Research ArticleResearch Article

A Conserved, Mg2+-Dependent Exonuclease Degrades Organelle DNA during Arabidopsis Pollen Development

Ryo Matsushima, Lay Yin Tang, Lingang Zhang, Hiroshi Yamada, David Twell, Wataru Sakamoto

Published April 2011. DOI: https://doi.org/10.1105/tpc.111.084012

Abstract

In plant cells, mitochondria and plastids contain their own genomes derived from the ancestral bacteria endosymbiont. Despite their limited genetic capacity, these multicopy organelle genomes account for a substantial fraction of total cellular DNA, raising the question of whether organelle DNA quantity is controlled spatially or temporally. In this study, we genetically dissected the organelle DNA decrease in pollen, a phenomenon that appears to be common in most angiosperm species. By staining mature pollen grains with fluorescent DNA dye, we screened Arabidopsis thaliana for mutants in which extrachromosomal DNAs had accumulated. Such a recessive mutant, termed defective in pollen organelle DNA degradation1 (dpd1), showing elevated levels of DNAs in both plastids and mitochondria, was isolated and characterized. DPD1 encodes a protein belonging to the exonuclease family, whose homologs appear to be found in angiosperms. Indeed, DPD1 has Mg2+-dependent exonuclease activity when expressed as a fusion protein and when assayed in vitro and is highly active in developing pollen. Consistent with the dpd phenotype, DPD1 is dual-targeted to plastids and mitochondria. Therefore, we provide evidence of active organelle DNA degradation in the angiosperm male gametophyte, primarily independent of maternal inheritance; the biological function of organellar DNA degradation in pollen is currently unclear.

INTRODUCTION

Mitochondria and plastids originate from the endosymbiosis of rickettsia-like α-proteobacteria and cyanobacteria-like photosynthetic bacteria, respectively (Gray et al., 1999; Dyall et al., 2004; Keeling, 2010). Most genes in the primitive endosymbionts were transferred to the plant nuclear genome, yet both organelles retain remnant genomes and carry out DNA replication, transcription, and translation (Gray, 1999; Kleine et al., 2009). For example, coordinated expression of nuclear and organelle genes is a central subject in the studies of chloroplast biogenesis.

Despite its smaller genome size and limited genetic information, organelle DNA sometimes account for a substantial amount of total DNA because it is present as multiple copies (for review, see Sakamoto et al., 2008). For example, total DNAs from leaf tissues in higher plants often contain >20% of plastid DNAs (ptDNAs) (Bennet and Smith, 1976; Lamppa and Bendich, 1979; Arabidopsis Genome Initiative, 2000; Rauwolf et al., 2010). The copy number of ptDNA appears to correlate with nuclear ploidy and appears to vary among species or even among different tissues and during developmental stages (Herrmann and Kowallik, 1970; Kowallik and Herrmann, 1972; Lamppa and Bendich, 1979; Scott and Possingham, 1980; Kuroiwa et al., 1981; Boffey and Leech, 1982; Tymms et al., 1983). Multiplication of plastids by division during leaf development further complicates the ptDNA amount per organelle. Such a complex polyploid nature of the plastid genome (also of mitochondrial genome) has raised the question of whether organelle DNA levels are controlled spatially or temporally. Although several proteins are known to play roles in maintaining the configuration of plant organelle genomes (Abdelnoor et al., 2003; Edmondson et al., 2005; Zaegel et al., 2006; Shedge et al., 2007; Maréchal et al., 2008, 2009; Rowan et al., 2010), very little is understood about DNA degradation at the molecular level.

ptDNAs (and also mitochondrial DNAs [mtDNAs]) are cytologically detectable as nucleoids by staining tissues with DNA fluorescent dye, such as 4′,6-diamidino-2-phenylindole (DAPI) or SYBR green I (SYBR) (Kuroiwa, 1991, 2010). The ptDNAs exist as a complex with proteins that constitute plastid nucleoids (Sato et al., 1998, 2001, 2003; Murakami et al., 2000; Jeong et al., 2003). The cytological detection of organelle DNAs is frequently used to estimate DNA levels together with DNA gel blot hybridization, quantitative PCR, and colorimetric detection of DNA hydrolysates. These methods enable us to investigate the number, morphologies, and behavior of plastid nucleoids during leaf development (Rauwolf et al., 2010 and references therein). In Arabidopsis thaliana, cytological observation demonstrated that amounts of ptDNA increase more than 10-fold during leaf development (Fujie et al., 1994). By contrast, contradictory results were reported for ptDNA levels in mature leaves: constant or declining ptDNA amounts have been reported by different laboratories (Rowan et al., 2004, 2009; Li et al., 2006; Zoschke et al., 2007). Studies of numbers of mtDNAs per cell have revealed that mtDNA levels also fluctuate during leaf development. More importantly, some mitochondria might lack a complete genome (Preuten et al., 2010). A decrease in mtDNAs has also been reported during pollen development (Wang et al., 2010). These circumstantial observations prompted us to study organelle DNA degradation by a forward genetic approach.

Here, we specifically examine organelle DNA levels in mature pollen of Arabidopsis. Much attention has been given to organelle DNA levels in male reproductive organs because the decrease of organelle DNAs in male tissues is suggested to correlate with their maternal inheritance: ptDNAs that are detectable by DAPI stains in male germ cells are often associated with biparental inheritance of ptDNAs (Hagemann and Schrödoer, 1989; Nagata et al., 1999; Birky, 2001; Hagemann, 2004; Kuroiwa, 2010). Irrespective of the inheritance mode, however, we noticed that organelle DNAs are cytologically absent in pollen vegetative cells (Matsushima et al., 2008a; Sakamoto et al., 2008). A survey of numerous mature pollen grains using DAPI staining has revealed that almost all angiosperm species lack cytologically detectable organelle DNAs in pollen vegetative cells (Corriveau and Coleman, 1988; Mogensen, 1996; Zhang et al., 2003; Wang et al., 2010). It is noteworthy that pollen vegetative cells do not contribute to fertilization, but they do contain numerous plastids and mitochondria, which might be necessary for a pollen tube to germinate, elongate, and deliver sperm cells into the embryo sac. Given that the lack of DAPI signals in pollen vegetative cells is so clear and consistent in many species (irrespective of inheritance mode), we reasoned that organelle DNA levels are strictly downregulated by a dominant mechanism. We also exploited the relative ease in examining organelle DNAs in mature pollen, rather than in leaf tissues, which require time-consuming sectioning or protoplast isolation.

For this study, we performed extensive forward genetic analysis and isolated Arabidopsis mutants in which organelle DNA was retained in mature pollen grains. Characterization of the gene responsible for the mutants led us to identify a nuclease that is expressed preferentially during pollen development. It is particularly interesting that this DNase is localized in both the plastid and mitochondria, providing evidence for the existence of an organelle nuclease in eukaryotes. Our data reveal an active mechanism of organelle DNA degradation in a tissue-specific manner, which is primarily independent of maternal inheritance.

RESULTS

Isolation of Arabidopsis Mutants Defective in Organelle DNA Degradation of Pollen Grains

Our cytological analysis of pollen grains in Arabidopsis revealed that when mature pollen was stained with DAPI, almost no signals corresponding to organelle DNAs were detected (Figures 1 and 2; see also Sakamoto et al., 2008). Numerous plastids and mitochondria exist in pollen vegetative and sperm cells. Therefore, we inferred that organelle DNAs decrease during pollen development and that it is feasible to screen mature pollen for mutants that exhibit altered levels of organelle DNAs. This strategy presents the additional advantage that the pollen phenotype will segregate in M1 pollen and thus can be found by screening mature pollen grains from M1 flowers (Chen and McCormick, 1996). In our screening method, pollen grains collected from M1 or M2 flowers were fixed briefly with glutaraldehyde with subsequent gentle squashing over a cover slip and DAPI staining, which allowed careful observation of DAPI signals (Figure 1A). Mature pollen grains of ~2000 individual ethyl methanesulfonate–mutagenized M1 and 2000 M2 Arabidopsis plants were screened using this method. As a consequence, we isolated five mutant lines that exhibited unusual DAPI signals within the cytoplasm of vegetative cells. These DAPI signals were distinct from those corresponding to vegetative and sperm nuclei and rather resembled organelle DNAs (Figure 1B). Mutants isolated in M1 population were further characterized to obtain M2 individuals where all pollen showed the phenotype. These mutants were designated as defective in pollen organelle DNA degradation (dpd). This dpd phenotype was characterized genetically in the subsequent generations. Overall, our screening and genetic analysis identified two recessive mutations, dpd1 and dpd2; this work specifically examined dpd1. All mutants except for dpd2 were the alleles of dpd1 (designated as dpd1-1 to dpd1-4; see below). The recessive nature of dpd mutations implied a dominant mechanism to decrease organelle DNAs in mature pollen, as we expected.

Figure 1.
Figure 1.

Experimental Strategy to Isolate Mutants Defective in Organelle DNA Degradation in Arabidopsis Pollen.

(A) Schematic representation of the experiment. Arabidopsis wild-type Col were mutagenized using ethyl methanesulfonate. Mature pollen grains from M1 or M2 plants were screened using the pollen squash method (middle) for easier observation of DAPI signals derived from organelle DNAs. Examples of DAPI-stained pollen grains before and after the squash are shown on the right. Bars = 20 μm.

(B) Isolation of dpd mutants. DAPI-stained squashed pollen from Col (wild type), dpd1, and dpd2 are shown.

Figure 2.
Figure 2.

Observation of Organelle DNAs in Developing Pollen and Leaf Mesophyll Cells of Col and dpd1-1.

(A) Schematic representation of pollen development in Arabidopsis.

(B) DAPI-stained Technovit sections of bicellular and tricellular pollens. Strong signals are indicated by arrows in tricellular pollen and sperm nuclei; other signals in dpd1 correspond to extrachromosomal DNAs. Bars = 5 μm.

(C) Electron micrographs of mature pollen grains from wild-type ecotype Col and dpd1-1. Bars = 2 μm.

(D) DAPI-stained mesophyll protoplasts from Col and dpd1-1. Protoplasts were gently squashed to detect DAPI signals within chloroplasts. Bars = 20 μm.

Organelle DNAs Do Not Decrease during dpd1 Pollen Development

To observe organelle DNA levels more carefully, we prepared Technovit-embedded thin sections (0.5 μm) of developing pollen grains from wild-type Columbia (Col) and dpd1 and stained them with DAPI. In Arabidopsis, mature pollen is formed after two characteristic mitoses (Figure 2A) (Borg et al., 2009). Asymmetric division of uninucleate microspores at pollen mitosis I produces bicellular pollen comprising a vegetative cell and a germ cell. Subsequent division of the germ cell at pollen mitosis II (PMII) produces a pair of sperm cells forming tricellular pollen. Our thin section analysis showed that, in Col, cytoplasmic DAPI signals start to decrease in the late bicellular stage. They disappear completely at the tricellular stage in Col, indicating that the decrease (or degradation) of organelle DNA normally occurs during PMII (Figure 2B). By contrast, dpd1 showed strong cytoplasmic DAPI signals in pollen, which were retained even at the tricellular stage. Segregation of the dpd phenotype in the F2 population from a cross between dpd1 and Col revealed that dpd1 behaved as a single recessive trait and that the dpd phenotype showed complete penetrance in the selfed progeny (see Supplemental Tables 1 and 2 online). No ultrastructural abnormality was observed in dpd1 organelles (Figure 2C), suggesting that the membrane integrity of both mitochondria and plastids is maintained. Examination of in vitro–germinated pollen by DAPI also revealed that organelle DNAs were retained even after germination in dpd1 (see Supplemental Figure 1 online). These results demonstrated that dpd1 appears to compromise DNA reduction during pollen development.

Vegetative and Reproductive Growth of dpd1

We next examined whether dpd1 displays any visible phenotypes not only in reproductive growth but also in vegetative growth. None of the dpd1 alleles showed differences in their vegetative growth under normal conditions (Figure 3A), suggesting that DPD1 primarily affects organelle DNA levels in pollen grains but not those in other tissues. To test this possibility, we examined organelle DNA levels by staining protoplasts derived from 6-week-old mature leaves with DAPI. Results showed that, unlike mature pollen grains, no apparent difference in DAPI-detectable organelle DNAs was detected in chloroplasts of leaf cells (Figure 2D). These results imply that, in dpd1, organelle DNA levels are increased in pollen grains but not in other somatic tissues. Subsequently, visible phenotypes in dpd1 male reproductive organs were also examined. Visual inspection showed that flowers and seed set appeared normal in dpd1 (Figures 3B to 3D). The dpd phenotype was completely penetrant in dpd1 selfed progeny and segregated normally in the F2 (see Supplemental Tables 1 and 2 online). Mature pollen grains in dpd1 exhibited morphology and viability (Alexander stain, Figure 3E; see Supplemental Table 3 online) that were indistinguishable from those of the wild type. No difference in pollen size was found between the wild type and dpd1 (see Supplemental Table 4 online). Collectively, we inferred that the dpd phenotype does not significantly influence overall plant growth and pollen morphology.

Figure 3.
Figure 3.

Plant Architecture and Reproductive Organs of dpd1 Mutants.

(A) Five-week-old dpd1-1. Bar = 1 cm.

(B) Immature seeds from wild-type Col and dpd1-1. Bars = 1 mm.

(C) Floral organs from Col and dpd1-1. Bars = 2 mm.

(D) Single flower from Col and dpd1-1. Bars = 2 mm.

(E) Single anther from Col and dpd1-1 stained with Alexander solution. Bars = 100 μm.

Both ptDNA and mtDNA Are Retained in dpd1 Pollen

The dpd phenotype in dpd1 raised the question of which DNA-containing organelles––plastids, mitochondria, or both––emitted the DAPI signals in pollen vegetative cells. To address this question, we visualized plastids and mitochondria in mature pollen by expressing organelle-targeted fluorescent proteins (green fluorescent protein in plastids [ptGFP] and red fluorescent protein in mitochondria [mtRFP]; Figure 4). We reported previously that these organelle-targeted GFP/RFPs, when expressed under the control of a vegetative cell-specific promoter LAT52 from tomato (Solanum lycopersicum; Twell et al., 1990), can show plastids or mitochondria in pollen vegetative cells (Matsushima et al., 2008b; Tang et al., 2009). We introduced the corresponding transgenes (Lat52pro:PTS:GFP and Lat52pro:MTS:RFP) into dpd1. Mature pollen grains from these transgenic lines were examined using DAPI or SYBR stain (depending on the fluorescent protein used) and simultaneous detection of fluorescent proteins. The results demonstrated that DAPI signals in dpd1 (Lat52pro:PTS:GFP) colocalized with ptGFP. Similarly, SYBR signals in dpd1 (Lat52pro:MTS:RFP) colocalized with mtRFP (Figure 4). These results indicate that the extrachromosomal DNA signals in dpd1 mutants were derived from both plastids and mitochondria.

Figure 4.
Figure 4.

Colocalization of DNA Signals with Plastids and Mitochondria in dpd1 Pollen.

A transgene that expresses plastid-targeted GFP or mitochondria-targeted RFP in pollen vegetative cells (presented on the left) was introduced into dpd1-1. Colocalization of plastidial GFP signals with DAPI signals (top panels) or of mitochondrial RFP signals with SYBR (bottom panels) was examined. Arrowheads indicate colocalization of fluorescent organelles and DAPI-stained or SYBR-stained signals. The asterisk denotes nuclear-derived DAPI signals. Bars = 5 μm.

Both ptDNA and mtDNA Increased in Pollen but Not in Somatic Cells

To confirm that both ptDNA and mtDNA levels were altered in pollen but not in somatic tissues of dpd1, we performed a PCR-based assay. Total DNAs were prepared from pollen and young seedlings and assayed by quantitative real-time PCR. Levels of ptDNAs (psbA) or mtDNAs (cox1) in these tissues of Col and dpd1 were normalized based on the level of nuclear DNAs (18S rDNA; see Methods and Rowan et al., 2009). As expected, ptDNAs and mtDNAs had significantly increased in dpd1-1 compared with Col (n = 3, Welch’s t test, P = 0.0041 and 0.0226, respectively) (Figures 5A and 5B). A large difference in mtDNA levels was detected, probably because of an extremely low level of mtDNA in Arabidopsis wild-type pollen grains (Wang et al., 2010). By contrast, young seedlings showed no significant differences between Col and dpd1-1 in the level of ptDNA and mtDNA (P = 0.2887 and 0.2692, respectively) (Figures 5C and 5D). Together, these results verified our cytological analysis of dpd1 pollen stained with DAPI. We concluded that both ptDNA and mtDNA levels are increased in dpd1 pollen.

Figure 5.
Figure 5.

Quantitative Analysis of Organelle DNA Levels in Pollen Grains and Young Seedlings.

Levels of organelle DNAs per nuclear DNAs in Col and dpd1-1 were determined using real-time PCR. Mean values of normalized organelle DNA levels of Col were determined as 1.0. The relative values of dpd1-1 were calculated (n = 3, sd as error bars).

(A) ptDNA in pollen grains.

(B) mtDNA in pollen grains.

(C) ptDNA in 17-d-old seedlings.

(D) mtDNA in 17-d-old seedlings.

Map-Based Cloning of the DPD1 Locus

We identified the DPD1 gene based on conventional map-based cloning. We mapped the dpd1-1 mutation within a 623-kb region on chromosome 5 (summarized in Supplemental Figure 2 online). A survey of genes that potentially encode proteins targeted to plastids and/or mitochondria allowed us to select possible genes responsible for the dpd1 phenotype. Subsequent sequencing of the candidate genes identified a base change in the At5g26940 gene, which results in an amino acid substitution (Figure 6A). Sequencing of this gene in three other dpd1 alleles (dpd1-2, dpd1-3, and dpd1-4; Figure 6B) revealed that all these alleles had base changes in At5g26940 (Figure 6). The base changes in dpd1-1, dpd1-2, and dpd1-3 caused amino acid substitutions (see Supplemental Figure 3 online), whereas dpd1-4 had a base change close to the border of intron1/exon2. Furthermore, we obtained two T-DNA insertion mutants (dpd1-5 and dpd1-6) in At5g26940 and examined whether they showed the dpd1 phenotype. As expected, they all exhibited the dpd phenotype that was indistinguishable from other dpd1 alleles.

Figure 6.
Figure 6.

Identification of the DPD1 Gene.

(A) Schematic representation of At5g26940 gene (DPD1) and the predicted protein encoded. Gray boxes show coding regions. Adenine of the translation start codon (ATG) is designated as +1. Arrows indicate positions of mutations in dpd1 alleles. Here, dpd1-1, dpd1-2, and dpd1-3 are located within the coding region; dpd1-4 is located at the splicing acceptor site of the first intron. In addition, dpd1-5 and dpd1-6 are T-DNA–inserted alleles. Red boxes (ExoI, ExoII, and ExoIII) are functional domains conserved among 3′-5′ exonucleases. a.a., amino acids.

(B) DAPI-stained squashed pollen grains in dpd1 mutant alleles other than dpd1-1. A part of the squashed pollen cytoplasm is shown. Bars = 10 μm.

(C) A phylogenetic tree of DPD1 and its homologs. Multiple alignment was performed using ClustalW as described in Methods (see Supplemental Figure 7 and Supplemental Data Set 1 online).

To prove that At5g26940 encodes DPD1, a genomic fragment encompassing At5g26540 was transformed into the dpd1-1 mutant, we generated five transgenic lines. One of the lines was homozygous for the transformed sequence. It fully complemented the dpd phenotype (Figure 7A) when mature pollen grains were examined using our squash method. Moreover, we conducted a PCR-based assay to verify the altered organelle DNA levels in dpd1 and the complemented line. PCR products corresponding to plastid and mitochondria DNAs were more abundant in dpd1-1 pollen than in Col and the complemented line. Collectively, we concluded that At5g26940 encodes DPD1.

Figure 7.
Figure 7.

Complementation of the dpd1 Phenotype.

(A) Squashed mature pollen stained with SYBR from dpd1, Col, and dpd1 complemented with At5g26940 genomic sequence (Comp.). A trace of the picture from dpd1 is shown at the top right. VN, vegetative nucleus; SN, sperm nucleus; PC, pollen coat; Ext, extrachromosomal DNAs. Bars = 10 μm.

(B) Comparison of organelle DNAs in Col, dpd1-1, and dpd1-1+At5g26940 pollen grains. Band intensities were quantified; each value is shown as the average of three biological repeats ± sd. Different letters denote significant differences between samples (P = 0.05, Tukey–Kramer’s HSD).

[See online article for color version of this figure.]

DPD1 Encodes a Protein Belonging to the Exonuclease Family and Dual Targeted to Plastids and Mitochondria

DPD1 is present in a single copy in Arabidopsis, where it encodes a protein (316 amino acids) belonging to the exonuclease family (Pfam: PF00929, ExonulX-T), which is included in the large ribonuclease H-like superfamily (Clan CL0219). Three domains included within the protein family members, ExoI, ExoII, and ExoIIIε, appeared to be conserved in DPD1 and other members (see Supplemental Figure 3 online). A BLAST search and systematic analysis of DPD1 homologs using the entire DPD1 amino acids and the SALAD Database (http://salad.dna.affrc.go.jp/salad/en/) revealed that DPD1 homologs are present in small green algae (Ostreococcus and Micromonas), moss (Physcomitrella), and higher plants (Figure 6C). By contrast, no such homolog was detectable in a green alga (Chlamydomonas), a red alga (Cyanidioschyzon), or a fungus (Saccharomyces). Although proteins detected as related to DPD1 in the green algae and moss had conserved exonuclease domains, they were much larger (>1260 amino acids) and apparently had additional DNA helicase domains, suggesting a role that is distinct from organelle DNA degradation. These results imply that DPD1 had evolved with the appearance of anisogamous male reproductive organs in the plant lineage.

It was tempting to speculate that DPD1 is targeted to both plastids and mitochondria. To examine the cellular localization of DPD1, we transiently expressed DPD1-GFP in Arabidopsis protoplasts prepared from mesophyll cells, as described previously (Miura et al., 2007). The DPD1-GFP fusion protein colocalized not only with chlorophyll autofluorescence, but also with the mitochondria-specific dye Mitotracker Red (see Supplemental Figure 4 online). These results indicate that DPD1 is localized in plastids and mitochondria, each of which shows defective organelle DNA degradation in dpd1 mutants.

DPD1 Is a Mg2+-Dependent Exonuclease

Sequence information and the dpd1 phenotype together strongly suggest that DPD1 has exonuclease activity (Figure 6A). To study this possibility in vitro, a recombinant DPD1-His protein (6 × histidine tagged at C terminus) was generated in Escherichia coli and affinity purified as described in Methods. In addition to the wild-type DPD1-His gene, we expressed two DPD1-His genes in which a point mutant was introduced (Figure 8A). Because of a nonsense mutation at the 186th amino acid, DPD1-His(Y186*) has DPD1 that is C-terminally truncated and lacks His. Similarly, DPD1-His(A236V) has an amino acid change equivalent to dpd1-1 (Figure 6A; see Supplemental Figure 3 online). A majority of these DPD1 proteins expressed in E. coli were detected as aggregated, although we purified DPD1-His and DPD1-His(A236V) from soluble fractions (Figures 8B to 8D). Purified DPD-His proteins were detected as two bands: The lower band was likely a degradation product. Actually, DPD1-His(Y186*) was not purified because of the absence of the His tag, but we used this fraction as a negative control.

Figure 8.
Figure 8.

DPD1-His Fusion Protein Has Exonuclease Activity in Vitro.

(A) Schematic view of the constructions employed in DPD1-His expression in E. coli. In order: bacteriophage T7 promoter, DPD1 cDNA, and His-tag sequence. Relative positions of the point mutations are indicated by vertical arrows (numbers denote corresponding amino acids).

(B) Expression of DPD1-His in E. coli. SDS-PAGE of cell lysate with (+) or without (−) isopropyl β-d-1-thiogalactopyranoside is shown along with molecular mass markers. Asterisks denote the bands corresponding to the fusion proteins.

(C) SDS-PAGE of DPD1-His protein purified by HiTrap Chelating HP column. TP, total soluble protein fraction; P, purified fraction. Bands corresponding to DPD1-His are indicated by an arrow.

(D) Immunoblot analysis of DPD1-His. Same protein fractions in (C) were probed with anti-His antibody.

(E) Nuclease assay of recombinant DPD1-His proteins. Effects of heat treatment and EDTA were tested simultaneously as negative controls.

(F) Requirements of bivalent cations: magnesium (Mg2+), copper (Ca2+), manganese (Mn2+), and zinc (Zn2+) were tested.

(G) Digestion of supercoiled and linearized plasmids with DPD1-His. Plasmid pGreen0229 (4454 bp), circular or linearized by digestion with restriction enzymes as indicated (EcoRI, EcoRV, and KpnI) was subjected to DPD1-His nuclear assay. Open and closed arrowheads indicate positions of linearized and supercoiled plasmids, respectively.

Data in (A) to (C) are representative of three independent experiments.

[See online article for color version of this figure.]

Nuclease activity of these recombinant proteins was assayed in a standard buffer with magnesium bivalent cation as a cofactor and DNA fragments (PCR-amplified ptDNA). The result showed that DNAs were degraded by DPD-His during the first 20 min of incubation (Figure 8E). By contrast, neither heat-denatured DPD1-His, DPD1-His(A236V), nor the negative control fraction degraded PCR fragments, even after 60 min. Also, EDTA inhibited nuclease activity of DPD1-His, indicating that magnesium is a necessary cofactor. By contrast, none of three other bivalent cations (copper, manganese, or zinc) was observed to act as a cofactor for DPD1 DNA degradation (Figure 8F). These data indicate that DPD1-His has magnesium-dependent nuclease activity. Complete degradation of PCR fragments indicates that DPD1-His has exonucleolytic activity, but its endonucleolytic activity was not ruled out completely. To test this possibility, we performed the same nuclease assays with circular plasmids. Intact plasmids were stable even after 60-min incubation with DPD1-His. However, once nicked or linearized by restriction enzymes (EcoRI, EcoRV, and KpnI), the plasmids became completely degradable by DPD1-His (Figure 8G). These data reflect that DPD1 is an exonuclease that requires accessible ends to degrade double-stranded DNAs.

Pollen-Enhanced Expression of DPD1

We next performed RT-PCR analysis to examine tissue-specific expression of DPD1 (Figure 9A). RNAs were prepared from seedlings, the shoot apex, flowers, and various stages of pollen development; RT-PCR was conducted as described previously (Honys and Twell, 2004). As expected, DPD1 was highly expressed in flowers compared with whole seedlings and the shoot apex. Further dissection of pollen development at the stage of unicellular microspore, bicellular pollen, tricellular pollen, and mature pollen grains revealed that DPD1 starts to express at the bicellular pollen stage. It reaches a peak at the tricellular pollen stage. This temporal expression of DPD1 in pollen coincides perfectly with the disappearance of DAPI-stained organelle DNAs (Figure 2B). It also coincides with the fact that organelle DNA levels in vegetative tissues are not altered in dpd1. To further characterize DPD1 expression in vivo, transgenic Arabidopsis plants were generated that expressed the previously described DPD1-GFP fusion under the DPD1 promoter. Characterization of these transgenic plants showed that the DPD1 promoter is highly active in mature pollen grains (Figures 9B and 9C). Furthermore, we crossed this transgenic plant with another line harboring Lat52pro:MTS:RFP, thereby expressing mtRFP in vegetative cells. Careful observation of this line revealed smaller GFP signals colocalized with mtRFP and additional larger signals corresponding to plastids (Figure 9D). This observation confirmed the results of our transient assay in mesophyll cells (see Supplemental Figure 4 online) that DPD1 is dual-targeted to both organelles. When characterized by confocal microscopy, we occasionally found smaller GFP mitochondrial signals that did not merge with mtRFP in a hollow area that likely corresponds to sperm cells (Figure 9E). The size and distribution of these small GFP signals resembled those observed in sperm mitochondria (Matsushima et al., 2008b), indicating that DPD1 is expressed not only in vegetative but also in sperm cells. Collectively, these results suggest that DPD1 is predominantly expressed in the male gametophyte.

Figure 9.
Figure 9.

Pollen-Specific Expression of DPD1.

(A) RT-PCR analysis to determine tissues and developmental stages expressing DPD1 gene. UNM, uninucleate microspore; BCP, bicellular pollen; TCP, immature tricellular pollen; MPG, mature pollen grain. Histone variant H3.3 gene (At4g40040) was used as a control.

(B) and (C) In vivo expression analysis of DPD1. DPD1 promoter was fused to the DPD1-GFP fusion protein used in Supplemental Figure 4 online. Transgenic lines expressing the transgene (DPD1pro-DPD1-GFP) were subjected to detection of GFP in flowers (B) and pollen grains (C).

(D) Detailed observation of DPD1-GFP in mature pollen. Pollen grains from a transgenic plant expressing mtRFP in the vegetative cell and DPD1pro-DPD1-GFP were examined using confocal microscopy. Signals corresponding to DPD1-GFP and mtRFP are shown along with the merged and differential interference contrast (DIC) images.

(E) A confocal section showing small GFP signals that do not merge with mtRFP in a hollow area (indicated by the arrow).

Bars = 1 mm in (B), 20 μm in (C), and 5 μm in (D) and (E).

Pollen Viability and Transmission Efficiency in dpd1

Our results thus far indicated that DPD1 is a pollen-specific exonuclease responsible for organelle DNA degradation. Supporting this assumption, dpd1 showed no detectable phenotype in plant vegetative growth (Figure 3). To characterize the effect of organelle DNA degradation in pollen vegetative cells, we further examined dpd1 pollen viability. We first examined the transmission efficiency (TE) of the dpd1 mutation. A heterozygous DPD1/dpd1-1 plant was subjected to a reciprocal cross with Col. The genotype of F1 seeds was determined using PCR-based genotyping, as described in Methods, to estimate TE of dpd1 in the next generation. This result indicated that TE of dpd1 through the male (TEmale) was not reduced to a statistically significant degree (Table 1). We also performed conditional TE tests to examine whether TEmale is affected by the position of seeds in fertilized siliques. After a reciprocal cross between Col and DPD1/dpd1-1, fertilized siliques were cut into upper and lower halves. Then, the genotype for DPD1 was determined separately (Table 2). Again, these results showed no significant difference in TEmale. We subsequently examined pollen germination ability in vitro. Despite the unaffected TE in dpd1, we found that the germination rate is slightly lower in dpd1-1 than in Col and the complemented line (Table 3). This reduction, which is likely to be too slight to affect pollen fertilization to a considerable degree, was the only phenotype we observed consistently in dpd1. By contrast, pollen tube growth and delivery into the embryo sac appeared to proceed normally in both Col and dpd1 (see Supplemental Figure 5 online). Based on these careful investigations, we concluded that organelle DNA degradation controlled by DPD1 in pollen vegetative cells has very limited influence on pollen viability, although we cannot completely rule out the possibility that it affects pollen grain germination.

View this table:
Table 1.

Genetic Transmission Analysis of dpd1-1 Mutations

View this table:
Table 2.

Conditional Genetic Transmission Analysis of dpd1-1 Mutations

View this table:
Table 3.

In Vitro Germination Rate of Wild-Type and dpd1-1 Pollen Grains

Inheritance Mode of Organelle DNAs in dpd1

Next, we raised the question of whether dpd1 exhibits altered organelle DNA inheritance. We therefore designed the following genetic analysis in which several other ecotypes were employed along with dpd1 to validate organelle DNAs transmitted from the male or female parent. First, we tested ptDNAs. When crossed with ecotype Cape Verde Islands (Cvi), with polymorphic ptDNA (Martínez et al., 1997), dpd1 (Col background) showed normal maternal inheritance of the plastid genome, as did the wild type (n = 65; see Supplemental Figure 6 online). Plastids have not been detected inside Arabidopsis sperm cells (Tang et al., 2009; Wang et al., 2010). Therefore, exclusion of plastids from the male gamete is perhaps the dominant mechanism for plastid maternal inheritance in Arabidopsis.

We subsequently investigated the mode of mitochondrial inheritance in dpd1. This experiment required careful investigation because the Arabidopsis nuclear genome contains sequences identical to mtDNA integrated in the pericentromeric region of chromosome 2 (Stupar et al., 2001). Therefore, we were compelled to identify a rare single-nucleotide polymorphism that is specific to mtDNAs, based on the available sequence data (Unseld et al., 1997). Our search identified one polymorphism between Col and C24 (Figure 10A). Based on this polymorphism, we created a degenerate cleaved amplified polymorphic sequence (dCAPS) marker. Our PCR analysis revealed that this polymorphism (C to A in C24 matR gene) was specific for the mitochondrial genome but not for the matR sequence integrated in chromosome 2 (Figures 10B and 10D). To test the paternal leakage of mtDNAs in dpd1, a tester dpd1 line that had a C24-derived cytoplasm was generated and crossed to Col female (Figures 10C and 10D). The result showed that no paternal transmission of mitochondria occurs in our experimental scale (n = 300). We conclude that DPD1 is independent of the inheritance mode of organelle DNA, at least under our experimental scale, highlighting the turnover of extrachromosomal DNAs as a unique biological process rather than a mechanism for maternal inheritance.

Figure 10.
Figure 10.

Genetic Analysis of mtDNA Transmission Confirming a Lack of Paternal mtDNA Transmission in dpd1.

(A) A rare single-nucleotide polymorphism identified between Col and C24 ecotypes of matR gene (highlighted in red). A dCAPS marker was generated to distinguish C or A polymorphism, which was detectable by digesting the PCR fragment with ScaI.

(B) Presence of the matR polymorphisms in different ecotypes and dpd1. Because of the presence of mtDNA sequence in the Arabidopsis nuclear genome, corresponding polymorphism was shown for nuclear DNA (n) and mtDNA (mt).

(C) A genetic strategy to test paternal leakage of mtDNA in dpd1. Because dpd1 is in the Col background, a cross was made between C24 (female) and dpd1 (male). The resulting F2 individual recessive for dpd1 (highlighted green) with mitochondria derived from C24 [dpd1(C24)] was used as a tester line. Three hundred F1 plants from a cross between Col (female) and the tester line were subjected to dCAPS analysis; no paternal leakage was detected.

(D) Examples of dCAPS analysis detecting A or C polymorphism. In crosses between Col and C24, the band corresponding to A polymorphism is only detected when C24 is a maternal parent.

DISCUSSION

Identification of Organelle Nuclease and Its Evolutional Implication

We conducted a genetic study of organelle DNA decrease of pollen grains. Isolation of recessive dpd mutations, which retained more organelle DNAs than Col, implies that some genes control organelle DNA degradation in the male gametophyte. Molecular cloning enabled us to identify DPD1, which resides in both plastids and mitochondria and has exonuclease activity in vitro. Enhanced expression in male reproductive organs, particularly at PMII, coincided perfectly with our cytological observations. Collectively, we concluded that DPD1 is a DNase controlling organelle DNA levels in a tissue-specific manner (in pollen development and particularly at PMII). The decline of organelle DNAs in pollen vegetative cells has been reported consistently, not only in Arabidopsis but in almost all angiosperm species (Mogensen, 1996; Nagata et al., 1999). Our data and circumstantial evidence strongly imply that organelle DNA levels decrease in pollen vegetative cells through a conserved mechanism that involves DPD1. Our expression analysis further implies that the DPD1 promoter is also active in sperm cells of Arabidopsis. Species showing biparental inheritance of ptDNAs (e.g., Medicago truncatula; Matsushima et al., 2008a) are known to have strong DAPI signals in male germ cells. In such species, altered DPD1 expression in male germ cells might account for biparental inheritance.

A database search for DPD1 homologs revealed several interesting aspects of how DPD1 emerged during evolution. First, a DPD1 homolog is not found in cyanobacteria, suggesting that DPD1 is not of endosymbiotic origin. Second, it is not found in lower eukaryotes, including fungi and green alga. Finally, a closely related homolog is not found in moss but is found specifically in angiosperm species. Based on these observations, we consider that DPD1 evolved along with the appearance of anisogamous male reproductive organs (i.e., male and female gametes that are not identical in morphology and the male gamete is smaller and contains vegetative cells). The exonuclease family to which DPD1 belongs includes various proteins from prokaryotes to eukaryotes. Examples include DnaQ protein in E. coli as a proofreading subunit of the DNA polymerase III holoenzyme (Scheuermann and Echols, 1984) and TREX1 that degrades retroelements and exogenous DNAs in mammalian cells (Lehtinen et al., 2008). It can be assumed that one of these nuclease members evolved into DPD1 to degrade organelle DNAs, along with the emergence of pollen vegetative cells in angiosperms. Conserved presence of DPD1 homologs in species with anisogamy is consistent with our result that DPD1 expression is limited predominantly to pollen. Collectively, these data demonstrate that organelle DNA degradation is common in angiosperm pollen and that it is controlled by the exonuclease DPD1.

Regulation of DPD1 Expression, Degradation, and Relevance to General Organelle DNA Degradation

In addition to the circumstances described previously, it is notable that most of the dpd mutants we isolated through our extensive screening were dpd1 alleles. We therefore consider that DPD1 plays a central role in DNA degradation in both plastids and mitochondria. Apparently, several questions arose from our findings. We first asked about whether the role of DPD1 in organelle DNA degradation can be generalized in other somatic tissues, such as those of leaves. This is apparently not the case because of (1) a lack of detectable phenotype in dpd1 vegetative growth, (2) pollen-specific expression of DPD1, and (3) a lack of detectable change of organelle DNA levels in dpd1 leaf tissues. Moreover, DPD1 does not appear in plastid proteome data prepared from nonpollen tissues (e.g., The Plant Protein Database, http://ppdb.tc.cornell.edu/). Actually, DPD1 has never been associated with plastid nucleoids (or transcriptionally active chromosomes) derived from nonpollen tissues (Pfalz et al., 2006). Based on these observations, we infer that DPD1 functions specifically in pollen and does not participate in general organelle DNA metabolism, including DNA replication and/or degradation. However, it is possible that misregulation of DPD1 expression affects organelle DNA levels in somatic cells. Additional investigations are necessary to elucidate the function of DPD1 in organelle degradation in tissues other than pollen.

We next examined the question of whether DPD1 acts on DNA degradation by itself or requires additional factor(s). Given its exonucleolytic activity, it is presumed that linear DNAs are the substrate of DPD1, but circular DNAs are not. Although predominant forms of both ptDNAs and mtDNAs are believed to consist of circular DNAs, nicked or linear molecules have been found in both plastid and mitochondria (Bendich, 2004; Oldenburg and Bendich, 2004); those molecules can be DPD1 substrates. It is therefore possible that DPD1 alone can degrade some, but not all, of the organelle DNA population. A significant difference in organelle DNA levels between Col and dpd1 pollen (Figure 5) implies that most of the organelle DNA population might in fact be nicked or linear molecules in pollen vegetative cells (Bendich, 2004). Alternatively, DPD1 degrades organelle DNAs cooperatively with an unidentified endonuclease. In either case, DPD1 nuclease requires magnesium as a cofactor. In a green alga, Chlamydomonas (showing isogamy with identical male and female gametes), sex-specific degradation of chloroplast genomes putatively correlates with a Ca2+-dependent nuclease (Nishimura et al., 1999, 2002). A Mg2+-dependent property distinguishes DPD1 from this Chlamydomonas nuclease. Again, our results obtained in this study are consistent with our presumption that DPD1 had emerged in angiosperms.

An important remaining question related to organelle DNA quantity in pollen is whether plastids and mitochondria that contain no DNAs exist. We have not performed detailed measurements of the organelle DNA quantity. Therefore, this question remains to be resolved in this study. Our work specifically examined the existence of a dual-targeted organelle DNase rather than the degree of DNA degradation. It is noteworthy, however, that several reports have described that the copy numbers of organelle genomes were less than the organelle numbers. For example, Wang et al. (2010) recently characterized mtDNA levels in both somatic and gametic tissues using quantitative PCR. Their results showed that in both tissues, mtDNA copy numbers were insufficient for each mitochondrion to have a complete copy of the genome. Preuten et al. (2010) also showed, based on quantitative PCR analysis, that mtDNA copy numbers are insufficient in various stages of leaf tissues. Accordingly, frequent fusion of mitochondria has been proposed to account for such genome insufficiency (Arimura et al., 2004; Sheahan et al., 2005). Consistent with our current study, Wang et al. (2010) revealed that mtDNA levels are much lower in pollen than in mesophyll cells. Based on these observations, we consider that mitochondria in pollen vegetative cells are functional even with low mtDNA levels to support pollen tube elongation and fertilization. Assessment of precise genome copy numbers in pollen vegetative cells, particularly for plastid genomes, awaits further study, which might require a new methodology.

Maternal Inheritance and Possible Role of Organelle DNA Degradation in Pollen Function

Because DPD1 is predominantly expressed in pollen, we carefully examined pollen viability in dpd1. The loss of organelle DNA degradation in dpd1 does not significantly affect pollen viability, although dpd1 showed lower TEmale than the wild type did. A difference was also observed in the pollen germination rate in vitro. Nevertheless, this difference was too little to influence TE to any great degree. Therefore, our results leave open the question of the relevance of DPD1 to pollen germination or other pollen functions. This work identified the molecule that governs the organelle DNA decrease in pollen through the dpd phenotype: upregulated organelle DNAs in pollen vegetative cells. To unravel the physiological significance of organelle DNA degradation, future analysis of dpd1 phenotypes is necessary under various conditions. Given that mature pollen grains are the only tissue that is physically isolated from parental plant bodies, it is presumed that organelle DNA can be salvaged for nucleotide recycling.

As described in the Introduction, the decline of organelle DNAs in pollen development has been documented in many species, particularly in relation to uniparental inheritance of organelle genomes. The cytological detection of ptDNAs in generative or sperm cells is well correlated with species showing biparental inheritance, suggesting that DNA quantity in male germs is relevant to organelle inheritance (Corriveau and Coleman, 1988; Mogensen, 1996; Nagata et al., 1999; Zhang et al., 2003). These observations raise the possibility that a DNase such as DPD1 excludes extra organelle DNAs in male germ cells and assures their uniparental inheritance. In Arabidopsis, no plastids are found in sperm cells (Tang et al., 2009; Wang et al., 2010), and DPD1 is unlikely to affect maternal ptDNA inheritance. Therefore, plastid maternal inheritance is determined primarily by the exclusion of plastids in germ cells rather than ptDNA amounts in Arabidopsis (Martínez et al., 1997; Azhagiri and Maliga, 2007). In fact, our F1 analysis between Cvi and dpd1 showed that DPD1 did not affect plastid inheritance (see Supplemental Figure 6 online).

In contrast with plastids, there are ~10 mitochondria in sperm cells that can be transmitted into fertilized egg and central cells (Matsushima et al., 2008b; Wang et al., 2010). The overwhelming number of mitochondria in egg cells (~800) appears to account for stochastic propagation of maternal mitochondria (Birky, 2001). In fact, results of our genetic analyses suggest that DPD1 has no effect on the inheritance mode of mtDNAs and ptDNAs (Figure 10). Again, we must emphasize that a large survey revealed that nonfertilizing pollen vegetative cells lack cytologically detectable organelle DNAs in many species, irrespective of the inheritance mode of plastids and mitochondria. We therefore consider that organelle DNA degradation by DPD1 is a priori independent of maternal inheritance in Arabidopsis. Given the common function of DPD1 at PMII, it is possible that the amplification of organelle DNAs, rather than their degradation, in generative cells is important for their inheritance mode. This possibility is in fact implied by Nagata et al. (1999). We propose that the control of organelle DNA degradation by DPD1 evolved with the formation of the anisogamous angiosperm male gametophyte and that it is primarily independent of organelle DNA inheritance in Arabidopsis. Future studies of DPD1 homologs in species showing biparental plastid inheritance can help us understand the role of organelle DNA degradation in pollen.

METHODS

Plant Materials, Mutant Screening, and Mapping

Arabidopsis thaliana ecotypes Col and Nossen were used as wild-type plant materials for mutagenesis. The T-DNA insertion alleles in DPD1 (dpd1-5 and dpd1-6) were obtained from the ABRC (Salk_091621 and Salk_015164, respectively). For detecting the inheritance mode of ptDNA and mtDNA, ecotypes Cvi and C24 were used, respectively. The tester line for mtDNA inheritance was generated by crossing dpd1-1 or dpd1-6 (Col background) to C24 female (Figure 10). For assessing paternal mtDNA leakage, pollen from this tester line was used to cross with Col female. Transgenic plants expressing ptGFP and mtRFP in pollen vegetative cells have been described in our previous works (Matsushima et al., 2008b; Tang et al., 2009).

For mutagenesis, wild-type seeds were mutagenized by soaking them initially for 16 h in 0.1 or 0.2% (v/v) methanesulfonic acid ethyl ester (Sigma-Aldrich). Approximately 2000 flowers from 857 M1 lines were collected, and pollen grains were examined using fluorescence microscopy. In addition, ~2000 M2 individual plants were used for screening pollen grains. Mature pollen grains were placed on a glass slide and immersed in a drop of deionized water that had been supplemented with 3% (w/v) glutaraldehyde and 1 to 10 μg mL−1 DAPI (Invitrogen) in TAN buffer (20 mM Tris-HCl, pH 7.65, 0.5 mM EDTA, 7 mM 2-mercaptoethanol, 0.4 mM phenylmethyl sulfonyl fluoride, and 1.2 mM spermidine). In some cases, 1:1000 diluted SYBR Green I (Invitrogen) was used also, instead of DAPI, especially for detecting mtDNAs. Pollen grains were squashed by putting gentle pressure on a cover slip that had been placed on the glass slide. They were examined using a fluorescence microscope (BX51; Olympus) and a confocal laser scanning microscope (FV1000; Olympus).

To determine the map position of the DPD1 locus, dpd1-1 and the wild-type ecotype Landsberg erecta were crossed. Pollen of F2 progenies was stained with DAPI to determine their genotype. Genomic DNAs from F2 progenies were isolated and analyzed using simple sequence length polymorphism markers with data obtained from The Arabidopsis Information Resource (http://www.Arabidopsis.org).

Thin Sections of Technovit 7100 Resin of Pollen Grains

Pollen grains were fixed in 2.5% (v/v) glutaraldehyde and 1% (w/v) paraformaldehyde in cacodylate buffer, pH 7.4, for at least 24 h at room temperature. Samples were subsequently dehydrated through a graded ethanol series (20% [v/v], 40%, 60%, 80%, and 100%) and then embedded in resin (Technovit 7100; Heraeus Kulzer). The embedded samples were cut into 0.5-μm sections using an ultramicrotome (Ultracut N; Reichert-Nissei) and diamond knives. Then they were dried on cover slips. Thin sections were stained with 1 μg mL−1 DAPI. To prevent fading, 1 mg mL−1 n-propyl gallate in 50% (v/v) glycerol was added to the samples before fluorescence microscopic examination.

For transmission electron microscopy, pollen grains were fixed in 4% glutaraldehyde and 5% paraformaldehyde in cacodylate buffer, pH 7.4, for at least 24 h at room temperature. A second fixation was performed in 5% (w/v) potassium permanganate solution at room temperature for 20 min. After rinsing in distilled water, the fixed pollen grains were dehydrated through a graded ethanol series (20%, 40%, 60%, 80%, and 100%) and embedded in Spurr’s resin (Polysciences). Ultrathin (70 to 90 nm) sections were stained in 1% (w/v) uranyl acetate and 0.5% (w/v) lead citrate and examined using an electron microscope (H-7100; Hitachi) operating at 75 kV.

Plasmid Construction and Nuclease Assay

To construct pG002926940 for complementation of dpd1, a DNA fragment containing the At5g26940 genomic sequence was amplified using PCR with primers: 5′-GTTunderline>GGTACC/underline>TTGTAGCTCTGTTTTGGCCTA-3′ (KpnI site underlined) and 5′-GCAunderline>GAGCTC/underline>ATGATGTTCCCTTATAATTAG-3′ (SacI site underlined). The fragment was cloned into the KpnI and SacI sites of pGreen0029. To construct p3526940TP55GFP, a DNA fragment corresponding to the putative transit peptide and the C-terminal 15 amino acids was amplified using PCR with primers: 5′-GAGunderline>CTCGAG/underline>ATGTGTATCTCAATCTCG-3′ (XhoI site underlined) and 5′-ACCTTGunderline>CATGGGAGACC/underline>ACACGTTACGTCT-3′ (BsaI site underlined). The fragment was digested with XhoI and BsaI and cloned into the SalI and NcoI sites of p35S-sGFP, as described previously (Sakamoto et al., 2003).

To construct pTopoCT26940w/o40 for expressing recombinant DPD1-His protein in Escherichia coli, a DPD1 cDNA containing the entire reading frame, except for the region corresponding to transit peptide (N-terminal 40 amino acids), was amplified by PCR using full-length DPD1 cDNA (U82439; ABRC) as a template with primers 5′-ATGGCTTCTTCTGTTGATGGTAAAGCA-3′ and 5′-GGCCTTCTTGTTCTTGGCCATGGC-3′. Mutagenesis of pTopoCT26940w/o40 to express DPD1-His(Y186*) and DPD1-His(A236V) was conducted using a QuickChange Multi site-directed mutagenesis kit (Stratagene).

To purify DPD1-His, pTopoCT26940w/o40 was transformed into E. coli strain BL21 (Invitrogen). Protein induction was performed with 0.8 mM isopropyl β-D-1-thiogalactopyranoside in 300 mL of culture. DPD1-His was purified using HiTrap Chelating HP (GE Healthcare) according to the manufacturer’s instructions. Anti-His antibody kit (Qiagen) was used to detect DPD1-His by immunoblotting. Purified DPD1-His was made imidazole-free and concentrated using a centrifugal filter (Centricon YM-10; Millipore). The protein concentration was determined using a kit (Bio-Rad protein assay; Bio-Rad Laboratories). The PCR fragments derived from ptDNA (1658 bp) were amplified using the following primers: 5′-GCTTCAGCGGCTGCAATTGCTAT-3′ and 5′-GCTTGTGAAGTATGTGTTCGAG-3′. The nuclease assay reaction (100 μL) consisted of 40 mM Tris-HCl, pH 7.5, 2 mM MgCl2, and 1 μg DNA substrate; purified DPD1-His (2.5 μg protein) was finally added to initiate the reaction. For inhibition analyses, DPD1-His were incubated at 98°C for 5 min prior to nuclease assay, or reactions were conducted with 10 mM EDTA. Reactions were completed by immediately adding stopping buffer (1% [w/v] SDS, 50% [v/v] glycerol, and 0.05% [w/v] bromophenol blue) and were subjected to 1% (w/v) agarose gel electrophoresis. After staining with ethidium bromide, remaining undigested PCR fragments were image captured and quantified using software (ImageJ; NIH).

Nucleic Acid Extraction and PCR Analysis

To prepare total DNAs, mature pollen grains were collected by rubbing a silicon bar (~2 mm width and 2 cm length) onto dehiscent anthers from seven flowers and by putting the bar into an Eppendorf tube containing 45 μL of distilled water. The pollen suspension was incubated at 95°C for 5 min and centrifuged for 5 min at 16,000g. Then, 9 μL of the supernatant was subjected to the PCR. Total DNAs from seedlings (17 d old) were isolated as described in a previous report (Miura et al., 2007).

For real-time PCR, the following primers were used according to Rowan et al. (2009): psbA (ptDNA), 5′-AGAGACGCGAAAGCGAAAG-3′ and 5′-CTGGAGGAGCAGCAATGAA-3′; cox1 (mtDNA), 5′-CCACGCATGTTGAAGATAGTTG-3′ and 5′-AGTAGGTAGCGGCACTGGGT-3′; and 18S rRNA (nuclear DNA), 5′-AAACGGCTACCACATCCAAG-3′ and 5′-ACTCGAAAGAGCCCGGTATT-3′. Amplification was conducted using THUNDERBIRD SYBR qPCR Mix kit (TOYOBO) and Light Cycler 2.0 (Roche Diagnostics), with 50 cycles of a denaturation at 95°C for 5 s and an extension at 60°C for 30 s. LightCycler Software (version 4.0; Roche Diagnostics) was used to quantify PCR reactions. The amount of organelle DNA was normalized by the value for 18S rRNA (n = 3).

To evaluate organelle DNA contents in mature dpd1 and wild-type pollen (Figure 7B), extracts from five pollen grains were subjected to PCR. Primers used to amplify ptDNA (region corresponding to ndhG) were 5′-GGCCCCCACATAAATAAGGAGTTG-3′ and 5′-TCACCTCAAACAAAAAATGGGGTAAA-3′. Primers used to amplify mtDNA (region corresponding to nad9) were 5′-ATGGAAAGATCGGAACATGGGAAT-3′ and 5′-GGGTCATCTCAATGGGTTCAGAA-3′. Both primer sets were designed according to the Arabidopsis chloroplast and mitochondrial genome sequences (AP000423 and NC_001284). For detecting a single nucleotide polymorphism between Col and C24 in the mitochondrial matR gene, a dCAPS primers were generated as follows: 5′-GTCAAGGCTGCCACTCGGTCCTAAGACG-3′ and 5′-CAACTCCTACGAGTCGTCCGGCGGAAAG-3′. The polymorphism (C or A at nucleotide +297) was estimated by ScaI digestion of the PCR fragment (Figure 10A).

Total RNAs were purified from isolated spores at four developmental stages, as described in an earlier study (Honys and Twell, 2004). The cDNA synthesis from 750 ng DNase-treated total RNA was primed with oligo(dT) in a 21-μL reaction using a SuperScript first-strand synthesis system for RT-PCR (Invitrogen). Histone variant H3.3 (At4g40040) was used as a control as described (Brownfield et al., 2009). The RT-PCR primers used for DPD1 were 5′-GCATCGGAAAAATGAGCGGAT-3′ and 5′-CACCCTCCCTTGTAAGACTATAG-3′ and for histone-H3 5′-AGCTCCCTTTCCAGAGGCTA-3′ and 5′-TCCAAGTCTCCTACACCCAAA-3′.

Characterization of Pollen and Other Phenotypes

For the pollen viability test, anthers were collected from stage 13 Arabidopsis flowers and stained with Alexander stain for at least 2 h. In vitro germination of pollen was performed as described previously (Matsushima et al., 2008b). For pollen tube guidance, pistils were fixed in 10% (v/v) acetic acid in ethanol overnight, incubated in 1 n NaOH overnight, washed three times with 50 mM potassium phosphate buffer, pH 7.5, and stained for at least 5 min in 0.05% (w/v) aniline blue. Pistils were supplemented with 50% (v/v) glycerol and squashed lightly under a cover slip. For observing organelle DNAs in mesophyll cells, protoplasts were prepared from mature leaves (7 weeks old) and stained with DAPI as described previously (Kato et al., 2007).

Phylogenetic Analysis

Multiple alignment was performed using ClustalW (gap open penalty, 10; gap extension penalty, 0.05; selected weight matrix, BLOSUM) and manually adjusted to optimize alignment (shown in Supplemental Data Set 1 online). The tree was generated as unrooted using the neighbor-joining method. The confidence of nodes in the tree was supported by the value from 1000 bootstrap replicates.

Accession Numbers

Sequence data from this article can be found in the Arabidopsis Genome Initiative or GenBank/EMBL databases under the following accession numbers: Arabidopsis DPD1, At5g26940; Zea mays, ACG42491; Oryza sativa, Os4g0623400; Ostreococcus tauri, CAL55462; Micromonas sp RCC299. XP002507456; Physcomitrella patens, XP001752327; Populus trichocarpa, XP002330239; Sorghum bicolor, XP002468000; Vitis vinihera, XP002282861; and Ricinus communis, XP002512483.

Supplemental Data

The following materials are available in the online version of this article.

Acknowledgments

We thank Sodmergen for assisting us with electron microscopy and for useful discussion. We also thank the ABRC for providing the T-DNA mutant lines, Mizuki Takenaka for providing information related to Arabidopsis mtDNA polymorphism, and Rie Hijiya, Yumiko Kaji, Nami Sakurai-Ozato, Chieko Hattori, and Said Hafidh for their technical assistance. This work was supported by Grants-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science, and Technology (No. 16085207 and No. 22112516 to W.S.) and by the Oohara Foundation (to W.S.).

Footnotes

  • The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) is: Wataru Sakamoto (saka{at}rib.okayama-u.ac.jp).

  • www.plantcell.org/cgi/doi/10.1105/tpc.111.084012

  • ↵1 These authors contributed equally to this work.

  • ↵[C] Some figures in this article are displayed in color online but in black and white in the print edition.

  • ↵[W] Online version contains Web-only data.

  • Received February 2, 2011.
  • Revised April 1, 2011.
  • Accepted April 11, 2011.
  • Published April 26, 2011.

References

  1. ↵
    1. Abdelnoor R.V.,
    2. Yule R.,
    3. Elo A.,
    4. Christensen A.C.,
    5. Meyer-Gauen G.,
    6. Mackenzie S.A.
    (2003). Substoichiometric shifting in the plant mitochondrial genome is influenced by a gene homologous to MutS. Proc. Natl. Acad. Sci. USA 100: 5968–5973.
    OpenUrlAbstract/FREE Full Text
  2. ↵
    Arabidopsis Genome Initiative (2000). Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408: 796–815.
    OpenUrlCrossRefPubMed
  3. ↵
    1. Arimura S.,
    2. Yamamoto J.,
    3. Aida G.P.,
    4. Nakazono M.,
    5. Tsutsumi N.
    (2004). Frequent fusion and fission of plant mitochondria with unequal nucleoid distribution. Proc. Natl. Acad. Sci. USA 101: 7805–7808.
    OpenUrlAbstract/FREE Full Text
  4. ↵
    1. Azhagiri A.K.,
    2. Maliga P.
    (2007). Exceptional paternal inheritance of plastids in Arabidopsis suggests that low-frequency leakage of plastids via pollen may be universal in plants. Plant J. 52: 817–823.
    OpenUrlCrossRefPubMed
  5. ↵
    1. Bendich A.J.
    (2004). Circular chloroplast chromosomes: The grand illusion. Plant Cell 16: 1661–1666.
    OpenUrlFREE Full Text
  6. ↵
    1. Bennet M.D.,
    2. Smith J.B.
    (1976). Nuclear DNA amounts in angiosperms. Philos. Trans. R. Soc. Lond. B Biol. Sci. 274: 222–274.
    OpenUrl
  7. ↵
    1. Birky C.W. Jr.
    (2001). The inheritance of genes in mitochondria and chloroplasts: Laws, mechanisms, and models. Annu. Rev. Genet. 35: 125–148.
    OpenUrlCrossRefPubMed
  8. ↵
    1. Boffey S.A.,
    2. Leech R.M.
    (1982). Chloroplast DNA levels and the control of chloroplast division in light-grown wheat leaves. Plant Physiol. 69: 1387–1391.
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Borg M.,
    2. Brownfield L.,
    3. Twell D.
    (2009). Male gametophyte development: A molecular perspective. J. Exp. Bot. 60: 1465–1478.
    OpenUrlAbstract/FREE Full Text
  10. ↵
    1. Brownfield L.,
    2. Hafidh S.,
    3. Durbarry A.,
    4. Khatab H.,
    5. Sidorova A.,
    6. Doerner P.,
    7. Twell D.
    (2009). Arabidopsis DUO POLLEN3 is a key regulator of male germline development and embryogenesis. Plant Cell 21: 1940–1956.
    OpenUrlAbstract/FREE Full Text
  11. ↵
    1. Chen Y.C.,
    2. McCormick S.
    (1996). sidecar pollen, an Arabidopsis thaliana male gametophytic mutant with aberrant cell divisions during pollen development. Development 122: 3243–3253.
    OpenUrlAbstract
  12. ↵
    1. Corriveau J.L.,
    2. Coleman A.W.
    (1988). Rapid screening method to detect potential biparental inheritance of plastid DNA and results for over 200 angiosperm species. Am. J. Bot. 75: 1443–1458.
    OpenUrlCrossRef
  13. ↵
    1. Dyall S.D.,
    2. Brown M.T.,
    3. Johnson P.J.
    (2004). Ancient invasions: From endosymbionts to organelles. Science 304: 253–257.
    OpenUrlAbstract/FREE Full Text
  14. ↵
    1. Edmondson A.C.,
    2. Song D.,
    3. Alvarez L.A.,
    4. Wall M.K.,
    5. Almond D.,
    6. McClellan D.A.,
    7. Maxwell A.,
    8. Nielsen B.L.
    (2005). Characterization of a mitochondrially targeted single-stranded DNA-binding protein in Arabidopsis thaliana. Mol. Genet. Genomics 273: 115–122.
    OpenUrlCrossRefPubMed
  15. ↵
    1. Fujie M.,
    2. Kuroiwa H.,
    3. Kawano S.,
    4. Mutoh S.,
    5. Kuroiwa T.
    (1994). Behavior of oeganelles and their nucleoids in the shoot apical meristem during leaf development in Arabidopsis thaliana L. Planta 194: 395–405.
    OpenUrl
  16. ↵
    1. Gray M.W.
    (1999). Evolution of organellar genomes. Curr. Opin. Genet. Dev. 9: 678–687.
    OpenUrlCrossRefPubMed
  17. ↵
    1. Gray M.W.,
    2. Burger G.,
    3. Lang B.F.
    (1999). Mitochondrial evolution. Science 283: 1476–1481.
    OpenUrlAbstract/FREE Full Text
  18. ↵
    1. Daniell H.,
    2. Chase C.
    1. Hagemann R.
    (2004). The sexual inheritance of plant organelles. In Molecular Biology and Biotechnology of Plant Organelles, Daniell H., Chase C., eds (Dortrecht, The Netherlands: Springer), pp. 93–113.
  19. ↵
    1. Hagemann R.,
    2. Schrödoer M.-B.
    (1989). The cytological basis of the plastid inheritance in angiosperms. Protoplasma 152: 57–64.
    OpenUrlCrossRef
  20. ↵
    1. Herrmann R.G.,
    2. Kowallik K.V.
    (1970). Multiple amounts of DNA related to the size of chloroplasts. II. Comparison of electron-microscopic and autoradiographic data. Protoplasma 69: 365–372.
    OpenUrlCrossRefPubMed
  21. ↵
    1. Honys D.,
    2. Twell D.
    (2004). Transcriptome analysis of haploid male gametophyte development in Arabidopsis. Genome Biol. 5: R85.
    OpenUrlCrossRefPubMed
  22. ↵
    1. Jeong S.Y.,
    2. Rose A.,
    3. Meier I.
    (2003). MFP1 is a thylakoid-associated, nucleoid-binding protein with a coiled-coil structure. Nucleic Acids Res. 31: 5175–5185.
    OpenUrlAbstract/FREE Full Text
  23. ↵
    1. Kato Y.,
    2. Miura E.,
    3. Matsushima R.,
    4. Sakamoto W.
    (2007). White leaf sectors in yellow variegated2 are formed by viable cells with undifferentiated plastids. Plant Physiol. 144: 952–960.
    OpenUrlAbstract/FREE Full Text
  24. ↵
    1. Keeling P.J.
    (2010). The endosymbiotic origin, diversification and fate of plastids. Philos. Trans. R. Soc. Lond. B Biol. Sci. 365: 729–748.
    OpenUrlAbstract/FREE Full Text
  25. ↵
    1. Kleine T.,
    2. Maier U.G.,
    3. Leister D.
    (2009). DNA transfer from organelles to the nucleus: The idiosyncratic genetics of endosymbiosis. Annu. Rev. Plant Biol. 60: 115–138.
    OpenUrlCrossRefPubMed
  26. ↵
    1. Kowallik K.V.,
    2. Herrmann R.G.
    (1972). Variable amounts of DNA related to the size of chloroplasts. IV. Three-dimensional arrangement of DNA in fully differentiated chloroplasts of Beta vulgaris L. J. Cell Sci. 11: 357–377.
    OpenUrlAbstract/FREE Full Text
  27. ↵
    1. Kuroiwa T.
    (1991). The replication, differentiation, and inheritance of plastids with emphasis on the concept of organelle nuclei. Int. Rev. Cytol. 128: 1–62.
    OpenUrlCrossRef
  28. ↵
    1. Kuroiwa T.
    (2010). Review of cytological studies on cellular and molecular mechanisms of uniparental (maternal or paternal) inheritance of plastid and mitochondrial genomes induced by active digestion of organelle nuclei (nucleoids). J. Plant Res. 123: 207–230.
    OpenUrlCrossRefPubMed
  29. ↵
    1. Kuroiwa T.,
    2. Suzuki T.,
    3. Ogawa K.,
    4. Kawano S.
    (1981). The chloroplast nucleus: Distribution, number, size, and shape, and a model for the multiplication of the chloroplast genome during chloroplast development. Plant Cell Physiol. 22: 381–396.
    OpenUrlAbstract/FREE Full Text
  30. ↵
    1. Lamppa G.K.,
    2. Bendich A.J.
    (1979). Changes in chloroplast DNA levels during development of pea (Pisum sativum). Plant Physiol. 64: 126–130.
    OpenUrlAbstract/FREE Full Text
  31. ↵
    1. Lehtinen D.A.,
    2. Harvey S.,
    3. Mulcahy M.J.,
    4. Hollis T.,
    5. Perrino F.W.
    (2008). The TREX1 double-stranded DNA degradation activity is defective in dominant mutations associated with autoimmune disease. J. Biol. Chem. 283: 31649–31656.
    OpenUrlAbstract/FREE Full Text
  32. ↵
    1. Li W.,
    2. Ruf S.,
    3. Bock R.
    (2006). Constancy of organellar genome copy numbers during leaf development and senescence in higher plants. Mol. Genet. Genomics 275: 185–192.
    OpenUrlCrossRefPubMed
  33. ↵
    1. Maréchal A.,
    2. Parent J.S.,
    3. Sabar M.,
    4. Véronneau-Lafortune F.,
    5. Abou-Rached C.,
    6. Brisson N.
    (2008). Overexpression of mtDNA-associated AtWhy2 compromises mitochondrial function. BMC Plant Biol. 8: 42.
    OpenUrlCrossRefPubMed
  34. ↵
    1. Maréchal A.,
    2. Parent J.S.,
    3. Véronneau-Lafortune F.,
    4. Joyeux A.,
    5. Lang B.F.,
    6. Brisson N.
    (2009). Whirly proteins maintain plastid genome stability in Arabidopsis. Proc. Natl. Acad. Sci. USA 106: 14693–14698.
    OpenUrlAbstract/FREE Full Text
  35. ↵
    1. Martínez P.,
    2. López C.,
    3. Roldán M.,
    4. Sabater B.,
    5. Martín M.
    (1997). Plastid DNA of five ecotypes of Arabidopsis thaliana: Sequence of ndhG gene and maternal inheritance. Plant Sci. 123: 113–122.
    OpenUrlCrossRef
  36. ↵
    1. Matsushima R.,
    2. Hamamura Y.,
    3. Higashiyama T.,
    4. Arimura S.,
    5. Sodmergen,
    6. Tsutsumi N.,
    7. Sakamoto W.
    (2008b). Mitochondrial dynamics in plant male gametophyte visualized by fluorescent live imaging. Plant Cell Physiol. 49: 1074–1083.
    OpenUrlAbstract/FREE Full Text
  37. ↵
    1. Matsushima R.,
    2. Hu Y.,
    3. Toyoda K.,
    4. Sodmergen,
    5. Sakamoto W.
    (2008a). The model plant Medicago truncatula exhibits biparental plastid inheritance. Plant Cell Physiol. 49: 81–91.
    OpenUrlAbstract/FREE Full Text
  38. ↵
    1. Miura E.,
    2. Kato Y.,
    3. Matsushima R.,
    4. Albrecht V.,
    5. Laalami S.,
    6. Sakamoto W.
    (2007). The balance between protein synthesis and degradation in chloroplasts determines leaf variegation in Arabidopsis yellow variegated mutants. Plant Cell 19: 1313–1328.
    OpenUrlAbstract/FREE Full Text
  39. ↵
    1. Mogensen H.L.
    (1996). The hows and whys of cytoplasmic inheritance in seed plants. Am. J. Bot. 83: 383–404.
    OpenUrlCrossRef
  40. ↵
    1. Murakami S.,
    2. Kondo Y.,
    3. Nakano T.,
    4. Sato F.
    (2000). Protease activity of CND41, a chloroplast nucleoid DNA-binding protein, isolated from cultured tobacco cells. FEBS Lett. 468: 15–18.
    OpenUrlCrossRefPubMed
  41. ↵
    1. Nagata N.,
    2. Saito C.,
    3. Sakai A.,
    4. Kuroiwa H.,
    5. Kuroiwa T.
    (1999). The selective increase or decrease of organellar DNA in generative cells just after pollen mitosis one controls cytoplasmic inheritance. Planta 209: 53–65.
    OpenUrlCrossRefPubMed
  42. ↵
    1. Nishimura Y.,
    2. Misumi O.,
    3. Kato K.,
    4. Inada N.,
    5. Higashiyama T.,
    6. Momoyama Y.,
    7. Kuroiwa T.
    (2002). An mt(+) gamete-specific nuclease that targets mt(-) chloroplasts during sexual reproduction in C. reinhardtii. Genes Dev. 16: 1116–1128.
    OpenUrlAbstract/FREE Full Text
  43. ↵
    1. Nishimura Y.,
    2. Misumi O.,
    3. Matsunaga S.,
    4. Higashiyama T.,
    5. Yokota A.,
    6. Kuroiwa T.
    (1999). The active digestion of uniparental chloroplast DNA in a single zygote of Chlamydomonas reinhardtii is revealed by using the optical tweezer. Proc. Natl. Acad. Sci. USA 96: 12577–12582.
    OpenUrlAbstract/FREE Full Text
  44. ↵
    1. Oldenburg D.J.,
    2. Bendich A.J.
    (2004). Most chloroplast DNA of maize seedlings in linear molecules with defined ends and branched forms. J. Mol. Biol. 335: 953–970.
    OpenUrlCrossRefPubMed
  45. ↵
    1. Pfalz J.,
    2. Liere K.,
    3. Kandlbinder A.,
    4. Dietz K.J.,
    5. Oelmüller R.
    (2006). pTAC2, -6, and -12 are components of the transcriptionally active plastid chromosome that are required for plastid gene expression. Plant Cell 18: 176–197.
    OpenUrlAbstract/FREE Full Text
  46. ↵
    1. Preuten T.,
    2. Cincu E.,
    3. Fuchs J.,
    4. Zoschke R.,
    5. Liere K.,
    6. Börner T.
    (2010). Fewer genes than organelles: Extremely low and variable gene copy numbers in mitochondria of somatic plant cells. Plant J. 64: 948–959.
    OpenUrlCrossRefPubMed
  47. ↵
    1. Rauwolf U.,
    2. Golczyk H.,
    3. Greiner S.,
    4. Herrmann R.G.
    (2010). Variable amounts of DNA related to the size of chloroplasts III. Biochemical determinations of DNA amounts per organelle. Mol. Genet. Genomics 283: 35–47.
    OpenUrlCrossRefPubMed
  48. ↵
    1. Rowan B.A.,
    2. Oldenburg D.J.,
    3. Bendich A.J.
    (2004). The demise of chloroplast DNA in Arabidopsis. Curr. Genet. 46: 176–181.
    OpenUrlCrossRefPubMed
  49. ↵
    1. Rowan B.A.,
    2. Oldenburg D.J.,
    3. Bendich A.J.
    (2009). A multiple-method approach reveals a declining amount of chloroplast DNA during development in Arabidopsis. BMC Plant Biol. 9: 3.
    OpenUrlCrossRefPubMed
  50. ↵
    1. Rowan B.A.,
    2. Oldenburg D.J.,
    3. Bendich A.J.
    (2010). RecA maintains the integrity of chloroplast DNA molecules in Arabidopsis. J. Exp. Bot. 61: 2575–2588.
    OpenUrlAbstract/FREE Full Text
  51. ↵
    1. Sakamoto W.,
    2. Miyagishima S.-y.,
    3. Jarvis P.
    (2008). Chloroplast biogenesis: Control of plastid development, protein import, division and inheritance. In The Arabidopsis Book 6: e0110, doi/10.1199/tab.0110.
  52. ↵
    1. Sakamoto W.,
    2. Zaltsman A.,
    3. Adam Z.,
    4. Takahashi Y.
    (2003). Coordinated regulation and complex formation of yellow variegated1 and yellow variegated2, chloroplastic FtsH metalloproteases involved in the repair cycle of photosystem II in Arabidopsis thylakoid membranes. Plant Cell 15: 2843–2855.
    OpenUrlAbstract/FREE Full Text
  53. ↵
    1. Sato N.,
    2. Nakayama M.,
    3. Hase T.
    (2001). The 70-kDa major DNA-compacting protein of the chloroplast nucleoid is sulfite reductase. FEBS Lett. 487: 347–350.
    OpenUrlCrossRefPubMed
  54. ↵
    1. Sato N.,
    2. Ohshima K.,
    3. Watanabe A.,
    4. Ohta N.,
    5. Nishiyama Y.,
    6. Joyard J.,
    7. Douce R.
    (1998). Molecular characterization of the PEND protein, a novel bZIP protein present in the envelope membrane that is the site of nucleoid replication in developing plastids. Plant Cell 10: 859–872.
    OpenUrlAbstract/FREE Full Text
  55. ↵
    1. Sato N.,
    2. Terasawa K.,
    3. Miyajima K.,
    4. Kabeya Y.
    (2003). Organization, developmental dynamics, and evolution of plastid nucleoids. Int. Rev. Cytol. 232: 217–262.
    OpenUrlCrossRefPubMed
  56. ↵
    1. Scheuermann R.H.,
    2. Echols H.
    (1984). A separate editing exonuclease for DNA replication: The epsilon subunit of Escherichia coli DNA polymerase III holoenzyme. Proc. Natl. Acad. Sci. USA 81: 7747–7751.
    OpenUrlAbstract/FREE Full Text
  57. ↵
    1. Scott N.S.,
    2. Possingham J.V.
    (1980). Chloroplast DNA in expanding spinach leaves. J. Exp. Bot. 31: 1081–1092.
    OpenUrlAbstract/FREE Full Text
  58. ↵
    1. Sheahan M.B.,
    2. McCurdy D.W.,
    3. Rose R.J.
    (2005). Mitochondria as a connected population: Ensuring continuity of the mitochondrial genome during plant cell dedifferentiation through massive mitochondrial fusion. Plant J. 44: 744–755.
    OpenUrlCrossRefPubMed
  59. ↵
    1. Shedge V.,
    2. Arrieta-Montiel M.,
    3. Christensen A.C.,
    4. Mackenzie S.A.
    (2007). Plant mitochondrial recombination surveillance requires unusual RecA and MutS homologs. Plant Cell 19: 1251–1264.
    OpenUrlAbstract/FREE Full Text
  60. ↵
    1. Stupar R.M.,
    2. Lilly J.W.,
    3. Town C.D.,
    4. Cheng Z.,
    5. Kaul S.,
    6. Buell C.R.,
    7. Jiang J.
    (2001). Complex mtDNA constitutes an approximate 620-kb insertion on Arabidopsis thaliana chromosome 2: Implication of potential sequencing errors caused by large-unit repeats. Proc. Natl. Acad. Sci. USA 98: 5099–5103.
    OpenUrlAbstract/FREE Full Text
  61. ↵
    1. Tang L.Y.,
    2. Nagata N.,
    3. Matsushima R.,
    4. Chen Y.,
    5. Yoshioka Y.,
    6. Sakamoto W.
    (2009). Visualization of plastids in pollen grains: involvement of FtsZ1 in pollen plastid division. Plant Cell Physiol. 50: 904–908.
    OpenUrlAbstract/FREE Full Text
  62. ↵
    1. Twell D.,
    2. Yamaguchi J.,
    3. McCormick S.
    (1990). Pollen-specific gene expression in transgenic plants: coordinate regulation of two different tomato gene promoters during microsporogenesis. Development 109: 705–713.
    OpenUrlAbstract
  63. ↵
    1. Tymms M.J.,
    2. Scott N.S.,
    3. Possingham J.V.
    (1983). DNA content of Beta vulgaris chloroplasts during leaf cell expansion. Plant Physiol. 71: 785–788.
    OpenUrlAbstract/FREE Full Text
  64. ↵
    1. Unseld M.,
    2. Marienfeld J.R.,
    3. Brandt P.,
    4. Brennicke A.
    (1997). The mitochondrial genome of Arabidopsis thaliana contains 57 genes in 366,924 nucleotides. Nat. Genet. 15: 57–61.
    OpenUrlCrossRefPubMed
  65. ↵
    1. Wang D.Y.,
    2. Zhang Q.,
    3. Liu Y.,
    4. Lin Z.F.,
    5. Zhang S.X.,
    6. Sun M.X.,
    7. Sodmergen
    (2010). The levels of male gametic mitochondrial DNA are highly regulated in angiosperms with regard to mitochondrial inheritance. Plant Cell 22: 2402–2416.
    OpenUrlAbstract/FREE Full Text
  66. ↵
    1. Zaegel V.,
    2. Guermann B.,
    3. Le Ret M.,
    4. Andrés C.,
    5. Meyer D.,
    6. Erhardt M.,
    7. Canaday J.,
    8. Gualberto J.M.,
    9. Imbault P.
    (2006). The plant-specific ssDNA binding protein OSB1 is involved in the stoichiometric transmission of mitochondrial DNA in Arabidopsis. Plant Cell 18: 3548–3563.
    OpenUrlAbstract/FREE Full Text
  67. ↵
    1. Zhang Q.,
    2. Liu Y.,
    3. Sodmergen
    (2003). Examination of the cytoplasmic DNA in male reproductive cells to determine the potential for cytoplasmic inheritance in 295 angiosperm species. Plant Cell Physiol. 44: 941–951.
    OpenUrlAbstract/FREE Full Text
  68. ↵
    1. Zoschke R.,
    2. Liere K.,
    3. Börner T.
    (2007). From seedling to mature plant: Arabidopsis plastidial genome copy number, RNA accumulation and transcription are differentially regulated during leaf development. Plant J. 50: 710–722.
    OpenUrlCrossRefPubMed
Back to top

Table of Contents

Print
Download PDF
Email Article
Citation Tools
Request Permissions
A Conserved, Mg2+-Dependent Exonuclease Degrades Organelle DNA during Arabidopsis Pollen Development
Ryo Matsushima, Lay Yin Tang, Lingang Zhang, Hiroshi Yamada, David Twell, Wataru Sakamoto
The Plant Cell Apr 2011, 23 (4) 1608-1624; DOI: 10.1105/tpc.111.084012
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo

Jump to section

In this issue

View this article with LENS