*Arabidopsis thaliana* has a single gyrase A (AtGYRA) that is localized to both plastids and mitochondria, and has two gyrase B enzymes that are localized to either chloroplasts (AtGYRB1) or mitochondria (AtGYRB2; Wall et al., 2004). T-DNA insertion mutation of *AtGYRA* leads to an embryo-lethal phenotype, whereas T-DNA insertion mutations of both plastidial *AtGYRB1* and mitochondrial *AtGYRB2* result in seedling-lethal phenotypes (Wall et al., 2004). In *Nicotiana benthamiana*, virusinduced silencing of the genes encoding GYRA and GYRB resulted in abnormal nucleoid content and structure of chloroplasts and mitochondria (Cho et al., 2004). *A. thaliana* also encodes a gyrase B-like gene, *GYRB3*; however, it is recently reported that *AtGYRB3* does not encode a gyrase subunit, as AtGYRB3 showed no supercoiling activity and did not interact with AtGYRA (Evans-Roberts et al., 2010). In addition to gyrases, plant organelles contain A-type topoisomerase I, which is a homolog of bacterial topoisomerase I (TopA). Based on localization analysis using a GFP-fusion protein, AtTOP1 was shown to be localized to both chloroplasts and mitochondria (Carrie et al., 2009).

The genome of *Cyanidioschyzon merolae* encodes genes for GYRA and GYRB, which are localized only to the plastid (Moriyama et al., 2014). Similarly, *Cyanidioschyzon merolae* TOP1 (type IA) is also localized only to the plastid. To search for mitochondrial topoisomerases, we examined the subcellular localization of topoisomerases encoded in the *Cyanidioschyzon merolae* genome, and showed that a homolog of eukaryotic TOP2 is targeted to mitochondria. To date, organellar localization of eukaryotic TOP2 has not been reported in plants. In *Cyanidioschyzon merolae*, the gyrase specific inhibitor nalidixic acid arrests not only replication of the plastid genome, but also that of the mitochondrial and nuclear genomes (Itoh et al., 1997; Kobayashi et al., 2009). The localization results of gyrases in *Cyanidioschyzon merolae* suggest that defective plastid replication leads to the arrest of mitochondrial and nuclear replication by a yet unknown mechanism.

#### **DNA LIGASE**

DNA ligase is required for DNA replication, repair, and recombination, as it seals nicked-DNA ends of single-stranded breaks or joins DNA ends after double-stranded breaks. Four DNA ligases have been identified in the *A. thaliana* genome. *A. thaliana* DNA ligase 1 (AtLIG1) is targeted to either the mitochondria or the nucleus when the gene transcript is translated from the first and second initiation codons, respectively (Sunderland et al., 2004, 2006). AtLIG1 is expressed in all tissues of *A. thaliana*, but higher transcript levels are found in young leaves and tissues containing meristem (Taylor et al., 1998). Plastid-targeting of AtLIG1 was not observed for any AtLIG1-GFP constructs translated from possible initiation codons, and the plastidial enzyme functioning as DNA ligase remains unclear. However, it has been noted that AtLIG6 has a putative plastid-targeting peptide at the *N*terminus and might therefore be targeted to plastids (Sunderland et al., 2006).

*Cyanidioschyzon merolae* has a single gene encoding DNA ligase. *Cyanidioschyzon merolae* DNA ligase 1 (CmLIG1) has two methionine residues in its *N*-terminal region and is targeted to both mitochondria and plastids when the transcript is translated from the first and second initiation codons (Moriyama et al., 2014). In our analysis, no nuclear localization was observed when the *N*-terminal peptide of CmLIG1 was fused with GFP. However, the protein subcellular localization prediction software WolfPSORT (http://wolfpsort.seq.cbrc.jp/) detected a nuclear localization signal in CmLIG1. Therefore, CmLIG1 appears to have triple localization in plastids, mitochondria, and the nucleus.

## **SINGLE-STRANDED DNA (ssDNA)-BINDING PROTEIN**

An SSB, AtSSB1, was identified in *A. thaliana* (Edmondson et al., 2005). AtSSB1 is localized to mitochondria, but was also reported to be localized to chloroplasts in the review by Cupp and Nielsen (2014). AtSSB1 binds to ssDNA, but not to dsDNA, and stimulates RecA-mediated strand exchange activity.

Organellar ssDNA-binding proteins (OSBs) comprise the second class of SSBs in plants (Zaegel et al., 2006). OSBs have an SSB-like domain in the central region and one, two, or three C-terminal PDF motifs, which consist of 50-amino acids and are responsible for ssDNA binding. PDF motifs are conserved only in green plants, including *Chlamydomonas reinhardtii*. *A. thaliana* has four OSBs: AtOSB1 and 2 are localized to mitochondria and chloroplasts, respectively, whereas AtOSB3 is localized to both chloroplasts and mitochondria. AtOSB1 and AtOSB2 have been purified as recombinant proteins that showed preferential binding activity to ssDNA, as compared to dsDNA or RNA. Expression analysis of *AtOSB1* using a βglucuronidase (GUS) assay demonstrated that *AtOSB1* is mainly expressed in gametophytic cells. T-DNA insertion mutation of the *OSB1*, *OSB2*, and *OSB3* genes revealed that *osb1* mutants accumulate homologous recombination products of mitochondrial DNA, whereas *osb2* and *osb3* mutants have no visible phenotype. These findings, together with the expression analysis for *AtOSB1*, indicate that AtOSB1 is involved in mitochondrial DNA recombination in gametophytic cells (Zaegel et al., 2006).

Replication protein A (RPA) is a nucleus-localized BBC in eukaryotes and is comprised of three subunits, RPA70, RPA32, and RPA14. The rice genome encodes three RPA70s, three RPA32s, and one RPA14. These RPA subunits combine in different variations to make three types of complexes: type A, B, and C. Among these RPA complexes, the type A complex is localized to chloroplasts in rice (Ishibashi et al., 2006).

The *Cyanidioschyzon merolae* genome encodes a single gene for SSB, but does not contain a gene for OSB. In our analysis, the SSB of *Cyanidioschyzon merolae* is localized only in the mitochondrion, unlike that of *A. thaliana* (Moriyama et al., 2014). We performed the localization analysis using a construct starting from the second methionine codon or starting from the ATA codon located upstream of the first methionine codon; however, none of the constructs showed plastid localization. We also examined the organellar localization of RPAs in *Cyanidioschyzon merolae*, and even though they have no extension sequence at the *N*-terminus, they were localized to the nucleus. Based on these findings, the plastidial SSB in red algae remains unidentified.

### **PRIMER REMOVAL ENZYME**

In *E. coli*, RNA primers are removed by nick translation with the 5- –3 exonuclease and polymerase activities of DNA polymerase I (Langston et al., 2009; Sanyal and Doig, 2012). In contrast, RNaseH1 performs this role in human mitochondria (Kasiviswanathan et al., 2012). Although there are no reports of RNA primer removal enzymes that are specific to the organelles in green plants, two 5- –3 exonucleases (5- –3- EXO1 and 2) having sequence homology to the 5- –3 exonuclease domain of *E. coli* PolI are predicted to be localized to chloroplasts or mitochondria (Sato et al., 2003).

*Cyanidioschyzon merolae* has a gene with high sequence homology to bacterial PolI (Moriyama et al., 2008). The corresponding protein, CmPolI, contains 5- –3 exonuclease and polymerase domains. We demonstrated the plastid localization of CmPolI by immunoblotting and observation of a CmPolI-GFP fusion protein (Moriyama et al., 2008, 2014). Because CmPolI has low processive polymerase activity and no 3- –5 exonuclease activity, the enzyme appears to function in repair and primer removal by nick translation, similar to PolI in *E. coli*. We also showed that *Cyanidioschyzon merolae* RNase HII is localized to the mitochondrion, and that DNA2 nuclease/helicase and FEN1 are localized to the nucleus (Moriyama et al., 2014).

## **PHYLOGENETIC DISTRIBUTION OF ORGANELLAR REPLICATIVE ENZYMES**

#### **ORIGIN OF ENZYMES RELATED TO ORGANELLAR GENOME REPLICATION**

Phylogenetic analyses of bacterial-type replicative enzymes have been performed (Moriyama et al., 2014). We previously suggested that POP did not originate from the PolI of α-proteobacteria or cyanobacteria (**Figure 1**, Moriyama et al., 2008). However, the analyses indicated that red algal DnaB helicase and DnaG primase originated from cyanobacteria (**Figure 3A**). Gyrases A and B also originated from cyanobacteria in both green plants and red algae (**Figure 3B**). Type IA topoisomerase was derived from α-proteobacteria in green plants and from cyanobacteria in red algae (**Figure 3C**). SSB and 5- –3 exonuclease/PolI originated from α-proteobacteria in green plants and red algae (**Figure 3D**). With regard to PolI, as green plants have enzymes with 5- –3 exonuclease domain, but lack 3- –5 exonuclease and DNA polymerase domains (**Figure 3F**), phylogenetic analysis of the 5- –3 exonuclease domain in bacteria and photosynthetic eukaryotes was

performed (**Figures 3D,E**). These results suggest that the organelle replication apparatus of both green plants and red algae is composed of enzymes of various origins, including α-proteobacteria, cyanobacteria, and eukaryotes.

## **REPERTOIRE OF ENZYMES RELATED TO ORGANELLAR GENOME REPLICATION IN PLANTS AND ALGAE**

The enzymes related to organellar DNA replication and recombination in a species of angiosperm, fern, moss, filamentous terrestrial alga, two green algae, and two red algae are listed in **Table 1**. The proteins conserved in the all examined species are POP, TWINKLE, Gyrases, type IA-TOP1, TOP2, and LIG1. DnaB and DnaG are conserved only in red algae. The retention of SSBs is highly variable in photosynthetic eukaryotes. Bacterial-type SSB proteins are conserved in land plants and *Cyanidioschyzon merolae*, whereas OSB proteins are conserved among land plants, including *A. thaliana*, *Physcomitrella patens*, and *Klebsormidium flaccidum*. It was reported that OSB proteins contain a few PDF motifs in addition to an SSB-like domain (Zaegel et al., 2006). According to this classification, *Physcomitrella patens* and *K. flaccidum* have a single OSB, which contains one and two PDF motifs, respectively, in addition to the SSB-like domain. RECA and Whirly (WHY) are recombination-related proteins and are also not uniformly conserved in photosynthetic eukaryotes. For example, *Selaginella moellendorffii* and *Porphyridium purpureum* do not have RECA, red algae do not encode WHY, and *Physcomitrella patens* has neither RECA nor WHY. Conservation of origin-binding protein (ODB) is more limited, as only land plants have this protein. PolI containing 5- –3 exonuclease and DNA polymerase domains is retained only in red algae. However, a protein with the 5- –3 exonuclease domain is found in photosynthetic eukaryotes, with the exception of *Cyanidioschyzon merolae*. Therefore, all photosynthetic eukaryotes contain proteobacteria-derived PolI. RNase H having high homology to RNase HII of *Cyanidioschyzon merolae* is conserved in most plants and algae, with the notable exception of *A. thaliana*. The observed distribution of enzymes that play key roles in replication indicates that they are essentially conserved in all plants and algae. In contrast, because recombination-related enzymes and SSBs are non-uniformly distributed among plants and algae, these enzymes are considered to exhibit high plasticity during evolution.

Based on the presence of enzymes related to organellar genome replication in plant genomes, we propose a model for the substitution of these enzymes in photosynthetic eukaryotes (**Figure 4**). In this model, the ancestor of photosynthetic eukaryotes, prior to its divergence into green and red lineages, contained POP, TWINKLE, DnaB, DnaG, gyrases, type-IA TOP1, TOP2, PolI, RNase HII, SSB, and LIG1. Several of these enzymes, including POP, TWINKLE, gyrases, type-IA TOP1, TOP2, LIG1, and PolI (or 5- –3 exonuclease), were retained by all plants. In the early green lineage, WHY was obtained, and mitochondrial TWINKLE became dually localized in mitochondria and plastids. However, the dual-localization of TWINKLE resulted in the loss of DnaB and DnaG in the green lineage. After the divergence of Chlorophyta, the ancestor of land plants obtained the SSBs, OSB, and ODB. The acquisition of these SSBs, in addition to WHY, in the

green lineage may have partially contributed to exploration of terrestrial habitats, as the high recombinant/repair activity of these enzymes would have potentially allowed the repair of DNA damaged by direct sunlight. The sets of RECA and WHY, and RECA and OSB were lost in Bryophyta and Pteridophyta, respectively, followed by the loss of RNase HII (homolog of *Cyanidioschyzon merolae* CMT626C) in Angiospermae. In red algae, most replication enzymes in the ancestor of photosynthetic eukaryotes are found in present-day species.

## **CONCLUDING REMARKS**

In the past decade, most enzymes related to plastid and mitochondrial DNA replication in plants and algae have been identified.

These studies have revealed that the core enzymes and components involved replication are identical in the plastids and mitochondria of land plants. Because the nuclear genomes of green plants and algae encode these core replicative enzymes, such as POP, TWIN-KLE, gyrases, TOP1, TOP2, LIG1, and 5- –3 exonuclease, which frequently contain putative dual-targeting sequences at the *N*terminus (**Table 1**), it is presumed that the green lineage contains a similar set of plastid and mitochondrial enzymes. In contrast, SSBs and recombination-related enzymes are not universally conserved in the green lineage, suggesting that these enzymes are possibly susceptible to exchange or loss during evolution, leading to the acquisition or creation of species-specific enzymes. Unlike the green lineage, red algae contain different replicative protein

**eukaryotes.**

**Table 1**

*(Continued)*

Moriyama and Sato Organellar genome replication in plants

**Table 1 |**

**Continued**

profiles in plastids and mitochondria. Red algal plastids contain numerous replication proteins that originated from cyanobacteria (Moriyama et al., 2014), suggesting that the mechanism of genome replication in these plastids might be similar to that found in bacteria.

To date, a number of organelle-localized enzymes have been identified. However, biochemical data are lacking for the majority of organellar replication enzymes in plants. The role of an enzyme predicted by homology searches against known enzymes might differ from its actual function or properties. For example, soybean plastid DNA replication ODB shares homology with bacterial and *A. thaliana* SSB, but only binds to dsDNA of the *oriA* sequence in plastid DNA, and not to ssDNA (Lassen et al., 2011).

The regulatory mechanisms controlling the initiation of plant organellar genome replication and the number of organellar DNA copies remains to be explored. Recently, chloroplast DNA replication was shown to be regulated by the cellular redox state in the green alga *Chlamydomonas reinhardtii* (Kabeya and Miyagishima, 2013). Specifically, chloroplast DNA replication was activated

and inactivated by the addition of reducing and oxidative agents, respectively, in both *in vivo* and *in vitro* assays. Light-dependent genome replication was also reported in cyanobacteria, in which DCMU [3-(3,4-dichlorophenyl)-1,1-dimethylurea], an inhibitor of electron transport between the PSII complex and plastoquinone pool, inhibits DNA replication initiation, and DBMIB (2,5 dibromo-3-isopropyl-6-methyl-p-benzoquinone), an inhibitor of electron transport between plastoquinone and cytochrome *b*6*f* complex, inhibits the initiation and elongation of replication (Watanabe et al., 2012; Ohbayashi et al., 2013). Thus, the lightmediated replication of plastid DNA in algae may have originated from cyanobacteria. However, organellar replication in land plants and multicellular plants appears to be regulated by other mechanisms. In land plants, the replication of organellar genomes is restricted to meristematic tissues, and is not associated with the cycle or organellar division (Hashimoto and Possingham, 1989; Fujie et al., 1993). These findings suggest that land plants have more complex regulatory mechanisms controlling the replication of organellar genomes than those operating in algae.

## **ACKNOWLEDGMENTS**

This work was supported in part by Core Research for Evolutional Science and Technology (CREST) from the Japan Science and Technology Agency (JST), and Grants-in-Aid for Young Scientists (B) from the Japan Society for the Promotion of Science (JSPS; no. 25870155).

## **REFERENCES**

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 22 July 2014; paper pending published: 16 August 2014; accepted: 30 August 2014; published online: 17 September 2014.*

*Citation: Moriyama T and Sato N (2014) Enzymes involved in organellar DNA replication in photosynthetic eukaryotes. Front. Plant Sci. 5:480. doi: 10.3389/fpls.2014.00480 This article was submitted to Plant Physiology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Moriyama and Sato. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## Nuclear-encoded factors associated with the chloroplast transcription machinery of higher plants

## *Qing-Bo Yu1,2, Chao Huang1,2 and Zhong-Nan Yang1,2\**

*1 Department of Biology, College of Life and Environmental Sciences, Shanghai Normal University, Shanghai, China 2 Institute for Plant Gene Function, Department of Biology, Shanghai Normal University, Shanghai, China*

#### *Edited by:*

*Thomas Pfannschmidt, University Joseph Fourier Grenoble, France*

#### *Reviewed by:*

*Jin Chen, Michigan State University, USA Takashi Shiina, Kyoto Prefectural University, Japan*

#### *\*Correspondence:*

*Zhong-Nan Yang, Department of Biology, College of Life and Environmental Sciences, Shanghai Normal University, No.100, Rd. GuiLin, Shanghai 200234, China e-mail<EMAIL_ADDRESS>
Plastid transcription is crucial for plant growth and development. There exist two types of RNA polymerases in plastids: a nuclear-encoded RNA polymerase (NEP) and plastid-encoded RNA polymerase (PEP). PEP is the major RNA polymerase activity in chloroplast. Its core subunits are encoded by the plastid genome, and these are embedded into a larger complex of nuclear-encoded subunits. Biochemical and genetics analysis identified at least 12 proteins are tightly associated with the core subunit, while about 34 further proteins are associated more loosely generating larger complexes such as the transcriptionally active chromosome (TAC) or a part of the nucleoid. Domain analyses and functional investigations suggested that these nuclear-encoded factors may form several functional modules that mediate regulation of plastid gene expression by light, redox, phosphorylation, and heat stress. Genetic analyses also identified that some nuclear-encoded proteins in the chloroplast that are important for plastid gene expression, although a physical association with the transcriptional machinery is not observed. This covers several PPR proteins including CLB19, PDM1/SEL1, OTP70, and YS1 which are involved in the processing of transcripts for PEP core subunit as well as AtECB2, Prin2, SVR4-Like, and NARA5 that are also important for plastid gene expression, although their functions are unclear.

**Keywords: plastid transcription, RNA polymerase, PEP, NEP, functional modules**

## **INTRODUCTION**

Plastids are specific organelles in plant and algal cells that are responsible for photosynthesis and some important metabolic pathways. They possess their own genetic material and are generally considered to be of endosymbiotic origin (McFadden and van Dooren, 2004). Similar to bacteria, the DNA is organized into dense particles, the nucleoids (Pfalz and Pfannschmidt, 2013). The genome size from plastids of vascular plants ranges from 120 to 180 kbp and the encoded gene set is highly conserved (Sugiura, 1992). They can be categorized into three groups according to their molecular function of the encoded components: (1) Components of the plastid gene expression machinery (RNA polymerase, ribosomal proteins, tRNAs, and rRNAs); (2) Subunits of photosynthesis-related complexes (Rubisco, PSII, the cytochrome b6f complex, PSI, NAPH dehydrogenase, and ATP synthase), and (3) a few proteins involved in other processes (e.g., ClpP1 and YCF3) (Sugiura, 1992). The chloroplast proteome is estimated to be between 2100 and 3600 proteins (Leister, 2003). Most of the chloroplast proteins are encoded by the nuclear genome and are imported from the cytosol (Li and Chiu, 2010), due to the limited coding capacity of the chloroplast genome. However, chloroplast gene expression is still essential for the development of chloroplasts and the maintenance of chloroplast functions. It involves the action of numerous nuclear-encoded factors, besides proteins encoded by the plastome.  In this paper, we focused on these nuclear-encoded factors for chloroplast transcription.

## **TWO TYPES OF PLASTID RNA POLYMERASES IN HIGHER PLANTS**

Plastid genes are transcribed by two RNA polymerases, the nuclear-encoded RNA polymerase (NEP) and the plastidencoded RNA polymerase (PEP). NEP is a phage-type RNA polymerase with a single subunit (Chang et al., 1999; Lerbs-Mache, 2011). In *Arabidopsis*, the nuclear genome encodes three NEPs. RpoTp is targeted to chloroplast, RpoTm is targeted to mitochondria, and RpoTmp is dually targeted to both organelles (Hess and Borner, 1999). NEP is important for plant development. Inactivation of RpoTp results in defects in plastid gene expression and leaf development (Hricová et al., 2006; Swiatecka-Hagenbruch et al., 2008) while plants with inactivated RpoTmp exhibit several defects, including a plastid gene expression defect, delayed greening and growth retardation of leaves and roots (Courtois et al., 2007). The dysfunction of both NEPs resulted in seedling lethality at a very early developmental stage (Hricová et al., 2006). Although NEP is generally considered to be a single subunit RNA polymerase, recent biochemical analysis revealed that RPOTmp interacts with a thylakoid RING-H2 protein. This protein might mediate the fixation of RPOTmp to thylakoid membranes in order to regulate the transcription of the plastid *rrn* genes (Azevedo et al., 2008).

PEP is composed of four core subunits encoded by the genes *rpoA, rpoB*, *rpoC1,* and *rpoC2* that are located on the plastid genome. PEP exhibits a certain sensitivity to inhibitors of bacterial transcription, such as tagetitoxin, and the group of rifampicin-related drugs, indicating a distinct degree of conservation of these eubacterial-type RNA polymerase during evolution (Liere et al., 2011). Like for bacterial RNA polymerases, the activity/specificity of the PEP core enzyme is regulated by sigma-like transcription factors that are encoded by the nuclear genome of higher plants. In *Arabidopsis*, there exist six chloroplast sigma factors (SIG1–SIG6). These sigma factors might have overlapping as well as specific functions for recognizing a specific set of promoters during chloroplast development (Schweer, 2010; Liere et al., 2011). Besides the sigma factors, however, the core subunits of PEP are associated also with additional proteins (see below) that mediate a number of additional functions to the PEP complex.

NEP and PEP play different roles in plastid gene transcription during plastid development and plant growth (Liere et al., 2011). Based on their transcription by the different RNA polymerases, plastid genes can be grouped into three classes (Hajdukiewicz et al., 1997; Ishizaki et al., 2005). Transcription of photosynthesisrelated genes (such as *psbA*, *psbD*, and *rbcL*) depend largely on PEP (class I), whereas a few house-keeping genes (mostly encoding components of the transcription/translation apparatus, such as *rpoB*) are exclusively transcribed by NEPs (class III). Most of plastid genes, however, are transcribed by both PEP and NEPs (class II). Generally, NEP is more active in the young, non-green tissues early in leaf development. It transcribes housekeeping genes including the four core subunits of PEP polymerase which primarily constitute the plastid gene expression machinery. Once PEP is formed in later developmental stages, it thereafter transcribes the photosynthesis-related genes (Hajdukiewicz et al., 1997; Lopez-Juez and Pyke, 2005; Schweer et al., 2010b) and plastid tRNAs (Williams-Carrier et al., 2014). In the mature chloroplast, the activity of NEP is barely detected, while PEP activity maintains high for chloroplast development and plant growth. Nevertheless, recent investigations demonstrated that both NEP and PEP are present in seeds, and PEP is also important for seed germination. This indicates that PEP exists also in non-photosynthetically active seed plastids (Demarsy et al., 2006).

#### **PEP IS ASSOCIATED WITH NUMEROUS NUCLEAR-ENCODED PROTEINS**

Early biochemical analysis demonstrated that two different forms of the PEP complex exist in higher plant, that is, PEP-A and PEP-B (Pfannschmidt and Link, 1994). PEP-B is composed only of the *rpo* core subunits and is present in both etioplasts and greening chloroplasts. During light-dependent chloroplast development, this PEP-B enzyme is reconfigured into an eukaryote-like enzyme complex, the PEP-A, by association of numerous proteins (Pfannschmidt and Link, 1997; Steiner et al., 2011; Pfalz and Pfannschmidt, 2013). PEP-A is the major RNA polymerase in matured chloroplast of higher plant. Attempts have been focused on the isolation of the plastid RNA polymerase complex and its associated proteins for many years (Pfalz and Pfannschmidt, 2013). Biochemical analyses uncovered that the core *rpo* subunits of PEP are present in both the insoluble RNA polymerase preparation called transcriptionally active chromosome (TAC), and the soluble RNA polymerase preparation (sRNAP) (Krause and Krupinska, 2000; Pfalz et al., 2006; Melonek et al., 2012). The TAC fraction was isolated from lysed plastids through one or two gel filtration chromatography steps and subsequent ultracentrifugation, while the soluble RNA polymerase (sRNAP) is prepared from isolated and lysed plastids *via* several chromatographic purification steps without precipitation by centrifugation (Pfalz and Pfannschmidt, 2013). Based on gel filtration and mass spectrometry analysis from different organisms, including *Nicotiana tabacum* (Suzuki et al., 2004), *spinach* (Melonek et al., 2012), *mustard* (*Sinapis alba*) (Pfannschmidt et al., 2000; Pfalz et al., 2006; Steiner et al., 2011), and *Arabidopsis* (Pfalz et al., 2006) it is estimated that the TAC complex contains 43 nuclear-encoded proteins (**Table 1**). Ten proteins were reproducibly found to be tightly associated with PEP core subunits in *mustard* seedlings and, therefore, were named polymeraseassociated proteins (PAPs) (Steiner et al., 2011). The other proteins were found in the previous reported TAC complex and might represent more loosely attached components of the transcription machinery (Pfalz et al., 2006). Two TAC components, pTAC7 (Yu et al., 2013) and MurE-like (Garcia et al., 2008), were not identified as PAPs in *mustard* (Steiner et al., 2011), however, based on their mutant phenotype in T-DNA inactivation mutants of *Arabidopsis* these two proteins were proposed to be PAPs (Pfalz and Pfannschmidt, 2013). One essential common feature of all PAPs is that they are essential for PEP activity. The *Arabidopsis* knock-out lines for the corresponding genes show all an albino/ivory or pale-green phenotype with severe defects in chloroplast development and PEP-dependent transcription (**Table 1**) (Pfalz et al., 2006; Garcia et al., 2008; Myouga et al., 2008; Arsova et al., 2010; Schröter et al., 2010; Gao et al., 2011; Steiner et al., 2011; Gilkerson et al., 2012; Yagi et al., 2012; Yu et al., 2013). The phenotype of these PAP mutants is identical to that of *rpo*-gene knock-out mutants in tobacco (Allison et al., 1996; Hajdukiewicz et al., 1997; De Santis-MacIossek et al., 1999). In the knockout mutants of AtECB1/SVR4/MRL7 (Qiao et al., 2011; Yu et al., 2014), PEP-Related Development Arrested 1 (PRDA1) (Qiao et al., 2013), and Delayed Greening 1 (DG1) (Chi et al., 2008), the expression of PEP-dependent chloroplast genes is also severely reduced. These proteins have not been identified in PEP complex by previous proteomic analyses (Krause and Krupinska, 2000; Suzuki et al., 2004; Pfalz et al., 2006; Steiner et al., 2011).

Based on proteomic analysis and protein interaction investigation, the TAC complex contains at least 50 proteins of which 46 are nuclear-encoded (**Tables 1, 2**). These nuclearencoded proteins can be classified into several groups including

*(Continued)*

**46**

*\*\*Localization information is from GFP- fusion data and or chloroplast proteomics data /immune analysis.*

*\*\*\*Protein domain information is from PPDB database.*

*\*\*\*\*Molecularfunctiondataisgivenbasedonthereference.N.A.meansthatitsdetailed*

 *molecular function remains unclear. \*\*\*\*\*ThephenotypesoftheknockoutlinesinArabidopsis are indicated.N.A.meansthatthephenotyperemainsunclear.*

 *aThese factors are essential for PEP activity.*

*bLocalizationinformationisfromindividual GFP-fusionexperimentorimmuneanalysis.*

DNA/RNA binding proteins, thioredoxin proteins, kinases, ribosome proteins and proteins with unknown function (**Table 1**). Yeast two-hybrid and other biochemical assays revealed the relationship of some proteins in the PEP complex (**Figure 1**). The interactions between these PAPs are consistent with the biochemical experiments that identified these proteins in the PEP complex under the stringent condition (Steiner et al., 2011). Currently, proteins directly interacting with the PEP core subunits have not been identified in the PEP complex. Immunoprecipitation analysis demonstrated that pTAC3 is associated with the *rpo* subunits (Yagi et al., 2012). However, the direct interaction between pTAC3 and PEP core subunits has not been verified.

## **PROTEINS IN THE PEP COMPLEX WITH DNA/RNA BINDING DOMAIN**

The eukaryotic transcriptional machinery consists of RNA polymerases and various DNA binding proteins, such as transcription factors. These DNA-binding proteins recognize the promoter to regulate downstream gene transcription. In the TAC complex, there are at least 14 proteins with DNA-binding domains (**Table 1**) (Pfalz et al., 2006; Steiner et al., 2011; Pfalz and Pfannschmidt, 2013). pTAC3 belongs to the SAP protein family. The *ptac3* mutant exhibits an albino phenotype with reduced PEP-dependent plastid transcription. It is unclear yet if pTAC3 can bind to a specific DNA region in order to regulate plastid gene transcription (Yagi et al., 2012). pTAC6 is essential for chloroplast transcription (Pfalz et al., 2006) since the expression of the *psbA* gene was barely detectable in the *ptac6* mutant, compared with that in *ptac2* and *ptac12* (Pfalz et al., 2006). It is likely that pTAC6 is a specific regulator for *psbA* (Pfalz et al., 2006), however, to date its function remains enigmatic. In bacteria, there exist two transcription termination mechanisms; Rho-independent transcription termination and Rho-dependent termination. The mitochondrial transcription termination factor (mTERF) family was identified to regulate mitochondrial gene expression including transcription termination (Kleine, 2012). pTAC15 is a member of the mTERF protein family (Pfalz et al., 2006). Whether it can terminate the transcription of PEP-dependent plastid genes needs to be verified.

The TAC complex contains at least six RNA-binding proteins including ZmWhy1, pTAC10, the elongation factor EF-Tu, and three ribosomal proteins, S3, L12-A, and L26 (**Table 1**). Whirly proteins belong to a small nuclear transcription factor family commonly found in plants. In *Arabidopsis*, pTAC1/AtWhy1 and pTAC11/AtWhy3 can bind DNA (Xiong et al., 2009). They are required to maintain the stability of the plastid genome (Maréchal et al., 2009). The whirly 1 ortholog in *maize* (ZmWHY1/pTAC1) can bind both RNA and DNA, and co-immuno-precipitated with chloroplast RNA splicing 1 (CRS1) (Prikryl et al., 2008). pTAC10 contains a S1 domain and has RNA binding activity in tobacco (Jeon et al., 2012), and it may be one substrate of chloroplast-target casein kinase 2 (cpCK2) (Reiland et al., 2009). The phosphorylation of pTAC10 may affect its RNA binding. The detailed function of the elongation factor EF-Tu and the ribosomal proteins S3, L12-A, and L26 in chloroplast is not reported. The existence of these RNA-binding proteins, however, suggests that there exists a translation subdomainin the TAC/nucleoid.

**Table 1 | Continued**

**2 | Individual mutant analysis identifies several factors which affect PEP-dependent chloroplast transcription in**

*\*\*Localization information is from GFP- fusion data.*

*\*\*\*Protein domain information is from PPDB database.* *\*\*\*\*Molecular function data is given based on the reference. U.K. means that its detailed molecular function remains unclear.*

*aThese factors are associated with the PAPs or the core subunits. bThesefactorsregulateplastidtranscriptionwithunknownmechanism.*

 *core subunits.*

 *cThesefactorsindirectlyaffectPEPactivitythroughregulating theprocessingofchloroplasttranscriptsencodingthe*

mechanisms are unclear.

## **CONNECTIONS OF REGULATORY MODULES WITH THE RNA POLYMERASE**

chloroplast transcription. Several PPR proteins including CLB19,

Light plays highly important roles in the regulation of plastid gene transcription. The majority of PAPs (Pfalz and Pfannschmidt, 2013) and most sigma factor genes of higher plants are lightinduced (Lerbs-Mache, 2011). Plastome-wide PEP-DNA association is also a light-dependent process (Finster et al., 2013). In plants, light plays an important role in almost every facet of plant growth and development through the action of photoreceptors. Interestingly, pTAC12 is an intrinsic subunit of the PEP complex (Pfalz et al., 2006; Steiner et al., 2011), but it was also identified as HEMERA and localized in both the nucleus and the chloroplast (Chen et al., 2010). pTAC12/HEMERA was considered as a proteolysis-related protein involved in phytochrome signaling in the nucleus (Chen et al., 2010). Its function in the PEP complex is unknown so far, but it was uncovered that pTAC12 interacts with pTAC14 in the yeast-two-hybrid system (Gao et al., 2011) suggesting that these two proteins might be also interaction partners in the native complex.

Chloroplasts are the site of photosynthesis that also produces reactive oxygen species (ROS). During photosynthesis, unbalanced excitation of the two photosystems affects the redox state of the electron transport chain which in turn serve as signals for plant acclimation responses. The PEP complex is a major target of such photosynthetic redox signals (Dietz and Pfannschmidt, 2011). Thioredoxin z (Trx Z) is a novel thioredoxin protein with disulfide reductase activity *in vitro*. It interacts with two fructokinase-like proteins FLN1 and FLN2 in the yeast two hybrid system and is also a component of the PEP complex (Pfalz et al., 2006; Steiner et al., 2011) (**Figure 1**). Trx-Z mediated redox change of FLN2 during light–dark transitions (Arsova et al., 2010). Recent studies identified AtECB1/MRL7 as a thioredoxinfold like protein with thioredoxin activity (Yu et al., 2014) that interacts with Trx Z in the PEP complex (Powikrowska et al., 2014; Yu et al., 2014). These two proteins thus may form a functional module to mediate redox signaling from thylakoids toward the RNA polymerase but the functional details of these interactions are completely unknown. Further redox mediators might be Fe Superoxide Dismutase 2 (FSD2) and FSD3, two iron superoxide dismutases, and PRDA1 is a chloroplast protein without any known domain. *prda1* and *fsd2 fsd3* knock out mutants are highly sensitive to oxidative stress (Myouga et al., 2008; Qiao et al., 2013). These proteins, therefore, may act as ROS scavengers in order to protect the PEP complex. The interactions between AtECB1 and PRDA1, FSD2, FSD3 suggest that the redox signaling pathway and ROS scavengers are eventually associated.

Protein phosphorylation is a very important post-translational modification in eukaryotic cells that regulates many cellular processes. In chloroplast, the phosphorylation of chloroplast proteins affects photosynthesis, metabolic functions and chloroplast transcription (Baginsky and Gruissem, 2009). The PEP complex appears to interact with a so-called plastid transcription kinase (PTK), named cpCK2 (Ogrzewalla et al., 2002). The *Arabidopsis* sigma factor 6 was reported to be phosphorylated by cpCK2 (Schweer et al., 2010a). Furthermore, pTAC5, pTAC10, and pTAC16, were also predicated to be phosphorylated by cpCK2 (Reiland et al., 2009). The enzyme activity of cpCK2 was inhibited by GSH, which suggests that cpCK2 is generally under SH-group redox regulation (Baginsky et al., 1999; Turkeri et al., 2012). Biochemical analyses of *mustard* seedlings during photosynthetic acclimation suggested that redox signals in chloroplasts are linked to chloroplast transcription *via* the combined action of phosphorylation and thiol-mediated regulation events (Steiner et al., 2009). Proteins related with phosphorylation and redox signaling are closely located in the PEP complex which is in agreement with the results of the physiological studies for plastid gene expression.

Heat stress is a major abiotic factor for plants, that leads to severe retardation in plant growth and development. To maintain the process of chloroplast transcription under heat stress and to support the survival of the plant, the chloroplast transcriptional machinery needs to deal with heat stress to a certain extent. The protein pTAC5 is a C4-type zinc finger DnaJ protein with disulfide isomerase activity. Its expression is induced by heat stress (Zhong et al., 2013) and, subsequently, pTAC5 and Heat Shock Protein 21 (HSP21) form a heterocomplex, although they are not PAP members of the PEP complex (Zhong et al., 2013). pTAC5 as well as HSP21 may protect chloroplast transcription under heat stress.

## **OTHER NUCLEAR ENCODED FACTORS THAT REGULATE PEP ACTIVITY**

In addition to the intrinsic components of PEP complex, multiple additional factors were identified to regulate the processing of PEP core subunit transcripts and PEP activity by individual mutant analysis. Both *Chloroplast Biogenesis19* (*CLB19*) (Chateigner-Boutin et al., 2008) and *Pigment-Deficient Mutant 1*(*PDM1*) (Wu and Zhang, 2010; Yin et al., 2012) genes encode pentatricopeptide repeat proteins. CLB19 is involved in the editing of the *rpoA* transcript (Chateigner-Boutin et al., 2008), while *PDM1* is associated with *rpoA* polycistronic for *rpoA* cleavage (Wu and Zhang, 2010; Yin et al., 2012). Recent investigations demonstrated that PDM1/Seedling Lethal1 (SEL1) was also involved in *accD* RNA editing (Pyo et al., 2013). The PPR protein OTP70 was reported to affect the splicing of the *rpoC1* transcript (Chateigner-Boutin et al., 2011). The gene *Yellow Seedling 1* (*YS1*) encoding a PPR-DYW protein is required for editing of *rpoB* transcripts (Zhou et al., 2009). The common feature of the *Arabidopsis* knockout lines for all these proteins is that the plastid expression pattern in these mutants is similar to that of *rpo*-gene knock-out mutants in tobacco (Chateigner-Boutin et al., 2008, 2011; Zhou et al., 2009; Wu and Zhang, 2010; Pyo et al., 2013).

Functional analyses revealed that several proteins including *Arabidopsis* Early Chloroplast Biogenesis 2 (AtECB2) (Yu et al., 2009), Plastid redox insensitive 2 (Prin2) (Kindgren et al., 2012), SVR4-Like (Powikrowska et al., 2014), and NARA5 (Ogawa et al., 2009), are also essential for PEP-dependent chloroplast transcription. However, it is unclear if they are directly associated with the PEP complex. AtECB2 encodes a pentatricopeptide repeat protein, and is involved in editing of *accD* and *ndhF* chloroplast transcripts (Yu et al., 2009; Tseng et al., 2010). The defective editing in *ecb2* is unlikely to affect PEP-dependent plastid gene expression. How AtECB2 affects plastid gene expression is still unclear. NARA5 encodes a chloroplast-localized phosphofructokinase B-type carbohydrate kinase family protein, which might be involved in massive expressions of plastid-encoded photosynthetic genes in *Arabidopsis* (Ogawa et al., 2009). The Prin2 is a small protein possibly involved in redox-mediated retrograde signaling in chloroplast (Kindgren et al., 2012) and the SVR4-like is a homolog of AtECB1/SVR4/MRL7, encoding a chloroplast protein essential for proper function of the chloroplast in *Arabidopsis* (Powikrowska et al., 2014). All these proteins may reversibly associate with the PEP complex but detailed studies are necessary to understand their functional roles and connections with the RNA polymerase. Alternatively, these proteins may act as signaling factors in order to mediate environmental stimuli and plastid gene expression.

## **CONCLUDING REMARKS**

Plants grow under very different environment conditions and photosynthesis is the major function of chloroplast which is important for plant growth and development. Plastid gene expression is essential for chloroplast development and normal functions including photosynthesis. The PEP complex is the major RNA polymerase activity in mature chloroplasts. Proteomic and genetic analyses identified that at least 50 nuclearencoded proteins in higher plant are important for PEP dependent plastid gene expression. These proteins may form several functional modules within the nucleoid or TAC in order to mediate plastid gene expression in response to light, redox changes, phosphorylation and heat stress or to protect the PEP complex from ROS damage. The large number of nuclear-encoded proteins reveals the complexity of plastid gene expression and regulation that is greatly different from the gene expression in the nucleus or in prokaryotes. However, the current knowledge about plastid transcription is quite limited and the investigation of the relationship between transcription, post-transcriptional processing as well as translation in the nucleoid could provide novel insights into chloroplast gene expression.

## **ACKNOWLEDGMENTS**

We are grateful to Prof. Thomas Pfannschmidt from Grenoble University for his help in editing the manuscript. This work was supported by grants from the National Science Foundation of China (Grant no. 31100965 and Grant no. 31370271).

## **REFERENCES**

transcription of chloroplast genes is compensated by a second phage-type RNA polymerase. *Nucleic Acids. Res.* 36, 785–792. doi: 10.1093/nar/gkm1111

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 31 March 2014; accepted: 14 June 2014; published online: 03 July 2014.*

*Citation: Yu Q-B, Huang C and Yang Z-N (2014) Nuclear-encoded factors associated with the chloroplast transcription machinery of higher plants. Front. Plant Sci. 5:316. doi: 10.3389/fpls.2014.00316*

*This article was submitted to Plant Physiology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Yu, Huang and Yang. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## Recent advances in the study of chloroplast gene expression and its evolution

## *YusukeYagi 1 and Takashi Shiina2 \**

*1 Faculty of Agriculture, Kyushu University, Fukuoka, Japan*

*2 Graduate School of Life and Environmental Sciences, Kyoto Prefectural University, Kyoto, Japan*

#### *Edited by:*

*Thomas Pfannschmidt, Joseph Fourier University, France*

#### *Reviewed by:*

*Frederik Börnke, Leibniz-Institute for Vegetable and Ornamental Crops, Germany Hannetz Roschzttardtz, University of Wisconsin–Madison, USA*

#### *\*Correspondence:*

*Takashi Shiina, Graduate School of Life and Environmental Sciences, Kyoto Prefectural University, Kyoto 606-8522, Japan e-mail<EMAIL_ADDRESS>
Chloroplasts are semiautonomous organelles which possess their own genome and gene expression system. However, extant chloroplasts contain only limited coding information, and are dependent on a large number of nucleus-encoded proteins. During plant evolution, chloroplasts have lost most of the prokaryotic DNA-binding proteins and transcription regulators that were present in the original endosymbiont. Thus, chloroplasts have a unique hybrid transcription system composed of the remaining prokaryotic components, such as a prokaryotic RNA polymerase as well as nucleus-encoded eukaryotic components. Recent proteomic and transcriptomic analyses have provided insights into chloroplast transcription systems and their evolution. Here, we review chloroplast-specific transcription systems, focusing on the multiple RNA polymerases, eukaryotic transcription regulators in chloroplasts, chloroplast promoters, and the dynamics of chloroplast nucleoids.

**Keywords: chloroplast, transcription, PEP, NEP, pTAC, nucleoid**

## **INTRODUCTION**

Chloroplasts are believed to have arisen from an endosymbiotic event between a photosynthetic cyanobacterium and the ancestral eukaryotic cell. Although chloroplasts of modern plants and algae have retained the genome of the symbiont, that genome has markedly shrunk over endosymbiotic evolution. Many chloroplast-encoded genes were lost or transferred to the nucleus soon after endosymbiosis. Thus, chloroplast genomes of extant land plants have only 50 protein-coding genes involved in photosynthesis, gene expression, lipid metabolism and other processes, 30 tRNA genes and full sets of rRNA genes. In spite of their small genomes (0.15 Mbp in land plant chloroplasts versus 3 Mbp in cyanobacteria), chloroplast gene expression is regulated by more complex systems compared to the simple prokaryotic regulatory system. Chloroplast gene expression is mediated by two distinct types of RNA polymerase (RNAP) and is highly dependent on post-transcriptional regulation, such as the processing of polycistronic transcripts, intron splicing and RNA editing. Moreover, recent RNA-seq analyses of chloroplast transcripts identified unexpected diversifications of RNA molecules, such as non-coding and antisense RNAs (Hotto et al., 2011; Zhelyazkova et al., 2012). However, the genes encoded in chloroplast genomes are insufficient to regulate their complicated gene expression, and so the chloroplast gene expression machinery includes various nucleus-encoded regulatory components.

Although basic chloroplast gene expression is mediated by prokaryotic machineries derived from the ancestral cyanobacterium, chloroplasts lost their homologs of bacterial regulatory elements such as transcription factors (TFs) and nucleoid proteins at an early stage of their evolution. Genomics and proteomics analyses of chloroplast proteins in *Arabidopsis thaliana* have suggested that 60% of the chloroplast proteome may have been newly acquired from the nuclear genome of host cells after the endosymbiotic event (Abdallah et al., 2000). Indeed, recent analyses of the chloroplast nucleoid proteins identified many non-bacterial components that play critical roles in chloroplast gene expression including transcription, post-transcriptional RNA processing, and translation. Here, we summarize the current knowledge regarding the chloroplast gene expression system.

## **TWO BASIC CHLOROPLAST TRANSCRIPTION MACHINERIES WITH DIFFERENT EVOLUTIONARY ORIGIN**

Chloroplast gene expression is largely dependent on prokaryotic machineries derived from the ancestral cyanobacterium. The bacterial multi-subunit RNAP is composed of a core Rpo complex, which has the catalytic enzyme activity, and a sigma factor, which recognizes promoter sequences (Ishihama, 2000). Chloroplasts contain the bacterial-type RNAP, called plastid-encoded plastid RNAP (PEP), which shares functional similarity with the bacterial RNAP (Igloi and Kossel, 1992; **Figure 1A**) However, all genes for chloroplast sigma factors have been transferred to the nuclear genome, whereas genes for core subunits are typically retained in the chloroplast genome as *rpoA*, *rpoB*, *rpoC1*, and *rpoC2*.

Early work demonstrated that almost all photosynthesis-related transcripts are significantly reduced in PEP-deficient plants, such as ribosome-deficient mutants of barley (*Hordeum vulgare*), *iojap* mutants of maize and tobacco mutants with disrupted *rpo* genes generated by gene targeting using chloroplast transformation (Han et al., 1992; Hess et al., 1993, 1994; Allison et al., 1996; De Santis-MacIossek et al., 1999), whereas a set of housekeeping genes are still active in these mutants. The inhibitor sensitivity of this transcription activity is similar to that of phage T7 RNAP, but not to that of bacterial RNAP (Kapoor et al., 1997; Sakai et al., 1998). In *Arabidopsis*, three phage-type RNAP genes

were identified and their subcellular localization was determined (*RpoTp*: *chloroplasts*, *RpoTm*: *mitochondria*, *RpoTmp*: *chloroplast and mitochondria*; Hedtke et al., 1997, 2000; Weihe and Borner, 1999; **Figure 1A**). RpoTmp and RpoTp likely represent nuclear encoded RNAP (NEP) enzyme in chloroplasts [reviewed in (Liere et al., 2011)], while RpoTmp has been identified in dicotyledonous plants such as *Arabidopsis* and tobacco but not in monocotyledonous plant genomes (Chang et al., 1999; Ikeda and Gray, 1999; Emanuel et al., 2004).

Only one *RpoT* gene has been identified in green algae, such as *Chlamydomonas reinhardtii*, *Ostreococcus tauri*, and *Thalassiosira pseudonana,* which likely encodes mitochondrial RNAP (Maier et al., 2008). Similarly, the genome of the lycophyte *Selaginella moellendorffii* contains only one *RpoT* gene, the product of which has been shown to target mitochondria (Yin et al., 2009). On the other hand, the moss *Physcomitrella patens* has three *RpoT* genes. However, all GFP-fused moss RpoTs were detected exclusively in mitochondria, suggesting that the moss *RpoT* genes also encode mitochondrial RNAP (Kabeya et al., 2002; Richter et al., 2002, 2013). Moreover, phylogenetic analysis of plant *RpoT* genes suggests that NEP appeared through the gene duplication of mitochondrial RNAP after the separation of angiosperms from gymnosperms (Yin et al., 2010).

## **SELECTIVE CHLOROPLAST TRANSCRIPTION BY PEP AND NEP**

Chloroplast genes can be categorized into three subgroups, classes I–III: class I photosynthesis-related genes are mainly transcribed by PEP; Class II includes many housekeeping genes (*clpP* and the *rrn* operon) that are transcribed by both PEP and NEP; class III genes (*accD* and the *rpoB* operon) are exclusively transcribed by NEP (Allison et al., 1996; Hajdukiewicz et al., 1997).

PEP recognizes standard chloroplast promoters resembling the bacterial σ70 type promoters with −10 and −35 consensus elements (Gatenby et al., 1981; Gruissem and Zurawski, 1985; Strittmatter et al., 1985; Shiina et al., 2005; **Figure 1A**). A genome-wide mapping of transcription start sites (TSSs) by RNA sequencing in barley green chloroplasts demonstrated that 89% of the mapped TSSs have a conserved −10 element (TAtaaT) at three to nine nucleotides upstream, while the −35 element was mapped upstream of the −10 element in only 70% of the TSSs (Zhelyazkova et al., 2012). These results suggest that most genes are transcribed from σ70-type promoters by PEP in green leaves.

Higher plants have multiple sigma factors that are expected to confer promoter specificity upon the PEP core complex (Shiina et al., 2005; Lerbs-Mache, 2011). Molecular genetic analyses revealed that SIG2 is responsible for the transcription of a group of tRNA genes, but not photosynthesis genes (Kanamaru et al., 2001), while SIG6 is essential for the transcription of a wide range of photosynthesis-related genes at an early stage of chloroplast development (Ishizaki et al., 2005). It seems likely that SIG2 and SIG6 work in cooperation during light-dependent chloroplast development (Hanaoka et al., 2003; Ishizaki et al., 2005). In addition, SIG3 and SIG4 have been shown to specifically target *psbN* and *ndhF* genes in *Arabidopsis* (Favory et al., 2005; Zghidi et al., 2007). Recently, ChIP analysis of SIG1 revealed the target genes (*psaAB*, *psbBT*, *psbEFLJ*,*rbcL,* and *clpP*;Hanaoka et al., 2012). SIG5 is a unique sigma factor whose expression is rapidly induced by various environmental stresses such as a high osmolarity, or salinity, a low temperature as well as high-light stress (Tsunoyama et al., 2002; Nagashima et al., 2004). SIG5 likely recognizes specific promoters, including the *psbD* light-responsive promoter (LRP), and mediates stress-induced transcription in chloroplasts (Nagashima et al., 2004; Tsunoyama et al., 2004). Taken together, it is likely that each chloroplasts sigma factor is responsible for the transcription of a distinct set of genes, and plays specific roles in transcriptional regulation in response to developmental and/or environmental cues.

Phylogenetic analysis revealed that chloroplast sigma factors are related to essential group 1 and non-essential group 2 sigma factors in bacteria. *Chlamydomonas*, a single-celled green alga, possesses a single sigma factor that is related to SIG2 in land plants, suggesting the absence of multiple sigma factor-mediated transcriptional regulation in chloroplasts. Endosymbiosis of ancestral cyanobacteria in plant cells may have reduced the need for transcriptional regulation in chloroplasts and caused the reduction of the number of sigma factors in green algae. On the other hand, in liverwort (*M. polymorpha* L*.*) and moss (*P. patens*), three sigma factors related to SIG1, SIG2, and SIG5 are encoded in the nucleus (Shiina et al., 2009; Ueda et al., 2013). The multiple sigma factors in bryophytes may show a promoter preference and play roles in tissue-specific and stress-responsive transcriptional regulation in chloroplasts (Hara et al., 2001; Ichikawa et al., 2004; Kanazawa et al., 2013; Ueda et al., 2013).

Most NEP promoters (*rpoB*, *rpoA,* and *accD*) share a core sequence, the YRTA motif (type-Ia; Liere and Maliga, 1999; Weihe and Borner, 1999; Hirata et al., 2004; **Figure 1A**). The YRTA motif is similar to motifs found in promoters of plant mitochondria (Binder and Brennicke, 2003; Kuhn et al., 2005). In addition, GAAbox has been identified upstream of the YRTA motif in a subclass of NEP promoters (type-Ib; Kapoor and Sugiura, 1999). In contrast to these standard NEP promoters, type-II NEP promoters mapped upstream of the dicot *clpP* gene lack the YRTA motif and are dependent on downstream sequences of the TSS (Weihe and Borner, 1999). Furthermore, it has been shown that the *rrn* operon and certain tRNAs are transcribed from other non-consensus-type NEP promoters [Reviewed by (Liere et al., 2011)].

Although the class I genes have been clarified as being exclusively transcribed by PEP, the genome-wide mapping of TSSs in barley revealed that most genes including photosynthesis genes have both PEP and NEP promoters. It seems likely that NEP supports transcription of photosynthesis genes at the early stage of seedling greening (Zhelyazkova et al., 2012). Interestingly, 73% of NEP-dependent TSSs possess the YRTA motif typical for type-Ia and -Ib NEP promoters, whereas GAA-boxes have been barely mapped upstream of the barley NEP promoters. These results suggest that type-Ia, but not type-Ib NEP promoters play a major role in transcription by NEP in barley chloroplasts. In contrast, type-II NEP promoters, which are dependent on downstream sequences of the TSSs, were identified in barley as well as tobacco.

## **THE LARGE TRANSCRIPTION COMPLEX IN HIGHER PLANT CHLOROPLASTS**

Two types of PEP-containing preparation have been biochemically isolated in mustard and *Arabidopsis*: soluble RNAP (sRNAP) and plastid transcriptionally active chromosome (pTAC) attached to chloroplast membranes (Hess and Borner, 1999). Transcription by sRNAP is dependent on exogenously added template DNA, whereas the pTAC can initiate transcription from the endogenous chloroplast DNA (Igloi and Kossel, 1992; Krause et al., 2000). Interestingly, protein compositions of highly purified sRNAP fractions are dependent on chloroplast development (Pfannschmidt and Link, 1994). The sRNAP of etioplasts in dark-grown leaves is a naked RNAP without additional subunits similar to the *E. coli* RNAP core complex. Etioplasts convert to photosynthetically active chloroplasts in the presence of light. During chloroplast development in mustard, the RNAP develops a more complex form that contains 13 additional polypeptides (Pfannschmidt and Link, 1994). It seems likely that the simple sRNAP in etioplasts converts to a more complex sRNAP in chloroplasts by recruiting additional components during chloroplast development.

Proteomic analyses of pTAC fractions isolated from mature chloroplasts of *Arabidopsis* and mustard have identified 35 polypeptides including 18 novel proteins termed pTAC1–pTAC18, in addition to PEP core subunits, DNA polymerase, DNA gyrase, Fe-dependent superoxide dismutases (FeS-ODs), phosphofructokinase–B type enzymes (PFKB1 and PFKB2), thioredoxin, and three ribosomal proteins (Pfalz et al., 2006). DNA- and/or RNA-binding domains, protein–protein interaction domains, or epitopes with other reported cellular functions have been identified in some of pTAC proteins. Most *Arabidopsis* knockout mutants of pTAC proteins exhibit seedling-lethal symptoms or chlorophyll-deficient phenotypes. PEP-dependent transcription is significantly impaired in the pTAC mutants, whereas NEP-dependent transcription is up-regulated. These phenotypes and chloroplast gene expression patterns are reminiscent of those of *rpo* mutants (Allison et al., 1996; Hajdukiewicz et al., 1997), suggesting a critical role for pTAC proteins in PEP transcription.

Affinity purification of the tobacco PEP (Suzuki et al., 2004) and more recent analysis of subunits of the PEP complex in mustard (Steiner et al., 2011) and tobacco complex identified at least 10 PEP-associated proteins (PAPs). Recently, chromatin immunoprecipitation assays were performed with one of the typical PAPs, pTAC3/PAP1. The results revealed that pTAC3/PAP1 associates with the PEP complex in all three steps of the transcription cycle including initiation, elongation and termination, suggesting that pTAC3/PAP1 is an essential component of the chloroplast PEP complex (Yagi et al., 2012). Several studies on protein–protein interactions among PAPs have been reported [reviewed in (Pfalz and Pfannschmidt, 2013)]. Almost all PAP genes, except for *Trxz*, are conserved among all land plants, but not in the green alga *Chlamydomonas*. It seems likely that terrestrial plants may have acquired non-cyanobacterial novel PEP components during land plant evolution to regulate plastid transcription (Pfalz and Pfannschmidt, 2013).

It has been suggested that a series of checkpoints control the establishment of the chloroplast transcription machinery (Steiner et al., 2011; Pfalz and Pfannschmidt, 2013). In imbibed seeds, predominant NEP is responsible for transcription of housekeeping genes. NEP also transcribes chloroplast-encoded *rpo* genes for PEP core subunits to produce a basic PEP-B complex (NEP– PEP cascade). PEP-B is responsible for the major activity in etioplasts and in an early stage of greening. This step may be the first checkpoint. Subsequently, PEP-B associates with PAPs and converts them into a larger PEP complex (PEP-A) during light-dependent chloroplast development. PEP-A formation is strictly dependent on light. Indeed, it has been reported that expression of the *pTAC3*/*PAP1* gene is induced by light during the greening process (Yagi et al., 2012). PAP mutants mostly show the aberrant development of chloroplasts and transcription of chloroplast-encoded genes, suggesting their essential roles in PEP-A. Furthermore, recent genome-wide analysis of the chloroplast transcriptome revealed reduced expressions of numerous chloroplast tRNAs in several PAPs mutants (pTAC2, pTAC12, MurE, PRIN2), suggesting that PAPs play a major role in tRNA transcription in chloroplasts (Williams-Carrier et al., 2014). Thus PAPs are also responsible for protein translation in chloroplasts. Therefore, the assembly of PAPs in the PEP-A complex may be the second checkpoint in the establishment of the chloroplast transcription machinery. To prevent uncontrolled chloroplast development under adverse conditions, these check points likely play critical roles in the control of chloroplast gene transcription.

## **THE PLASTID NUCLEOIDS: DYNAMICS AND UNIQUE COMPONENTS**

The plastid DNA exists as large protein-DNA complexes named the plastid nucleoid. Plastid nucleoids contain an average of 10–20 copies of the plastid DNA (Kuroiwa, 1991), and their size, shape, and distribution vary depending on the plastid type (Miyamura

et al., 1986; Sato et al., 1997). Each chloroplast contains ∼20 nucleoids that are randomly located on the thylakoid membranes. Immature proplastids in seeds contain only one nucleoid that is located at the center of the organelle. The plastid nucleoids divide into a few small dots and redistribute to the inner envelope membranes during early chloroplast development. At a later stage of chloroplast development, nucleoids are relocated to the thylakoid membranes. It has been suggested that plastid nucleoid organization and dynamics are involved in the regulation of plastid function, gene expression and differentiation. Two DNA-binding proteins, PEND and MFP1, are likely responsible for the association of nucleoids with chloroplast membranes (Sato et al., 1998; Jeong et al., 2003; **Figure 1B**).

In *E. coli*, chromosome DNA packaging patterns affect gene expression, and are regulated by nucleoid-associated proteins (NAPs) such as HU, H-NS, and FIS [reviewed by (Dillon and Dorman, 2010)]. Among bacterial NAPs, HU is one of the major DNA-binding proteins and is involved in chromosome DNA packaging. HU-like proteins (HLPs) are conserved in cyanobacteria, the red alga *Cyanidioschyzon merolae* (Kobayashi et al., 2002), and the green alga *Chlamydomonas* (Karcher et al., 2009). The HLP in *Chlamydomonas* has roles in nucleoid maintenance and gene expression, indicating conserved roles of HU during chloroplast evolution (Karcher et al., 2009). However, land plants including mosses and flowering plants have not only lost the HU genes, but also other prokaryotic DNA-binding proteins (**Figure 2**). Nevertheless, atomic force microscopy observations revealed that plastid nucleoids are highly organized and form a beads-on-astring structure similar to that observed in bacterial nucleoids, suggesting that another host cell-derived DNA-binding protein took over the functions of HU (Melonek et al., 2012). Recently, eukaryotic SWIB (SWI/SNF complex B) domain containing proteins have been identified from the proteome of a further-enriched pTAC fraction (TAC-II) of spinach chloroplasts (Melonek et al., 2012). SWIB4 that has a histone H1 motif, can functionally complement an *E. coli* mutant lacking the histone-like nucleoid structuring protein H-NS, indicating that SWIB4 is the most likely counterpart of the bacterial NAPs in chloroplasts. EM observation of isolated pTAC identified chromatin-like beaded structures with several protruding DNA loops, suggesting that pTACs represent a subdomain of the chloroplast nucleoid (Yoshida et al., 1978; Briat et al., 1982). These findings suggest that pTAC forms a central core of the plastid nucleoid and a transcription factory (**Figure 1B**).

The proteomes of highly enriched nucleoid fractions have been characterized in maize proplastids and mature chloroplasts (Majeran et al., 2012). As expected, the chloroplast nucleoids contain all PEP core Rpos and PAPs, and almost all other pTAC proteins. Furthermore, additional proteins involved in post-transcriptional processes, such as pentatricopeptide repeat proteins (PPR proteins), mitochondrial transcription factor (mTERF)-domain proteins, 70S ribosomes and ribosome assembly factors have been identified in the proteome of the chloroplast nucleoids, suggesting that several post-transcriptional events including RNA processing, splicing and editing, and translation, occur in nucleoids, and that these processes are co-regulated with transcription (**Figure 1B**). Human mitochondrial nucleoids have

NAPs.

been shown to form layered structures, the central core involved in replication and transcription, and the peripheral region where translation and complex assembly may occur (Bogenhagen et al., 2008). By analogy, the further characterization of plastid nucleoids will provide insights into the structural specialization of plastid nucleoids; DNA maintenance and transcription in a core domain and various aspects of RNA metabolism in several subdomains.

endosymbiosis, the primary chloroplast lost its sigma factors, except the

## **PERSPECTIVE**

Recent proteomic and transcriptomic researches and the development of novel ChIP and imaging technologies have advanced the understanding of the molecular basis of RNAP complexes and nucleoid architecture. In land plants, neither the nuclear nor chloroplast genome encodes prokaryotic transcription factors and nucleoid proteins, whereas chloroplasts retain prokaryotic-type RNAP (**Figure 2**). In fact, land plants have a number of novel host cell-derived transcription regulators and DNA-binding proteins that are involved in the regulation of chloroplast transcription. Thus, it seems likely that chloroplast transcription is mediated by a hybrid system of prokaryotic and eukaryotic origin. Further molecular characterization of pTACs and plastid nucleoid proteins would provide novel insights into the unique plastid gene expression system and as yet known mechanisms of plastid differentiation.

## **ACKNOWLEDGMENTS**

We would like to thank Y. Ishizaki for critical reading of the manuscript. This work was supported by JSPS and MEXT Grantsin-Aid for Scientific Research (24657036, 25291065, 25120723) and a grant from the Mitsubishi Foundation to Takashi Shiina, and a Grant-in-Aid for JSPS Fellows to Yusuke Yagi.

## **REFERENCES**

Abdallah, F., Salamini, F., and Leister, D. (2000). A prediction of the size and evolutionary origin of the proteome of chloroplasts of *Arabidopsis*. *Trends Plant Sci.* 5, 141–142. doi: 10.1016/S1360-1385(00)01574-0

transcription extracts from cultured tobacco BY-2 cells. *Plant Cell* 11, 1799–1810. doi: 10.1105/tpc.11.9.1799

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 30 December 2013; paper pending published: 31 January 2014; accepted: 06 February 2014; published online: 25 February 2014.*

*Citation: Yagi Y and Shiina T (2014) Recent advances in the study of chloroplast gene expression and its evolution. Front. Plant Sci. 5:61. doi: 10.3389/fpls.2014.00061*

*This article was submitted to Plant Physiology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Yagi and Shiina. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## *AtSIG6* and other members of the sigma gene family jointly but differentially determine plastid target gene expression in *Arabidopsis thaliana*

## *Sylvia Bock , Jennifer Ortelt and Gerhard Link\**

*Department of Biology and Biotechnology, University of Bochum, Bochum, Germany*

#### *Edited by:*

*Thomas Pfannschmidt, University Joseph Fourier Grenoble, France*

#### *Reviewed by:*

*David G. Oppenheimer, University of Florida, USA Stefan Gleissberg, Ohio University, USA*

#### *\*Correspondence:*

*Gerhard Link, Department of Biology and Biotechnology, University of Bochum, Universitaetsstr. 150, D-44780 Bochum, Germany e-mail<EMAIL_ADDRESS>
Plants contain a nuclear gene family for plastid sigma factors, i.e., proteins that associate with the "bacterial-type" organellar RNA polymerase and confer the ability for correct promoter binding and transcription initiation. Questions that are still unresolved relate to the "division of labor" among members of the sigma family, both in terms of their range of target genes and their temporal and spatial activity during development. Clues to the *in vivo* role of individual sigma genes have mainly come from studies of sigma knockout lines. Despite its obvious strengths, however, this strategy does not necessarily trace-down causal relationships between mutant phenotype and a single sigma gene, if other family members act in a redundant and/or compensatory manner. We made efforts to reduce the complexity by genetic crosses of Arabidopsis single mutants (with focus on a chlorophyll-deficient *sig6* line) to generate double knockout lines. The latter typically had a similar visible phenotype as the parental lines, but tended to be more strongly affected in the transcript patterns of both plastid and sigma genes. Because triple mutants were lethal under our growth conditions, we exploited a strategy of transformation of single and double mutants with RNAi constructs that contained sequences from the unconserved sigma region (UCR). These RNAi/knockout lines phenotypically resembled their parental lines, but were even more strongly affected in their plastid transcript patterns. Expression patterns of sigma genes revealed both similarities and differences compared to the parental lines, with transcripts at reduced or unchanged amounts and others that were found to be present in higher (perhaps compensatory) amounts. Together, our results reveal considerable flexibility of gene activity at the levels of both sigma and plastid gene expression. A (still viable) "basal state" seems to be reached, if 2–3 of the 6 Arabidopsis sigma genes are functionally compromised.

**Keywords: chloroplast transcription, plant sigma factors, nuclear gene family, knockout mutants, RNA interference, plastid target gene expression**

## **INTRODUCTION**

Despite the small number of genes in the chloroplast genome (Sugiura, 1992), plastid transcription is a surprisingly complex process. It involves two different RNA polymerases commonly named NEP (nuclear-encoded polymerase) and PEP (plastid-encoded polymerase) (Hedtke et al., 1997; Maliga, 1998). The latter is surrounded by multiple transcription factors (Shiina et al., 2005), important representatives of which are the nuclear-encoded sigma factors. Like their bacterial counterparts (Ishihama, 1988; Burgess and Anthony, 2001), these plant factors are thought to direct the (PEP) polymerase complex to its cognate promoters and ensure faithful transcription initiation. Again, as is the case in bacteria, the plastids of higher plants typically contain more than a single sigma factor species (e.g., a family comprising six proteins ATSIG1 - 6 in *Arabidopsis thaliana*) (Isono et al., 1997; Tanaka et al., 1997; Fujiwara et al., 2000; Shiina et al., 2009). This obvious analogy has therefore stimulated research addressing the role of individual members of the plant sigma factor family.

 A readily noticeable phenotype is evident for instance in the case of AtSIG6, where mutant lines tend to have strong chlorophyll deficiency and altered plastid target gene expression patterns, yet only in seedlings but not, e.g., in plants during the subsequent rosette leaf stages (Ishizaki et al., 2005; Loschelder et al., 2006; Schweer et al., 2006, 2009).

Unlike the situation in bacteria (Ishihama, 1988), none of the plastid sigma factors seems to have a "primary" essential role in a sense that its loss would confer a lethal or seriously compromised phenotype (Ortelt and Link, 2014). What then might be the reasons for the variable (non-lethal) phenotypes noticeable in plant sigma knockout lines? A perhaps most direct explanation would be that the plastid factors function in a partially overlapping manner, yet in a highly flexible way at different developmental stages, in different organs, and under variable environmental conditions. Clues supporting this idea come, e.g., from the modular architecture of the plant sigma factors, each of which has a C-terminal conserved region (CR) responsible for basal sigma activity and a N-terminal unconserved region (UCR) of regulatory function (Ortelt and Link, 2014).

Nevertheless, it seems likely that the current functional description of the underlying network is not yet complete. For instance, knocking out one single sigma gene may or may not have the consequence that a second (or third, fourth etc.) member of the factor family is functionally recruited in a specific developmental context. To reduce the complexity of the Arabidopsis sigma family, we took advantage of single and double mutant lines and also adopted RNAi (RNA interference) techniques (Fire et al., 1998) for that purpose. We then analyzed the gene expression situation (at RNA level) both for plastid target genes as well as for the members of sigma gene family themselves. We reasoned that such work could add novel information to help explain the flexibility of the plant sigma factor network and its phenotypic consequences.

## **MATERIALS AND METHODS**

## **ARABIDOPSIS MUTANT AND RNAi LINES, GROWTH CONDITIONS, SCREENING**

Single mutant lines *sig1-2* ("*sig1*"), *sig3-2* ("*sig3*"), and *sig6- 2* ("*sig6*") in the Col-0 (ecotype Columbia) background of *Arabidopsis thaliana* were obtained from the GABI-Kat collection of T-DNA insertion lines (www.gabi-kat.de) (Rosso et al., 2003). Double knockouts *sig1 sig6* and *sig6 sig3* were generated from the single mutants by genetic crosses. Both single and double knockouts were transformed with sigma UCR sequences cloned into RNAi vector pHELLSGATE12 (Wesley et al., 2001) (www*.*csiro*.* au) using the Gateway system (Life Technologies). PCR-amplified sigma cDNA representing the full-size UCR (555 bp after the start codon for AtSIG1, 768 bp for AtSIG3, and 696 bp for AtSIG6) was inserted into the donor vector pDONR/zeo (Life Technologies) and mobilized to the destination vector as described in the Gateway manual (www*.*invitrogen*.*com). It was then introduced into Arabidopsis by Agrobacterium-mediated floral dip transformation (Clough and Bent, 1998) and progeny were screened using antibiotic selection, PCR and Southern blot analyses as described (Loschelder et al., 2006). The criteria for successful generation of double mutants, i.e., absence of an amplified PCR product using primers that flank the T-DNA insertion and presence of a product using a primer pair across the junction between T-DNA and sigma gene sequence, were tested for each candidate line (signal*.* salk*.*edu/tdnaprimers*.*2*.*html). Only those lines that fulfilled these requirements were further propagated and subsequent experiments were carried out with at least three independent T3 lines for each construct. Seeds were sown on MS medium (Murashige and Skoog, 1962) containing 0.4% (w/v) gelrite and 1% (w/v) sucrose, stratified at 4◦C for 2 d, and then transferred to 24◦C for germination and growth under short-day conditions (8 h light/16 h dark, 60 mmol m−2 s −1). Seedlings were harvested at day 6 or 10, or growth was continued until day 14, at which time plantlets were transferred to sterile soil for another 6 d until day 20. Tissue samples were immediately frozen in liquid nitrogen and stored at −85◦C until use.

## **PHYSIOLOGICAL PARAMETERS (CHLOROPHYLL CONTENT, ROOT LENGTH MEASUREMENTS)**

Chlorophyll content was measured using 8 replicates of 10 seedlings or young plantlets at each time-point (6 d, 10 d, 20 d after sowing) and measurements were repeated twice using independently grown plant material. Following weighing of seedlings, they were ground in 80% (v/v) acetone. The extract was centrifuged at 10,000 g for 10 min and the supernatant was used for photometric chlorophyll determination at 663 and 645 nm. For root lengths measurements, seedlings were grown as described above, yet using vertical positioning of Petri dishes. Length determination was carried out using ImageJ (http://imagej*.*nih*.*gov/ij) followed by graphical presentation. Values were means of three independent replicates each obtained from 20 seedlings.

## **RT-qPCR DETECTION OF TRANSCRIPTS**

Total seedling RNA (2μg) was prepared and reverse-transcribed into cDNA as described by Loschelder et al. (2006), yet using oligo-(dT)-primers and the Moloney Murine Leukemia Virus Reverse Transcriptase (M-MLV; Promega). Quantitative real-time RT-PCR (RT-qPCR) was in an Illumina Eco System (www*.* illumina*.*com) with KAPA SybrFast qPCR MasterMix Universal (Peqlab) in a volume of 20μl. Upon amplification (50◦C for 2 min, 95◦C for 10 min, followed by 40 cycles at 95◦C for 10 s, 60◦C for 30 s, and 72◦C for 15 s), melting curve analysis was carried out, using the *Actin2 C*t value as a reference for normalization. For each primer pair, the real-time experiments were carried out with at least three biological and technical replicate samples. Primers (**Table 1**) were selected on the basis of minimal sequence homology among the members of the Arabidopsis sigma gene family (Schweer et al., 2009).

## **NORTHERN BLOT HYBRIDIZATION**

Total RNA was isolated as described by Chomczynski and Sacchi (1987). RNA (2μg per lane) was fractionated, blotted, and hybridized, using gene-specific RNA probes (**Table 1**). The latter were generated by cloning of PCR-amplified gene segments into pGEM-T Easy (Promega), followed by *in vitro* transcription of constructs using T7 RNA polymerase (Promega) as described (Loschelder et al., 2006; Schweer et al., 2006). DIG (digoxigenin) labeling conditions and immunological detection using anti-digoxigenin antibody (Roche) were as detailed in the DIG user's manual (www.roche-applied-science.com).

## **RESULTS**

## **CHARACTERIZATION OF SIGMA-DEFICIENT SINGLE AND DOUBLE MUTANTS**

To raise information on the fine-tuning and complex regulation within the sigma gene family from Arabidopsis, we analyzed single and double sigma mutants. Starting material were the three single mutant lines *sig1-2*, *sig3-2*, and *sig6-2* defective

in *AtSIG1*, *AtSIG3*, and *AtSIG6*, respectively (**Figure 1A**). These lines were then used in crosses to generate double mutants and selfed progeny lines were tested by genomic PCR for presence of the T-DNA insertion. As shown in **Figure 1B**, the amplification products detectable with wildtype DNA (using the primer pairs detailed in **Table 1**) were absent in either the single or double mutant lines, indicating a loss of sigma gene function in all these lines. This was confirmed by using primer pairs across the junction between T-DNA and sigma gene sequence, which generated amplification products (data not shown).

The single mutant lines *sig1-2* and *sig3-2* each have a visible phenotype resembling that of the wildtype (**Figure 1C**, panels 2 and 3). In contrast, the two double mutants *sig1 sig6* and *sig6 sig3*, each resulting from crosses with the *sig6* mutant line (panels 4), both reveal yellowish to white cotyledons, with only minimal light-green leaf primordia recognizable at day 10 (panels 5 and 6). Previous work (Ishizaki et al., 2005; Loschelder et al., 2006; Schweer et al., 2006) had shown that mutant alleles of the Arabidopsis sigma gene *AtSIG6* account for reduced chlorophyll content during seedling development. As is evident from **Figure 1D**, the *sig6-2* line (in comparison with the wildtype) reveals approximately 80% loss of chlorophyll at the 6 d and 10 d stages, respectively, but only 20% loss in 20 day old rosette plants. Chlorophyll quantification of the double mutants likewise shows highly reduced amounts, which (in *sig1 sig6*) are similar to or (in *sig6 sig3*) are even more pronounced than those of the *sig6* parental line at the 6 d and 10 d time-points. At day 20, there is an even further reduction by 15% in *sig1 sig6* and by almost 30% in *sig6 sig3* beyond the *sig6-2* level.

Root length (**Figure 1E**) is likewise reduced in the single mutants compared to wildtype, with a stronger effect noticeable for *sig6* than for *sig1-2* and *sig3-2*. The double mutant lines again show similar (*sig1 sig6*) or even greater (*sig6 sig3*) length reduction compared to *sig6-2*.

## **PLASTID GENE EXPRESSION AT RNA LEVEL IN WILDTYPE, SINGLE AND DOUBLE KNOCKOUT MUTANTS**

To assess consequences of sigma gene inactivation on plastid transcript patterns, we carried out northern blot analyses using total RNA from 6- and 10-day old single and double mutant lines. Since many plastid genes give rise to distinct (single or multiple) transcripts of relatively high abundance, this technique was selected to rapidly reveal stage-specific RNA expression patterns of organellar target genes of sigma-dependent transcription. Representatives of all three major classes of plastid genes, i.e., those for proteins (*atpB* and *rbcL*, the genes for the β subunit of the organellar ATP synthase and the large subunit of ribulose-1,5-bisphospate carboxylase-oxygenase or abbreviated Rubisco, respectively), ribosomal RNAs (*rrn16*, the gene for 16S rRNA) and transfer RNAs (*trnE*, the gene for glutamic acid-specific tRNA), were included in this analysis (for map position and sequence see www*.*ncbi*.*nlm*.*nih*.*gov/nuccore/ 7525012).

Using plastid RNA probes *rrn16*, *trnE*, and *rbcL* (**Figure 2**), no appreciable deviation from the Col-0 wildtype pattern (lane 1 and 7) was noticeable at 6 or 10 days for either the *sig3-2* or *sig1-2* mutant lines (lane 2, 3 and 8, 9), except for a relative increase in intensity of the *rrn16* signal at 6 d in the *sig3-2* line (lane 3). The *atpB* probe (top panel) revealed another difference, i.e., preferential loss of the 2.6 kb (PEP-dependent) transcript (Schweer et al., 2006) in the *sig1-2* line (lane 2 and 8). Unlike both the *sig3-2* and *sig1-2* lines, the *sig6-2* line shows overall less intense hybridization bands compared to wildtype (lane 4 and 10), although to a variable extent depending on the gene investigated. This is most evident for *atpB* (Loschelder et al., 2006), with a loss of the 2.6 kb (PEP-dependent) transcript and concomitant appearance of the 4.8 kb (NEP-dependent) "SOS" transcript (Schweer et al., 2006), particularly at the 6 day stage. Another gene showing dramatic down-regulation of transcript intensity in the *sig6* line is *trnE*, whereas the *rbcL* (Loschelder et al., 2006) and *rrn16* gene expression appears somewhat less affected and that of a nuclear (*RbcS*) control gene remains substantially unaffected (lane 4 and 10).

The double mutant lines *sig1 sig6* and *sig6 sig3* (lane 5, 6 and 11, 12) show a reduction in transcript intensities, which however only partly reflects the patterns of their parental single mutant lines. Following hybridization with the *atpB* probe (top panels), both double mutants show highly reduced signal intensity at the position of the 2.6 kb transcript (lane 2, 5, 6 and 8, 11, 12). In addition, also the 2.0 kb (NEP-dependent) signal is reduced at day 6 but reappears at day 10. The transcript detected with the *rrn16* probe (second panels) is present in reduced amounts in the *sig1 sig6* mutant (lane 5, 11) and is virtually absent in *sig6 sig3* (lane 6 and 12). The *trnE* signal (third panels) is not detectable in both double mutants at either time-point. In the *sig6 sig3* mutant (lane 6 and 12), the intensity of the *rbcL* hybridization signal (forth

and *AtSIG6* (At2g36990). Regions colinear with the mature messenger RNA are boxed, including protein-coding (gray) and non-coding regions (white), while regions corresponding to introns of the RNA precursor are depicted as a single line. T-DNA insertion sites (in exon 9 of *AtSIG1*, exon 7 of *AtSIG3*, and exon 5 of *AtSIG6*, respectively) are marked by triangles. Genomic sequence, without T-DNA, is drawn to scale (scale bar on top). **(B)** RT-PCR detection of sigma factor transcripts in single mutant lines. RT-PCR detection seedlings, reverse transcribed, and cDNA was amplified using gene-specific full-length primer pairs (**Table 1**) as described in "Materials and Methods." **(C)** Visible phenotype. Arabidopsis wildtype (Col-0) as well as sigma single and double knockout lines were photographed at three different time-points during seedling and rosette leaf development (6, 10, and 20 d after sowing). Scale bars: 2 mm. **(D)** Chlorophyll content of 6 d, 10 d, and 20 d seedlings. **(E)** Root lengths measuments (for details, see "Materials and Methods").

panels) resembles that of the parental *sig6-2* single mutant (lane 4, 10) and is diminished compared to the *sig3-2* signal (lane 3 and 9). The *sig1 sig6* signal (lane 5 and 11), however, is increased relative to that for *sig6-2* (lane 4 and 10) at both time points. It is comparable to that for *sig1-2* (lane 2) at 6 d (lane 5) but shows some relative decrease at 10 d (lane 11 vs. lane 8). Finally, the nuclear (*RbcS*) control reveals more uniform transcript intensity in all tested lines at 6 d (lane 1–6) than at 10 d (lane 7–12). At the latter time-point, the most notable effect is the relative increase in signal intensity for the sig1 sig6 line (lane 11) compared to the parental mutants and even the wildtype. This may reflect the known plastid to nuclear signaling (Woodson et al., 2012) in response to altered sigma-dependent chloroplast transcription (see "Discussion").

## **SIGMA GENE EXPRESSION NETWORK IN SIGMA SINGLE AND DOUBLE KNOCKOUT LINES**

To assess correlations between plastid target gene expression and sigma gene activity, transcript levels for all members of the sigma family were determined. To detect and quantitate these lowabundant transcripts, real-time qPCR rather than northern-blot hybridization was used. Whole-cell RNA preparations from 10 day-old homozygous single and double knockout lines as well as from the Col-0 wildtype were reverse-transcribed and subjected to real-time qPCR. Data were normalized to *Actin2* RNA expression levels (see "Materials and Methods"). As the RNA patterns of plastid target genes (**Figure 2**) indicated a close similarity of double mutant lines primarily with the *sig6-2* single mutant line, we tested these lines for correlation of target gene expression

with sigma gene expression patterns themselves. As shown in **Figure 3A** for the *sig6-2* single mutant line, most sigma transcripts are down-regulated compared to wildtype, while the *SIG1* transcript seems to be strongly enhanced. In both double mutant lines, however, all six sigma transcripts including that of *SIG1* are reduced. Hence, except for *SIG1*, the expression phenotype of the sigma gene family in each double knockout substantially reflects that of the *sig6-2* parental line.

To test for possible similarity with other parental mutants, we also investigated *sig1-2* and *sig3-2* lines. In terms of their real-time transcript patterns (**Figure 3A**), these two lines each can be clearly set apart from *sig6-2* as well as from the two double mutants. In the *sig3-2* single knockout line, the only strong down-regulation is noticeable for transcripts detected by the *SIG3* primers, which is consistent with a major or full loss of *SIG3*-related transcripts in this mutant. While *SIG4* transcript levels are almost at wildtype level, those of *SIG2* and *SIG5* are moderately elevated and transcript levels of *SIG1* and *SIG6* are even strongly enhanced.

The *sig1-2* knockout line reveals transcript levels that are decreased in the case of *SIG3* and *SIG5*, but are increased for *SIG2*, *SIG4*, and *SIG6*. In addition, despite the T-DNA insertion interrupting the *SIG1* gene (**Figure 1A**) and the concomitant loss of a full-size *SIG1* transcript (**Figure 1B**), the RT-qPCR data for the *sig1-2* line (**Figure 3A**) suggested the existence of significant amounts of *SIG1*-related transcripts. The latter may represent partial (non-functional) transcripts upstream of the T-DNA insertion site. To test this, RT-PCR was carried out with primers that flank the distance from the ATG of the first exon to a site directly in front of the T-DNA insertion (**Figure 3B**). Both the wildtype (lane 2) and the *sig1-2* single mutant (lane 3) showed a PCR signal consistent with a transcript spanning the entire region defined by this primer pair. In contrast, such signal was not detectable in the double mutant line (*sig1 sig6*) (lane 4).

### **CHARACTERIZATION OF RNAi-MODIFIED SIGMA KNOCKOUT LINES**

Attempts to further reduce the complexity of the sigma family were initially hampered by our inability to generate sigma triple-mutant lines, possibly because of their lethality. We therefore chose RNAi in combination with the existing single and double sigma mutants as an alternative strategy. "Combined" RNAi/knockout lines were created by transformation of the *sig6-2* and *sig6 sig3* mutants using constructs that contain the complete sequence for the unconserved sigma region (UCR) in pHELLS-GATE12 (Wesley et al., 2001). Lines that could be stably maintained included those with the UCR sequences of *SIG2* or *SIG4* in the *sig6-2* single mutant background as well as those with the UCR sequences of *SIG1* or *SIG4* in the *sig6 sig3* double mutant background.

At each time-point (6, 10, and 20 days after sowing) in **Figure 4A**, the combined RNAi/knockout line *sig6::UCRSIG4* (panels 3) has a pigment-deficient visible phenotype which resembles that of the parental *sig6* line (panels 1). The same is true for *sig6::UCRSIG2* (panels 2) as well as for combined lines *sig6 sig3::UCRSIG1 (*panels 5) and *sig6 sig3::UCRSIG4* (panels 6), and their parental double mutant line *sig6 sig3* (panels 4) (see also **Figure 1C**). The pigment-deficient visible appearance of all investigated RNAi/knockout lines is reflected by their chlorophyll content (**Figure 4C**). Likewise, root length measurements (**Figure 4D**) establish growth deficiency similar to that noticeable for the parental single (*sig6-2*) and double mutant lines (*sig6 sig3*) (see **Figures 1D,E**).

As shown in **Figure 4B**, transcripts of plastid target genes were found to be even more compromised than those of the (single and double mutant) parental lines (see also **Figure 2**), although differentially and to a variable extent. For instance, except for the loss of the 4.8 kb *atpB* transcript, the relative intensity of hybridization signals of *sig6::UCRSIG4* (panels 5 and 6) is comparable to that of *sig6-2* (panels 1 and 2) both at 6 d and 10 d, respectively. In contrast, *sig6::UCRSIG2* (panels 3 and 4) shows a weak but discernible signal (especially at 6 d) at the position of the 4.8 kb *atpB* transcript, while all other signals are virtually absent or highly reduced at both time-points in comparison with *sig6* (panels 1 and 2). This also includes the 2.0 kb band at the position of the NEP-dependent *atpB* transcript (Schweer et al., 2006), whereas the PEP-dependent 2.6 kb *atpB* transcript is absent both in *sig6* itself and in all *sig6*-derived lines. The transcript patterns of *sig6 sig3::UCRSIG1* (lanes 9 and 10) and *sig6 sig3::UCRSIG4* (panels 11 and 12) are similar to, but are more strongly affected than, that of their parental line *sig6 sig3* (panels 7 and 8). While all three lines show the 4.8 kb *atpB* transcript (at day 10 but not day 6), none of them reveals the 2.6 kb (PEP-dependent) *atpB* transcript and the 2.0 kb (NEP-dependent) transcript seems to be diminished in the combined RNAi/double knockout lines (panels 9–12). The *trnE* transcript is absent in all three lines (panels 7–12), and the *rbcL* transcript is highly reduced in the "combined" lines (lanes 9–12) compared to the parental double knockout (panels 7 and 8).

The same RNAi/knockout lines were also tested for their sigma transcript patterns in comparison with those of the *sig6* and *sig6 sig3* parental lines (**Figure 5**). In the case of the *sig6::UCRSIG4* line, the steady-state transcript concentrations of all sigma genes are further reduced compared to those of *sig6* itself. In contrast, *sig6::UCRSIG2* shows a more diverse pattern, with a moderate further reduction of the *SIG2*, *SIG3*, and *SIG6* transcripts compared to the *sig6-2* line, substantially unchanged levels of the *SIG1* and *SIG5* transcripts, and strongly increased concentration of the *SIG4* transcript (**Figure 5A**). The RNAi/double knockout lines *sig6 sig3::UCRSIG1* and *sig6 sig3::UCRSIG4* (**Figure 5B**) both reveal moderate to strong further reduction of the *SIG2*–*SIG6* transcripts but enhanced levels of the *SIG1* transcript compared to the parental *sig6 sig3* line. Their transcript patterns thus seem more similar to one another than to those shown in **Figure 5A**.

## **DISCUSSION**

This work was carried out to help reach a fuller understanding of the plant sigma genes, i.e., the small nuclear gene family for chloroplast transcription factors resembling the bacterial sigma transcription initiation factors. We sought to analyze phenotypic and molecular consequences of altered patterns of sigma gene activity in *Arabidopsis thaliana*. Such changes can be "homeostatic," i.e., balanced and compensatory, which can be anticipated during developmental and/or physiological transitions. In more extreme situations, however, the sigma network can be thought to become "disrupted," i.e., (irreversibly) imbalanced and rendered non-functional, as might be expected in lethal or heavily compromised mutants and/or under strong environmental stress. To narrow down the limits of homeostatic vs. imbalanced states of the sigma gene family, we have chosen strategies to differentially affect the sigma network, including single and double sigma mutants in combination with RNAi.

The parental single mutants, *sig1-2*, *sig3-2*, and *sig6-2*, were chosen for reasons of their specific phenotypic characteristics. *SIG6* single mutant lines reveal a clear-cut chlorophyll-deficient

#### **FIGURE 4 | Continued**

during development (6, 10, and 20 d after sowing). Scale bars: 2 mm. **(B)** Northern blot analysis of plastid gene expression in RNAi/knockout lines and their parental lines. Total RNA (2μg/lane) from 6 and 10 d old seedlings was fractionated, blotted and hybridized with *atpB*, *trnE*, or *rbcL* probes as indicated in the left margin. Ethidium bromide-stained loading controls (25S rRNA) are shown at the bottom. These experiments were carried out at least in triplicate with RNA from independent preparations. Right margin: transcript sizes (kb). **(C)** Chlorophyll content of

and developmental-stage-specific (seedling) phenotype, indicating a specialized and/or functionally dominating role of this factor (Ishizaki et al., 2005; Loschelder et al., 2006; Schweer et al., 2006, 2009). In contrast, *sig3* lines do not show pronounced pigment deficiency and the corresponding factor SIG3 has been assigned a functionally redundant role, perhaps as a possible safeguard in case of loss of other sigma factor(s) (Schweer, 2010; Lerbs-Mache, 2011). Although *sig1* knockout mutants of Arabidopsis have not yet been presented in terms of their gene expression characteristics, such mutants are of considerable interest in view of recent findings that sigma factor 1 (SIG1) is subject to phosphorylation control (Shimizu et al., 2010), as is known for sigma factor 6 (Schweer et al., 2010a,b; Türkeri et al., 2012).

To reduce the number of functional sigma genes, the *sig6-2* knockout line (Loschelder et al., 2006; Schweer et al., 2006, 2009) was crossed with either of two other single mutant lines, *sig1-2* and *sig3-2,* giving rise to the double mutants sig*1 sig6* and *sig6 sig3*. The latter reveal a growth-retarded and highly chlorophyll-deficient phenotype at seedling stage, even exceeding that of the parental *sig6-2* knockout. Plastid target gene seedlings and young plantlets 6 d, 10 d, or 20 d after sowing. Measurements involved 8 replicates of 10 samples each from three series of independently grown plant material. Weighed samples were ground in 80% (v/v) acetone and photometric chlorophyll determination at 663 and 645 nm was carried out. **(D)** Root lengths. Following growth of seedlings and young plantlets on vertically positioned Petri dishes, length measurements were carried out using ImageJ (http://imagej*.*nih*.*gov/ij). Values were means of three independent replicates using samples representing 20 seedlings or plantlets each.

expression at RNA level was strongly compromised in both double mutants. Assessment of sigma gene expression itself using RT-qPCR showed both losses but also increases in transcript frequency for individual members of the gene family in the single mutants. In contrast, however, a global decrease of sigma transcripts was noticeable in the double mutants. Hence, due to functional redundancy and compensation of sigma family members, a homeostatic balance seems still to prevail in the single knockout lines, while the balance may be strongly shifted or completely lost in the double mutants.

Attempts to further reduce the complexity of the sigma family by construction of triple mutants were unsuccessful, likely because of lethality of the progeny from these crosses. As an alternative, we therefore combined knockout mutant with RNAi technology. Using various sigma-specific unconserved regions (UCRs) in the pHELLSGATE12 RNAi vector, single (*sig6*) and double (*sig6 sig3*) mutant lines were transformed by constructs based on this vector. Resulting progeny lines generated by transformation of *sig6* showed a chlorophyll-deficient phenotype similar to or even stronger than that of the parental knockout line. Those generated from *sig6 sig3* did not reveal a further enhanced phenotype compared to the parental double knockout line, suggesting that a "basal" (minimal) state which cannot be further reduced without loss of viability may have been reached already in the latter.

An argument against this notion, however, comes from results of the target gene expression studies, showing that the *rbcL* transcript is readily detectable in *sig6 sig3* but is highly reduced or absent in the "combined" RNAi/double knockout lines (**Figure 4B**). Furthermore, the RT-qPCR analysis (**Figure 5B**) shows enhanced transcript levels of the *SIG1* transcript in *sig6 sig3::UCRSIG1* and *sig6 sig3::UCRSIG4* as compared to the parental line *sig6 sig3*, which may reflect a still balanced functional state of the sigma gene family in these lines. In any case, it is notable that down-regulation of the expression of a single sigma gene can both negatively or positively affect that of another family member. For instance, the expression of *SIG1* and *SIG6* seems to be regulated in an opposite manner in single knockout lines (**Figure 6**), indicating functional redundancy and mutual compensation.

A perhaps unexpected finding is the partial or even dramatic loss of the 2.0 kb (NEP-dependent) *atpB* transcript (Schweer et al., 2006) in all RNAi/knockout lines (**Figure 4B**). Sigma factor SIG6 was previously implicated in retrograde signaling from the chloroplast to nucleus (Woodson et al., 2012), i.e., a mechanism that can affect the expression of nuclear genes in response to altered sigma factor function and chloroplast transcription

(Pfannschmidt, 2010). It can be envisaged that the nuclearencoded plastid polymerase might be regulated via this route, which in turn could explain the loss of the 2.0 kb NEP-dependent *atpB* transcript (Hanaoka et al., 2005).

Concerted regulation of both the PEP and NEP transcription systems via functional alterations of one or several sigma factors can be considered as efficient and flexible mechanism to achieve interorganellar integration. For instance, it might be interesting to investigate if the increased *RbcS* transcript level seen in the sig1 sig6 double mutant (**Figure 2**) is primarily due to an altered sigma network in this mutant and/or involves NEP-dependent regulation. Clearly, differential and compensatory expression of sigma genes as studied here is only one of several control levels. Posttranslational modification such as phosphorylation (Schweer et al., 2010a,b; Shimizu et al., 2010) as well as interactions of sigma facors with other regulatory proteins are equally important (Morikawa et al., 2002; Chi et al., 2010), as is the topology of the plastid transcriptome (Yagi et al., 2012; Zhelyazkova et al., 2012). In any case, our current work points to a causal relationship between the expression status of the sigma gene family and responses at the level of target gene expression.

Finally, it should be recalled that all RNAi lines described here were generated using the "constitutive" pHELLSGATE12 silencing vector (Wesley et al., 2001). A somewhat similar picture also emerges from initial recent work with chemically inducible dexamethasone (DEX)-responsive) RNAi lines based on the pOpOff2 vector (Wielopolska et al., 2005), providing proof of principle for DEX-responses that are visible at both target gene and sigma expression patterns (data not shown). Usage of such inducible "knock-down" system, also including, e.g., virus-induced gene silencing (VIGS) (Ratcliff et al., 1997), can be expected to open up new avenues in studies on temporal and spatial activities of the gene-containing plant cell organelles. This way, it should become possible to successfully analyze stages throughout the entire Arabidopsis life cycle.

## **AUTHOR CONTRIBUTIONS**

Sylvia Bock carried out planning, performance and presentation of most experiments. Jennifer Ortelt assisted in all aspects of experimental analyses and manuscript preparation. Gerhard Link provided advice and assistance throughout this work.

## **ACKNOWLEDGMENTS**

We are indepted to Prof. Bernd Weisshaar, University of Bielefeld, and the GABI-Kat team at the Max-Planck-Institute fuer Zuechtungsforschung, Cologne, for the supply of the sigma factor mutant lines, to Prof. Peter Michael Waterhouse and colleagues at the CSIRO, Canberra, for the RNAi vectors, and to Prof. Minou Nowrousian for guidance on RT-qPCR. We gratefully acknowledge the excellent technical assistence of Brigitte Link. This work was funded by the Deutsche Forschungsgemeinschaft (LI261/21-1).

## **REFERENCES**

of cysteinyl SH-groups in regulatory phosphorylation of plastid sigma factors. *FEBS J.* 279, 395–409. doi: 10.1111/j.1742-4658.2011.08433.x

noncoding RNAs and the dominating role of the plastid-encoded RNA polymerase. *Plant Cell* 24, 123–136. doi: 10.1105/tpc.111.089441

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 24 July 2014; accepted: 09 November 2014; published online: 25 November 2014.*

*Citation: Bock S, Ortelt J and Link G (2014) AtSIG6 and other members of the sigma gene family jointly but differentially determine plastid target gene expression in Arabidopsis thaliana. Front. Plant Sci. 5:667. doi: 10.3389/fpls.2014.00667*

*This article was submitted to Plant Physiology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Bock, Ortelt and Link. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## Plastid encoded RNA polymerase activity and expression of photosynthesis genes required for embryo and seed development in *Arabidopsis*

## *Dmitry Kremnev and Åsa Strand\**

*Umeå Plant Science Centre, Department of Plant Physiology, Umeå University, Umeå, Sweden*

#### *Edited by:*

*Thomas Pfannschmidt, University Joseph Fourier Grenoble, France*

#### *Reviewed by:*

*Tatjana Kleine, Ludwig-Maximilians-Universität München, Germany Silva Mache, Laboratoire de Physiologie Cellulaire Végétale, France*

#### *\*Correspondence:*

*Åsa Strand, Umeå Plant Science Centre, Department of Plant Physiology, Umeå University, S-901 87 Umeå, Sweden e-mail<EMAIL_ADDRESS>
Chloroplast biogenesis and function is essential for proper plant embryo and seed development but the molecular mechanisms underlying the role of plastids during embryogenesis are poorly understood. Expression of plastid encoded genes is dependent on two different transcription machineries; a plastid-encoded bacterial-type RNA polymerase (PEP) and a nuclear-encoded phage-type RNA polymerase (NEP), which recognize distinct types of promoters. However, the division of labor between PEP and NEP during plastid development and in mature chloroplasts is unclear. We show here that PLASTID REDOX INSENSITIVE 2 (PRIN2) and CHLOROPLAST STEM-LOOP BINDING PROTEIN 41 kDa (CSP41b), two proteins identified in plastid nucleoid preparations, are essential for proper plant embryo development. Using Co-IP assays and native PAGE we have shown a direct physical interaction between PRIN2 and CSP41b. Moreover, PRIN2 and CSP41b form a distinct protein complex *in vitro* that binds DNA. The *prin2.2* and *csp41b-2* single mutants displayed pale phenotypes, abnormal chloroplasts with reduced transcript levels of photosynthesis genes and defects in embryo development.The respective *csp41b-2prin2.2* homo/heterozygote double mutants produced abnormal white colored ovules and shrunken seeds. Thus, the *csp41b-2prin2.2* double mutant is embryo lethal. *In silico* analysis of available array data showed that a large number of genes traditionally classified as PEP dependent genes are transcribed during early embryo development from the pre-globular stage to the mature-green-stage. Taken together, our results suggest that PEP activity and consequently the switch from NEP to PEP activity, is essential during embryo development and that the PRIN2-CSP41b DNA binding protein complex possibly is important for full PEP activity during this process.

**Keywords: chloroplast, PEP, NEP, embryo development, photosynthesis, PRIN2, CSP41b**

## **INTRODUCTION**

The chloroplasts house the photosynthetic light reactions where sunlight is converted into chemical energy. Plastids are also the location of a number of vital metabolic pathways, including primary carbon metabolism and the biosynthesis of fatty acids, amino acids, and tetrapyrroles. Chloroplast function is required throughout the life cycle of the plant and compromised activity can result in embryo lethality. Abortion of developing embryos is known to occur when amino acid, nucleotide or fatty acid biosynthesis is impaired, or when import of chloroplast proteins and translation are disrupted (McElver et al., 2001; Tzafrir et al., 2003; Hsu et al., 2010; Bryant et al., 2011). In contrast, disrupting components of the photosynthetic apparatus leads to reduced pigmentation and changed physiology rather than embryo lethality (Tzafrir et al., 2003; Bryant et al., 2011). Chloroplasts are detected as early as at the globular stage of the embryo (Tejos et al., 2010) and transcript profiling during embryo development showed a significant increase in the expression of nuclear genes encoding components involved in energy production, carbon fixation and photosynthesis already from the globular embryogenic stage (Spencer et al., 2007; Le et al., 2010; Belmonte et al., 2013). Furthermore, it has been indicated that chloroplasts can provide the embryo with energy and O2 required for biosynthesis and respiration (Rolletschek et al., 2003). Additionally, it was shown that *Brassica* embryos were able to fix CO2 and contributing to embryo growth rate and biomass (Goffman et al., 2005).

Chloroplasts, like mitochondria, evolved from free-living prokaryotic organisms that entered the eukaryotic cell through endosymbiosis. The gradual conversion from endosymbiont to organelle during the course of evolution has clearly been accompanied by a dramatic reduction in genome size as the chloroplasts lost most of their genes to the nucleus. The genes remaining in the chloroplast genome are related to photosynthesis or encode components of the plastid gene expression machinery (Wakasugi et al., 2001; Martin et al., 2002). The chloroplast genes of higher plants are transcribed by at least two types of RNA polymerases; the nuclear encoded plastid RNA polymerase (NEP), a T3-T7 bacteriophage type that predominantly mediates the transcription of the house keeping genes (Hedtke et al., 1997, 2000; Puthiyaveetil et al., 2010). The other type, plastid encoded RNA polymerase (PEP) is a bacterial-type multi-subunit enzyme that predominantly mediates the transcription of photosynthesis-related genes (Allison et al., 1996; De Santis-MacIossek et al., 1999). Most chloroplast genes can be transcribed by both polymerases but they utilize different promoter elements (Hajdukiewicz et al., 1997; Pfannschmidt and Liere, 2005). The PEP enzyme recognizes the −10 and −35 *cis*elements, similar to those found in bacterial promoters whereas the NEP enzyme recognizes the YRTA-motif, which can also be found upstream of several genes with PEP promoters indicating that these genes can be transcribed by both polymerases (Pfannschmidt and Liere, 2005). Transcription of the plastid encoded photosynthesis genes during chloroplast development and the activation of the photosynthetic reactions are accompanied by a switch from NEP to PEP activity (Hanaoka et al., 2005). However, the mechanisms underlying this change in major RNA polymerase activity and the division of labor between NEP and PEP in the chloroplast are unknown (Zhelyazkova et al., 2012).

A large number of proteins have been shown to be associated with the PEP complex and the components associated with PEP changes in response to developmental signals and changes in the environment (Pfalz and Pfannschmidt, 2013). The variation in the composition of the PEP complex suggests that regulation of plastid gene expression is both complex and sophisticated. PLASTID REDOX INSENSITIVE 2 (PRIN2) is a novel plant protein localized to the plastid nucleoids and plastid transcriptome analyses demonstrated that PRIN2 is required for full expression of genes transcribed by PEP (Kindgren et al., 2012). The role of the PEP associated proteins is unclear and whether these proteins contribute to the regulation of plastid gene expression by environmental and developmental cues remains to be determined. In order to shed light on the complex regulation of PEP activity and to understand the function of PRIN2 we pursued an assay to identify interacting protein partners of PRIN2. We report here a direct interaction between PRIN2 and CHLOROPLAST STEM-LOOP BINDING PROTEIN 41 kDa (CSP41b) using three different biochemical methods. CSP41b was described as an RNA binding protein identified in nucleoid preparation and was attributed numerous plastid functions, for example CSP41b was suggested to stimulate both transcription and translation in the chloroplast (Pfannschmidt et al., 2000; Yamaguchi et al., 2003; Suzuki et al., 2004; Hassidim et al., 2007; Bollenbach et al., 2009). We also demonstrate that PRIN2 and CSP41b form a distinct DNA binding protein complex *in vitro* and that the *csp41b-2prin2.2* double mutant is embryo lethal. Taken together, our results suggest that PEP activity and consequently the switch from NEP to PEP activity, is essential also during embryo development and that the PRIN2-CSP41b protein complex potentially is important for full PEP activity during this process.

## **MATERIALS AND METHODS**

#### **PLANT MATERIAL AND GROWTH CONDITIONS**

Seedlings of Arabidopsis thaliana were grown on phytoagar plates containing 1 × Murashige and Skoog salt mixture supplemented with vitamins (Duchefa) and 2% sucrose. The T-DNA insertion lines: prin2.2 (GK-772D07-024643) and csp41b-2

#### **MORPHOLOGICAL ANALYSIS**

Embryo isolation was done according to (Perry and Wang, 2003). Briefly, The siliques at the desired stage of DAP were dissected under a stereo microscope and the seeds were collected. Embryos were gently extruded from the seed coat by applying pressure on a glass plate covering them. Embryo/seed coat mixture was loaded a 25% Percoll gradient in isolation buffer (10 mM potassium phosphate, pH 7.0, 50 mM NaCl, 0.1 M sucrose) and centrifuged for 10 min at 800 g. The pellet was subjected to another round of purification using the 25% Percoll gradient. The pellet was re-suspended in isolation buffer and used for subsequent analysis. For transmission electron microscopy (TEM) pictures, 3-weeks-old plants grown on soil and the isolated seed embryos were prepared according to (Barajas-Lopez Jde et al., 2013).

#### **RNA ISOLATION, cDNA SYNTHESIS AND REAL-TIME PCR**

Total RNA was isolated using Plant RNA Mini Kit (EZNA) and genomic DNA contamination was removed by DNase treatment (Fermentas). cDNA was synthesized using the iScript cDNA kit (Bio-Rad) according to the manufacturer's instructions and 10 × diluted. Real-Time PCR was performed using iQSYBR Green Supermix (Bio-Rad), with a final volume of 10 μL. The PCR amplification was done using twostep protocol using the CFX96 Real-Time system (C1000 Thermal Cycler; Bio-Rad). All experiments were performed with three biological and three technical replicates, the relative gene expression was normalized to the expression of RCE1 (At4g36800) and PP2AA3 (At1g13320). Data analysis was done by CFX manager (Bio-Rad) and LinRegPCR software.

#### **EXPRESSION AND PURIFICATION OF RECOMBINANT PROTEINS**

The coding sequences of PRIN2 and CSP41b were amplified with PCR. The PCR products were cloned using the NcoI–AccI sites into pET\_His1a vector. BL21 *Escherichia coli* cells were transformed with the expression constructs and induced for 6 h with 1 mM IPTG. Overexpressed proteins were affinity purified on Ni2+- NTA agarose resin (Qiagen). The pET-His1a expression vector was kindly provided by Günter Stier, Umeå University, Sweden.

#### **EMSA**

The 197 bp probe containing −196 to +1 PsaA promoter region was PCR amplified and labeled at the 3 -end with biotin-14 dCTP using biotin labeling kit (Invitrogen) according to the manufacturer's instructions. DNA–protein interactions were performed in 25 mcL reactions containing following reagents: 2.5 mcL of × 10 binding buffer (100 mM Tris HCl, 250 mM KCl, and 10 mM DTT), 1 mcg poly dIdC (Sigma–Aldrich), 2,5% glycerol, 0.05% Triton X-100, 5 mM MgCl2, 10 mM EDTA. The reaction mixture was incubated with DNA and protein at room temperature for 30min and was run on 6% native TBE-PAGE in x0,5 TBE buffer at 100 V. DNA was transferred to nylon + membrane (Amersham) and was UV cross-linked to the membrane, incubated with Streptavidin-HRP and detected by Chemoluminescence Nucleic Acid Detection Module (Pierce) according to manufacturer's instructions (Shaikhali et al., 2012a,b).

## **Co-IP**

To identify PRIN2 interacting partners, a 35S promoter linked to the full length PRIN2 coding sequence was cloned into the pGWB16\_myc expression construct. Col-0 plants were then transformed with 35S\_pGWB16myc\_PRIN2 using the floraldip method (Clough and Bent, 1998). Two weeks old stable transformants overexpressing PRIN2 were used for chloroplast isolation in a two-step 50–25% Percoll gradient as described previously (Aronsson and Jarvis, 2002). Chloroplast proteins were extracted and incubated with 3 mg of anticMYC monoclonal antibody (Bio-Site) bound to the protein G-coated magnetic beads (Dynabeads Protein G Immunoprecipitation, Invitrogen) for 1 h at 4◦C. All the washing steps were performed at 4◦C according to the manufacturer's instructions. Immunoprecipitated PRIN2 protein complexes were eluted in SDS loading buffer containing 100 mM β-mercaptoethanol and proteins were separated on 6–12% SDS PAGE. Non-transformed Col-0 plants were used as negative control. For direct protein Co-IP assay, full length PRIN2 and CSP41b coding sequences were cloned into *NcoI–NcoI* sites of pRT104\_3myc and into *SacI–NotI* sites of pRT104\_3HA vector, respectively. Protoplasts obtained from *Arabidopsis Ler-0* cell culture were transformed with pRT104\_PRIN2\_myc and pRT104\_CSP41b\_HA constructs as described previously by (Doelling and Pikaard, 1993).

#### **ISOLATION OF THYLAKOID COMPLEXES AND BLUE NATIVE PAGE**

Chloroplasts were isolated on a two-step 50–25% Percoll gradient from 4 weeks old rosette plants grown in short day as described previously (Aronsson and Jarvis, 2002). Thylakoid membranes were then purified (Hall et al., 2011) and protein complexes were solubilized in BN-solubilization buffer (30 mM HEPES, pH 7.4; 150 mM potassium acetate; 10% glycerole, 4% digitonin (SIGMA); 1% *b*-Dodecylmaltoside (SIGMA) for 40 min, 4◦C. DM and digitonin in a mixture was shown to be suitable for better preservation of megacomplexes and at the same time good for solubilization (Järvi et al., 2011). 35 micro gram protein was loaded onto the 4–12% Bis-Tris Gel (NuPAGEH Novex 1.0 mm, Invitrogen) from each genotype.

### **RESULTS**

## **PRIN2 AND CSP41B FORM A DISTINCT PROTEIN COMPLEX THAT BINDS DNA**

To understand the function of PRIN2 we wanted to identify proteins interacting with PRIN2 *in vivo*. To achieve this a co-IP approach was used. Full length PRIN2 protein fused to a cMyc-tag was expressed in *Arabidopsis* plants, intact chloroplasts were isolated and PRIN2-containing protein complexes were precipitated with anti-cMyc antibody, proteins were separated on SDS PAGE and distinct bands identified using mass spectrometry (**Figure 1A**). The Co-IP experiment was performed twice and as

many as 17 bands absent from the negative controls were in total cut from the gels. Most of the identified proteins were only found in one experiment and most likely represented unspecific interactions. However, one protein, CSP41b was identified in both experiments and several peptides with significant scores corresponding to CSP41b were identified in each sample (Table S1 in Supplementary Material). CSP41b is a conserved chloroplast protein and the *csb41b-2* mutant displayed impaired chloroplast transcription and plant development (Pfannschmidt et al., 2000; Suzuki et al., 2004; Bollenbach et al., 2009; Qi et al., 2012). The phenotype of the *csb41b-2* mutant was very similar to the phenotype of the *prin2* mutant and it is possible that CSP41b and PRIN2 are involved in the same process. Thus, the identified interaction between PRIN2 and CSP41b was chosen for further analysis.

First, to really confirm the interaction between CSP41b and PRIN2 purified recombinant proteins were separated on 2D electrophoresis (**Figure 1B**; Table S2). PRIN2 forms protein complexes ranging from 20–66 kDa while CSP41b is present in high molecular weight complexes ranging from 40–700 kDa. When PRIN2 and CSP41 are incubated together, PRIN2 seems to break the high molecular weight complexes of CSP41b and form a distinct protein complex with CSP41b suggesting heteromerization. To further confirm *in vivo* that PRIN2 and CSP41b directly interact with each other we used a third method by transiently expressing the full length proteins PRIN2 and CSP41b fused to cMycand HA-tags, respectively, in *Arabidopsis* protoplasts. Two bands of approximately 50 kDa in size correspond to CSP41b in the CSP41b-HA transformed protoplasts and these could be cytoplasmic/transite peptide-processed forms of the protein and/or proteolytic fragment of the protein. The PRIN2 protein exists in monomeric, dimeric and oligomeric forms (unpublished data) and migrates under these conditions as two bands of approximately 20 and 40 kDa, respectively, most likely corresponding to monomer and dimer. CSP41b was detected in the Co-IP fraction confirming interaction with PRIN2 (**Figure 1C**). Thus, using three independent methods CSP41b and PRIN2 were shown to directly interact.

Both PRIN2 and CSP41b were suggested to regulate transcription of PEP dependent chloroplast genes (Bollenbach et al., 2009; Kindgren et al., 2012). To investigate if PRIN2 and CSP41b interact upon DNA binding *in vitro*, the proteins were incubated with a 197 bp DNA probe containing −196 to +1 region of the *psaA* promoter from *Arabidopsis*. In the electrophoretic mobility shift assay (EMSA) both PRIN2 and CSP41b bound the labeled probe, PRIN2 formed at least two distinct complexes with DNA, while CSP41b formed only one DNA-protein complex (**Figure 1D**). PRIN2 and CSP41b are about 15 and 40 kDa, respectively, and the difference in the migration of the protein/DNA complexes of PRIN2 and CSP41b suggests that PRIN2 forms higher molecular weight oligomeric complexes and/or binds to several regions of the DNA probe. When PRIN2 and CSP41b were incubated together with DNA, a new band, intermediate in size to what was observed for the individual proteins, was detected that most likely corresponded to a *psaA*197-PRIN2/CSP41b heteromeric complex. The competition reactions with unlabeled *psaA*-198 bp confirm the DNA binding capacity of PRIN2 and CSP41b. A similar

**FIGURE 1 | PRIN2 interacts with CSP41b and forms a DNA-binding complex. (A)** Native PRIN2-cMyc protein complex from *Arabidopsis* chloroplasts was immunoprecipitated with anti-cMyc antibody and proteins were separated on the SDS-PAGE. Non-transformed *Col-0* chloroplasts were used as a negative control. Protein bands were cut from the gel and analyzed by mass spectrometry. An arrow indicates the position of the band where CSP41b was identified. **(B)** PRIN2 interaction with CSP41b detected by 2D Native PAGE/SDS PAGE gel. Recombinant PRIN2, CSP41b and 1:1 mixture of both proteins were incubated in the binding buffer and run on the 4–12% Native Bis-Tris Gel. Individual stripes were then cut and loaded on the 12% SDS PAGE gel. Proteins were detected with anti-His peroxidase. **(C)** Interaction between PRIN2 and CSP41b *in vivo* Co-IP assay. Full length PRIN2-cMyc and CSP41b-HA fusion proteins were expressed in *Arabidopsis* protoplasts and Co-IP was performed using anti-cMyc antibody linked to protein G-covered magnetic beads.

Immunoblotted samples from input and Co-IP probes were detected with either anti-cMyc chicken IgY fraction and rabbit HRP-linked anti-chicken IgY (H+L) or with anti-HA peroxidase. Arrowheads indicate PRIN2 protein and arrows show CSP41b protein bands detected by anti-HA antibody. **(D)** DNA binding of PRIN2 and CSP41b and their heteromerization in EMSA assay. Signal from *psaA*-198 bp biotin labeled probe was detected by chemoluminescence nucleic acid detection module. 3 μg of each purified protein was used in every reaction, the DNA/protein molar ratio was 1:100. DNA/protein complexes are marked with asterisks, free DNA probe with an arrow. Competition was done with unlabeled *psaA*-198 bp probe with 50-fold excess over the labeled probe. 3 μg of BSA was used as a negative control. (1) free probe, (2) probe + PRIN2, (3) probe + CSP41b, (4) probe + PRIN2 + CSP41b, (5) probe + PRIN2 + unlabeled probe, (6) probe + CSP41b + unlabeled probe, (7) probe + PRIN2 + CSP41b + unlabeled probe, (8) probe + BSA.

pattern of DNA-binding could also be observed when using the *psbA*-198 bp fragment as a probe, where PRIN2 and CSP41b proteins form heteromeric protein/DNA complexes (Supplementary Figure S1). Thus, both PRIN2 and CSP41b are able to interact with DNA in the EMSA assay and the formation of the heteromeric PRIN2/CSP41b complex also occurs upon DNA binding *in vitro*.

## **THE** *prin2.2* **AND** *csp41b-2* **MUTANTS SHOW DISTINCT PHENOTYPES AND IMPAIRED EXPRESSION OF CHLOROPLAST ENCODED GENES**

The phenotypes of the *prin2.2* and *csp41b-2* mutants are similar to mutants of defined components of the PEP complex (De Santis-MacIossek et al., 1999; Krause et al., 2000; Legen et al., 2002; Pfalz et al., 2006). As has been demonstrated before *prin2.2* and *csp41b-2* showed a clear reduction in growth rate and pale leaves compared to wild type (**Figure 2A**). The *prin2.2* plants have impaired chloroplast structure with reduced thylakoid membranes and grana stacks (**Figure 2B**). In contrast, the *csp41b-2* mutant showed an increased number of thylakoid membranes organized in grana stacks (**Figure 2B**). Moreover, large areas of the chloroplasts (depicted with an arrowhead) were devoid of any thylakoid membranes in the *csp41b-2* mutant. Correlated with aberrant chloroplast structure is the observed defect in the organization of the photosynthetic complexes in the *prin2.2* and *csp41b-2* mutants. The mutants showed reduced amounts of PSI, PSII, ATPase, and antenna complexes compared to wild type (**Figure 2C**). The defect in photosynthetic complex

stoichiometry and organization was especially strong for the *prin2.2* mutant.

Expression analyses were performed for genes categorized as genes primarily transcribed by PEP and NEP, respectively. The *psaA*, *psbA, psbD,* and *rbcL* genes belong to class I genes transcribed by the PEP polymerase. Consistent with previous studies (Bollenbach et al., 2009; Kindgren et al., 2012; Qi et al., 2012), both *prin2.2* and *csp41b-2* mutants showed decreased *psaA*, *psbA, psbD,* and *rbcL* expression levels compared to wild type. In contrast, the expression levels of the NEP genes, *accD, rpoB, ycf2* were elevated compared to wild type (**Figure 2D**). The chloroplast transcriptional machinery