminy S, Chang CC, Lagrange T, BaruahWolff J, Stern DB, Lerbs-Mache S. 2000. Regulation of plastidrDNA transcription by interaction of CDF2 with two different RNA polymerases. EMBO Journal 19, 1851–1860. Borisjuk L, Neuberger T, Schwend J, et al. 2013 Seed architecture shapes embryo metabolism in oil seed rape. The Plant Cell 25, 1625–1640. Börner T, Aleynikova AY, Zubo YO, Kusnetsov VV. 2015. Chloroplast RNA polymerases: Role in chloroplast biogenesis. Biochim Biophys Acta 1847, 761–769. Bülow S, Link G. 1988. Sigma-like activity from mustard (Sinapis alba L.) chloroplasts conferring DNA-binding and transcription specificity to E. coli core RNA polymerase. Plant Molecular Biology 10, 349–357. Cahoon AB, Harris FM, Stern DB. 2004. Analysis of developing maize plastids reveals two mRNA stability classes correlating with RNA polymerase type. EMBO Reports 5, 801–806. Casal JJ, Yanovsky MJ. 2005. Regulation of gene expression by light. International Journal of Developmental Biology 49, 501–511. Cavalier-Smith T. 1999. Principles of protein and lipid targeting in secondary symbiogenesis: euglenoid, dinoflagellate, and sporozoan plastid origins and the eukaryote family tree. Journal of Eukaryotic Microbiology 46, 347–366. Charuvi D, Kiss V, Nevo R, Shimoni E, Adam Z, Reich Z. 2012. Gain and loss of photosynthetic membranes during plastid differentiation in the shoot apex of Arabidopsis. The Plant Cell 24, 1143–1157. Chateigner-Boutin AL, Colas des Francs-Small C, Delannoy E, Kahlau S, Tanz SK, Falcon de Longevialle A, Fujii S, Small I. 2011. OTP70 is a pentatricopeptide repeat protein of the E subgroup involved in splicing of the plastid transcript rpoC1. The Plant Journal 65, 532–542. Chateigner-Boutin AL, Ramos-Vega M, Guevara-Garcia A, et al. 2008. CLB19, a pentatricopeptide repeat protein required for editing of rpoA and clpP chloroplast transcripts. The Plant Journal 56, 590–602. Chen M, Chory J. 2011. Phytochrome signaling mechanisms and the control of plant development. Trends in Cell Biology 21, 664–671. Chen M, Galvao RM, Li MN, Burger B, Bugea J, Bolado J, Chory J. 2010. Arabidopsis HEMERA/pTAC12 initiates photomorphogenesis by phytochromes. Cell 141, 1230–1240. Chi W, He B, Mao J, Jiang J, Zhang L. 2015. Plastid sigma factors: their individual functions and regulation in transcription. Biochimica et Biophysica Acta 1847, 770–708. Courtois F, Merendino L, Demarsy E, Mache R, Lerbs-Mache S. 2007. Phage-type RNA polymerase RPOTmp transcribes the rrn operon from the PC promoter at early developmental stages in Arabidopsis. Plant Physiology 145, 712–721. De Santis-Maciossek G, Kofer W, Bock A, Schoch S, Maier RM, Wanner G, Rudiger W, Koop HU, Herrmann RG. 1999. Targeted disruption of the plastid RNA polymerase genes rpoA, B and C1: molecular biology, biochemistry and ultrastructure. The Plant Journal 18, 477–489. Delannoy E, Stanley WA, Bond CS, Small ID. 2007. Pentatricopeptide repeat (PPR) proteins as sequence-specificity factors in posttranscriptional processes in organelles. Biochemical Society Transactions 35, 1643–1647. Delwiche C. 1999. Tracing the thread of plastid diversity through the tapestry of life. American Naturalist 154, S164–S177. Demarsy E, Buhr F, Lambert E, Lerbs-Mache S. 2012. Characterization of the plastid-specific germination and seedling Plastid RNA polymerases govern chloroplast biogenesis | 6971 establishment transcriptional programme. Journal of Experimental Botany 63, 925–939. Demarsy E, Courtois F, Azevedo J, Buhot L, Lerbs-Mache S. 2006. Building up of the plastid transcriptional machinery during germination and early plant development. Plant Physiology 142, 993–1003. Dietzel L, Steiner S, Schröter Y, Pfannschmidt T. 2008. Plastidto-nucleus communication in plant cells: retrograde signalling. Berlin: Springer. Eberhard S, Drapier D, Wollman F-A. 2002. Searching limiting steps in the expression of chloroplast-encoded proteins: relations between gene copy number, transcription, transcript abundance and translation rate in the chloroplast of Chlamydomonas reinhardtii. The Plant Journal 31, 149–160. Favory J-J, Kobayshi M, Tanaka K, Peltier G, Kreis M, Valay J-G, Lerbs-Mache S. 2005. Specific function of a plastid sigma factor for ndhF gene transcription. Nucleic Acids Research 33, 5991–5999. Ferro M, Brugière S, Salvi D, et al. 2010. AT_CHLORO, a comprehensive chloroplast proteome database with subplastidial localization and curated information on envelope proteins. Molecular & Cellular Proteomics 9, 1063–1084. Galvao RM, Li M, Kothadia SM, Haskel JD, Decker PV, Van Buskirk EK, Chen M. 2012. Photoactivated phytochromes interact with HEMERA and promote its accumulation to establish photomorphogenesis in Arabidopsis. Genes & Development 26, 1851–1863. Gao ZP, Yu QB, Zhao TT, Ma Q, Chen GX, Yang ZN. 2011. A functional component of the transcriptionally active chromosome complex, Arabidopsis pTAC14, interacts with pTAC12/HEMERA and regulates plastid gene expression. Plant Physiology 157, 1733–1745. Garcia M, Myouga F, Takechi K, Sato H, Nabeshima K, Nagata N, Takio S, Shinozaki K, Takano H. 2008. An Arabidopsis homolog of the bacterial peptidoglycan synthesis enzyme MurE has an essential role in chloroplast development. The Plant Journal 53, 924–934. Gilkerson J, Perez-Ruiz JM, Chory J, Callis J. 2012. The plastidlocalized pfkB-type carbohydrate kinases FRUCTOKINASE-LIKE 1 and 2 are essential for growth and development of Arabidopsis thaliana. BMC Plant Biology 12, 102. Green BR. 2011. Chloroplast genomes of photosynthetic eukaryotes. The Plant Journal 66, 34–44. Guisinger MM, Kuehl JV, Boore JL, Jansen RK. 2008. Genome-wide analyses of Geraniaceae plastid DNA reveal unprecedented patterns of increased nucleotide substitutions. Proceedings of the National Academy of Sciences, USA 105, 18424–18429. Hajdukiewicz PTJ, Allison LA, Maliga P. 1997. The two RNA polymerases encoded by the nuclear and the plastid compartments transcribe distinct groups of genes in tobacco plastids. EMBO Journal 16, 4041–4048. Hakimi MA, Privat I, Valay J-G, Lerbs-Mache S. 2000. Evolutionary conservation of C-terminal domains of primary sigma70-type transcription factors between plants and bacteria. Journal of Biological Chemistry 275, 9215–9221. Hara K, Morita M, Takahashi R, Sugita M, Kato S, Aoki S. 2001. Characterization of two genes, Sig1 and Sig2, encoding distinct sigma factors in the moss Physcomitrella patens: phylogenetic relationship to plastid sigma factors in higher plants. FEBS Letters 499, 87–91. Hedtke B, Börner T, Weihe A. 1997. Mitochondrial and chloroplast phage-type RNA polymerases in Arabidopsis. Science 277, 809–811. Hedtke B, Börner T, Weihe A. 2000. One RNA polymerase serving two genomes. EMBO Reports 1, 435–440. Hess WR, Prombona A, Fieder B, Subramanian AR, Borner T. 1993. Chloroplast Rps15 and the Rpob/C1/C2 gene cluster are strongly transcribed in ribosome-deficient plastids: evidence for a functioning nonchloroplast-encoded RNA polymerase. EMBO Journal 12, 563–571. Hudson GS, Holton TA, Whitfeld PR, Bottomley W. 1988. Spinach chloroplast rpoBC genes encode three subunits of the chloroplast RNA polymerase. Journal of Molecular Biology 200, 639–654. Ichikawa K, Sugita M, Imaizumi T, Wada M, Aoki S. 2004. Differential expression on a daily basis of plastid sigma factor genes from the moss Physcomitrella patens. Regulatory interactions among PpSig5, the circadian clock, and blue light signalling mediated by cryptochromes. Plant Physiology 136, 4282–4298. Igloi GL, Kossel H. 1992. The transcriptional apparatus of chloroplasts. Critical Reviews in Plant Sciences 10, 525–558. Igloi GL, Meinke A, Dory I, Kossel H. 1990. Nucleotide sequence of the maize chloroplast rpo B/C1/C2 operon: comparison between the derived protein primary structures from various organisms with respect to functional domains. Molecular & General Genetics 221, 379–394. Isemer R, Mulisch M, Schäfer A, Kirchner S, Koop HU, Krupinska K. 2012. Recombinant Whirly1 translocates from transplastomic chloroplasts to the nucleus. FEBS Letters 586, 85–88. Iyer ML, Koonin EV, Aravind L. 2004. Evolution of bacterial RNA polymerase: implications for large-scale bacterial phylogeny, domain accretion, and horizontal gene transfer. Gene 335, 72–88. Jarvis P, Lopez-Juez E. 2013. Biogenesis and homeostasis of chloroplasts and other plastids. Nature Reviews 14, 787–802. Jeon Y, Jung HJ, Kang H, Park YI, Lee SH, Pai HS. 2012. S1 domaincontaining STF modulates plastid transcription and chloroplast biogenesis in Nicotiana benthamiana. New Phytologist 193, 349–363. Jiao YL, Lau OS, Deng XW. 2007. Light-regulated transcriptional networks in higher plants. Nature Reviews Genetics 8, 217–230. Johnson X, Wostrikoff K, Finazzi G, Kuras R, Schwarz C, Bujaldon S, Nickelsen J, Stern DJ, Wollman F-A, Vallon O. 2010. MRL1, a conserved pentatricopeptide repeat protein, is required for stabilization of rbcL mRNA in Chlamydomonas and Arabidopsis. The Plant Cell 22, 234–248. Kahlau S, Bock R. 2008. Plastid transcriptomics and translatomics of tomato fruit development and chloroplast-to-chromoplast differentiation: chromoplast gene expression largely serves the production of a single protein. The Plant Cell 20, 856–874. Kanazawa T, Ishizaki K, Kohchi T, Hanaoka M, Tanaka K. 2013. Characterization of four nuclear-encoded plastid RNA polymerase sigma factor genes in the liverworth Marchantia polymorpha: blue-light- and multiple stress-responsive SIG5 was acquired early in the emergence of terrestrial plants. Plant Cell Physiology 54, 1736–1748. Kanesaki Y, Imamura S, Minoda A, Tanaka K. 2012. External light conditions and internal cell cycle phases coordinate accumulation of chloroplast and mitochondrial transcripts in the red alga Cyanidioschyzon merolae. DNA Research 19, 289–303. Kindgren P, Strand A. 2015. Chloroplast transcription: untangling the Gordian knot. New Phytologist 206, 889–891. Kirk JTO. 1964. DNA-dependent RNA synthesis in chloroplast preparations. Biochemical and Biophysical Research Communications 14, 393–397. Kobayashi K, Baba S, Obayashi T, et al. 2012. Regulation of root greening by light and auxin/cytokinin signaling in Arabidopsis. The Plant Cell 24, 1081–1095. Krause K, Krupinska K. 2009. Nuclear regulators with a second home in organelles. Trends in Plant Science 14, 194–199. Kremnev D, Strand A. 2014. Plastid encoded RNA polymerase activity and expression of photosynthesis genes required for embryo and seed development in Arabidopsis. Frontiers in Plant Science 5, 385. Krupinska K, Oetke S, Desel C, Mulisch M, Schäfer A, Hollmann J, Kumlehn J, Hensel G. 2014. WHIRLY1 is a major organizer of chloroplast nucleoids. Frontiers in Plant Science 5, 432. Kühn K, Bohne AV, Liere K, Weihe A, Börner T. 2007. Arabidopsis phage-type RNA polymerases: accurate in vitro transcription of organellar genes. The Plant Cell 19, 959–971. Kühn K, Richter U, Meyer EH, Delannoy E, Falcon de Longevialle A, O’Toole N, Börner T, Millar AH, Small ID, Whelan J. 2009. Phagetype RNA polymerase RPOTmp performs gene-specific transcription in mitochondria of Arabidopsis thaliana. The Plant Cell 21, 2762–2779. Lane WJ, Darst SA. 2010. Molecular evolution of multisubunit RNA polymerases: sequence analysis. Journal of Molecular Biology 395, 671–685. Le BH, Cheng C, Bui AQ, et al. 2010. Global analysis of gene activity during Arabidopsis seed development and identification of seed-specific transcription factors. Proceedings of the National Academy of Sciences, USA 107, 8063–8070. Legen J, Kemp S, Krause K, Profanter B, Herrmann RG, Maier RM. 2002. Comparative analysis of plastid transcription profiles of entire plastid 6972 | Pfannschmidt et al. chromosomes from tobacco attributed to wild-type and PEP-deficient transcription machineries. The Plant Journal 31, 171–188. nucleoids against oxidative stress and is essential for chloroplast development in Arabidopsis. The Plant Cell 20, 3148–3162. Lerbs S, Bräutigam E, Mache R. 1988. DNA-dependent RNA polymerase of spinach chloroplasts: characterization of α-like and σ-like polypeptides. Molecular Genetics and Genomics 211, 459–464. Nagashima A, Hanaoka M, Shikanai T, Fujiwara M, Kanamaru K, Takahashi H, Tanaka K. 2004. The multiple-stress responsive plastid sigma factor, SIG5, directs activation of the psbD blue light-responsive promoter (BLRP) in Arabidopsis thaliana. Plant Cell Physiology 45, 357–368. Pfalz J, Bayraktar OA, Prikryl J, Barkan A. 2009. Site-specific binding of a PPR protein defines and stabilizes 5’ and 3’ mRNA termini in chloroplasts. EMBO Journal 28, 2042–2052. Pfalz J, Holtzegel U, Barkan A, Weisheit W, Mittag M, Pfannschmidt T. 2015. ZmpTAC12 binds single-stranded nucleic acids and is essential for accumulation of the plastid-encoded RNA polymerase complex in maize. New Phytologist 206, 1024–1037. Pfalz J, Liere K, Kandlbinder A, Dietz KJ, Oelmuller R. 2006. PTAC2,-6, and-12 are components of the transcriptionally active plastid chromosome that are required for plastid gene expression. The Plant Cell 18, 176–197. Pfalz J, Pfannschmidt T. 2013. Essential nucleoid proteins in early chloroplast development. Trends in Plant Sciences 18, 186–194. Pfannschmidt T, Link G. 1994. Separation of two classes of plastid DNA-dependent RNA polymerases that are differentially expressed in mustard (Sinapis alba L) seedlings. Plant Molecular Biology 25, 69–81. Pfannschmidt T, Ogrzewalla K, Baginsky S, Sickmann A, Meyer HE, Link G. 2000. The multisubunit chloroplast RNA polymerase A from mustard (Sinapis alba L.): integration of a prokaryotic core into a larger complex with organelle-specific functions. European Journal of Biochemistry 267, 253–261. Pfannschmidt T, Schutze K, Fey V, Sherameti I, Oelmuller R. 2003. Chloroplast redox control of nuclear gene expression—a new class of plastid signals in interorganellar communication. Antioxidants and Redox Signaling 5, 95–101. Pogson BJ, Albrecht V. 2011. Genetic dissection of chloroplast biogenesis and development: an overview. Plant Physiology 155, 1545–1551. Pogson BJ, Ganguly D, Albrecht-Borth V. 2015. Insights into chloroplast biogenesis and development. Biochimica et Biophysica Acta 1847, 1017–1024. Pogson BJ, Woo NS, Förster B, Small ID. 2008. Plastid signalling to the nucleus and beyond. Trends in Plant Science 13, 602–609. Powikrowska M, Oetke S, Jensen PE, Krupinska K. 2014. Dynamic composition, shaping and organization of plastid nucleoids. Frontiers in Plant Science 5, 424. Qiu Y, Li M, Pasoreck EK, et al. 2015. HEMERA couples the proteolysis and transcriptional activity of PHYTOCHROME INTERACTING FACTORs in Arabidopsis photomorphogenesis. The Plant Cell 27, 1409–1427. Rajasekhar VK, Sun E, Meeker R, Wu BW, Tewari KK. 1991. Highly purified pea chloroplast RNA polymerase transcribes both rRNA and mRNA genes. European Journal of Biochemistry 195, 215–228. Reyes-Prieto A, Hackett JD, Soares MB, Bonaldo MF, Bhattacharya D. 2006. Cyanobacterial contribution to algal nuclear genomes is primarily limited to plastid functions. Current Biology 16, 2320–2325. Reyes-Prieto A, Weber APM, Bhattacharya D. 2007. The origin and establishment of the plastid in algae and plants. Annual Reviews in Genetics 41, 147–158. Richter U, Kühn K, Okada S, Brennicke A, Weihe A, Börner T. 2010. A mitochondrial rRNA dimethyladenosine methyltransferase in Arabidopsis. The Plant Journal 61, 558–569. Rodermel S. 2001. Pathways of plastid-to-nucleus signaling. Trends in Plant Science 6, 471–478. Rüdiger W, Grimm B. 2006. Chlorophyll metabolism, an overview. In: Grimm B, Porra RJ, Rüdiger W, Scheer H, eds. Advances in photosynthesis and respiration. Dordrecht: Springer, 133–146. Sakamoto W. 2003. Leaf-variegated mutations and their responsible genes in Arabidopsis thaliana. Genes & Genetic Systems 78, 1–9. Sato M. 2009. Bacterial alarmone, guanosine 5’-diphosphate 3’-diphosphate (ppGpp), predominantly binds the β’ subunit of plastidencoded plastid RNA polymerase in chloroplasts. ChemBioChem 10, 1227–1233. Lerbs S, Bräutigam E, Parthier B. 1985. Polypeptides of DNAdependent RNA polymerase of spinach chloroplasts: characterisation by antibody-linked polymerase assay and determination of sites of synthesis. EMBO Journal 4, 1661–1668. Lerbs-Mache S. 1993. The 110-kDa polypeptide of spinach plastid DNAdependent RNA polymerase: single-subunit enzyme or catalytic core of multimeric enzyme complexes? Proceedings of the National Academy of Sciences, USA 90, 5509–5513. Lerbs-Mache S. 2011. Function of plastid sigma factors in higher plants: regulation of gene expression or just preservation of constitutive transcription? Plant Molecular Biology 76, 235–249. Liere K, Kaden D, Maliga P, Börner T. 2004. Overexpression of phage-type RNA polymerase RpoTp in tobacco demonstrates its role in chloroplast transcription by recognizing a distinct promoter type. Nucleic Acids Research 32, 1159–1165. Liere K, Maliga P. 1999. In vitro characterization of the tobacco rpoB promoter reveals a core sequence motif conserved between phagetype plastid and plant mitochondrial promoters. EMBO Journal 18, 249–257. Liere K, Weihe A, Borner T. 2011. The transcription machineries of plant mitochondria and chloroplasts: composition, function, and regulation. Journal of Plant Physiology 168, 1345–1360. Liu B, Troxler RF. 1996. Molecular characterization of a positively photoregulated nuclear gene for a chloroplast RNA polymerase sigma factor in Cyanidium caldarium. Proceedings of the National Academy of Sciences, USA 93, 3313–3318. Lopez-Juez E, Pyke KA. 2005. Plastids unleashed: their development and their integration in plant development. International Journal of Developmental Biology 49, 557–577. Lysenko EA, Kusnetsov VV. 2005. Plastid RNA polymerases. Molecular Biology 39, 762–775. Mache R, Cottet A, Imberty A, Hakimi MA, Lerbs-Mache S. 2002. The plant sigma factors: structure and phylogenetic origin. Genome Letters 1, 71–76. Majeran W, Friso G, Asakura Y, Qu X, Huang MS, Ponnala L, Watkins KP, Barkan A, van Wijk KJ. 2012. Nucleoid-enriched proteomes in developing plastids and chloroplasts from maize leaves: a new conceptual framework for nucleoid functions. Plant Physiology 158, 156–189. Malik Ghulam M, Zghidi-Abouzid O, Lambert E, Lerbs-Mache S, Merendino L. 2012. Transcriptional organization of the large and the small ATP synthase operon, atpI/H/F/A and atpB/E, in Arabidopsis thaliana chloroplasts. Plant Molecular Biology 79, 259–272. Martin W, Rujan T, Richly E, Hansen A, Cornelsen S, Lins T, Leister D, Stoebe B, Hasegawa M, Penny D. 2002. Evolutionary analysis of Arabidopsis, cyanobacterial, and chloroplast genomes reveals plastid phylogeny and thousands of cyanobacterial genes in the nucleus. Proceedings of the National Academy of Sciences, USA 99, 12246–12251. Maul JE, Lilly JW, Cui L, dePamphilis CW, Miller W, Harris EH, Stern DB. 2002. The Chlamydomonas reinhardtii plastid chromosome: islands of genes in a sea of repeats. The Plant Cell 14, 2659–2679. Melonek J, Matros A, Trösch M, Mock HP, Krupinska K. 2012. The core of chloroplast nucleoids contains architectural SWIB domain proteins. The Plant Cell 24, 3060–3073. Monde RA, Schuster G, Stern DB. 2000. Processing and degradation of chloroplast mRNA. Biochimie 82, 573–582. Morden CW, Wolfe KH, dePamphilis CW, Palmer JD. 1991. Plastid translation and transcription genes in a non-photosynthetic plant: intact, missing and pseudo genes. EMBO Journal 10, 3281–3288. Mullet JE. 1993. Dynamic regulation ofchloroplast transcription. Plant Physiology 103, 309–313. Myouga F, Hosoda C, Umezawa T, Iizumi H, Kuromori T, Motohashi R, Shono Y, Nagata N, Ikeuchi M, Shinozaki K. 2008. A heterocomplex of iron superoxide dismutases defends chloroplast Plastid RNA polymerases govern chloroplast biogenesis | 6973 Schmitz-Linneweber C, Small I. 2008. Pentatricopeptide repeat proteins: a socket set for organelle gene expression. Trends in Plant Science 13, 663–670. Schröter Y, Steiner S, Matthai K, Pfannschmidt T. 2010. Analysis of oligomeric protein complexes in the chloroplast sub-proteome of nucleic acid-binding proteins from mustard reveals potential redox regulators of plastid gene expression. Proteomics 10, 2191–2204. Schweer J, Geimer S, Meurer J, G. L. 2009. Arabidopsis mutants carrying chimeric sigma factor genes reveal regulatory determinants for plastid gene expression. Plant Cell Physiology 50, 1382–1386. Schweer J, Türkeri H, Kolpack A, Link G. 2010. Role and regulation of plastid sigma factors and their functional interactors during chloroplast transcription: recent lessons from Arabidopsis thaliana. European Journal of Cell Biology 89, 940–946. Sharwood RE, Halpert M, Luro S, Schuster G, Stern DB. 2011. Chloroplast RNase J compensates for inefficient transcription termination by removal of antisense RNA. RNA 17, 2165–2176. Shiina T, Tsunoyama Y, Nakahira Y, Khan MS. 2005. Plastid RNA polymerases, promoters, and transcription regulators in higher plants. International Review of Cytology 244, 1–68. Shimizu M, Kato H, Ogawa T, Kurachi A, Nakagawa Y, Kobayashi H. 2010. Sigma factor phosphorylation in the photosynthetic control of photosystem stoichiometry. Proceedings of the National Academy of Sciences, USA 107, 10760–10764. Solymosi K, Schoefs B. 2010. Etioplast and etio-chloroplast formation under natural conditions: the dark side of chlorophyll biosynthesis in angiosperms. Photosynthesis Research 105, 143–166. Steiner S, Schröter Y, Pfalz J, Pfannschmidt T. 2011. Identification of essential subunits in the plastid-encoded RNA polymerase complex reveals building blocks for proper plastid development. Plant Physiology 157, 1–13. Stoebe B, Maier UG. 2002. One, two, three: nature’s tool box for building plastids. Protoplasma 219, 123–130. Sugiura M. 1992. The chloroplast genome. Plant Molecular Biology 19, 149–168. Sun Q, Zybailov B, Majeran W, Friso G, Olinares PDB, van Wijk KJ. 2009. PPDB, the Plant Proteomics Database at Cornell. Nucleic Acids Research 37, D969–D974. Suzuki JY, Ytterberg AJ, Beardslee TA, Allison LA, Wijk KJ, Maliga P. 2004. Affinity purification of the tobacco plastid RNA polymerase and in vitro reconstitution of the holoenzyme. The Plant Journal 40, 164–172. Taillardat-Bisch A-V, Raoult D, Drancourt M. 2003. RNA polymerase β-subunit-based phylogeny of Ehrlichia spp., Anaplasma spp., Neorickettsia spp. and Wolbachia pipientis. International Journal of Systematic and Evolutionary Microbiology 53, 455–458. Takahashi K, Kasai K, Ochi K. 2004. Identification of the bacterial alarmone guanosine 5’-diphosphate 3’-diphosphate (ppGpp) in plants. Proceedings of the National Academy of Sciences, USA 101, 4320–4324. Tanaka K, Oikawa K, Ohta N, Kuroiwa H, Kuroiwa T, Takahashi H. 1996. Nuclear encoding of a chloroplast RNA polymerase sigma subunit in a red alga. Science 272, 1932–1935. Tanaka R, Tanaka A. 2007. Tetrapyrrole biosynthesis in higher plants. Annual Review of Plant Biology 58, 321–346. Timmis JN, Ayliffe MA, Huang CY, Martin W. 2004. Endosymbiotic gene transfer: organelle genomes forge eukaryotic chromosomes. Nature Reviews Genetics 5, 123–135. Türkeri H, Schweer J, Link G. 2012. Phylogenetic and functional features of the plastid transcription kinase cpCK2 from Arabidopsis signify a role of cysteinyl SH-groups in regulatory phosphorylation of plastid sigma factors. FEBS Journal 279, 395–409. Udy DB, Belcher S, Williams-Carrier R, Gualberto JM, Barkan A. 2012. Effects of reduced chloroplast gene copy number on chloroplast gene expression in maize. Plant Physiology 160, 1420–1431. Ueda M, Takami T, Peng L, Ishizaki K, Kohchi T, Shikanai T, Nishimura Y. 2013. Subfunctionalization of sigma factors during the evolution of land plants based on mutational analysis of liverworth (Marchantia polymorpha L.) MpSIG1.. Genome Biology and Evolution 5, 1836–1848. Valkov VT, Scotti N, Kahlau S, Maaclean D, Grillo S, Gray JC, Bock R, Cardi T. 2009. Genome-wide analysis of plastid gene expression in potato leaf chloroplasts and tuber amyloplasts: transcriptional and posttranscriptional control. Plant Physiology 150, 2030–2044. van der Biezen EA, Sun J, Coleman MJ, Bibb MJ, Jones JD. 2000. Arabidopsis RelA/SpoT homologs implicate (p)ppGpp in plant signaling. Proceedings of the National Academy of Sciences, USA 97, 3747–3752. Van Wijk KJ, Baginsky S. 2011. Plastid proteomics in higher plants: current state and future goals. Plant Physiology 155, 1578–1588. Wagner R, Pfannschmidt T. 2006. Eukaryotic transcription factors in plastids – bioinformatic assessment and implications for the evolution of gene expression machineries in plants. Gene 381, 62–70. Weihe A, Börner T. 1999. Transcription and the architecture of promoters in chloroplasts. Trends in Plant Sciences 4, 169–170. Weng M-L, Blazier JC, Govindu M, Jansen RK. 2014. Reconstruction of the ancestral plastid genome in Geraniaceae reveals a correlation between genome rearrangements, repeats, and nucleotide substitution rates. Molecular Biology and Evolution 31, 645–659. Weng M-L, Ruhlman TA, Gibby M, Jansen RK. 2012. Phylogeny, rate variation, and genome size evolution of Pelargonium (Geraniaceae). Molecular Phylogenetics and Evolution 64, 654–670. Wetzel CM, Jiang CZ, Meehan LJ, Voytas DF, Rodermel SR. 1994. Nuclear-organelle interactions: the immutans variegation mutant of Arabidopsis is plastid autonomous and impaired in carotenoid biosynthesis. The Plant Journal 6, 161–175. Williams-Carrier R, Zoschke R, Belcher S, Pfalz J, Barkan A. 2014. A major role for the plastid-encoded RNA polymerase complex in the expression of plastid transfer RNAs. Plant Physiology 164, 239–248. Wimmelbacher M, Börnke F. 2014. Redoxactivity of thioredoxin z and fructo-kinase-like protein 1 is dispensable for autotrophic growth of Arabidopsis thaliana. Journal of Experimental Botany 65, 2405–2413. Woodson JD, Chory J. 2008. Coordination of gene expression between organellar and nuclear genomes. Nature Reviews Genetics 9, 383–395. Yagi Y, Ishizaki Y, Nakahira Y, Tozawa Y, Shiina T. 2012. Eukaryotictype plastid nucleoid protein pTAC3 is essential for transcription by the bacterial-type plastid RNA polymerase. Proceedings of the National Academy of Sciences, USA 109, 7541–7546. Yamburenko MV, Zubo YO, Börner T. 2015. Abscisic acid affects transcription of chloroplast genes via protein phosphatase 2C-dependent activation of nuclear genes: repression by guanosine-3’5’-bisdiphosphate and activation by sigma factor 5. The Plant Journal 82, 1030–1041. Yu QB, Huang C, Yang ZN. 2014. Nuclear encoded factors associated with the chloroplast transcription machinery of higher plants. Frontiers in Plant Science 5, 316. Yu QB, Lu Y, Ma Q, Zhao T-T, Huang C, Zhao H-F, Zhang X-L, Lv R-H, Yang ZN. 2013. TAC7, an essential component of the plastid transcriptionally chromosome complex, interacts with FLN1, TAC10, TAC12 and TAC14 to regulate chloroplast gene expression in Arabidopsis thaliana. Physiologia Plantarum 148, 408–421. Zghidi W, Merendino L, Cottet A, Mache R, Lerbs-Mache S. 2007. Nucleus-encoded plastid sigma factor SIG3 transcribes specifically the psbN gene in plastids. Nucleic Acids Research 35, 455–464. Zghidi-Abouzid O, Merendino L, Buhr F, Malik Ghulam M, LerbsMache S. 2011. Characterization of plastid psbT sense and antisense RNAs. Nucleic Acids Research 39, 5379–5387. Zhang J, Ruhlman TA, Sabir J, Blazier C, Jansen RK. 2015. Coordinated rates of evolution between interacting plastid and nuclear genes in Geraniaceae. The Plant Cell 27, 563–573. Zhelyazkova P, Sharma CM, Forstner KU, Liere K, Vogel J, Borner T. 2012. The primary transcriptome of barley chloroplasts: numerous noncoding RNAs and the dominating role of the plastid-encoded RNA polymerase. The Plant Cell 24, 123–136. Zhou WB, Cheng YX, Yap A, Chateigner-Boutin AL, Delannoy E, Hammani K, Small I, Huang JR. 2009. The Arabidopsis gene YS1 encoding a DYW protein is required for editing of rpoB transcripts and the rapid development of chloroplasts during early growth. The Plant Journal 58, 82–96.