were grown on soil under long day conditions (16 h light/8 h darkness) to characterize *prin2.2* and *csp41b-2* mutants **(A)** Representative images of plants grown on soil for 3 weeks. **(B)** Representative TEM images of chloroplasts from the Col-0, *csp41b-2,* and *prin2.2* rosette plants. Bars represent 1 μm **(C)** Blue Native PAGE of thylakoid membrane complexes expression levels of chloroplast encoded genes in *prin2.2* and *csp41b-2* mutant plants. Expression levels were compared to the respective Col-0 samples and calculated using Ubiquitin-protein ligase (At4g36800) as a reference gene. Data represents the mean from three independent biological replicates.

that primarily utilizes PEP polymerase is clearly impaired in the *prin2.2* and *csp41b-2* mutants and when the two mutants were compared side by side the effect was stronger in the *prin2.2* mutant.

## **THE** *prin2.2* **AND** *csp41b-2* **MUTANTS SHOW DEFECTS IN EMBRYO DEVELOPMENT AND THE** *csp41b-2 prin2.2* **DOUBLE MUTANT IS EMBRYO LETHAL**

In order to investigate the genetic interaction between PRIN2 and CSP41b we attempted to generate a *csp41b-2prin2.2* double mutant (Supplementary Figure S2). However, the *csp41b-2prin2.2* double mutant was embryo lethal and the *CSP41b-2prin2.2/csp41b-2prin2.2* mutant produced siliques where 18% (green:albino ovule = 240:51, Chi-square 8,67 for *p* < 0,05) of all ovules appeared opaque (**Figure 3A**, arrows). Those impaired ovules finally turned into shrunken, dark colored seeds unable to germinate on MS media. A few ovules were also aborted at very early developmental stages (**Figure 3A**, arrowheads). The fact that theoretical 3:1 segregation green/albino is not supported by our statistics could be explained by the observed range in the stage at which the embryo development is arrested. Given the embryo lethality of the double mutant we investigated if there were any effects during embryo development in the *csp41b-2* and *prin2.2* single mutants. The *csp41b-2* embryos were undistinguishable from the wild type at the heart stage (**Figure 3B**). However, at the linear cotyledon and mature green (MG) stages the *csp41b-2* embryos were not as uniformly green as the wild type embryos. As has been shown before wild type embryos displayed a specific pattern of chlorophyll autofluorescence during embryogenesis (Tejos et al., 2010). This pattern was significantly altered at the linear cotyledon stage (LC), where the chloroplast containing tissue was mostly localized to the epidermal layers, in the *csp41b-2* embryos (**Figure 3C**). Distribution of chloroplast containing tissue was even more altered in the *prin2.2* embryos (**Figure 3C**). Consistent with these findings are the light microscopy pictures demonstrating that the *prin2.2* embryos were paler than wild type embryos at all developmental stages (**Figure 3B**).

To investigate the potential role of PRIN2 and CSP41b during embryo development, we used TEM to examine morphological differences in the embryos at the MG stage. Plastids of wild type embryos develop normally and have numerous thylakoids organized in grana stacks indicating that the plastids can be photosynthetically functional at this stage of embryo development (**Figure 4**). However, *csp41b-2* chloroplasts developed less thylakoid membranes and fewer grana stacks and exhibit quite often chloroplasts with large areas completely devoid of membranes (**Figure 4**). This specific defect in chloroplast structure of *csp41b-2* was also observed for the chloroplasts from 3-week-old plants (**Figure 2B**). The *prin2.2* showed an even stronger impairment in chloroplast development, the *prin2-2* plastids showed numerous vesicles and few thylakoid membranes and grana stacks. Thylakoid membranes were also often mis-oriented (**Figure 4**). Similarly to what was observed for the *csp41b-2* mutant the chloroplasts from the embryos of *prin2.2* showed similar defects to what was also seen in the adult plants (**Figure 2B**). Taken together our results indicate that chloroplast development in the embryo is impaired

in both *prin2.2* and *csp41b-2* single mutants and that the ovules are arrested at early developmental stages in the *csp41b*-2*prin2.2* double mutant.

## **EXPRESSION OF PHOTOSYNTHESIS GENES ESSENTIAL DURING EMBRYO DEVELOPMENT**

We performed *in silico* analysis of available array data where gene activity was profiled genome-wide in every organ, tissue, and cell type of *Arabidopsis* seeds from fertilization through maturity (Belmonte et al., 2013). We specifically investigated the expression of plastid encoded genes from preglobular (PG) to MG embryo stage. Overall transcription of chloroplastencoded photosynthesis associated genes was activated from globular to MG embryo stage (**Figure 5A**). The highest fold change in expression level was observed for the genes encoding PSI and PSII core subunits, ATPase, and the ribosomal subunits (**Figure 5A**). Thus, from the analysis of the plastid transcriptome it is clear that photosynthesis associated components are highly expressed during embryo development and that therefore PEP mediated transcription most likely is activated during this process. Expression of the nuclear encoded components associated with the PEP complex was demonstrated to increase from PG to LC (**Figure 5B**). Especially the genes encoding the PTAC proteins showed a very strong upregulation during embryo development. In addition, transcript levels of *TRXZ* were significantly up-regulated during the transition from PG to LC stage. Also transcription of *PRIN2* and *CSP41b* were observed during the LC stage of embryo development (**Figure 5B**).

The observed embryo phenotype in the *prin2.2* and *csp41b-2* mutants encouraged us to study expression of *psaA, psbA,* and *psbD* at the MG embryo stage. Expression of *psaA, psbA,* and *psbD* was strongly down regulated both in *prin2.2* and *csp41b-2* mutants compared to wild type (**Figure 5B**; Supplementary Figure S3). Thus, these results suggest that also under embryo development PRIN2 and CSP41b are important for proper transcription of PEP genes. Taken together, these results suggest that the PEP component of the chloroplast transcription machinery is active during embryo development and that its activity is essential for proper embryo and seed development.

## **DISCUSSION**

In leaf tissue, the initiation of chloroplast development in the light and the activation of the photosynthetic reactions are accompanied by repression of NEP activity and an increase of PEP-mediated plastid transcription (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). Expression of plastid-encoded photosynthetic components, thought to be mediated exclusively by PEP, was recently shown both in tobacco and barley to be driven by NEP in the absence of functional PEP, suggesting a less strict division of target genes between NEP and PEP (Legen et al., 2002; Lyubetsky et al., 2011; Zhelyazkova et al., 2012). NEP is particularly active in non-green tissues and in very young leaves (Emanuel et al., 2006). Similarly, the expression of the previously described

NEP-dependent genes was distinctly higher in developing compared with mature chloroplasts (Zoschke et al., 2007), indicating that NEP is very active during the early stages of chloroplast biogenesis. However, we have demonstrated that PEP activity is essential during embryo development in *Arabidopsis*. During embryo development NEP was unable to compensate for impaired PEP activity in the *prin2.2* and *csp41b-2* mutants. Thus, the switch from NEP- to PEP-dependent transcription of the plastid-encoded genes is essential for embryo and seed development in *Arabidopsis*.

*In silico* analyses of available microarray data (Belmonte et al., 2013) showed a gradual increase in expression of genes encoding

from the Col-0, *csp41b-2,* and *prin2.2* mature green (MG) embryos. Bars represent 1 μm.

components involved in photosynthesis and energy production from the PG to the MG stage of embryo development (**Figure 5A**). During embryogenesis, expression of *RPOT* encoding the plastid NEP polymerase was induced and the maximum expression level was reached as early as at the PG and globular stages (G; **Figure 5B**). The *RPOT* expression then diminished already at the heart stage (H; **Figure 5B**). The NEP promoter is the only promoter in the *rpoBC* operon, and expression of *rpoA, rpoB,* and *rpoC1/C2* followed the induction of *RPOT*. The core components of PEP were transcribed from the globular stage but in contrast to *RPOT*, expression was maintained all through the different embryo developmental stages (**Figure 5A**). The expression profile of the nuclear encoded sigma factors, especially SIGB (SIG2) was similar to the profiles of *rpoA, rpoB,* and *rpoC1/C2* (**Figure 5B**). Correlated with the induction of the genes encoding the core components of PEP was a very strong induction of the plastid encoded photosynthesis genes. Expression of *psbA,B,C,D,E,* and *psaA,B,J*, for example, increased approximately 10-fold when the MG embryo was compared to the PG suggesting that PEP transcription is activated during this developmental transition. The strong induction of photosynthesis related mRNAs during the PG to MG transition was also shown in a study where three periods of seed formation was investigated (Allorent et al., 2013). The embryo lethality of the

*csp41b-2prin2.2* double mutant further indicates that PEP driven transcription is required for effective transcription of photosynthesis genes to sustain the embryo with energy. It was also shown previously that genes associated with photosynthesis and carbon metabolism, are active in all embryo and endosperm sub-regions during early seed development (Belmonte et al., 2013) strongly suggesting there is a need for photosynthetic activity to contribute to embryo growth rate and biomass. Interestingly, at the MG embryo stage transcription of the nuclear encoded components of PEP was halted, suggesting that there is no need for further expression of chloroplast genes when the seeds enter the post-MG stage. In contrast, the chloroplast encoded photosynthesis genes showed a delayed response and maintained high expression also at the MG embryo stage. Possibly chloroplast transcription is subject to later anterograde regulation from the nucleus to repress expression.

True to its cyanobacterial origin, the core subunits of PEP are homologous to the cyanobacterial RNAP components (Martin et al., 2002). However, PEP also requires additional nuclearencoded factors for its function (Pfannschmidt et al., 2000; Pfalz et al., 2006). As many as 40–60 proteins appear to be present in the TAC from chloroplasts. From Arabidopsis and mustard TACs 35 components were identified and 18 of those components, called pTACs, were novel proteins (Pfalz et al., 2006).

In addition, TAC and sRNAP preparations from pro-plastids, chloroplasts and etioplasts have different protein compositions, suggesting a multifaceted regulation of PEP activity (Reiss and Link, 1985; Pfannschmidt and Link, 1994; Suck et al., 1996). Very high expression levels were observed for genes encoding the pTAC components PTAC3, PTAC10, PTAC12, FLN1, PTAC6, and TRXZ

during the globular (G) to LC stages. Expression of *PRIN2* and *CSP41B* was also detected during the LC stage. The expression of all these additional PEP components suggests that PEP activity requires many different components already during embryo development and that regulation of plastid transcription is complex and sophisticated from the very early stages of plant development.

PRIN2 and CSP41b were both identified in nucleoid preparations (Majeran et al., 2012) and our results suggest that these two components and possibly the PRIN2-CSP41b complex are essential for PEP dependent transcription during early embryo development. Using three independent methods CSP41b and PRIN2 were shown to directly interact (**Figure 1**). PRIN2 have two conserved Cys residues that possibly are responsible for monomer/dimer/oligomer formation upon oxidation. Consistent with this hypothesis is the observation of two bands of 20 and 40 kDa that might correspond to PRIN2 monomer and dimer (**Figure 1C**). Interestingly, the CSP41b protein migrated as continuous multimeric complex ranging from 40 to ∼700 kDa on the BN-PAGE. The presence of high molecular weight complexes was previously described for the native CSP41b protein in chloroplasts (Qi et al., 2012). Moreover, the CSP41b protein contains redox active Cys residues suggested to be putative targets of TRX (Stroher and Dietz, 2008) and the formation of high molecular weight complexes were also shown to be enhanced by oxidized conditions in the chloroplast stroma (Qi et al., 2012). When PRIN2 and CSP41b were incubated together they migrated on the gel as a distinct band (∼66 kDa; **Figure 1B**) suggesting the formation of a defined heteromeric protein complex containing both proteins. PRIN2 and CSP41b were shown to independently bind to promoter fragments of *psaA* and *psbA* in EMSA assays. However, when PRIN2 and CSP41b were incubated together, a new band of intermediate size was observed (**Figure 1D**, Supplementary data Figure S1), suggesting an interaction between PRIN2 and CSP41b also upon DNA binding *in vitro*. CSP41b was previously shown in RIP-chip analysis to be an RNA binding protein with specificity toward photosynthesis-related transcripts mainly expressed by PEP (Qi et al., 2012). However, many RNA binding proteins have also been described to have DNA binding properties, including TFIIIA, p53, STAT1, β/β subunits of RNAP and σ70, factors well known to regulate transcription (Sakonju et al., 1980; Clemens et al., 1993; Cassiday and Maher, 2002; Suswam et al., 2005). Another plant specific protein that binds both DNA and RNA is GUN1, a key component in retrograde communication between chloroplasts and nucleus (Koussevitzky et al., 2007). Possibly, the observed DNA binding of the PRIN2-CSP41b protein complex is required for full PEP activity as indicated by the embryo lethality of the *csp41b-2prin2.2* double mutant (**Figure 3A**).

Correlated with the embryo phenotype observed in *prin2.2* and *csp41b-2* single mutants was the impaired expression of chloroplast photosynthesis genes. The expression levels of *psaA, psbA,* and *psbD* were significantly lower in both mutants compared to wild type in MG embryos (**Figure 5C**), suggesting that the previously described defect in PEP-mediated gene expression in rosette plants is maintained in the mutants also during embryo development. Thus, our results establish a link between PEP activity and embryo development. Previously compromised translational and post-translational activities in the chloroplasts, such as mutations in elongation factor G and PPR proteins, have been shown to lead to embryo lethality (Ruppel and Hangarter, 2007; Khrouchtchova et al., 2012; Sosso et al., 2012). Interestingly, in tobacco and barley neither of the knockouts completely lacking PEP activity exhibits an embryo lethal phenotype (Allison

et al., 1996; De Santis-MacIossek et al., 1999; Zhelyazkova et al., 2012). However, it should be emphasized that an essential role of plastid activity and embryo greening during embryogenesis has so far only been reported for oil-seed plants such as *Arabidopsis* and *Brassica* (He and Wu, 2009; Hsu et al., 2010). In *Arabidopsis*, PEP activity appears essential during embryo development and during this process NEP is unable to compensate for the impaired PEP activity. Recently maize mutants lacking several PEP-associated proteins such as PTACs and PRIN2 demonstrated deficiency of numerous plastid tRNAs. Thus, a role for PEP, and for the PEP associated proteins, was demonstrated for the expression of plastid transfer RNA (Williams-Carrier et al., 2014). These results emphasize the complex division of labor between NEP and PEP during the initiation of chloroplast development and that PEP, and its associated proteins, are also essential for the translation of the photosynthetic components.

## **AUTHOR CONTRIBUTIONS**

Dmitry Kremnev carried out the experiments and analyzed the data. Dmitry Kremnev and Åsa Strand planned the study and wrote the manuscript. Both authors read and approved the final manuscript.

## **ACKNOWLEDGMENT**

This work was supported by grants from the Swedish research foundation, VR (ÅS).

### **SUPPLEMENTARY MATERIAL**

The Supplementary Material for this article can be found online at: http://www.frontiersin.org/journal/10.3389/fpls.2014.00385/ abstract

## **REFERENCES**

to excess light is mediated through the transcriptional activators ZINC FIN-GER PROTEIN EXPRESSED IN INFLORESCENCE MERISTEM LIKE1 and 2 in Arabidopsis thaliana. *Plant Cell* 24, 3009–3025. doi: 10.1105/tpc.112. 100099

**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: 14 March 2014; accepted: 19 July 2014; published online: 12 August 2014. Citation: Kremnev D and Strand Å (2014) Plastid encoded RNA polymerase activity and expression of photosynthesis genes required for embryo and seed development in Arabidopsis. Front. Plant Sci. 5:385. doi: 10.3389/fpls.2014.00385*

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

*Copyright © 2014 Kremnev and Strand. 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.*

## Establishment of the chloroplast genetic system in rice during early leaf development and at low temperatures

## *Kensuke Kusumi\* and Koh Iba*

*Department of Biology, Faculty of Sciences, Kyushu University, Fukuoka, Japan*

#### *Edited by:*

*Thomas Pfannschmidt, University Joseph Fourier Grenoble, France*

#### *Reviewed by:*

*Wataru Sakamoto, Okayama University, Japan Ben Field, Centre National de la Recherche Scientifique, France*

#### *\*Correspondence:*

*Kensuke Kusumi, Department of Biology, Faculty of Sciences, Kyushu University, Fukuoka 812-8581, Japan e-mail<EMAIL_ADDRESS>Chloroplasts are the central nodes of the metabolic network in leaf cells of higher plants, and the conversion of proplastids into chloroplasts is tightly coupled to leaf development. During early leaf development, the structure and function of the chloroplasts differ greatly from those in a mature leaf, suggesting the existence of a stage-specific mechanism regulating chloroplast development during this period. Here, we discuss the identification of the genes affected in low temperature-conditional mutants of rice (*Oryza sativa*). These genes encode factors involved in chloroplast rRNA regulation (*NUS1*), and nucleotide metabolism in mitochondria, chloroplasts, and cytosol (*V*2, *V*3, *ST1*). These genes are all preferentially expressed in the early leaf developmental stage P4, and depleting them causes altered chloroplast transcription and translation, and ultimately leaf chlorosis.Therefore, it is suggested that regulation of cellular nucleotide pools and nucleotide metabolism is indispensable for chloroplast development under low temperatures at this stage. This review summarizes the current understanding of these factors and discusses their roles in chloroplast biogenesis.

**Keywords: chloroplast transcription, translation,** *Oryza sativa***, low temperature, nucleotide metabolism**

## **INTRODUCTION**

Low temperature is a major abiotic constraint to plant growth. In rice, two stages of development are known to be the most sensitive to low temperatures the young seedling stage and the booting stage (Kaneda and Beachell, 1974; Cruz et al., 2013). At the booting stage, pollen sterility caused by low temperatures decreases the final grain yield. At the seedling stage, low temperatures reduce germination and delay leaf emergence and greening. Leaf chlorosis and yellowing are common symptoms when a low temperature prevails during this stage (Cruz et al., 2013), which suggests the low temperature arrests chloroplast development and functioning.

The effect of a low-temperature environment on chloroplast functions has been studied extensively (Berry and Björkman, 1980; Hasanuzzaman et al., 2013). A low temperature causes swelling of the thylakoid lamellae, vesiculation of the thylakoid, and ultimately breakdown of the entire chloroplast. A low temperature also inhibits electron transport and the carbon assimilation apparatus such as the Calvin cycle, ATP synthase, and ribulose 1,5-bisphosphate carboxylase/oxygenase (RuBisCO; Demmig-Adams and Adams, 1992; Asada, 1999; Yamori et al., 2011; Hasanuzzaman et al., 2013). However, these physiological symptoms have been mainly investigated in mature leaves containing functionally established chloroplasts. The molecular mechanisms underlying early chloroplast development under low temperatures have not yet been extensively studied.

*virescent* is a chlorotic mutant of higher plants causing young leaves to have a reduced chlorophyll content, but the chlorophyll levels recover as they grow (Archer and Bonnett, 1987). In contrast with other chlorotic mutants showing lethality such as *albino*, *chlorina*, and *xantha*, the *virescent* mutants are not terminal, and can reach maturity and produce seeds. Certain classes of the *virescent* mutations that have been reported are low-temperature conditional. They develop chlorotic leaves under low temperatures, but not under higher temperatures, suggesting a temporal aberration in a factor governing chloroplast development under low-temperature conditions. During the past decade, numerous genes responsible for *virescent* mutations have been identified in rice, and they have been shown to be involved in the chloroplast genetic system, including transcription, translation, and nucleotide metabolism. Because many of these genes are expressed temporally during early leaf development, they are probably involved in the establishment of the plastid genetic system at this phase under low temperatures. Here, we introduce four factors involved in chloroplast biogenesis under low-temperature conditions (NUS1, GKpm, RNRS1, and RNRL1) that have been identified through genetic and functional analysis of *virescent* mutants of rice (*v*1, *v*2, *v*3, and *st1*).

## **TEMPERATURE-SENSITIVE PHASE OF** *virescent* **MUTANTS DURING EARLY LEAF DEVELOPMENT**

*virescent-1*, *-2*, and *-3* (*v*1, *v*2, *v*3) were the first *virescent* mutants reported in rice and the mutations have been used as classical genetic markers (Omura et al., 1977). They develop chlorotic leaves at a restrictive low temperature (20◦C) but nearly normal green leaves at a permissive higher temperature (30◦C; **Figure 1A**). They are often hard to distinguish from each other, showing similar phenotypes. An important characteristic of these *virescent* mutants is that the leaf phenotype is not influenced by growth temperature after its emergence (Iba et al., 1991). This indicates that the leaf phenotype is irreversibly determined by the environmental temperature at a certain developmental stage before emergence. Furthermore, this phenotype can be useful for determining the temperature-sensitive period (TSP), by shifting the temperature from restrictive to permissive during leaf development, or vice versa. This technique was originally performed with *Drosophila* (Suzuki, 1970), in which the TSP for conditional mutants was limited to particular stages of development. Temperature-shift experiments showed that the TSPs of all *v*1, *v*2, and *v*<sup>3</sup> mutants was at stage P4 of leaf development (Iba et al., 1991).

Rice has the striking feature of leaf primordia production (plastochron) that is synchronized with leaf emergence (phyllochron) in shoot development (Nemoto and Yamazaki, 1993; Itoh et al., 2005). This regularity of leaf development enables a series of successive stages to be defined, starting with P0 (leaf founder cells), through P1 (youngest primordium), P2, P3, P4, and P5, to P6 (a fully expanded leaf; **Figure 1B**). Anatomical studies have shown that in rice the P4 stage is characterized by rapid leaf blade elongation (Itoh et al., 2005). Leaves at the P4 stage have an initial length of 3–5 mm and reach a final size of about 8–10 cm. Chlorophyll concentration per unit of fresh weight is negligible in the early P4 stage, and increases to about 40% in a mature leaf (Kusumi et al., 2010a). Electron microscopic observations have indicated that chloroplasts in the leaves at the early P4 stage have a small spherical shape (below 1 μm) and poor internal thylakoid structures. After the mid-P4 stage, thylakoid extension and grana formation in chloroplasts has been observed within the mid-portion of the leaf (Kusumi et al., 2010b).

The process of chloroplast development is divided roughly into three steps: (i) plastid division and DNA replication; (ii) establishment of the plastid genetic system; and (iii) activation of the photosynthetic apparatus (Jarvis and Lopez-Juez, 2013; Yagi and Shiina, 2014). These stepwise processes are partially achieved by two plastidial RNA polymerases; a nucleus-encoded phage-type RNA polymerase (NEP), and a plastid-encoded bacterial-type RNA polymerase (PEP; Hajdukiewicz et al., 1997; Yagi and Shiina, 2014). Plastid genes involved in the second step are known to be mainly transcribed by NEP, and those involved in the third step are transcribed by PEP (Yagi and Shiina, 2014). Analyses for chloroplast transcript accumulation revealed that the first step of chloroplast differentiation is likely to start in the leaves at the P0–P3 stages, and will largely finish during the early P4 stage (**Figure 1C**; Kusumi et al., 2010b). The second step occurs significantly in the leaves at mid-P4, and the decline of the second step and onset of the third step take place during late P4 (Kusumi et al., 2010b). The accumulation of tRNAGlu, a bifunctional molecule mediating the early steps of chlorophyll synthesis, and the switching of transcription from NEP to PEP (Hanaoka et al., 2005), showed two peaks (late-P4 and P5). The first activation of tRNAGlu can be related to the NEP–PEP transition. Therefore, the TSP of *v*1, *v*2, and *v*<sup>3</sup> mutants at the P4 stage suggests that they may be related to the establishment of the chloroplast genetic system, which is the major process occurring at this stage.

## **REGULATION OF CELLULAR NUCLEOTIDE POOLS INVOLVED IN THE CHLOROPLAST DEVELOPMENT**

*Virescent-2* (*V*2) was the first gene isolated from *virescent* mutants of rice (Sugimoto et al., 2007). Functional analyses showed that *V*<sup>2</sup> encoded guanylate kinase (GK), a key enzyme in guanine nucleotide biosynthesis that catalyzes the conversion of GMP to

*v*2, and *v*3) grown at a restrictive temperature (20◦C). **(B)** Schematic of a rice seedling with a fully expanded third leaf. L1, L2, L3, and L4 indicate the first, second, third, and fourth leaf, respectively. Developmental stages (P0–P6) are bottom of the shoot and contains pre-emerged leaves at P0–P3 stages. **(C)** Plastid gene expression patterns during leaf development (Kusumi et al., 2010a). Horizontal bars indicate the leaf developmental stages.

GDP (**Figure 2**). In bacterial and animal species, GK is localized in the cytoplasm and participates in maintenance of the guanine nucleotide pools. Plants possess two types of GK; cytosolic GK (GKc) and plastid/mitochondrial GK (GKpm; Sugimoto et al., 2007). Analysis of RNAi knockdown plants showed that GKc is essential for the growth and development of plants, but not for chloroplast development (Sugimoto et al., 2007). *V*<sup>2</sup> is a single-copy gene encoding the GKpm protein. *V*2*-*encoded GKpm predominantly accumulates in developing leaves at the P0–P4 stages (Sugimoto et al., 2007), which is consistent with a temperature-shift experiment in which the *V*<sup>2</sup> gene product was shown to be necessary at the P4 stage. A chloroplast possesses its own nucleoside diphosphate kinase that catalyzes subsequent GDP to GTP conversion (**Figure 2**; Sugimoto et al., 2007; Kihara et al., 2011; Nomura et al., 2014). Therefore, GKpm can limit the GDP/GTP pool in the chloroplast. Reduction of GKpm activity will cause a shortage of the GTP necessary for the assembly and function of the plastid translation machinery. In the *v*<sup>2</sup> mutant, Val162 has been substituted with Ile, which caused a 20-fold reduction in specific GMP activity (Sugimoto et al., 2007), and severely suppresses chloroplast translation (Sugimoto et al., 2004). Similarly, bacterial GTPases have important roles in ribosome biogenesis and protein translation (Verstraeten et al., 2011). In *Arabidopsis* and tobacco, plastidial GTPases have been reported to be involved in chloroplast rRNA processing and ribosome biogenesis in higher plants (Jeon et al., 2014). It has also been reported that an *Arabidopsis* mutant deficient in GTP-dependent chloroplast elongation factor G developed pale cotyledons and greenish true leaves, as observed in the GKpmdeficient *Arabidopsis* (Albrecht et al., 2006; Sugimoto et al., 2007).

This phenotypic similarity suggests the involvement of GKpm in the regulation of plastid translation, via limitation of the GTP pool.

Additionally, it was recently reported that GKpm is a target of regulation by guanosine 3 ,5 -bisdiphosphate (ppGpp) in chloroplasts of rice, as well as those of peas and *Arabidopsis* (Nomura et al., 2014). In bacteria, ppGpp is a key regulatory molecule that controls the stringency of responses through direct interaction with protein factors involved in gene expression such as RNA polymerase, translation factors, and DNA primase (Potrykus and Cashel, 2008; Tozawa and Nomura, 2011). In higher plants, ppGpp is synthesized in chloroplasts from GTP (GDP) and ATP (**Figure 2**). Major ppGpp synthase/hydrolase enzymes, named RSH (*RelA/SpoT homolog*), are localized to chloroplasts (Mizusawa et al., 2008). It has also been reported that ppGpp can negatively regulate chloroplast RNA polymerase (Sato et al., 2009) and the elongation cycle of translation (Nomura et al., 2012). This suggests that ppGpp functions as a regulatory molecule in chloroplasts, and interaction between GKpm and ppGpp may limit the GTP (and ATP) pool, which will subsequently retard chloroplast transcription and translation.

Nucleotide biosynthesis in the cytosol is also involved in the regulation of chloroplast differentiation at early leaf development under cold stress. Yoo et al. (2009) showed that the genes responsible for the *v*<sup>3</sup> and *st1* mutants of rice encoded the large and small subunits of ribonucleotide reductase (RNR), RNRL1 and RNRS1, respectively (Yoo et al., 2009). RNR is constructed from large RNR (α) and small RNR (β) subunits, which associate to form an active heterodimer complex (α2β2) and catalyze conversion of nucleotide diphosphates (NDPs) to deoxyribonucleotide diphosphates (dNDPs; **Figure 2**). Synthesized dNDPs are rapidly converted into dNTPs for DNA replication and repair. Therefore, the RNR activity affects the entire *de novo* nucleotide synthesis pathway *in vivo* (Elledge et al., 1992). As observed in the *v*<sup>3</sup> mutant, *st1* also caused low-temperature-dependent leaf chlorosis. RNRL1 and RNRS1 abundantly accumulated in the leaves at the P0–P4 stages, and this was enhanced by low temperature (Yoo et al., 2009). Both the *v*<sup>2</sup> and *st1* mutations caused missense mutations resulting in reduction of the first ab dimerization, which correlated with the degree of chloroplast disruption (Yoo et al., 2009). This suggests that a threshold level of RNR activity plays an important role in regulating nucleotide flow from the cytosol to chloroplasts. The involvement of cytosolic RNR in plastid nucleotide metabolism is further supported by the report that RNR deficiency causes plastid DNA degradation in pollen in *Arabidopsis* (Tang et al., 2012). Balancing chloroplast biogenesis and cell division during early leaf development would be achieved through optimization of the nucleotide pool in the cellular compartments.

## **NUS1 REQUIRED FOR rRNA MATURATION AT LOW TEMPERATURES**

In bacteria, synthesis of ribosomes requires a Rho-dependent antitermination system for the efficient transcription of 16S, 23S, and 5S rRNA from *rrn* operons (Santangelo and Artsimovitch, 2011). All *rrn* operons have anti-terminator sequences in their leader and spacer regions, referred to as BoxB, BoxA, and BoxC, that allow RNA polymerase, modified with protein factors, to transcribe rRNA operons. Previously known protein factors that interact with the anti-terminator include NusA, NusB, NusE, and NusG (Santangelo and Artsimovitch, 2011). Recently, *Virescent-1* (*V*1) was identified from a *v*<sup>1</sup> mutant of rice and shown to encode a novel chloroplast RNA binding protein, named NUS1 (Kusumi et al., 2011). The C-terminal region of NUS1 has a structural similarity to the RNA-binding domain of the bacterial NusB, which is classified as alpha helical with seven helices. Accumulation of NUS1 specifically occurs in the developing leaves at the P4 stage, and is enhanced by low-temperature treatment (Kusumi et al., 2011). Although there are no regions identical to bacterial Box regions within the chloroplast *rrn* operon in rice, the gene order of 16S–23S–4.5S–5S and their coding sequences are highly conserved with those of the bacterial *rrn* operon (Bollenbach et al., 2007). RNA-immunoprecipitation and gel mobility shift assays indicated that NUS1 binds to the upstream leader region of the 16S rRNA precursor (Kusumi et al., 2011). The *v*<sup>1</sup> mutant had a nonsense mutation in the helical domain and failed to accumulate the NUS1 protein, and therefore probably represents the null phenotype. In the *v*<sup>1</sup> seedlings grown at low temperatures, the processing and accumulation of chloroplast rRNA and chloroplast translation/transcription was severely suppressed (Kusumi et al., 1997, 2011). Additionally, *Arabidopsis* seedlings deficient in a *NUS1* ortholog also exhibited a similar phenotype (Kusumi et al., 2011). Therefore, NUS1 is likely to be involved in the regulation of rRNA maturation, which occurs at the P4 stage. Bacterial NusB is involved in the protein complex that interacts with RNA polymerase, nascent mRNA, and ribosomes (Santangelo and Artsimovitch, 2011; Bubunenko et al., 2013). Recent proteomics-based techniques have allowed the identification of previously uncharacterized proteins that contribute to the chloroplast genetic system (Majeran et al., 2012; Pfalz and Pfannschmidt, 2013). Majeran et al. (2012) showed that in maize, NUS1 and other factors structurally similar to the bacterial Nus-related factors, such as NusG and Rho, were included in nucleoid-enriched fractions. Examination of the physical interactions among these proteins, and identification of other factors interacting with the NUS1 protein, will be vital for elucidating the role of NUS1 in the regulation of the chloroplast genetic system.

## **CONCLUDING REMARKS**

Compared with other cereals such as wheat and barley, rice is susceptible to cold stress, probably because of its tropical origin (Cruz et al., 2013). The degrees of low-temperature sensitivity and damage vary according to the growth stage. Yoshida (1981) showed that temperature sensitivity varies between stages and that rice plants have a lower threshold temperature for cold damage during the early young seedling stage (10–13◦C) than during the reproductive stage (18–20◦C), making them less sensitive to low temperature as young seedlings. In field conditions, sudden low-temperature phases often occur during the early seedling development in spring. Therefore, it is reasonable to infer that rice developed this mechanism to protect leaf and internal chloroplast development against low temperature-induced retardation.

It has been known that processes of chloroplast translation are sensitive to cold stress. Environmental low temperature arrests protein synthesis by causing ribosomal pausing (Grennan and Ort, 2007) or retardation of ribosomal biogenesis and RNA processing (Millerd et al., 1969; Hopkins and Elfman, 1984; Barkan, 1993). Furthermore, loss of translational factors such as ribosomal protein (Rogalski et al., 2008; Fleischmann et al., 2011; Ehrnthaler et al., 2014; Song et al., 2014), translation elongation factor (Liu et al., 2010), rRNA methylase (Tokuhisa et al., 1998) and the RNA binding protein required for RNA processing (Kupsch et al., 2012) leads to sensitivity to low temperatures. These reports suggest the existence of a particular mechanism that protects chloroplast translation against cold stress, which can be expected to be associated with *NUS1*, *V*2, *V*3, and *ST1*.

The observed involvement of control of translation and nucleotide metabolism in low-temperature tolerance/adaptation has also been reported in bacteria. For example, in *Escherichia coli*, a temperature downshift hampers ribosome function, and ribosomes change their composition to function properly (Akanuma et al., 2012). It has also been reported that cold-shock proteins are often induced not only by low temperatures but also by translational inhibitors, such as chloramphenicol and tetracycline (Weber and Marahiel, 2003). Therefore, a reduction in translational capacity may be interpreted as a cellular signal triggering the cold adaptation response. Furthermore, analyses of bacterial mutants deficient in ppGpp synthesis showed that artificially induced high levels of ppGpp diminish the expression of cold-shock proteins, while low levels increase their production (Potrykus and Cashel, 2008). ppGpp synthesis is triggered by occupation of the ribosomal A-site by an uncharged tRNA (Potrykus and Cashel, 2008). Considering the hampered ribosomalfunction at low temperature, it is possible that a decrease in cellular ppGpp levels following a temperature downshift plays a physiological role in the regulation of gene expression and adaptation to growth at low temperature. The bacterial NusB protein has also been reported to be involved in cold tolerance. Cells containing a disrupted *nusB* gene are viable under standard growth conditions, but are cold sensitive (Quan et al., 2005). They are defective in rRNA synthesis and have a decreased peptide elongation rate at low temperatures. NusA, another host factor of the Nus complex, is also induced by cold treatment (Phadtare and Severinov, 2010), suggesting the importance of Nus and the anti-termination system in the cold response in bacteria. These similar properties between the chloroplast and the bacterial low-temperature response imply that higher plants have taken over the bacterial protective system in response to low temperature.

Recently, several other genes have been isolated from low temperature-conditional, chloroplast-deficient mutants of rice, such as *OsV4* (*virescent 4*), *wlp1* (*white leaf and panicles 1*), and *tcd9* (*thermo-sensitive chloroplast development 9*; Gong et al., 2014; Jiang et al., 2014; Song et al., 2014). The corresponding genes in *OsV4*, *wlp1*, and *tcd9* mutants encode plastidial pentatricopeptide repeat (PPR) protein, plastid ribosomal protein L13, and a subunit of chaperonin 60 (CP60α) required for chloroplast division, respectively. Similarly to *NUS1*, *V*2, *V*3, and *ST1*, their functions are speculated to be involved in early chloroplast development at low temperatures (Gong et al., 2014; Jiang et al., 2014; Song et al., 2014). It is possible that these factors are involved in a closely related mechanism to chloroplast protein expression and assembly, which is required at low temperatures, but not essential for chloroplast development during early leaf development at higher temperatures. Therefore, the maintenance of the developing plastid genetic system will be crucial for tolerance of cold at the seedling stage in rice.

#### **ACKNOWLEDGMENTS**

We are grateful to Dr. Yuzuru Tozawa (Ehime University) for helpful discussion. This work was supported by the Ministry of Education, Culture, Sports, Science, and Technology of Japan (Nos. 21114002, 22570045).

#### **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: 17 May 2014; accepted: 20 July 2014; published online: 11 August 2014. Citation: Kusumi K and Iba K (2014) Establishment of the chloroplast genetic system in rice during early leaf development and at low temperatures. Front. Plant Sci. 5:386. doi: 10.3389/fpls.2014.00386*

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

*Copyright © 2014 Kusumi and Iba. 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.*

## The nucleoid as a site of rRNA processing and ribosome assembly

## *Alexandra-Viola Bohne\**

*Department of Molecular Plant Sciences, Ludwig-Maximilians-University Munich, Planegg-Martinsried, Germany \*Correspondence<EMAIL_ADDRESS>
#### *Edited by:*

*Thomas Pfannschmidt, University Joseph Fourier Grenoble, France*

#### *Reviewed by:*

*Alice Barkan, University of Oregon, USA Katharine A. Howell, The University of Western Australia, Australia*

**Keywords: rRNA processing, ribosome assembly, nucleoid, plastid, mitochondria**

Protein biosynthesis is one of the key elements of gene expression in living cells. All proteins are synthesized on ribonucleoprotein complexes, the ribosomes. In bacteria and their derivatives eukaryotic mitochondria and plastids ribosomes consist of a small and a large subunit which together comprise more than 50 ribosomal proteins and usually two to four ribosomal RNAs (rRNAs). Synthesis of the required rRNAs and proteins, and their correct folding, maturation/modification and assembly into functional particles are highly coordinated. However, whereas ribosomal composition and many mechanistic aspects of their biogenesis are well understood, little is known about the spatial organization of the procedure in bacteria and organelles. In eukaryotes, the individual processes involved occur in defined regions of the cell: ribosomal proteins are synthesized in the cytosol, but most rRNAs are transcribed, processed and modified in the nucleolus, a distinct subnuclear compartment (Lafontaine and Tollervey, 2001; Boisvert et al., 2007). Only the transcription of the small *5S* rRNA occurs in the nucleoplasm. After their synthesis, ribosomal proteins and assembly factors are imported into the nucleus, where they are combined with the appropriate rRNAs. Subsequently, small and large ribosomal subunits are exported into the cytosol, where they pair up to form functional ribosomes.

This Opinion Article focuses on recent findings which support the idea that, as in eukaryotes, ribosomal biogenesis in bacteria, mitochondria, and plastids is spatially organized. In these systems, there is growing evidence that rRNA processing and ribosome assembly most likely take place in association with the nucleoid.

## **rRNA PROCESSING, MATURATION AND RIBOSOME ASSEMBLY IN BACTERIA**

The processes of rRNA maturation and ribosome assembly are probably best understood in bacteria (reviewed in Kaczanowska and Rydén-Aulin, 2007; Shajani et al., 2011). In *E. coli*, the small *30S* ribosomal subunit contains 21 ribosomal proteins and a *16S* rRNA, while the large *50S* subunit consists of 33 proteins and two rRNAs, the *23S* and *5S* rRNAs (reviewed in Melnikov et al., 2012). All rRNAs are encoded in a polycistronic gene cluster and transcribed as a single precursor, which undergoes extensive processing and maturation to generate the mature rRNAs (reviewed in Deutscher, 2009; Shajani et al., 2011). The processing reactions are carried out by the exoor endonucleolytic activities of at least five ribonucleases (RNases), including RNases III, E, G, T, and YbeY (reviewed in Deutscher, 2009; Davies et al., 2010). Furthermore, the *23S* and *16S* rRNAs undergo methylations and pseudouridylations at several positions, which are assumed to influence ribosomal structure and function (reviewed in Shajani et al., 2011).

Many of these maturation steps are coupled and take place on nascent rRNAs. The final rRNA folding pattern is probably determined by an ordered sequence of interactions with ribosomal proteins and assembly factors, which induce conformational changes and stabilize the proper rRNA structures (reviewed in Shajani et al., 2011).

It was long believed that DNAassociated bacterial ribosomes translate nascent mRNAs while these are being synthesized by the RNA polymerase (co-transcriptional translation). Striking evidence for this hypothesis came from early electron microscopy studies showing ribosome arrays (polysomes) attached to mRNA strands that were being transcribed from DNA by multiple RNA polymerase (RNAP) molecules (Miller et al., 1970). However, recent research suggests that, at least in *E. coli* and *B. subtilis*, most translation is not coupled to transcription, and that cotranscriptional translation may be limited to mRNAs encoding membrane proteins. By tracking the distribution of RNA polymerases (as markers for the nucleoid DNA) and ribosomes by means of fluorescence microscopy, a clear segregation of the nucleoid from ribosome-rich regions of the cytoplasm was observed (Lewis et al., 2000; Bakshi et al., 2012; Chai et al., 2014). Bakshi et al. (2012) found approximately 85% of the ribosomes in the ribosome-rich regions, while only 10 to 15% were detected in close proximity to the nucleoid. The majority fraction probably comprises actively translating "protein factories." The nucleoid-associated particles are thought to be in various stages of assembly, as several rRNA maturation steps occur in a cotranscriptional and assembly-assisted manner (reviewed in Shajani et al., 2011).

Further data support the identification of the nucleoid as the site of early rRNA processing events. RNase III, which cotranscriptionally cleaves the primary rRNA transcript to yield *16S*, *23S*, and *5S* precursors, has been shown to be required for localization of the *pre*-*16S* rRNA 5 leader region to the nucleoid (Malagon, 2013). In RNA-FISH assays for single-cell visualization, Malagon (2013) found that the nucleoid localization of the *16S* 5 leader was dependent on the catalytic activity of RNase III, which provides indirect evidence for the presence of the enzyme in this region. Based on evidence for interplay between the Nus transcription elongation factors and RNase III in the modulation of pre-rRNA biogenesis (Bubunenko et al., 2013), Malagon further speculated that Nus proteins might serve to localize pre-rRNAs to the nucleoid.

## **LOCALIZED rRNA PROCESSING AND RIBOSOME ASSEMBLY IN ORGANELLES**

Eukaryotic plastids and mitochondria are descended from endosymbiotically acquired bacteria, and have retained their own genomes and machineries for gene expression during evolution. It is therefore not surprising that many aspects of these genetic systems, including genome organization and ribosome structure, resemble those of bacterial rather than eukaryotic systems. Similarly, evidence for sublocalization of rRNA processing and ribosome assembly events to the organellar nucleoids is now emerging.

### **MITOCHONDRIA**

Evidence for rRNA maturation and ribosome assembly in association with the mitochondrial nucleoid mainly derives from studies on mammalian mitochondria. Affinity purification of established protein components of the mitochondrial nucleoid results in copurification of ribosomal proteins and the ribosome assembly factor ERAL1 (He et al., 2012b). ERAL1, a GTP-binding protein with RNA-binding activity, which had previously been linked with a number of nucleoid proteins, interacts both with proteins of the small mitoribosomal subunit and its rRNA component (Dennerlein et al., 2010; Uchiumi et al., 2010). ERAL1 was proposed to act as a chaperone for the small ribosomal RNA, as its depletion leads to destabilization of the small subunit (Dennerlein et al., 2010).

Similarly, another mitoribosome assembly factor, the human GTPase NOA1 (C4orf14), a homolog of the plant Rif1 protein, has been found together with the small ribosomal subunit and translation factors in affinity-purified nucleoids (Flores-Pérez et al., 2008; He et al., 2012a). This finding prompted the suggestion that assembly of the small subunit in the mitochondrial nucleoid enables the direct transfer of newly transcribed mRNAs to the ribosome.

Like bacterial rRNAs, mammalian mitochondrial rRNAs undergo sitespecific methylations, although to a lesser extent (reviewed in Rorbach and Minczuk, 2012). The enzymes responsible belong to a family of rRNA methyltransferases, some of whose members have been reported to modify nascent rRNAs in association with the nucleoid (Lee et al., 2013). A more detailed analysis of one of these methyltransferases (named RNMTL1) revealed an additional interaction between this protein and the large ribosomal subunit, suggesting that assembly of mitochondrial ribosomes begins before rRNA transcription is complete (Lee et al., 2013).

Moreover, a very recent study by Bogenhagen et al. (2014) provides evidence that mammalian mitochondrial RNA processing enzymes, like RNase P and ELAC2 (tRNaseZL), as well as a number of nascent mitochondrial ribosomal proteins associate with nucleoids to initiate RNA processing and ribosome assembly. The authors therefore propose the mtDNA nucleoid as a critical control center for mitochondrial biogenesis.

### **PLASTIDS**

An extensive body of evidence for localized ribosome biogenesis comes from plastids. Besides numerous ribosomal proteins, the majority of proteins with known roles in ribosome biogenesis have been identified in a comprehensive proteomic analysis of the maize chloroplast nucleoid (Majeran et al., 2012; reviewed in Germain et al., 2013). Many of these nucleoid-enriched ribosome biogenesis factors function in rRNA processing, maturation and modification, as well as in ribosome assembly (summarized in **Table 1**).

Among the processing and splicing factors identified are many plant homologs of bacterial RNases that have been reported to be responsible for exo- and endonucleolytic cleavage of the large rRNA precursor and maturation of cotranscribed tRNAs (reviewed in Stoppel and Meurer, 2012; Germain et al., 2013). Most of the nucleoid-enriched proteins involved in ribosome assembly and rRNA modification are classified as either GTPases, RNA helicases or rRNA methylases (**Table 1**).

Two of the nucleoid-enriched plastid proteins listed in **Table 1** are homologous to the mitochondrial enzymes ERAL1 and NOA1, for which a nucleoid localization has also been reported (see above). For several others, links with the nucleoid are further supported by alternative approaches as exemplified in the following.

An immunological analysis of maize chloroplast subfractions revealed that the DEAD-box RNA helicase RH3, thought to function in assembly of the *50S* subunit, localizes to the chloroplast stroma and thylakoids, as well as to nucleoids (Asakura et al., 2012). For two recently characterized plant proteins required for ribosome biogenesis, DER and RAP, cytological evidence for nucleoid localization comes from GFP fusion experiments (Jeon et al., 2014; Kleinknecht et al., 2014). DER is a Double Era-like GTPase, whose bacterial homolog (also known as EngA) acts as a ribosome assembly factor in *E. coli*, was found to bind to the *50S* ribosomal subunit and to play a role in pre-rRNA processing in tobacco (Hwang and Inouye, 2006; Jeon et al., 2014). The *Arabidopsis* protein RAP is a member of the Octotricopeptide Repeat (OPR) protein family, and binds to the 5- leader sequence of the *16S* rRNA precursor (Kleinknecht et al., 2014). Depletion of RAP specifically affected the trimming/processing of the chloroplast *16S* rRNA precursor, which supports the identification of the nucleoid as the site of *16S* rRNA processing in chloroplasts. Preliminary data imply that RAP has no intrinsic RNase activity, and might influence *16S* rRNA maturation by conferring sequence specificity on an RNase or by modulating RNA secondary structures to control the accessibility of an RNase recognition site within the *16S* precursor (Kleinknecht et al., unpublished results).

However, in the case of many chloroplast proteins reported to be involved in various steps of ribosome biogenesis, GFP-based fusion studies provide no

**Table 1 | Nucleoid-localized proteins with proposed functions in rRNA processing, maturation and ribosome assembly in plant chloroplasts.**

*All proteins listed were identified as nucleoid proteins by Majeran et al. (2012). Proteins known to be involved in ribosome biogenesis but not identified in their nucleoid fraction, as well as factors that probably have only indirect effects on rRNA maturation and ribosome assembly, are not included in this list.*

*\*Nucleoid localization demonstrated by groups other than (Majeran et al., 2012) (see text).*

*aOnly most relevant publications referring to a function in ribosome biogenesis or nucleoid localization are listed.*

support for specific localization to the plastid nucleoid. One possible reason for this may be the use of transit peptide-GFP instead of full-length protein fusions, as determinants of nucleoid localization are unlikely to be encoded in the transit peptide. Accordingly, Park et al. (2011) also reported the localization of the protein PRBP, which likely functions in *4.5S* rRNA processing, to distinct spots within chloroplasts, which probably represent nucleoids. This sublocalization was only observed when the full-length protein was fused to GFP and not with a transit peptide-GFP fusion. Nevertheless, other proteins for which full length-GFP fusions were used could not be clearly assigned to the nucleoid (e.g., Flores-Pérez et al., 2008; Yu et al., 2008; Chi et al., 2012). This might be due to weak or transient association of the respective protein with the nucleoid. Alternatively, some of these proteins may perform further functions in other organellar subcompartments leading to ambiguous localization signals.

## **CONCLUSION AND FUTURE PERSPECTIVES**

Localization of rRNA processing and ribosome assembly to organellar nucleoids seems to be a general phenomenon derived from a bacterial ancestor. Unlike the case of the eukaryotic nucleus, no physical barrier intervenes between bacterial/organellar RNA synthesis and translation. The nucleoid might therefore provide a scaffold for an intra-organellar microenvironment which enables coupling of rRNA transcription to ribosome assembly. This might not only enhance the efficiency of ribosome assembly by substrate channeling, but also largely prevent the precocious association of mRNAs with immature *30S* ribosomal subunits. However, it remains to be shown whether the colocalization of rRNA processing and ribosome assembly with nucleoids has functional significance or simply reflects the rapid kinetics with which ribosome biogenesis factors bind to nascent rRNA targets.

Nonetheless, the growing awareness of the requirement for sublocalized cellular processes in apparently less organized bacteria and organelles, and the availability of more sensitive detection techniques including super-resolution imaging, will undoubtedly lead to new insights into the spatiotemporal organization of ribosome biogenesis in the future.

#### **ACKNOWLEDGEMENT**

I thank Joerg Nickelsen for critical reading of this article.

### **REFERENCES**

Asakura, Y., Galarneau, E., Watkins, K. P., Barkan, A., and van Wijk, K. J. (2012). Chloroplast RH3 DEAD Box RNA helicases in maize and Arabidopsis function in splicing of specific group II introns and affect chloroplast ribosome biogenesis. *Plant Physiol.* 159, 961–974. doi: 10.1104/pp.112.197525

phytochrome-regulated gene whose protein product binds to plastid ribosomal RNAs. *Planta* 236, 677–690. doi: 10.1007/s00425-012-1638-6

and for optimal induction of rRNA synthesis in *E. coli. RNA* 19, 1200–1207. doi: 10.1261/rna.038 588.113

plastid gene expression. *J. Exp. Bot.* 63, 1663–1673. doi: 10.1093/jxb/err401

**Conflict of Interest Statement:** The author declares 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: 27 March 2014; accepted: 19 May 2014; published online: 05 June 2014.*

*Citation: Bohne A-V (2014) The nucleoid as a site of rRNA processing and ribosome assembly. Front. Plant Sci. 5:257. doi: 10.3389/fpls.2014.00257*

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

*Copyright © 2014 Bohne. 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.*

## Complex(iti)es of the ubiquitous RNA-binding CSP41 proteins

## *Dario Leister 1,2\**

*<sup>1</sup> Department Biology I, Plant Molecular Biology (Botany), Ludwig-Maximilians-University Munich, Martinsried, Germany*

*<sup>2</sup> Department of Plant and Environmental Sciences, Copenhagen Plant Science Centre, University of Copenhagen, Copenhagen, Denmark*

*\*Correspondence<EMAIL_ADDRESS>
### *Edited by:*

*Thomas Pfannschmidt, University Joseph Fourier Grenoble, France*

#### *Reviewed by:*

*Zhong-Nan Yang, Shanghai Normal University, China*

**Keywords: Arabidopsis, chloroplast, gene expression, RNA, RNA-binding protein, transcription, translation**

Photosynthetic eukaryotes encode two copies of the CSP41 (Chloroplast Stemloop binding Protein of 41 kDa) protein that are of cyanobacterial origin. In *Arabidopsis thaliana*, the two CSP41 proteins belong to the group of mostabundant chloroplast proteins. Multiple functions have been described for CSP41 proteins, including roles in chloroplast rRNA metabolism and transcription. CSP41a and CSP41b interact physically. Recent data show that CSP41b is an essential and major component of highmolecular weight complexes that form in the dark, disassemble in the light, and bind chloroplast mRNAs coding for photosynthetic proteins and some ribosomal RNAs, but not the plastid-encoded RNA polymerase (PEP). This, together with the effects seen in leaves of plants lacking CSP41b, implies that complexes containing CSP41 proteins stabilize untranslated mRNAs and precursor rRNAs. This occurs in a redox-dependent manner and seems to be important in the absence light when the translation is less active. In this scenario, translation and transcription is secondarily affected by the decreased transcript stability.

## **CSP41 PROTEINS ARE ABUNDANT AND CONSTITUENTS OF COMPLEXES**

CSP41 proteins are highly abundant chloroplast proteins. Zybailov et al. (2008) grouped chloroplast stromal proteins into seven abundance classes and CSP41b is found in the group of highest abundance together with the Calvin cycle enzymes for instance. CSP41b is more abundant than CSP41a, which is found in the group of second highest abundance together with most chloroplast ribosomal proteins. Therefore, it does not come as surprise that CSP41 proteins have been detected in several stromal complexes, but not all of these associations are necessarily of physiological significance in the light of the ubiquity of these proteins.

The first report on CSP41a function described its binding *in vitro* to the 3- end of the *petD* mRNA (Yang et al., 1995, 1996). Subsequently, CSP41 proteins were also found in preparations that were enriched for the plastid-encoded RNA PEP (Pfannschmidt et al., 2000) or plastid ribosomes (Yamaguchi et al., 2003). However, later studies could not confirm that CSP41 proteins are part of the PEP complex (Suzuki et al., 2004; Pfalz et al., 2006). A ribosome association of CSP41a and b was also observed by Peltier et al. (2006) in their analysis of the stromal proteome in its oligomeric state extracted from highly purified chloroplasts of *Arabidopsis thaliana*. The two CSP41 proteins were each found at three different locations of the stromal colorless native (CN)-PAGE native gels: (i) in a complex larger than 950 kDa most likely associated with 70 S ribosomes, (ii) at 224 kDa, and (iii) at 106– 126 kDa. At 224 kDa, the only obvious potential partners are the ribosomal proteins L5 and L31. At 106–126 kDa, CSP41a and b possibly form a heterotrimer. A further proteomics analysis found CSP41b mostly in 0.8–2 MDa fractions of stromal high-molecular-weight (HMW) complexes, together with other proteins like subunits of the 30S part of the plastid ribosome and subunit E2 of the plastid pyruvate decarboxylase (LTA2) (Olinares et al., 2010).

Recently, Qi et al. (2012) found by co-immunoprecipitation experiments that the major interactor of CSP41b is the CSP41a protein. The majority of both CSP41 proteins comigrates in several distinct spots during 2D BN/SDS PAGE, implying that they are present in multimeric protein complexes mainly comprised of these two subunits. Because the HMW CSP41 complexes are disrupted by treatment with RNase, they should be associated with RNAs (Qi et al., 2012). RIP-chip analysis points to chloroplast mRNAs coding for photosynthetic proteins and some ribosomal RNAs, but no tRNAs or mRNAs for ribosomal proteins, as putative ligands of CSP41 complexes (see below). The only other protein, besides CSP41a, found in immunoprecipitates of tagged CSP41b was LTA2 (Qi et al., 2012), corroborating the results of the proteomic study of Olinares et al. (2010). Interestingly, LTA2 is not a highly abundant stromal protein (Zybailov et al., 2008); therefore, this interaction might be specific and not due to contamination.

Taken together, because of their high abundance, CSP41 proteins can be found as contaminants in preparations of several stromal complexes. However, the results obtained by Qi et al. (2012) imply that CSP41 does not functionally interact with PEP or the plastid ribosome, as proposed before. Indeed, CSP41 complexes appear to contain chloroplast mRNAs coding for photosynthetic proteins and some ribosomal RNAs.

## **CSP41 FUNCTIONS AT THE BIOCHEMICAL LEVEL**

Multiple functions have been assigned to CSP41 proteins. (1) RNase activity, with a preference for 3stem-loops (Yang and Stern, 1997; Bollenbach and Stern, 2003a,b). (2) Ribosomal biogenesis, because of the association of CSP41b with pre-ribosomal particles (Beligni and Mayfield, 2008). (3) Plastid transcription (Bollenbach et al., 2009). (4) Cytosolic functions based on its interaction with heteroglycans in the cytosol (Fettke et al., 2011).

Not all of the four tentative functions of CSP41 proteins described above might be physiologically relevant and rather represent artifacts similar to the multiple presences of CP41 proteins in HMW complexes. Moreover, the high abundance of CSP41 proteins argues against a specific catalytic function but points in the direction of a more general function, requiring relative large quantities of the protein. Recently, Qi et al. (2012) demonstrated by RIP-chip analysis that CSP41 can bind various chloroplast RNAs. This includes transcripts for the large Rubisco subunit (*rbcL*), PSI (*psaA*, *psaB*), and PSII (*psbA*, *psbC*, *psbD*) core proteins, and 16S and 23S rRNAs. Therefore, Qi et al. (2012) concluded that the CSP41 proteins might serve to stabilize RNAs. Indeed, in their *in-organello* assay the stability of two tentative target RNAs (one of them 23S rRNA) was found to be decreased in mutants lacking CSP41b. Consequently, such destabilization of the precursors of 23S and 16S rRNA might result in fewer functional ribosomes, and in turn in a decrease of the rate of chloroplast translation. As a further consequence of a reduced translation rate, a decline of the levels of PEP synthesis can be expected and, in turn, of the rate of transcription. Nevertheless, a direct effect on transcription/translation through binding of CSP41 to target transcripts cannot be entirely ruled out yet.

The findings that CSP41 displays endonuclease activity *in vitro* (Yang et al., 1996; Yang and Stern, 1997; Bollenbach and Stern, 2003a,b) and CSP41 proteins stabilize target RNAs *in vivo* are not necessarily mutually exclusive, because the endoribonuclease activity of CSP41 could be highly regulated *in vivo*, e.g., by phosphorylation (Qi et al., 2012). Thus, complexes of CSP41 in its inactive state (without endonuclease activity) might stabilize RNAs by binding to protect them against degradation. Certain conditions could then activate the ribonucleolytic activity of CSP41, leading to the degradation of the target transcripts; however, it remains to be clarified whether CSP41 actually plays a role as RNase *in vivo*. In this context it is interesting to note that changes in the pIs of CSP41b species between dark and light conditions suggests that redox-dependent post-translational modifications of CSP41 might regulate the capacity of CSP41 complexes to bind RNA (Qi et al., 2012).

Immunoprecipitates of tagged CSP41b contain also LTA2, the E2 subunit of the plastid pyruvate decarboxylase (Qi et al., 2012). Interestingly, the counterpart of LTA2 in the green alga *Chlamydomonas reinhardtii*, DLA2, binds *psbA* mRNA and has been implicated in the reciprocal regulation of protein synthesis and carbon metabolism for thylakoid membrane biogenesis (Bohne et al., 2013). Therefore, their *psbA* transcript binding activity might bring together CSP41 proteins and LTA2 in the same complex.

## **MUTANT ANALYSES OF CSP41 FUNCTIONS IN PLANTS**

The *csp41b* mutation affects the morphology of chloroplasts, photosynthesis and circadian rhythms (Hassidim et al., 2007). Based on the observation that *A. thaliana* mutants without both CSP41 proteins are not viable, Beligni and Mayfield (2008) proposed that CSP41a and CSP41b have redundant functions. However, recent data by Qi et al. (2012) argue in favor of the notion that CSP41a and CSP41b do not have entirely redundant functions and that CSP41b is functionally more important than CSP41a. (1) While loss of CSP41a does not result in obvious phenotypic effects, chloroplast RNA levels and plant performance are impaired when CSP41b is inactive. Moreover, the *csp41ab* double mutant behaves like *csp41b* mutant plants (Qi et al., 2012). (2) Although CSP41 protein complexes seem to contain both proteins in wild-type plants, only CSP41b is essential for their formation. (3) Phylogenetic analysis of CSP41 sequences

**FIGURE 1 | Model for action of CSP41 protein complexes.** In the dark, CSP41 protein complexes associate with various mRNAs and some pre-rRNAs and protect them from nucleolytic cleavage. Untranslated RNAs not stabilized by CSP41 protein complexes are degraded by ribonucleases. Newly synthesized precursors of rRNAs are rapidly incorporated in the light into functional ribosomes, which in turn stabilize plastid transcripts during translation. In the absence of CSP41b, protection of mRNAs and pre-rRNAs in the dark is impaired and HMW RNA-CSP41 complexes are not formed. In consequence less functional ribosomes and mRNAs are available in the light. Therefore, less photosynthetic subunits are synthesized and the translational capacity is generally decreased. The latter can explain the pleiotropic effects seen in *csp41b* plants. Modified from Qi et al. (2012).

from *A. thaliana, C. reinhardtii* and the cyanobacterium *Synechocystis* suggest that CSP41a might be less constrained evolutionarily (Qi et al., 2012).

The major CSP41 protein, CSP41b, accumulates predominantly in mature leaves (Fettke et al., 2011). Accordingly, the function of the CSP41 proteins appears to be particularly required in mature leaves, as determined by measurements of translation rate and photosynthetic activity (Qi et al., 2012). Interestingly, mutants with reduced PEP levels show on opposite behavior compared to *csp41b* mutant plants with normal mature leaves but compromised younger leaves (Chi et al., 2008; Chateigner-Boutin et al., 2011). Moreover, the sets of mRNAs that are bound by CSP41 complexes or which are transcribed by the PEP overlap. Therefore, it can be concluded that in young leaves sufficient transcripts are synthesized by PEP such that the function of CSP41 complexes is not required. In older leaves, however, chloroplast gene expression can only be maintained by transcript stabilization through CSP41 complexes. In line with this, it has been described that the stability of chloroplast transcripts increases with the leaf age (Klaff and Gruissem, 1991). In fact, the post-translational modification of CSP41 proteins could represent a development-dependent regulatory mechanism by which the function of CSP41 is controlled.

In the light with highest activity of the chloroplast translational machinery the CSP41 proteins fail to form significant amounts of HMW complexes. On the contrary, darkness induces formation of HMW CSP41 complexes that are sensitive to treatment with RNase (Qi et al., 2012). This, together with the assumption that increased polysomal association of transcripts serves to stabilize them in the light (Qi et al., 2012), is the basis for our working model for the mechanism by which CSP41 complexes stabilize mRNAs in the dark (**Figure 1**). In this model, CSP41 proteins bind in the dark to non-translated mRNAs and rRNA precursors (which are not incorporated in ribosomes) to protect them against degradation. As soon as translation is activated in the light, this would allow the rapid initiation of translation and elongation from these stabilized transcripts, as well as *de-novo* assembly of ribosomes. In the mutants that lack CSP41 complexes, untranslated target mRNAs and precursors of rRNAs are prone to increased degradation. A role of the chloroplast redox state in the regulation of the association of CSP41 proteins with their RNA targets in the dark (and their light-induced dissociation) became evident when studying a photosynthetic mutant line with a more oxidized redox state of the stroma (Qi et al., 2012): here, HMW CSP41complexes can persist also in the light.

## **CONCLUSIONS**

The key to understand the multiple functions of CSP41 proteins probably lies in their abundance. Actually, CSP41 proteins are the most abundant RNA-binding proteins in the chloroplast stroma. Their function becomes critical in mature leaves when transcripts produced by PEP might become limiting and need to be stabilized and protected. Therefore, lack of CSP41 proteins decreases transcripts for photosynthetic proteins and of some ribosomal RNAs, which in turn, appears to result in pleiotropic effects due to a decrease in the translational activity of chloroplasts.

## **REFERENCES**

Proteomic characterization of the *Chlamydomonas reinhardtii* chloroplast ribosome. Identification of proteins unique to the 70S ribosome. *J. Biol. Chem*. 278, 33774–33785. doi: 10.1074/jbc.M301 934200

**Conflict of Interest Statement:** The author declares 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 March 2014; accepted: 19 May 2014; published online: 06 June 2014.*

*Citation: Leister D (2014) Complex(iti)es of the ubiquitous RNA-binding CSP41 proteins. Front. Plant Sci. 5:255. doi: 10.3389/fpls.2014.00255*

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

*Copyright © 2014 Leister. 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.*

## A purification strategy for analysis of the DNA/RNA-associated sub-proteome from chloroplasts of mustard cotyledons

## *Yvonne Schröter 1, Sebastian Steiner 1,2, Wolfram Weisheit 1,3, Maria Mittag1,3 and Thomas Pfannschmidt 1,4,5,6,7\**

*<sup>1</sup> Lehrstuhl für Pflanzenphysiologie, Institut für Allgemeine Botanik und Pflanzenphysiologie, Friedrich-Schiller-Universität Jena, Jena, Germany*

*<sup>2</sup> KWS SAAT AG, Einbeck, Germany*

*<sup>3</sup> Department of General Botany, Institute of General Botany and Plant Physiology, Friedrich Schiller University Jena, Jena, Germany*

*<sup>4</sup> University of Grenoble-Alpes, Grenoble, France*

*<sup>5</sup> CNRS, UMR5168, Grenoble, France*

*<sup>6</sup> Commissariat a L'energie Atomique (CEA), iRTSV, Laboratoire de Physiologie Cellulaire & Végétale, Grenoble, France*

*<sup>7</sup> INRA, USC1359, Grenoble, France*

#### *Edited by:*

*Steven Carl Huber, United States Department of Agriculture - Agricultural Research Service, USA*

#### *Reviewed by:*

*Frederik Börnke, Leibniz-Institute for Vegetable and Ornamental Crops (IGZ), Germany Karin Krupinska, Christian-Albrechts University of Kiel, Germany*

#### *\*Correspondence:*

*Thomas Pfannschmidt, Commissariat a L'energie Atomique (CEA), iRTSV, Laboratoire de Physiologie Cellulaire & Végétale, 17 Rue des Martyrs, 38000 Grenoble, France e-mail: Thomas.Pfannschmidt@ ujf-grenoble.fr*

Plant cotyledons are a tissue that is particularly active in plastid gene expression in order to develop functional chloroplasts from pro-plastids, the plastid precursor stage in plant embryos. Cotyledons, therefore, represent a material being ideal for the study of composition, function and regulation of protein complexes involved in plastid gene expression. Here, we present a pilot study that uses heparin-Sepharose and phospho-cellulose chromatography in combination with isoelectric focussing and denaturing SDS gel electrophoresis (two-dimensional gel electrophoresis) for investigating the nucleic acids binding sub-proteome of mustard chloroplasts purified from cotyledons. We describe the technical requirements for a highly resolved biochemical purification of several hundreds of protein spots obtained from such samples. Subsequent mass spectrometry of peptides isolated out of cut spots that had been treated with trypsin identified 58 different proteins within 180 distinct spots. Our analyses indicate a high enrichment of proteins involved in transcription and translation and, in addition, the presence of massive post-translational modification of this plastid protein sub-fraction. The study provides an extended catalog of plastid proteins from mustard being involved in gene expression and its regulation and describes a suitable purification strategy for further analysis of low abundant gene expression related proteins.

**Keywords:** *Sinapis alba***, cotyledon, chloroplast, nucleic acids binding protein, post-translational modification, mass spectrometry**

## **INTRODUCTION**

Plant chloroplasts are semiautonomous cell organelles of endosymbiotic origin that emerged from a cyanobacteria-like ancestor (Lopez-Juez and Pyke, 2005). One evolutionary remnant of this origin is their own genome (called plastome) comprising 100–120 genes and a pre-dominantly bacteria-like geneexpression machinery being essential for its proper expression. The plastome gene set in vascular plants is highly conserved and encodes mainly proteins with a function in photosynthesis and the gene expression machinery (Sugiura, 1992). However, for full functionality plastids require the import of many proteins that are encoded by the nuclear compartment since during evolution the endosymbiotic ancestor lost most of its genes to the nucleus of the host cell *via* horizontal gene transfer (Martin et al., 2002; Stoebe and Maier, 2002). These nuclear-encoded plastid proteins are translated in the cytoplasm as precursor molecules that are subsequently imported into plastids with the help of N-terminal transit peptides directing them to their correct sub-compartment (Soll and Schleiff, 2004). After removal of the transit peptide the mature proteins are then assembled into their final configuration together with the plastid-expressed proteins and, therefore, all major multi-subunit complexes (such as photosystems, ribosomes, or metabolic enzyme complexes) represent a patchwork of nuclear as well as plastid expressed proteins (Allen et al., 2011).

Based on the prediction of transit peptides and genome-scale proteomics it was estimated that plastids may contain around 1500–4000 different proteins (Abdallah et al., 2000; Baerenfaller et al., 2008; Ferro et al., 2010; van Wijk and Baginsky, 2011). Reference proteomes generated for maize and *Arabidopsis* cover 1564 and 1559 proteins, respectively, so far (Huang et al., 2013) indicating that a large part of the predicted plastid proteome has yet not been detected. This might be caused by the fact that plastids from different tissues (for instance roots, cotyledons, leaves, flowers, and fruits) likely contain different protein compositions, but also from the fact that especially regulatory proteins are present in only trace amounts that are difficult to detect in a matrix of highly abundant proteins, e.g., from the photosynthetic apparatus (Huang et al., 2013). Further complexity in the plastid protein complement may derive from the occurrence of multiple post-translational modifications that are essential for regulatory events.

Cotyledons display a high activity in plastid transcription and translation being essential for the light-induced development of chloroplasts out of the embryonic pro-plastids (Baumgartner et al., 1989, 1993). Thus, the proteome of cotyledon plastids comprises a high amount of proteins implicated in gene expression providing a useful source material for the characterization of the nucleic acids binding proteome. The chloroplast proteome of the dicotyledonous model organism *Arabidopsis thaliana* is well studied in adult leaves, however, an analysis of that of cotyledons is lacking mainly because the small size of the cotyledons is not very suitable for the isolation of chloroplasts and subsequent analyses of their proteins *via* chromatography. In recent investigations, the fast growing cruciferous plant mustard (*Sinapis alba*) demonstrated a high suitability for performing biochemical and physiological analyses of plastid gene expression in cotyledons since the seedlings and their cotyledons are much larger than that of *Arabidopsis* (Oelmuller et al., 1986; Tiller and Link, 1993; Pfannschmidt and Link, 1994; Link, 1996; Baginsky et al., 1997). Isolation of cotyledons in the order of kilograms is easily achieved after just 5 days of growth and provides enough material even for the biochemical analysis of low-abundant proteins by chromatography followed by mass spectrometry. Since *Sinapis* is a close relative of *Arabidopsis*, peptide data evaluation for the identification of mustard plastid proteins was found to be applicable for well conserved proteins by using the *A. thaliana* or *Brassicales* protein databases (Schröter et al., 2010; Steiner et al., 2011). Thus, the use of mustard as a source for cotyledons combines the advantages of mustard chloroplast preparation with the availability of protein data of well studied organisms like *A. thaliana* or some *Brassica* species.

In recent studies, proteins implicated in plastid gene expression in mustard have been isolated by a number of different purification schemes. These include the isolation of the membrane bound insoluble transcriptionally active chromosome (TAC) by ultracentrifugation and gel filtration (Hallick et al., 1976; Bülow et al., 1987; Pfalz et al., 2006) and the isolation of soluble proteins such as RNA polymerases, kinases, RNA binding proteins and sigma factors by various chromatographic steps (Tiller et al., 1991; Nickelsen and Link, 1993; Tiller and Link, 1993; Pfannschmidt and Link, 1994; Liere and Link, 1995; Baginsky et al., 1999). Recently, we applied the purification scheme of plastid isolation followed by protein enrichment *via* heparin-Sepharose (HS) chromatography and visualization by two-dimensional (2D) blue native (BN)-PAGE to isolate protein complexes such as the RNA polymerase complex as well as a number of gene expression related proteins (Schröter et al., 2010; Steiner et al., 2011). However, these HS purified fractions still included a number of metabolic enzymes which exacerbate the analysis of the nucleic acids binding sub-proteome as they tend to cover low abundant proteins or even hinder their visualization and identification. Here, we present a pilot characterization of the nucleic acids binding sub-proteome of chloroplasts from mustard cotyledons. To this end we used HS chromatography followed by a second chromatographic step with phosphocellulose (PC) which was shown to be very effective for isolating nucleic acids binding enzymes like RNA polymerases (Bottomley et al., 1970; Tiller and Link, 1993). This was followed by isoelectric focussing (IF) and 2D gel electrophoresis that allowed us to estimate the size of the nucleic acids binding sub-proteome and the ideal IF range for its visualization and protein determination using mass spectrometry. The use of 2D gel electrophoresis also revealed massive post-translational modifications of the sub-proteome.

## **RESULTS**

## **ENRICHMENT OF NUCLEIC ACIDS BINDING PROTEINS FROM MUSTARD CHLOROPLASTS**

In previous studies we analyzed gene expression related protein complexes from isolated mustard chloroplasts using a combination of HS chromatography followed by a two dimensional BN/SDS polyacrylamid gel-electrophoresis (2D BN-PAGE) and electro-spray ionization-tandem mass spectrometry (ESI-MS/MS). Besides the plastid-encoded RNA polymerase, various CSP41 complexes and translation related proteins, we identified several metabolic enzyme complexes such as GAP-dehydrogenase, ATPases, or RubisCO that co-purify in this affinity chromatography. These abundant proteins exacerbated the identification of further low-abundant proteins (Schröter et al., 2010; Steiner et al., 2011). In addition, these studies were focussed on the analysis of large native protein complexes using a BN-PAGE approach. This limited the characterization of gene expression related proteins that may occur in small complexes or as individual proteins. In this study, we aimed a deeper investigation of the size, composition and complexity of the nucleic acids binding subproteome of mustard chloroplasts. To this end, we performed chloroplast isolation and HS chromatography from mustard cotyledons precisely as described before (Schröter et al., 2010). Bound proteins were eluted with a high-salt step, concentrated by dialysis and, for further enrichment of gene expression related proteins, applied to a cation exchange column with PC as matrix as described earlier (see above). Proteins were eluted by a second high-salt step and dialyzed against a low-salt storage buffer for analysis and further use (see Materials and Methods). A first comparison of peak fractions with equal protein amounts of both purification steps was done by SDS-PAGE and silver staining (**Figure 1A**). The PC fraction exhibited a selective enrichment of many protein bands between 5 and 75 kDa and a strong exclusion of proteins larger than 75–80 kDa. For a more detailed resolution of this protein fraction, we performed 2D gel electrophoresis with an IF as first dimension followed by a SDS-PAGE (**Figures 1B,C**) as second dimension. Using IPG stripes with a non-linear (NL) pH range from 3 to 11 for the IF and a gradient polyacrylamide gel, we could obtain an overview of the total protein content leading to the identification of around 600 individual spots. We observed two major areas where multiple proteins accumulated on the gel which were located between approximately pH 4.5–7 and pH 9–11. Because of the non-linearity of the IF gradient, proteins at the outer ranges of the IF stripe were poorly resolved which became mainly evident at the basic pH values. Therefore, linear IPG gels were used in addition, overlapping with the first one between pH 3–10 and pH 6–11. The latter gradient resolved the problem with spot accumulation especially observed

gels using three different pH gradients in the first dimension. The pH gradient used is indicated in the upper left corner of each gel and the pH range is given in detail below each gel. Marker sizes are given in the right margin. Dotted lines indicate the overlapping pH areas. Four hundred microgram of total protein separated in each gel. **(D)** Numbering of protein spots visualized in the 2D gel with pH 3–11NL for first dimension as shown in **(C)**. Marker sizes and pH range are given in the margin or below the gel, respectively.

at the cathode. The higher resolution led to the identification of further proteins leading to a total count of 1079 individual protein spots within the PC fraction which could be distinguished between the different gels. We regard this as the nucleic acids

Protein pattern of PC peak fractions in silver stained 7.5–20% SDS acrylamide

binding sub-proteome of mustard plastids. Our data indicate a significant higher complexity of this specific sub-proteome as it was estimated earlier from the HS fractions (Schröter et al., 2010).

## **IDENTIFICATION OF PROTEINS FROM THE PC FRACTION BY LC-ESI-MS/MS**

All 1079 spots were cut out and proteins were subjected to an ingel tryptic digest. In 153 cases, selected spots were pooled from duplicate gels in order to increase the protein amount for the subsequent measurements. Since a database from *S. alba* is currently not available, protein identification was performed by comparing the determined mass spectrometry data to the *Brassicales* and *A. thaliana* databases (compare Materials and Methods). By this means 225 proteins were reliably identified with at least two different peptides in 180 spots indicating that several spots contained more than one protein. In addition, 36 particular proteins were identified in more than one spot (up to 40 different ones) suggesting post-translational modification of these proteins (**Table 1**). In total, 58 different proteins were identified. In further analyses, the identified gene models were checked for presence of a plastid transit peptide using TargetP (Emanuelsson et al., 2000) (**Figure 2**). Plastid-directing transit peptides could be predicted for 36 of these proteins, ten of them exhibit an additional luminal transit peptide and four plastid-encoded proteins were identified. Considering a detection probability of 73% for a transit peptide, we estimated the percentage of true plastid proteins within the PC fraction to be around 94%. Some of the identified proteins were found before in mustard (Pfannschmidt et al., 2000; Pfalz et al., 2006; Schröter et al., 2010), but 36 were identified here for the first time (**Table 1**).

Based on functional similarities and structural homologies, a categorization of proteins into protein families or subgroups was conducted (**Figure 3**). A practical classification mode is given by the modified MapMan bin system (Thimm et al., 2004) of the Plant Proteomics Data Base (PPDB) (Sun et al., 2009). In **Table 1**, proteins were listed following the PPDB bin grouping as given in column 2. For further comparison, we summarized identified proteins into five major groups. The first group comprises transcription and transcript related proteins, namely subunits of the plastid encoded RNA polymerase (PEPs) and PEP associated proteins (PAPs) as defined in Steiner et al. (2011), other pTACs (pTAC proteins not belonging to the PAPs) and RNA and DNA related proteins (bin 27 and 28, not belonging to PAPs and pTACs). A second large group comprises translation related proteins (bin 29.2 and 29.5). Three further groups cover proteins involved in protein homeostasis (bin 29 and 21 not belonging to PEPs and PAPs), photosynthesis (bin 1) and a miscellaneous group called "others" including various enzymes catalyzing metabolic reactions or protein modifications.

## **PEPs, PAPs, AND OTHER pTACs**

We detected most subunits of the soluble PEP complex including PAP3, PAP4, PAP5, PAP6, PAP8, PAP10, PAP11, PAP12 as well as the PEP core subunit RpoA (Pfalz and Pfannschmidt, 2013). Other PEP core subunits (RpoB, RpoC1, RpoB) and PAP1, PAP2, PAP7, and PAP9 were not identifiable in spots of these gels. Most of the identified proteins of this group became visible as single isolated spots in the acidic range (pH 3–6) on the gel (**Figure 4**) and at their expected molecular weight. An exception was PAP6 representing the protein fructokinase-like 1 (FLN1) that contains a protein domain of the pfkB-carbohydrate kinase family (Arsova et al., 2010; Steiner et al., 2011). This protein appeared in a chain of five spots of the same apparent molecular weight but with slightly varying isoelectric points from which the two strongest spots were identified as PAP6 here. This observation suggests post-translational modification of this kinase. In addition, for PAP6 but also for PAP3 and PAP11 one or two spots of lower molecular weight, respectively, were detected suggesting a targeted degradation or proteolytic modification of these two proteins (**Figure 4**). For PAP4 and PAP12 only a degradation product was detectable, while a spot of the full length protein was not identified.

Besides PEP and PAP proteins, we identified two proteins described as component of the TAC in mustard, PTAC4 and PTAC18 (Pfalz et al., 2006). PTAC4 is the vesicle-inducing protein in plastids 1 (VIPP1) which plays a crucial role in membrane stability (Zhang et al., 2012). The PTAC18 protein belongs to the cupin superfamily that merges proteins with a conserved βbarrel fold, giving this type of protein a strong thermal stability. It represents a family of very diverse members including enzymes and seed storage proteins, but also transcription factors (Dunwell et al., 2001). However, the exact function of pTAC18 is largely unknown. PTAC18 was identified in spot 255 being smaller and more in the acidic range as expected from the predicted protein representing likely a fragment. PTAC4 was identified in spots 243, 278, 280, and 295. Two hundred and seventy-eight and 280 are on the same size but with slightly different IPs suggesting post-translational modification of the protein.

An exceptional constituent of the PC protein fraction represents the protein CSP41 that appears in two forms, CSP41a and CSP41b. Originally described as the chloroplast stem-loop binding protein of 41 kDa (Yang et al., 1996) it has been discussed to be involved in RNA processing and stabilization as well as in RNA protection (Qi et al., 2012). As described for the HS fractions it represents a dominant protein of the nucleic acids binding proteome of plastids being present in multiple multimeric complexes of highly variable sizes (Schröter et al., 2010; Qi et al., 2012). In the PC fractions, the two forms of CSP41 appear to be especially enriched as they can be detected in 10 spots of the same apparent molecular weight of around 34 kDa but with different IPs (three for CSP41a and seven for CSP41b). The main accumulation is visible in the middle of the gels between pH 5.5 and 7. The CSP41a spots are by far the strongest spots observed in the whole gel followed by the spots for CSP41b. Roughly estimated they account for 30–40% of the total protein content in this fraction making a precise estimate difficult. In addition, the proteins are detectable in 34 less stained and smaller spots of different sizes suggesting massive post-translational modifications as well as multiple degradation or targeted proteolytic events acting on both protein forms. These smaller protein spots of CSP41a/b appear to contain not only random fragments of the proteins but could be observed as reproducible spot pattern in all replicates of nucleic acids binding sub-proteome preparations from mustard.

## **TRANSLATION ASSOCIATED PROTEINS**

Numerous proteins identified in this work are directly or indirectly related to translation. In total 12 ribosomal proteins of the large 50S subunit of plastid ribosomes (PRPL) were identified, **Table 1 | Functional categorization and characterization of proteins from the phosphocellulose fraction identified by LC-ESI-MS/MS.**

*(Continued)*

### **Table 1 | Continued**

*Identified proteins are named in the first column according to the annotation of the respective gene at NCBI. They are grouped into different classes written bold at the beginning of each group as defined in results. Proteins within each group are sorted alphabetically. Spots: number of the spots containing the respective protein. MapMan bin: classification groups for proteins according to the modified MapMan system of the plant proteome database (ppdb) [(http:// ppdb.tc.cornell.edu/ dbsearch/ mapman.aspx) ©Klaas J. van Wijk Lab, Cornell University; Sun et al., 2009] based on the MapManBins of Thimm et al. (2004); Accession: gi identification number and At gene accession number; cTP: possibility of a plastid transit peptide and the respective reliability class (RC); References: first identification of the protein in mustard by mass spectrometry.*

namely PRPL1, -4, -5, -6, -10, -12, -14, -15, -18, -21,-24, -29. The solely identified protein of the small 30S ribosomal subunit (PRPS) is PRPS5. We also identified two ribosomal subunits that belong to the large subunit of the cytosolic 80S ribosomes (CRPL), CRPL11 and -22-2. *S. alba* proteins of PRPL12-1 and PRPL29 were formerly identified by Pfalz et al. (2006) and PRPL6 by Schröter et al. (2010). The remaining ribosomal proteins listed in **Table 1** are identified in mustard plastid protein samples here for the first time.

Beside the ribosomal subunits a number of translation initiation factors (IF) were present in the fractions and were detected here for the first time in *S. alba*. Except of eIF1A (a subunit of the cytosolic translation initiation complex) all of them contain a predicted plastid transit peptide. This accounts also to eIF3 which is known as a subunit of a eukaryotic IF (eIF). IF2 and IF3 represent plastid translation IF while elongation factors (EF) EF-Tu and the eukaryotic EF1alpha4 are involved in translation elongation. eIF1A, EF-Tu, and EF1-alpha4 appear as single spots while the others were found in several spots suggesting post-translational modifications here, too.

Furthermore, we identified a SpoU methylase that belongs to the class of SPOUT enzymes and introduces a methylation of 2- -OH groups of tRNA or rRNA riboses (Cavaillé et al., 1999; Tkaczuk et al., 2007), and two proteins that are subunits of

**PC fractions. (A)** Distribution of the identified proteins of the HS fractions analyzed in Schröter et al. (2010) and classification into groups in correlation to the recent work. **(B)** Percentage of identified proteins of PC peak fractions analyzed in this work and classified into groups as shown in **Table 1**. **(C)** Distribution of solely the plastid proteins of the recent PC fractions to functional groups according to **Table 1** but with an aggregation of "PEPs and PAPs" with "Other pTACs" and a part of "DNA and RNA" to one bin "Transcription."

the nascent polypeptide associated complex (NAC). This dimeric complex is composed of an alpha- and beta-chain and may reversibly bind to ribosomes (Wiedmann et al., 1994). The alpha-NAC-like proteins identified during this work are encoded by different genes in *Arabidopsis* but exhibit a strong similarity within their amino acid sequence. The α-NAC like protein 1 and 3 were determined in the same two spots on the gels representing double spots.

## **PROTEINS INVOLVED IN PROTEIN HOMEOSTASIS, PHOTOSYNTHESIS, AND METABOLISM**

We identified the chloroplast heat shock cognate protein 70-2 (cpHsc70-2) which is the analog of one of only two stromal Hsp70s in *A. thaliana* plastids (Su and Li, 2008). In addition, we

found a TCP-1/cpn60 family chaperonin and a protein disulfide isomerase like 2-1 (PDIL 2-1) belonging to the thioredoxin superfamily and acting as folding catalyst. All proteins are identified in mustard fractions here for the first time and likely function in protein stability or formation. The correct folding of proteins is the last but essential step of gene expression.

The group of photosynthesis related proteins contains four proteins. The alpha and beta subunits of the plastid ATP synthase were formerly identified in *S. alba* (Schröter et al., 2010). Another ATPase, the RubisCO activase and the Rieske cluster of the cytochrome b6/f complex were detected here first by mass spectrometry in the mustard plastid proteome. These proteins are most likely not involved in gene expression but co-purify in the column chromatography because of their substrate affinities. This is also true for the group of the miscellaneous proteins including the malate dehydrogenases (MDH) and the malate synthase (MLS), both identified in several spots.

Proteins involved in fatty acid metabolism were identified as well. These include acetyl-coenzyme A carboxylase carboxyl transferase subunit alpha (CAC3) and FabZ, a beta-hydroxyacylacyl carrierprotein (ACP) dehydratase. An earlier study on the purification of the acetyl-CoA carboxylase multienzyme complex also resulted in the enrichment of nucleoid-associated proteins (Phinney and Thelen, 2005) suggesting a potential physical link between these two larger protein associations.

A third protein found (MFP2) is involved in lipid degradation. It was already identified in the HS-fractions in former experiments (Schröter et al., 2010). We found also a cystein synthase and phosphoserine aminotransferase (PSAT) as well as a pyrroline-5-carboxylate reductase (P5CR) known to be essential for amino acid metabolism and a serine hydroxymethyltransferase (SHMT) being essential for photorespiration. The mustard protein in the PC fractions matches to mitochondrial SHMT1 and 2 peptides of several *Brassicales.* The exact affiliation to one of these SHMTs remains unclear since the matching peptides fit to both proteins (**Table 1**). The PC fractions contain also the myrosinase MB3 (involved in glucosinolate degradation) and a cruciferin fitting best to *A. thaliana* CRU3 (**Table 1**). Finally, also actin was detected in one spot, although mustard peptides of PC fractions match to different actin types of different *Brassicales*.

## **DISCUSSION**

## **THE PLASTID NUCLEIC ACIDS BINDING PROTEOME OF MUSTARD**

Goal of our study was the establishment of a purification scheme allowing the estimation of size and composition of the plastid nucleic acids binding sub-proteome from mustard. By using HS and PC chromatography coupled to IF and SDS-PAGE we could reproducibly isolate 1079 protein spots from which we could identify 180 protein spots by mass spectrometry. However, to our surprise these 180 protein spots were found to represent just 58 individual proteins indicating a high degree of posttranslational modification of this specific sub-proteome which in part might be caused by differential phosphorylation (Reiland et al., 2009, 2011). Since we used NaF as phosphatase inhibitor in all preparation steps, the differential phosphorylation states of the analyzed proteins should be well conserved. In contrast, different redox states of thiol groups were not maintained during our purification procedure since reducing agents were included in all steps. Detection of a differential redox state in these fractions will require more specific methods such as redox difference gel electrophoresis (redox-DIGE) (Hurd et al., 2007, 2009). We also observed numerous smaller fragments from several proteins indicating degradation events. These, however, were not random as the spot pattern was reproducible between different preparations suggesting that it is not caused by action of proteases during purification, but by targeted events in the chloroplast. Whether these products represent intermediate steps of protein degradation or whether these fragments perform distinct functions remains to be determined. In summary, this high degree of post-translational modification indicates that the size of the sub-proteome is certainly smaller than the 1079 spots detected. If we assume a similar percentage of individual proteins as within the identified spots (32.2%) for the complete fraction then we estimate 347 proteins for the total nucleic acids binding subproteome. Since we identified a number of co-purifying proteins involved in metabolic processes (29.3%), we had to reduce this number to 236 proteins. However, our mass spectrometry determination has a certain bias since we could detect only the fraction of sufficiently abundant proteins which likely is enriched in metabolic enzymes. In addition, a significant part of posttranslational modification detected in our fractions is focussed on only two proteins, CSP41a and b which partly compromise our estimate. Without these two proteins, we estimate 314 proteins for the chloroplast nucleic acids binding sub-proteome. This appears a reasonable number taking into account the proteins that are already known to be involved in the regulation of plastid gene expression such as NEP, PEP, PAPs, pTACs, PPRs, ribosomal proteins and so on. It, however, leaves still some space for the discovery of as yet unidentified regulators that might appear only in trace amounts such as eukaryotic transcription factors (Wagner and Pfannschmidt, 2006).

## **SPECIFIC FEATURES OF THE PROTEIN FRACTION AFTER PC CHROMATOGRAPHY**

PC chromatography is a well established purification step for nucleic acids binding proteins from chloroplasts (Bottomley et al., 1970; Tiller and Link, 1993). Crucial for the quality of these fractions, however, are a thorough chloroplast preparation *via* sucrose gradient centrifugation and a pre-purification step of the chloroplast lysate using HS chromatography. In comparison to results from earlier work using just HS fractions (Schröter et al., 2010) we observed a high enrichment of translation associated proteins and especially of CSP41 proteins. Co-purification of metabolic enzymes as well as components from other cell compartments was clearly reduced. We obtained a good coverage of the subunits for the plastid RNA polymerase complex PEP; however, surprisingly the larger subunits of this complex were not detectable. We observed a significant reduction of proteins above 80 kDa in size within the PC fractions (**Figure 1**), however, this might be not the reason for the failure of detection since all other components of the complex were identified in the fractions and especially RpoC2 and RpoB are known to bind DNA/RNA. Since these large subunits are highly conserved and have been successfully detected earlier in HS fractions (Steiner et al., 2011) it is likely that they are not well separated on the IEF. Further analyses using additional enrichment methodologies before the IEF step such as size-exclusion chromatography might help to target this problem in the future.

The largest amount of all identified proteins in the PC fractions is dedicated to translational processes with 43% of all proteins (**Figure 3C**). The 50S subunit of plastid ribosomes contains 33 subunits with 31 orthologs to *Escherichia coli* and the two plastid specific subunits PRPL5 and PRPL6 (Yamaguchi and Subramanian, 2000). The 30S subunit is composed of 21 *E. coli* orthologs and four plastid specific proteins with no homologs in other ribosomes (Yamaguchi et al., 2000). Most ribosomal proteins have contact to RNA in various ways, either they are structural components or directly involved in the translational process. Thus, ribosomal proteins contain nucleic acids binding structures which adhere to the used column materials and represent one main component of the nucleic acids binding subproteome of plastids. On the 2D-gels most of them accumulate at the higher pH-ranges and the use of the basic IPG-gels of pH 6–11 led to a good resolution of this group of proteins. The identification of 80S ribosomal proteins in plastid fractions is likely caused by the co-purification of particles attached to the outer chloroplast membrane, like known for tonoplast membrane fragments (Schröter et al., 2010). The main regulation of translation occurs at the level of initiation which is performed by initiation factors (IF). In eukaryotes this process is assured *via* 12 eIFs comprised by 23 polypeptides, whereas in prokaryotes three IFs are sufficient (Kapp and Lorsch, 2004). In plastids orthologs for all bacteria-type translation factors can be found, but the translational complex contains additional proteins not present in bacteria (Beligni et al., 2004). Three of the four IFs identified in this study contain a cTP although only IF2 and IF3 are plastid IFs with a prokaryotic origin. The third one, eIF3f, is a subunit of the eIF3 and is important for the basic cell growth and development and influences the expression of about 3000 genes in *A. thaliana* also in interaction with two other eIF3 subunits (Xia et al., 2010). ChloroP predicts a plastid transit peptide of 40 amino acids for eIF3f of *A. thaliana* and it was previously also identified in fractions enriched in plastid nucleoids (Huang et al., 2013). Thus, it seems to be a true plastid protein and not a co-purification of the cytosolic translational apparatus. However, it might be also possible that this protein possesses a dual localization both in nucleus and plastids contributing to the coordination of gene expression between the two genetic compartments as proposed for other plant cell proteins (Krause and Krupinska, 2009). The elucidation of the precise role of eIF3f in plastids and whether it is involved in the regulation of plastid gene expression will be an interesting field of future research.

The dominant proteins in the PC fractions are the two proteins named CSP41a and CSP41b (Yang et al., 1996; Yang and Stern, 1997). CSP41a and b were also detected in isolates of the PEPcomplex as one of the most abundant component (Pfannschmidt et al., 2000; Suzuki et al., 2004; Schröter et al., 2010) but they appear not to belong to the PAPs but co-purify with these fractions because of the enormous size of their largest conglomerates (Peltier et al., 2006; Schröter et al., 2010; Qi et al., 2012). Here, we identified CSP41a in 8 and CSP41b in 40 spots of diverse sizes and isoelectric points. Thereby, both form a defined spot pattern which was congruent in most replicates of the 2D-gels prepared for this work. This suggests that not only a multimerization of CSP41a/b occurs but maybe also an integration of defined fragment species of the proteins that might be important for specific functions. In addition to targeted fragmentation, the spot pattern after 2D SDS-PAGE suggests also a strong post-translational modification of the two proteins. Indeed, phosphorylation and lysine acetylation have been reported for the corresponding *Arabidopsis* proteins (Reiland et al., 2009, 2011; Finkemeier et al., 2011). The spot pattern as well as the positions of the two proteins in the 2d-gels is highly reminiscent to those recently reported for *Arabidopsis* (Qi et al., 2012). The only difference occurs in the number of identified spots which were 6 Csp41a and 5 Csp41b in *Arabidopsis* while in mustard we observed 3 Csp41a and 7 Csp41b variants (besides the fragmented versions) (**Figure 5**). This suggests the action of at least some species-specific modifications of the proteins.

## **CONCLUSION**

Here, we describe the technical requirements for a highly resolved biochemical purification of several hundreds of protein spots representing the nucleic acids binding sub-proteome of plastids. Our analyses indicate a high enrichment of proteins involved in transcription and translation and, in addition, the presence of massive post-translational modification of this plastid protein sub-fraction. Furthermore, our study provides an extended catalog of plastid proteins from mustard being involved in gene

expression and its regulation and describes a suitable purification strategy for further analysis of low abundant gene expression related proteins.

## **MATERIALS AND METHODS**

## **PLANT GROWTH AND ISOLATION OF PLASTIDS**

Mustard seedlings (*Sinapis alba* L., var. Albatros) were cultivated under permanent white light illumination at 20◦C and 60% humidity. Cotyledons were harvested under the respective light and stored on ice before homogenization in ice-cold isolation buffer in a Waring Blender and filtering through muslin and nylon. Chloroplast isolation by differential centrifugation and sucrose gradient centrifugation in a gradient between 30 and 55% sucrose was conducted as described earlier (Schröter et al., 2010).

## **ISOLATION OF NUCLEIC ACIDS BINDING PROTEINS BY HS- AND PC-CHROMATOGRAPHY**

Lysis of plastids and the chromatography at HS CL-6B was performed according to (Tiller and Link, 1993; Steiner et al., 2009). Proteins were washed, eluted with 1.2 M (NH4)2SO4 and the peak fractions detected via protein quantification assays (RC DC™, Bio-Rad Laboratories, Inc., Hercules, CA, USA) (Schröter et al., 2010). For PC chromatography pooled HS peak fractions were diluted to 10% (v/v) glycerol with dilution buffer [50 mM Tris/HCl, pH 7.6, 0.1 mM EDTA, 0.1% (v/v) TritonX-100, 10 mM sodium fluoride, 62.5 mM (NH4)2SO4, 6 mM 2-mercaptoethanol]. Activation and equilibration of PC (cellulose phosphate ion-exchanger P11, Whatman™ GE healthcare UK Limited, Little Chalfont, UK) to pH 7.6 occurred following the distributor's instructions. Diluted HS proteins were applied to disposable PD-10 columns (Amersham™ GE healthcare UK Limited, Little Chalfont, UK) filled with activated PC, closed carefully and rotated gently for 60 min at 4◦C. After fixing the column on a stand and washing with washing buffer [50 mM Tris/HCl, pH 7.6, 0.1 mM EDTA, 0.1% (v/v) TritonX-100, 10 mM sodium fluoride, 50 mM (NH4)2SO4, 5 mM 2 mercaptoethanol, 10% (v/v) glycerol] proteins were eluted in 3 ml fractions with elution buffer [50 mM Tris/HCl, pH 7.6, 0.1 mM EDTA, 0.1% (v/v) TritonX-100, 10 mM sodium fluoride, 1.2 M (NH4)2SO4, 5 mM 2-mercaptoethanol, 10% (v/v) glycerol] and dialyzed against storage buffer [50 mM Tris/HCl, pH 7.6, 0.1 mM EDTA, 0.1% (v/v) TritonX-100, 10 mM sodium fluoride, 50 mM (NH4)2SO4, 5 mM 2-mercaptoethanol, 50% (v/v) glycerol]. Peak fractions were determined by a protein quantification assay (RC DC™, Bio-Rad Laboratories, Inc., Hercules, CA, USA), pooled and stored at −20◦C.

## **2D GEL ELECTROPHORESIS**

For 2D gel electrophoresis an acetone precipitation of dialyzed proteins from PC chromatography was used to remove the storage buffer following manuals instruction (2-D Electrophoresis principles and methods, 2004, GE healthcare UK Limited, Little Chalfont, UK). For first dimension 18 cm IPG-stripes pH 3–11NL, pH 6–11, and pH3–10 were used (GE Healthcare UK Limited, Little Chalfont, UK). IPG-stripes were rehydrated in rehydration buffer [8 M urea, 0.5% (w/v) chaps, 0.2% (w/v) DTT, 0.5% (v/v) IPG-Buffer, 0.002% (w/v) bromophenol blue] for about 14–16 h. An amount of 400μg precipitated and dried protein per stripe was resolved in rehydration buffer and applicated on the IPG-stripes as cup-loading procedure following manufacturer's instructions (2-D Electrophoresis principles and methods, 2004, GE Healthcare UK Limited, Little Chalfont, UK). For focussing of proteins the following protocol was used on IPGphor (Amersham™ GE Healthcare UK Limited, Buckinghamshire, UK) 6 h step and hold 150 V, 3 h step and hold 300 V, 6 h gradient 1200 V, 3 h gradient 8000 V, 3 h step and hold 8000 V. After IF IPG-strips were equilibrated twice in equilibration solution [50 mM Tris-HCl, ph 8.8, 6 M urea, 30% (v/v) glycerol, 2% (w/v) SDS, 0.002% (w/v) bromophenol blue] first with addition of 2% (w/v) DTT for 15 min under gentle agitation and after removing the first solution second with addition of 2.5% (w/v) iodacetamide (IAA, for alkylation of thiol groups) again for 15 min and gently agitated as described (2-D Electrophoresis principles and methods, 2004, GE Healthcare UK Limited, Little Chalfont, UK). As second dimension a SDS PAGE in gradient gels of 7.5–20% acrylamide with Rhinohide™ gel strengthener (Molecular Probes, Inc., Eugene, OR, USA) was used following manual instructions. Afterwards gels were stained with silver according to manufacturer's instruction (Amersham™ GE Healthcare UK Limited, Buckinghamshire, UK).

## **TRYPTIC DIGEST, LC/ESI-MS/MS AND DATA ANALYSIS**

The spot pattern of the different gels was compared. Matching low abundant spots were pooled (as indicated in **Supplemental Table 1**) to increase the detectable protein amount. Tryptic digest of protein spots was conducted after destaining as referred (Mørtz et al., 1994; Stauber et al., 2003). Mass spectrometry was carried out at LCQ™-DecaXP ion trap mass spectrometer (Thermo Finnigan, San Jose, CA, USA) using a data-dependent scan procedure with four cyclic scan events as described in Schröter et al. (2010). The first cycle, a full MS scan of the mass range m/z 450–1200, was followed by three dependent MS/MS scans of the three most abundant ions. Sample run and data acquisition was performed using the Xcalibur™ software (Version1.3 © Thermo Finnigan 1998–2001). Seventy-six of the low abundant spots were measured at a Finnigan LTQ linear ion trap mass spectrometer (Thermo Finnigan, Thermo Fisher Scientific Inc., Waltham, MA, USA) coupled online after a nano HPLC Ultimate 3000 (Dionex, Thermo Fisher Scientific Inc., Waltham, MA, USA) (Schmidt et al., 2006). After one full MS the instrument was set to measure the collision induced dissociation pattern of the four most abundant ions and exclude the measured once for 10 s from newly measuring.

The resulting spectra were analyzed using the Proteome Discoverer vs. 1.0 (Thermo Fisher Scientific Inc., Waltham, MA, USA) with the implemented Sequest algorithm (Link et al., 1999). Therefore, a database of all RefSeq (reference sequence) sequences of *A. thaliana* and *Arabidopsis lyrata* as well as the complete *Brassica napus* and *Capsella rubella* and the remaining *Brassicales* proteins of NCBI was created [NCBI 2012.03.19 109146 sequences: *Arabidopsis* RefSeq 67924 sequences (35375 *A. thaliana*, 32549 *A. lyrata*) + *B. napus* 10622 sequences + *C. rubella* 4246 sequences + other *brassicales* 26354 sequences]. The Proteome Discoverer Software was set to adjust the Xcorr to reach a false discovery rate of ≤ 1% (Veith et al., 2009). All proteins with at least two unique peptides were taken for further analysis.

For transit peptide prediction the web-based tools TargetP (http://www*.*cbs*.*dtu*.*dk/services/TargetP/) (Emanuelsson et al., 2000) was used and for prediction of the transit peptide length the web-tool ChloroP (http://www*.*cbs*.*dtu*.*dk/services/ ChloroP/) (Emanuelsson et al., 1999). For further analyses identified proteins were grouped into bins according to the modified MapMan system of the plant proteome database (ppdb) (http://ppdb*.*tc*.*cornell*.*edu/dbsearch/mapman*.*aspx) (Sun et al., 2009) based on the MapManBins of Thimm et al. (2004) (**Supplemental Table 2**).

## **ACKNOWLEDGMENT**

This work was supported by the Deutsche Forschungsgemeinschaft (Grants Pf323/4, Mi373/11-1, and Mi373/15-1).

## **SUPPLEMENTARY MATERIAL**

The Supplementary Material for this article can be found online at: http://www.frontiersin.org/journal/10.3389/fpls.2014. 00557/abstract

**Supplemental Figure S1 | Silver stained 2D-gels of the PC fractions with isoelectric focusing for the first dimension in pH gradients between 6–11, 3–11 NL, and 3–10 indicated in the upper left corner of each gel.** The second dimension is performed in a 7.5–20% SDS polyacrylamide gel. Spots are marked and numbered in yellow. Marker sizes and pH range are given right beside and below the gel, respectively.

**Supplemental Table 1 | Identified peptides from the PC fraction.** Spots are listed in numerical order. Accession numbers of proteins belonging to the same spot are listed in an order starting with the highest peptide coverage. Spot nr., identification number of the protein containing spot on the 2D-gels (see **Supplemental Figure S1**). Descriptions of depicted proteins are given as stated in the databases (see Materials and Methods). Coverage, coverage of the depicted proteins by the identified peptides; calc. pI, calculated pI of the depicted proteins based on the protein sequences in the database; MW, calculated molecular weight based on the protein sequences in the databases; z, peptide ion charge;

lower case "m" in the peptide sequence, oxidized form of methionine; lower case "w" oxidized form of tryptophane; lower case "c" cystein with carbamidomethylation; lower case "k" acetylation of lysine.

**Supplemental Table 2 | Detailed characterization of proteins from the PC fraction identified by LC-ESI-MS/MS.** Identified proteins are given in the first column according to the annotation of the respective gene at NCBI. Proteins were sorted according to the MapMan bin numbering in the second column representing classification groups for proteins according to the modified MapMan system of the plant proteome database (ppdb) [(http://ppdb*.*tc*.*cornell*.*edu/dbsearch/mapman*.*aspx) ©Klaas J. van Wijk Lab, Cornell University (Sun et al., 2009) based on the MapManBins of Thimm et al. (2004)]. Spots: number of spots containing the respective protein. Spot nr: identification number of the protein containing spot on the 2D-gels. NCBI accession: gi identification number at NCBI (The National Center for Biotechnology Information http://www*.*ncbi*.*nlm*.*nih*.* gov/); ATG: gene accession of the first matching Arabidopsis thaliana hit or (if no A. thaliana protein was matching) the best matching other organism and the respective A. thaliana gene accession determined by a protein-protein blast at NCBI in brackets; cTP: probability of a plastid transit peptide; MW [kDa]: calculated theoretical molecular weight in kilodalton; MW(-cTP) [kDa]: calculated theoretical molecular weight without chloroplast transit peptide (cTP) in kilodalton; MW(-lTP) [kDa]: calculated theoretical molecular weight without cTP and luminal transit peptide (lTP) in kilodalton; PI: calculated isoelectric point; PI (-cTP): calculated isoelectric point without cTP; PI (-lTP): calculated isoelectric point without cTP and lTP. The last six parameter were provided at the related accession entry by ppdb (http://ppdb*.*tc*.*cornell*.*edu/dbsearch/ searchacc*.*aspx).

## **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: 11 July 2014; accepted: 29 September 2014; published online: 29 October 2014.*

*Citation: Schröter Y, Steiner S, Weisheit W, Mittag M and Pfannschmidt T (2014) A purification strategy for analysis of the DNA/RNA-associated sub-proteome from chloroplasts of mustard cotyledons. Front. Plant Sci. 5:557. doi: 10.3389/fpls. 2014.00557*

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

*Copyright © 2014 Schröter, Steiner, Weisheit, Mittag and Pfannschmidt. 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.*