Download Spontaneous chloroplast mutants mostly occur by

Plant Cell Advance Publication. Published on April 6, 2016, doi:10.1105/tpc.15.00879
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Spontaneous chloroplast mutants mostly occur by replication slippage and
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show a biased pattern in the plastome of Oenothera
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Authors: Amid Massouh1), Julia Schubert1), Liliya Yaneva-Roder1), Elena S.
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Ulbricht-Jones1), Arkadiusz Zupok1), Marc T. J. Johnson2), Stephen I. Wright3),
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Tommaso Pellizzer1), Johanna Sobanski1), Ralph Bock1), Stephan Greiner1,4)
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1)
Max-Planck-Institut für Molekulare Pflanzenphysiologie, Am Mühlenberg 1, D-
14476 Potsdam-Golm, Germany
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2)
Department of Biology, University of Toronto-Mississauga, Mississauga, ON, L5L
1C6, Canada
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3)
Department of Ecology and Evolutionary Biology, University of Toronto, Toronto,
ON, M5S 3B2, Canada
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4)
Corresponding author:
Stephan Greiner
[email protected]
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The author responsible for distribution of materials integral to the findings presented
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in this article in accordance with the policy described in the Instructions for Authors
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(www.plantcell.org) is: Stephan Greiner ([email protected]).
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Short title: Spontaneous plastome mutants in Oenothera
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Synopsis: Spontaneous chloroplast mutants mostly occur by replication slippage
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and display a mutation spectrum very different from that in the eukaryotic nucleus
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and that of bacteria.
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©2016 American Society of Plant Biologists. All Rights Reserved.
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Abstract
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Spontaneous plastome mutants have been used as a research tool since the
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beginnings of genetics. However, technical restrictions have severely limited their
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contributions to research in physiology and molecular biology. Here, we used full
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plastome sequencing to systematically characterize a collection of 51 spontaneous
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chloroplast mutants in Oenothera (evening primrose). Most mutants carry only a
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single mutation. Unexpectedly, the vast majority of mutations do not represent single
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nucleotide polymorphisms, but are insertions/deletions originating from DNA
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replication slippage events. Only very few mutations appear to be caused by
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imprecise double-strand break repair, nucleotide misincorporation during replication,
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or incorrect nucleotide excision repair following oxidative damage. U-turn inversions
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were not detected. Replication slippage is induced at repetitive sequences that can
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be very small and tend to have high A/T content. Interestingly, the mutations are not
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distributed randomly in the genome. The underrepresentation of mutations caused
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by faulty double-strand break repair might explain the high structural conservation of
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seed plant plastomes throughout evolution. In addition to providing a fully
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characterized mutant collection for future research on plastid genetics, gene
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expression and photosynthesis, our work identified the spectrum of spontaneous
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mutations in plastids and reveals that this spectrum is very different from that in the
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nucleus.
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Introduction
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Mutations in the chloroplast genome (plastome) are typically recognized by pale or
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bleached leaf areas in variegated plants (Kirk and Tilney-Bassett, 1978; Greiner,
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2012; Figure 1A). Such material has been used as a genetic tool since the beginning
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of genetics. For example, the use of plastome mutants by Correns and Baur formed
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the foundation of cytoplasmic genetics (Baur, 1909; Correns, 1909; Baur, 1910;
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Hagemann, 2010). Plastome mutants continue to play an important role in
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elucidating the rules of non-Mendelian inheritance (Chiu and Sears, 1985; Azhagiri
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and Maliga, 2007; Ruf et al., 2007) and in studying chloroplast gene expression and
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photosynthesis (Schaffner et al., 1995; Landau et al., 2009). Moreover, plastid
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mutants provide selectable markers for antibiotic or herbicide resistance that are
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widely used in molecular biology, biotechnology, and agriculture (e.g., Schmitz-
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Linneweber et al., 2005; Powles and Yu, 2010).
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Over several decades, a substantial number of chlorotic chloroplast mutants
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have been identified and characterized from multiple plant species (Kirk and Tilney-
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Bassett, 1978; Börner and Sears, 1986; Somerville, 1986; Hagemann, 1992;
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Greiner, 2012). Importantly, phenotypes of plastome mutants go beyond simple
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bleaching.
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developmentally impaired or sensitive to environmental factors, show mottled
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phenotypes and/or bleach reversibly (e.g., Kirk and Tilney-Bassett, 1978; Stubbe
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and Herrmann, 1982; Chia et al., 1986; Archer and Bonnett, 1987; Colombo et al.,
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2008; Hirao et al., 2009; Landau et al., 2009; Greiner, 2012; Figure 1B-D, Table 1). A
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number of mutant lines have been described that display unexpected phenotypes,
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such as mutant plastids having a phenotypic effect on wild-type plastids in the same
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cell (Michaelis, 1957), or mutations that confer drought and temperature tolerance
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(Usatov et al., 2004; Mashkina et al., 2010). Collectively these works show that
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although chlorosis is a common feature of plastome mutants, plastid mutations can
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have diverse effects on the plant phenotype.
Some
plastome
mutants
display
mild
chlorotic
effects,
are
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Although chloroplast mutants can provide insight into chloroplast gene
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function and regulation, technical limitations have limited wider application in plant
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research. Disregarding mutants resistant to herbicides or antibiotics, only 12
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plastome mutations have been identified in vascular plants (Greiner, 2012). Since
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such mutations cannot be identified by conventional linkage mapping, direct
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sequence analysis is required to pinpoint a mutated locus. Past approaches relied on
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physiological and biochemical analyses of the mutant, which allowed predictions
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about the mutated gene. This strategy was applied to most of the plastome
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mutations that have been identified by molecular approaches (Greiner, 2012), and is
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still successfully used in screens of plastid-encoded herbicide resistance (Thiel et al.,
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2010). However, the approach of combining physiological characterization with local
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sequence analysis suffers from limitations: First, the candidate gene needs to be
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predictable, which implies that its biochemical and physiological attributes are
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already well characterized. Consequently, identifying mutations in genes of unknown
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function is very difficult. Second, the presence of a second site mutation cannot be
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ruled out. Full chloroplast genome sequencing by next-generation sequencing can
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overcome these technical constraints.
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Another reason why chloroplast mutants are currently understudied is the
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challenge of maintaining plastome mutants in standard model species with
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uniparental plastid inheritance. Many plastome mutants are not viable on soil. They
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must therefore be kept in as variegated plants during propagation, because wild-type
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plastomes are essential to nourish the mutant tissue (Figure 1A). Variegation occurs
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because of random chloroplast sorting during cell division, which gives rise to
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homoplastic mutants and wild-type tissue. Sorting-out is a statistical process and
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very rapid (Michaelis 1955, Kirk and Tilney-Bassett, 1978). It is typically completed
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early in leaf development (Schötz 1954). Hence, in most plant species, variegation
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cannot be maintained through sexual reproduction because it is challenging to
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identify egg cells that are heteroplasmic and could give raise to variegated plants in
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the next generation (Kirk and Tilney-Bassett, 1978; Greiner 2012). Therefore,
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chloroplast mutants are typically maintained by cutting variegated stems or in tissue
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culture, but these approaches are infeasible for many vascular plants. Using a model
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organism displaying biparental plastid inheritance, like Oenothera, overcomes the
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technical limitation.
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The genus Oenothera (evening primrose, Onagraceae) is an excellent model
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for the study of chloroplast mutants, including their mutation spectrum, their
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molecular basis and phenotypic effects. Past research shows that spontaneous
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chloroplast mutants occur with a frequency of 0.3% in Oenothera (Kutzelnigg and
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Stubbe, 1974). Biparental transmission allows the directed generation of variegated
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plants by crossing, using individuals with mutated and wild-type plastomes as
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crossing partners. Thus, plastome mutants can be reliably propagated from seeds
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and kept as variegated plants in soil (Figure 1 A, Methods). This approach has
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allowed the generation of a large collection of spontaneous chloroplast mutants.
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Based on chlorotic phenotypes, a total of 51 spontaneous plastome mutants were
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isolated and maintained through efforts of Wilfried Stubbe during the past century
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(Kutzelnigg and Stubbe, 1974; Stubbe and Herrmann, 1982; Table 1; Supplemental
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Table 1). All mutations arose in the shoot apical meristem, since they entered the
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germline and are sexually transmitted. Hence, the collection represents the spectrum
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of heritable mutations that lead to visibly impaired chloroplast function. Although the
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collection has been characterized extensively at the phenotypic level, technical
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limitations have prevented molecular analysis (Greiner, 2012).
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Here we describe the mutation spectrum of disruptive spontaneous
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chloroplast mutations using the Oenothera plastome mutant collection. Based on full
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plastome sequencing we suggest a molecular mechanism of their occurrence.
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Employing statistical methods, we test to what extent mutation events and recovery
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are biased toward the mutated target gene, the local sequence context, and the
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location of a mutation on the chloroplast genome.
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Results
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Phenotypes of chloroplast mutants in homoplasmic leaf tissue
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The 51 mutants of the collection show various types of chlorophyll deficiency, which
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typically occurred in an age-dependent manner. Some of the phenotypes are
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sensitive to environmental factors (Figure 1, Table 1). Non-uniform leaf phenotypes
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in homoplasmic mutant tissue occur either constitutively or are restricted to a certain
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developmental stage in 16 mutants. Three of those display a pigmentation gradient
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from the leaf base to the tip (Figure 1B). In mature rosette leaves, 40 mutants fully
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bleach to a nearly white or yellowish creamed-colored phenotype, whereas light
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green tissue remains throughout development in eleven mutants. Two of those are
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viable in soil and need no nourishment from wild-type tissue (Table 1, cf. Methods).
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Particularly interesting phenotypes are seen in the mutants I-tau and I-iota. In the
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mutant I-iota, an age-dependent mosaic pattern of pigmentation is observed where
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young leaves are white and slightly mottled. The green spots increase in size and
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number during development, leading to nearly green mature rosette leaves (Figure
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1C). I-tau displays a reversible bleaching (virescent) phenotype (Figure 1D). It can
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grow photoautotrophically and forms bleached and green leaves in an alternating
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order.
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Wild-type plastomes
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For identification of the underlying mutations, the full sequence of the corresponding
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wild-type reference plastome was obtained. The mutants of the collection are derived
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from 13 wild-type strains (Supplemental Tables 1 and 2). The complete plastome
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sequences of four of the wild-type lines had been determined previously (Greiner et
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al., 2008a). In this study, we re-sequenced all five previously determined Oenothera
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plastomes
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(EMBL/GenBank accessions: AJ271079.4, EU262887.2, EU262889.2, EU262890.2,
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EU262891.2). In addition to correcting a few sequencing errors (mostly single
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nucleotides) we detected two larger insertions of 147 and 24 bps compared to the
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previously published chloroplast genome of Oenothera elata ssp. hookeri strain
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johansen Standard. This likely reflects genetic variation in our seed stocks within
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unstable highly repetitive genomic regions (Figure 2, Greiner et al. 2008a, and
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below). Newly determined were the complete chloroplast genome sequence of eight
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reference strains and an additional line to address plastome stability (see below;
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GenBank accession numbers KT881169.2 - KT881177.1; for summary see
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Supplemental Table 3). The wild-type reference strain of one mutant (V-alpha) is no
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longer present in our collections (see Table 1 for details). Overall, the Oenothera
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chloroplast genome is well conserved in structure and size (spanning 163,367 -
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166,545 bp) and harbors an identical gene set of 113 unique genes, as described
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earlier (Greiner et al., 2008a; Greiner et al., 2008b).
using
Illumina
next-generation
DNA
sequencing
technology
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Identification of mutated alleles and second-site mutations in the plastome mutant
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collection
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From the 51 spontaneous mutants, 46 unique chloroplast genomes were recovered
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(Table 1), and five mutant pairs share an identical sequence. It is conceivable that
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contamination during propagation of the collection is responsible for these cases of
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seemingly identical chloroplast genomes (the collection was maintained for decades
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without the benefit of molecular markers for tracking). However, since the distribution
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of mutated alleles was found to be biased within the Oenothera plastome (Figure 3
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and below), we cannot exclude the possibility that at least some of the identical
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mutations might have arisen independently. This view is supported by the fact that in
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lines with different wild-type plastomes, the same locus sometimes experienced
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independent mutations. However, those resulted in distinct mutated alleles (mutant
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pairs I-iota/III-delta and I-xi/III-alpha; Supplemental Figure 1 - 3). All mutants sharing
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an identical mutation also share the same wild-type background. Also, in previous
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analyses, their phenotypic descriptions were essentially identical (Tables 1 and
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Methods for details). In this context, it is noteworthy that the oldest viable seed lots
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that we could investigate for potential propagation errors date back to 1985.
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Within the 46 unique mutated chloroplast genomes of the collection, 37
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genomes carry a mutation at a single site (Table 1). The identification of a single
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mutation in V-alpha is tentative, since the original wild-type reference is no longer
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present in our strain collection (see above; Table 1). Two mutated sites are present
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in each of the remaining nine mutants. Two of these mutants represent known
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second-site mutations in that they were isolated based on a phenotypic change
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arising in a pre-existing chloroplast mutant. Thus, in total 53 (37 + 9 + 7) unique
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mutated sites could be detected in the entire collection (Table 1 and Supplemental
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Data Set 1). The second-site mutations of the remaining seven mutants are located
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in regions of intrinsic instability associated with larger tandem repeats or oligo(N)
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stretches in intergenic regions (Figure 2). Their phenotypic relevance is discussed
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below.
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Types of mutations in the collection
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The 53 mutant sites in the collection essentially fall into five classes: (i) single-bp
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transversions (two mutations), (ii) single-bp, double-bp and micro insertions/deletions
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(indels, < 10 bp) at oligo(N) stretches (six mutations), (iii) micro indels at micro
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repeats (< 10 bp; 32 mutations; 21 duplications, eleven deletions), (iv) macro indels
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(mostly between 12 bp and 70 bp, one exceptional deletion of 153 bp) at macro
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repeats (> 10 bp; twelve mutations; five duplications, six deletions; one mutation
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event most likely caused by an insertion and a deletion), and (v) one large deletion
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(1,364 bp), which could not be associated with a repetitive element (see
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Supplemental Data Set 1 and Supplemental Figures 1 - 6 for detailed description of
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the individual mutations). Within the micro indels (< 10 bp), the major class of
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mutations detected in this study, a bias towards duplications is not present (χ2: p =
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0.3113). However, a clear bias to indels conferred by tandem/direct repeats is
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present (χ2: p = 0.0003; 32 mutations, 30 associated with tandem/direct repeats, two
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with palindrome/inverted repeats; Supplemental Data Set 1). In summary, nearly all
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chloroplast mutations are associated with repetitive elements. These repeats can be
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very small, in some cases with two repetitive elements of 3 bp (Supplemental
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Figures 1 - 6; Supplemental Data Set 1).
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Molecular cause of mutations and phenotypic relevance of the second site mutations
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The detected chloroplast mutations have a bias toward coding sequence. Most
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mutations are located in protein-coding regions (44 of 53) of which 37 induce
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frameshifts and six represent in-frame indels. One mutation is conferred by a
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transversion, leading to a single amino acid change (Table 1). At least five of the
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mutations affect chloroplast mRNA metabolism. Three of these occur in introns
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where they result in splicing defects. The mutations in I-psi and II-kappa are located
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in intron 1 and intron 2 of ycf3, respectively, and affect splicing of these introns (cf.
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Landau et al., 2009). The gene is involved in photosystem I assembly (Ruf et al.,
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1997). A similar situation is found for the mutant II-theta where the mutation is
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located in the intron of petB. The pet genes encode subunits of the cytochrome b6f
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complex. Here, unspliced transcripts overaccumulate. The mutant II-delta occurs in
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the intergenic region of the ndhE-psaC spacer (ndh genes encode subunits of the
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chloroplast NADH dehydrogenase complex; psa genes encode subunits of
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photosystem I). The mutation in the ndhE-psaC spacer is the sole mutation in this
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line and it is not located in a known chloroplast promoter (see Greiner et al., 2008a
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for searched motifs). The mutant has an mRNA processing defect in the ndhH-ndhA-
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ndhI-ndhG-ndhE-psaC-ndhD operon (Figure 4). Interestingly, defects in mRNA
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metabolism were not only found in the four mutants where one might have expected
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them (i.e., mutants with mutations in introns or intergenic regions). The mutant II-eta,
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harboring a mutation in exon 2 of petB, was found to be affected in processing
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and/or stability of the psbB (petB) operon transcripts. The primary transcripts from
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this operon undergo a series of processing and maturation steps, resulting in a
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highly complex pattern of partially processed transcripts and mature mRNAs
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(Westhoff and Herrmann, 1988). As judged from transcript sizes and the previous
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characterization of RNA species (Westhoff and Herrmann, 1988), a major spliced
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RNA species comprising psbH, petB, and petD is hardly detectable in II-eta (psbH
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encodes a subunit of photosystem II), whereas a transcript likely containing the
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spliced petB/petD dicistron overaccumulates (Figure 4).
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The remaining five mutations of the collection are in non-coding regions and
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are second-site mutations. Two are located within oligo(N) stretches in intergenic
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regions; three are upstream of ycf1 (tic214) or accD in regions of intrinsic instability
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due to the presence of large complex tandem repeats (Figure 2 and below). The
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accD gene encodes a subunit of the plastidic acetyl-CoA carboxylase; ycf1 (tic214)
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is currently considered to encode a subunit of the chloroplast protein import
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machinery (Kikuchi et al., 2013).
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Among the mutations in genic regions, three are associated with ATP
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synthase, twelve with photosystem I (nine of them within psaA, exclusively
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representing duplications; Figure 3), eleven with photosystem II, and six within the
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cytochrome b6f complex. Further, four mutations were found in ycf3 and one in ycf4
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(as ycf3 involved in photosystem I assembly; Ruf et al., 1997; Krech et al., 2012),
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three within rbcL (encoding the large subunit of RuBisCO), and four within ccsA (a
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gene involved in cytochrome c biogenesis; Xie and Merchant, 1996). These
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functional gene classes affect photosynthesis and are considered to be non-
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essential for cell viability (Scharff and Bock, 2014). An additional mutation occurred
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in rps7, which is likely an essential gene in dicots encoding a protein for the small
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(30S) subunit of the plastid ribosome (Scharff and Bock, 2014; Tiller and Bock,
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2014). Viability of this mutation is likely due to the particular mutated allele, which
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modifies the very C-terminal sequence of the protein. Additional second site
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mutations include in-frame indels in the intrinsically unstable regions of accD and
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ycf1 (tic214). Those are again associated with large complex tandem repeats (Figure
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2).
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The phenotypic relevance of the second site mutations is not entirely clear.
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Mutations within oligo(N) stretches in intergenic regions are likely to be
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phenotypically neutral. By contrast, the contribution to the phenotype is uncertain for
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the mutations in the intrinsically unstable repetitive regions: The location of these
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mutations is upstream or in the coding regions of accD and ycf1 (tic214),
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respectively, which are both essential in dicots (Drescher et al., 2000; Kode et al.,
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2005; Supplemental Data Set 2). However, the phenotype of all mutants carrying
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such mutations can be fully explained by the second mutated allele present in these
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lines (Table 1 and Supplemental Data Set 2). This is likely also the case with
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mutated sites in repetitive regions that represent indels within the reading frames of
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accD and ycf1 (tic214; Figure 2). These indels are in frame and the amino acid
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sequences in these regions are poorly conserved among Oenothera species
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(Greiner et al., 2008a). In addition, very similar polymorphisms, including in-frame
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indels within the coding region of ycf1 (tic214), were also detected upon comparison
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of very closely related wild-type strains, as described below.
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Statistical distribution of mutant alleles within the chloroplast genome
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An important question addressed by our study is whether mutations are randomly or
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non-randomly distributed within the plastome, and whether these patterns reflect real
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mutational biases or biases related to our selection process. We tested for biological
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and methodological biases by assessing the influence of the expected phenotype,
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gene length, and the location of the gene in the plastome on the probability of
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isolating a mutation. We assigned the genes of the Oenothera plastome to one of
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three classes based on phenotypes observed in systematic knock-out analyses of
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chloroplast genes in Nicotiana tabacum (Supplemental Data Set 2). “Class I” genes
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included those in which spontaneous mutations are easily detected. This class
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combines genes that are non-essential for heterotrophic growth and whose knock-
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outs display either white or pale phenotypes (subgroups: “White” and “Pale”).
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Examples for such genes are psaA and rbcL. “Class II” genes do not display a visible
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phenotype (e.g., ndhB and all other subunits of the plastid NDH complex) and
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unsurprisingly the collection does not contain any mutants in that class, which
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excludes it from the following analyses. “Class III” genes are essential for cellular
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viability [e.g., rps7 or ycf1 (tic214)]. Only a limited number of specific (non-lethal)
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mutations could be isolated from this class (see above). Class I genes are
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exclusively located in the two single-copy regions of the plastome [large single-copy
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(LSC) or small single-copy (SSC) region]. Class III genes, however, are also located
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in the inverted repeat (IR; Figure 2). The IR is a typical structure of chloroplast
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genomes and contains exclusively Class II and Class III genes. Due to
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recombination and gene conversion between the two large inverted repeats (Palmer,
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1983), mutations in the IR are constantly purged. Hence, Class III genes need to be
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subdivided based on their physical location on the plastome (LSC and SSC vs. IR). It
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also should be noted that regions of intrinsic instability are found only in Class III
320
genes (Figure 2).
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The first assumption we tested was if mutations in non-genic regions (52,922
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bp in the genome; six mutations in the collection) were more common than mutations
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within genes (112,977 bp in the genome; 47 mutations in the collection).
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Unsurprisingly, this was the case (χ2: p = 0.0022). Second, we confirmed our
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expectation that mutations in Class I genes were more common than mutations in
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Class III genes (Class I: 44 mutations within 37,080 bp of genic sequence; Class III:
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three mutations within 56,920 bp; χ2: p < 0.0001). We found no difference in the
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abundance of mutations between the two subgroups of Class III genes (IR: one
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mutation within 33,672 bp; LSC and SSC: two mutations within 23,248 bp; χ2: p =
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0.7470), but this non-significance may reflect the small sample size.
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Next, we asked if gene length affected the recovery of mutations in Class I
332
genes, represented by 44 mutations within 37,080 bp of genic sequence. χ2 test
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yielded p = 0.0002 for the likelihood of accumulating nine mutations in psaA (2,253
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bp), p = 0.0254 for four mutations in ccsA (960 bp), but p = 0.5287 for three
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mutations in rbcL (1,428 bp). This strongly suggests a gene-specific bias for
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mutation recovery beyond the differences observed between gene classes (Figure
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3). The lack of relationship with gene length is further supported by a large cluster of
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Class I genes in the LSC (13,969 bp) that lack any mutation. The cluster contains
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ATP synthase genes (atpA, F, H and I) and genes for the chloroplast bacterial-type
340
RNA polymerase (rpoB, C1, C2; Figure 2, Supplemental Data Set 2). The joint
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random probability that among 44 independent mutations not a single one is located
342
in this cluster is p = 9.2 x 10-10.
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Finally, we assessed whether bias in mutant recovery can be explained by the
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strength of the chlorotic effect of a mutation, since strong (white) phenotypes might
345
be easier to detect by eye during the selection procedure. Of the 51 mutant lines, 40
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displayed an essentially white phenotype and eleven a pale phenotype
347
(Supplemental Table 4). Surprisingly, compared to the combined lengths of the
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coding sequences of “White” and “Pale” genes (35,119 bp and 1,961 bp,
349
respectively), white phenotypes are significantly under-represented whereas pale
350
phenotypes are over-represented (χ2: p < 0.0001). This difference is most likely
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explained by many mutations not representing full knock-outs (see Discussion).
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Addressing plastome stability
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354
The finding that chloroplast mutations are biased towards replication slippage and
355
spatially clustered in the genome raises the question: To what extent is the natural
356
mutation spectrum of the chloroplast represented by our collection? To address this
357
point, we sequenced and compared four wild-type Oenothera plastomes from O.
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glazioviana (=O. lamarckiana; Cleland, 1972; Harte, 1994; GenBank accessions:
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EU262890.2, KT881171.1, KT881172.1, and KT881174.1). This is a young species
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that arose in Europe in the mid-19th century by hybridization (Cleland, 1972; Dietrich
361
et al., 1997). We investigated the Swedish strain of O. lamarckiana, a line that has
362
been cultivated since 1906, as well as three independent descendants of O.
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lamarckiana de Vries (strains blandina, deserens and vetaurea). Those were isolated
364
in 1908, 1913, and 1921 from the original line of de Vries, collected in the
365
Netherlands in 1886 (see Supplemental Table 2 for details on the O. lamarckiana
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lines). Sequence comparison of the plastome of O. lamarckiana Sweden with those
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of the three descendants of O. lamarckiana de Vries revealed one transition in the
368
intergenic spacer between trnL-UAA and trnT-UGU, one 48 bp indel in the region of
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larger complex repeats upstream of accD, and a single bp insertion at a tandem
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oligo(C+T) stretch between trnL-UAG and ccsA. The three O. lamarckiana de Vries
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plastomes differ by two 21 bp indels in the repetitive region of the coding frame of
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ycf1 (tic214). These arose independently at different sites in blandina and deserens.
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Only the vetaurea strain fully resembles the ycf1 (tic214) sequence of O.
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lamarckiana Sweden. Lastly, the deserens strain has a 6-bp deletion in the
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intergenic spacer between rbcL and atpB that is associated with a micro repeat. The
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mutation is not present in the corresponding plastome mutant III-alpha. Hence, it
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must have occurred in the deserens plastome after isolation of the mutant in 1954
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(cf. Table 1).
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Further evidence that certain regions in the chloroplast genome of Oenothera
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can evolve very rapidly was provided by comparison of plastome sequences of O.
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elata ssp. hookeri strain johansen Standard (assembled in 2007; Greiner et al.,
382
2008a) with that obtained in this work (EMBL accessions: AJ271079.3 vs.
383
AJ271079.4). A 147-bp insertion was detected in the region of large complex repeats
384
upstream of accD, and a 24-bp insertion was identified at the 3’ end of rpoA,
385
associated with a larger tandem repeat. Inspection of the original Sanger sequencing
386
chromatograms revealed that our 2007 sequence lacked these features. The current
387
version of our johansen Standard plastome is based on multiple sequencing
12
388
methods (Illumina, 454, PacBio, and Sanger) and about 50 individuals. It is further
389
supported by the independent chloroplast mutants I-zeta, I-xi and I-fa, which had
390
been isolated in 1962, 1968, and 2011, respectively. It is therefore clear that the
391
particular material sequenced in 2007 must have been polymorphic for the two
392
plastomes.
393
In summary, all but one mutations detected in the closely related Oenothera
394
plastome under study have arisen by replication slippage. They are found at very
395
similar locations and/or sequence contexts as the detect mutations of the collection
396
and are often located in regions of intrinsic instability associated with oligo(N)
397
stretches or large repetitive elements (Figure 2).
398
399
Discussion
400
Spontaneous mutations are largely limited to replication slippage-based events
401
Strikingly, nearly all detected mutations could be associated with repetitive elements
402
(Supplemental Data Set 1). Therefore they must have occurred by a replication
403
slippage-based mechanism, conferred by misalignment or formation of hairpin
404
structures (Levinson and Gutman, 1987; Wolfson et al., 1991; Leach, 1994; Lovett,
405
2004; Figure 5). Consistent with this reasoning is the observation that, with only one
406
exception, all duplications are perfect tandem duplications, which strongly implicates
407
replication slippage as a template-based mechanism responsible for the occurrence
408
of spontaneous chloroplast mutations. Although tandem, direct, inverted, or
409
palindrome repeats can induce indels in the chloroplast genome (cf. Lovett, 2004;
410
Darmon and Leach, 2014), tandem/direct repeats dominated as mutation-inducing
411
sequence motives. Interestingly these repeats can be very small, spanning only two
412
repetitive elements, sometimes no larger than 3 bp (cf. Kondrashov and Rogozin,
413
2004; Tanay and Siggia, 2008; Supplemental Figures 1 - 6; Supplemental Data Set 1
414
and Results).
415
We only rarely detected mutations suggestive of DNA damage by reactive
416
oxygen species (ROS; resulting in transversions; Imlay, 2008) or ultraviolet (UV) light
417
(likely causing transitions; Pfeifer et al., 2005). This also holds true for the types of
418
mutations that are caused by double-strand breaks (DSB) followed by non-
419
homologous end-joining (NHEJ), microhomology-mediated end-joining (MMEJ) or
420
microhomology-mediated break-induced replication (MMBIR) pathways (McVey and
13
421
Lee, 2008; Hastings et al., 2009). NHEJ typically results in small indels of 1 to 4 bp
422
and require 0 - 5 bp microhomologies at the joining ends. Deletions of variable size
423
are common for MMEJ (between 3.1 kb and 4.6 kb, as determined for the chloroplast
424
genome) and microhomologies are utilized for end joining (6 - 12 bp for perfect and
425
10 - 16 bp for imperfect homology in the chloroplast; Kwon et al., 2010). However,
426
such signatures are not found in the indels identified in this study (Supplemental
427
Data Set 1). Furthermore, no plastome rearrangements suggesting MMBIR repair
428
(Cappadocia et al., 2010) or U-turn-like inversions (Zampini et al. 2015) were found.
429
One possible exception is the large 1,364 bp deletion in the mutant V-delta, which is
430
not associated with a repetitive element (Supplemental Figure 6). It might have
431
arisen from a NHEJ-like pathway (cf. Kwon et al., 2010).
432
Our data strongly suggest that mutations caused by processes other than
433
replication slippage are rare in the germline and largely absent from our collection.
434
This observation might reflect protection of the plastid DNA (ptDNA) from ROS and
435
UV light mutagenesis in the shoot apical meristem (Greiner et al., 2015). Harder to
436
explain is the nearly complete lack of signatures resulting from the repairing of DSB
437
in the chloroplast genome. Appropriate pathways exist for DSB repair (Maréchal et
438
al., 2009; Cappadocia et al., 2010; Kwon et al., 2010), but they are likely of minor
439
biological relevance. Similar arguments apply to U-turn-like rearrangements. NHEJ-
440
like repair mechanisms, if present in chloroplasts at all, occur at very low frequencies
441
(Kohl and Bock, 2009; Cappadocia et al., 2010; Kwon et al., 2010). However, neither
442
MMEJ nor MMBRI repair nor U-turn-like rearrangements appear to have produced
443
mutations in our material. This is in line with the observation that extensive X-ray
444
radiation experiments failed to cause recoverable chloroplast mutations (Greiner,
445
2012). Therefore, it seems reasonable to propose an integrity-sensing mechanism
446
for ptDNA in the germline. This could explain the strong structural conservation of
447
higher plant chloroplast genomes during evolution, as reflected in the rarity of large
448
sequence inversions or rearrangements (Ruhlman and Jansen, 2014).
449
450
Mutation spectrum of the chloroplast mutant collection
451
The mutation spectrum in the Oenothera plastome is in sharp contrast to that
452
detected in the nuclear genome of Arabidopsis (Ossowski et al., 2010; Yang et al.,
453
2015) or Escherichia coli (Lee et al., 2012). Mutations in nuclear genomes are
454
biased towards single base pair G:C to A:T transitions (with a lower frequency of
14
455
transversions) that are explained by deamination of methylated cytosine and UV
456
light-induced DNA damage. If single base pair substitutions were common in plastid
457
genomes we would expect more than just the two single-nucleotide mutations
458
present in our collection. The paucity of single-nucleotide mutations in the plastome
459
compared to the nuclear genome may reflect that substitution rates in chloroplast
460
DNA are substantially lower than in the nuclear genome (Wolfe et al., 1987; Drouin
461
et al., 2008). Our results clearly show that chloroplast ptDNA replication machinery is
462
more prone to mutations from replication slippage than misincorporation of single
463
nucleotides. Small indels in E. coli also arise in repetitive sequences but they
464
represent only 10% of spontaneous mutations (Lee et al., 2012). Hence, the
465
chloroplast DNA replication and/or repair machinery differs in its error spectrum from
466
that of bacteria. Although the chloroplast originated from an endosymbiotic event, the
467
difference mutation spectrum likely reflects the fact that the DNA replication system
468
is a mosaic of proteobacterial, cyanobacterial and eukaryotic components (Moriyama
469
and Sato, 2014). The fact that indels in chloroplast genomes are generated with
470
much higher frequency than single base substitutions should be taken into account
471
when performing phylogenetic analyses based on chloroplast sequences. The two
472
classes of mutations cannot be weighted equally as phylogenetically informative
473
characters.
474
475
Reasons for biased mutation recovery
476
Some regions in the chloroplast genome of Oenothera are more prone to mutation
477
than others. Although oligo(N) stretches in coding regions are rare, they are
478
preferred targets of mutation (Supplemental Figure 1, Supplemental Data Set 1).
479
Other hotspots reside in large tandem repeats (Figure 2). Strikingly, compared to the
480
second site mutations found in the plastome mutant collection, very similar mutations
481
could be detected between the closely related wild type plastomes of O. glazioviana.
482
We therefore conclude that specific repetitive regions in the Oenothera chloroplast
483
genomes are intrinsically unstable and prone to mutation. This finding is in line with
484
earlier studies on Oenothera plastome organization, since these reparative regions
485
are major sources of phylogenetic divergence in evening primrose (cf. Gordon et al.,
486
1982; Greiner et al., 2008a; Greiner et al., 2008b). Many mutations in these regions
487
might be phenotypically neutral and can be fixed over experimental time frames, at
488
least in cultured evening primroses.
15
489
Our work also shows a strong mutation bias towards certain non-essential
490
genes. Most mutations appear to be mediated by micro repeats, but several lines of
491
evidence point to additional factors contributing to the unequal distribution of mutated
492
alleles. First, many of the micro repeats are very small (Supplemental Data Set 1),
493
and they are abundant throughout the chloroplast DNA. From a statistical
494
perspective, they might occur at random. For example, the nested micro tandem
495
repeat TATAT independently induced an insertion and a deletion at exactly the same
496
site in the mutants I-xi and III-alpha (Supplemental Figures 2 and 3). However, this is
497
the only TATAT site in the genome where mutations were found, although the same
498
motif is present altogether 114 times in the sequences of Class I genes. Within the
499
atp-rpo cluster of Class I genes, this motif occurs 45 times and again with no
500
mutations. In psaA, nine mutations could be recovered independently of this element
501
(Figure 3, Supplemental Data Set 1, Supplemental Figure 2 and 4). It is present
502
there three times. Hence, although this element is distributed randomly in the
503
genome (χ2: p = 0.4942 for the full set of Class I genes vs. the atp-rpo subset
504
cluster), its potential to induce a recoverable mutation seems to be confined to
505
particular positions in the plastome. It seems conceivable that the surrounding
506
sequence context governs whether TATAT motifs are susceptible to DNA
507
polymerase slippage (cf. Figure 3).
508
We were not able to deduce a consensus sequence for the mutation-inducing
509
micro repeats. The 32 identified sites with mutations consist of many different types
510
of repeats [perfect or imperfect, (nested) tandem, direct or inverted/palindromic
511
repeats]. Also, in a few cases, a combination of repeat types is present. The
512
individual repeat elements are 3 - 9 bp long and the spacer regions, if present, are
513
between 1 bp and 9 bp (Supplemental Data Set 1). A much larger dataset would be
514
needed to deduce any consensus that may exist. Nevertheless, we noticed that
515
mutation-inducing micro repeats tend to be more A/T rich (mutation-inducing sites in
516
total: 364 bp, with 252 bp A+T; “Class I genes”: 37,080 bp, with 22,031 bp A+T; χ2: p
517
= 0.0694; cf. Tanay and Siggia, 2008). However, this tendency cannot explain the
518
lack of mutations in the atp-rpo cluster, when compared to Class I genes (13,969 bp,
519
with 8,424 bp A+T; χ2: p = 0.3631).
520
Beside the sequence context, the relative positions of genes to each other or
521
within the chloroplast genome might generate mutational hotspots. For example,
522
DNA polymerase processivity could vary with distance relative to the origins of
16
523
replication. Furthermore, collision of the replication with the transcription complexes,
524
or other DNA-binding proteins might differ depending on gene position, or the usage
525
of the nuclear- or plastid-encoded RNA polymerase (Börner et al. 2014). Whereas
526
the interference of the DNA replication complex with other DNA binding proteins is a
527
matter of speculation, the relative position of a gene to an origin of replication does
528
not seem to have an influence on a mutational hotspot. For example, the hotspot in
529
ccsA is rather close to the origin of replication, where the one in psaA is not. On the
530
other hand, the atp-rpo gene cluster, fully lacking mutations, is most distantly located
531
from the replication origins (Figure 2).
532
Differences in gene expression and mutation rate have been reported to be
533
correlated in E. coli (Martincorena et al., 2012; but also see Lee et al., 2012).
534
However, this observation is unlikely to explain the patterns of mutation in our
535
material. Since all chloroplast mutations are present in the germ line, they must have
536
arisen in the meristem of the plants. Although gene expression data from proplastids
537
in meristematic tissues are largely lacking, it is extremely unlikely that the expression
538
rates of photosynthesis genes differ substantially in meristems. We, therefore, favor
539
the alternative explanation that mutations in the atp-rpo gene cluster are more
540
difficult to recover than genes with elevated mutation rates such as psaA or ccsA
541
(see above). Possible reasons include biased gene conversion (Khakhlova and
542
Bock, 2006)
543
recombination repair by homologous recombination with an unmutated genome
544
copy. Such hotspots of recombination were reported for the chloroplast DNA of the
545
green alga Chlamydomonas (Newman et al., 1992). It seems conceivable that
546
molecular mechanisms like biased gene conversion or biased recombination repair
547
eliminate de novo mutations in some plastome regions more efficiently than in
548
others.
or
recombinational hotspots
that increase the efficiency of
549
550
Mutation rates in the chloroplast genome
551
Comparison of the closely related O. glazioviana plastomes allows a rough
552
estimation of the mutation rates for chloroplast genomes. We used the reasonable
553
assumptions that O. glazioviana is monophyletic. The species arose by hybridization
554
in the middle of the 19th century (Cleland, 1972; Dietrich et al., 1997). Hence, we can
555
estimate the maximum number of generations separating the Swedish strain from
556
the lines of Hugo de Vries to be about 150 generations with annual reproduction.
17
557
Since three to five polymorphisms distinguish these lines (see Results), we can
558
assume one fixed mutation per 30 - 50 generations, equivalent to 2.0 – 3.3 x 10-2
559
mutations per generation and genome. Per site (~165,000, with each bp
560
representing a site), the mutation rate would be 1.2 - 2.0 x 10-7 per site and
561
generation. Interestingly, compared to the nuclear genome of Arabidopsis thaliana (7
562
x 10-9; Ossowski et al., 2010), and the genome of E. coli (2.2 × 10-10; Lee et al.,
563
2012), this rate is rather high. Nevertheless, taking into account the small genome
564
size of the plastid, the actual number of mutations that effectively accumulate is still
565
small.
566
These approximations should be treated with caution for three reasons. First,
567
the number of mutations is low and thus the precision of our estimate is
568
compromised. Second, a previous study assessing intraspecific plastome variation in
569
Solanum lycopersicum (tomato) over the timespan of approximately one century
570
found no polymorphisms (Kahlau et al., 2006). Third, substitution rates calculated
571
based on phylogenetic data suggest a lower mutation rate in chloroplast genomes
572
than in plant nuclear genomes (Wolfe et al., 1987; Drouin et al., 2008). However, it is
573
also known that organellar mutation rates can vary significantly in different lineages
574
of seed plant evolution, even within a genus (e.g., Cho et al., 2004; Guisinger et al.,
575
2008; Sloan et al., 2009). Last, it should be noted that three of the mutations in the
576
O. glazioviana lines are associated with large repetitive elements repeats. These
577
regions are a major source of phylogenetic divergence (Greiner et al. 2008a) and
578
evolve relatively quickly. Therefore, an accurate mutation rate for plastid DNA in
579
Oenothera remains to be determined by genome sequencing at a larger scale.
580
581
Chloroplast mutants as a tool to understand chloroplast gene functions and gene
582
regulation
583
The finding that most chloroplast mutants in our collection show only one mutated
584
site makes them a valuable resource to understand chloroplast gene function and
585
regulation. Although not all genes in the genome are covered, chloroplast mutants
586
for all photosynthetic complexes and also for the chloroplast translation machinery
587
are represented. For the psaA, psbA, and ccsA genes, allelic series are available,
588
with nine, five, and four mutants, respectively (Table 1, Figure 3). Although rare, the
589
collection also contains mutations in essential genes, such as the mutant I-tau, which
590
carries a defect in rps7. Since this mutant displays a reversible bleaching (virescent)
18
591
phenotype (Figure 1D) its characterization should provide new insights into the
592
developmental regulation of chloroplast translation. Also of particular interest are the
593
five mutants identified so far that display defects in chloroplast RNA metabolism
594
(Figure 4). Such mutants are not readily obtainable by targeted approaches. A
595
particularly interesting finding is this respect was that mutations in the protein-coding
596
region can influence transcript processing, as discovered in the mutant II-eta (Figure
597
4). The molecular basis of these effects needs to be further investigated.
598
It is also important to note that many of the mutants (I-alpha, I-delta, I-iota, I-
599
tau, I-omega, I-do, II-delta, II-theta, II-kappa, II-lambda, III-alpha, III-beta, IV-beta,
600
and V-alpha) do not harbor full gene knock-outs, as judged by their phenotype and
601
genotype (Table 1). This renders them a rich platform for functional analyses. Most
602
null mutants of photosynthesis genes are essentially white (Schaffner et al., 1995;
603
Ruf et al., 1997; Hager et al., 1999; Supplemental Data Set 2 and Supplemental
604
Table 4), lacking any photosynthetic activity and thus of limited value for functional
605
studies. The mutations in our Oenothera collection that affect chloroplast mRNA
606
metabolism are located in photosynthesis genes. At least two of them (II-kappa and
607
II-delta) display mild phenotypes. This is due to reduced abundance of the mature
608
transcript (Figure 4), leading to a less severe phenotype than a frameshift mutation
609
in the coding region (Table 1). An additional set of five mutants that are affected in
610
the coding region of photosynthesis genes (I-alpha, I-delta, I-iota, II-lambda and III-
611
beta) are pale green rather than white. With one exception (II-lambda), the mutations
612
cause a frameshift. In some of these cases, the pale phenotype of the mutants are
613
likely caused by the position of the frame-shifting indel within the gene. For example,
614
the mutants I-alpha and I-delta carry a 5-bp duplication just downstream of the
615
translation initiation codon of psbD (Table 1). We hypothesize that these mutations
616
are compensated by an alternative in-frame start codon present downstream of the
617
mutation. By contrast, in the mutant III-beta, a 5 bp duplication exists in the middle of
618
psaB (encoding an essential reaction center protein of photosystem I) (Table 1),
619
which makes its pale green phenotype more difficult to understand (cf. Table 1). A
620
possible clue may be provided by one of the most striking mutant phenotypes
621
observed in the collection: the mottled phenotype of the mutant I-iota, which largely
622
disappears with age, giving rise to a nearly green phenotype (Figure 1C). The
623
underlying mutation is a single A insertion located in a oligo(A) stretch very early in
624
atpB (Table 1; Supplemental Data Set 1). The recovery of the mutant over time
19
625
strongly points to the presence of a frame-correcting mechanism in chloroplasts,
626
such as ribosomal frameshifting (Kohl and Bock, 2009).
627
In summary, the mild chlorotic and developmental-stage dependent
628
phenotypes of several mutants in the collection provide interesting new avenues for
629
the study of plastid gene function and expression.
630
631
Methods
632
Plant material
633
Throughout this work, the terms Oenothera or evening primrose(s) refer to
634
subsection Oenothera (genus Oenothera section Oenothera; Dietrich et al., 1997).
635
Plant material used here is derived from the Oenothera germplasm collection
636
harbored at the Max Planck Institute of Molecular Plant Physiology, Potsdam-Golm,
637
Germany (Greiner and Köhl, 2014). Part of this collection is represented by the 51
638
spontaneous plastome mutants that were collected systematically during the past
639
century in Germany (Kutzelnigg and Stubbe, 1974; Stubbe and Herrmann, 1982;
640
Stubbe, 1989; Supplemental Table 1). The plastome mutants were mostly compiled
641
by Wilfried Stubbe and co-workers, maintained by Reinhold G. Herrmann since the
642
end of the 1980s and later taken over by the authors. The mutants are designated
643
according to their basic plastome genotype (I - V) and Greek letters, in order of the
644
respective isolation date (i.e., I-alpha, I-beta, II-alpha, II-beta, etc.). In the case of
645
basic plastome I, of which more mutants were isolated than Greek letters exist,
646
further mutants are designated by the seven tone syllables. To maintain the material,
647
a substitution of the mutant chloroplast by introgression into the suitable johansen
648
Standard strain of O. elata ssp. hookeri was performed by W. Stubbe. Except for the
649
lines containing plastome V, which are still maintained in the background of O.
650
argillicola, the collection has been propagated in the johansen Standard background
651
since then (Greiner 2012). Due to the introgression of most of the lines into the
652
johansen Standard strain, chloroplast substitution lines with this nuclear background
653
and the wild-type plastomes of the investigated mutant chloroplasts were used for
654
RNA gel blot analyses as wild-type controls. For a summary of the genetic stocks,
655
see Supplemental Tables 1 and 2. For further details see Kutzelnigg and Stubbe
656
(1974), Stubbe (1989), Rauwolf et al. (2008), and Greiner (2012).
657
20
658
Plant cultivation, maintenance, tissue harvest, and phenotyping of chloroplast
659
mutants
660
Chloroplast mutants, wild type, and introgression strains were cultivated as
661
described previously (Greiner and Köhl, 2014). In Oenothera, photosynthetically
662
incompetent chloroplast mutants can be kept as variegated plants, containing green
663
wild type as well as mutated chloroplasts (Figure 1A). These plants are viable in soil,
664
because the green tissue feeds the bleached tissue. The combination of two
665
chloroplast types in one plant is possible by taking advantage of biparental
666
chloroplast transmission, which is the rule in evening primroses. In such
667
experiments, a maintainer strain containing a green nursery plastome is crossed to
668
the chloroplast mutant. This results in variegated offspring, because sorting-out of
669
the two chloroplast types leads to tissues homoplasmic for either wild-type or mutant
670
chloroplasts (Figure 1A,B; also see Introduction). Homoplasmic mutant tissue
671
generated this way was used for sequence and molecular analysis, as well as
672
phenotyping in this work. For details on the genetics, see Kutzelnigg and Stubbe
673
(1974), Kirk and Tilney-Bassett (1978), Stubbe and Herrmann (1982) and Greiner
674
(2012).
675
Phenotyping of mutants was performed in an unheated polyethylene
676
greenhouse. Plants were analysed after the appearance of the first two true leaves
677
(young), during the early rosette stage (medium) and during the mature rosette stage
678
(old). Observed phenotypes largely resembled those reported in the laboratory
679
documentation of W. Stubbe that was based on field-grown material. However, minor
680
discrepancies were observed. Different degrees of bleaching were assigned to a
681
numerical code (1 = nearly white, 2 = yellowish cream-colored, 3 = light green, 4 =
682
nearly green). Numbers in parentheses in Table 1 indicate mottled or heterogeneous
683
chlorotic phenotypes.
684
685
Chloroplast isolation
686
An improved chloroplast isolation protocol for Oenothera was developed for this
687
study. It overcomes severe technical limitations caused by the high amounts of
688
mucilage present in Oenothera leaf tissue and does not require ultra-centrifugation
689
steps. As judged from sequencing analysis, ptDNA can be enriched to up to 90% by
690
employing the following procedure at 4°C: 50 g of leaf material were homogenized in
691
a blender adding 1 L of BoutHomX buffer [0.4 M sucrose, 50 mM Tris, 25 mM boric
21
692
acid, 10 mM EGTA, 10 mM KH2PO4; 1% (w/v) BSA, 0.1% (w/v) PVP-40; pH 7.6 with
693
KOH; 5 mM freshly added β-mercaptoethanol]. Aliquots of the homogenate were
694
filtered through a double layer of cheese cloth (Hartmann, Heidenheim, Germany),
695
followed by filtering through a single layer of Miracloth (Merck, Darmstadt, Germany).
696
The filtrate was then centrifuged for 15 min at 5,000 g. The chloroplast pellet was
697
resuspended in 150 ml ChloroWash (2 mM EDTA, 1 mM MgCl2, 1 mM MnCl2, 50
698
mM HEPES, 330 mM sorbitol; pH 7.6 with KOH; 5 mM freshly added sodium
699
ascorbate) using a 30 cm3 Potter homogenizer (mill chamber tolerance: 0.15 - 0.25
700
mm; VWR, Darmstadt, Germany). The suspension was again filtered in aliquots
701
through a double layer of Miracloth and adjusted with ChloroWash to a volume of
702
about 700 mL followed by another filtering step. Next, chloroplasts were pelleted at
703
2,400 g, re-suspended in 9 mL of ChloroWash and transferred to three 85%/45%
704
Percoll step gradients [PBF-Percoll stock solution: 3% (w/v) PEG 6000, 1% (w/v)
705
BSA, 1% (w/v) Ficoll 400, dissolved in Percoll; 45%/85% Percoll: 45%/85% PBF-
706
Percoll stock solution, 330 mM sorbitol, 50 mM HEPES, 2 mM EDTA, 1 mM MgCl2;
707
pH 7.6 with KOH; Percoll step gradient: 8 ml 85% Percoll, 16 ml 45% Percoll in a 30
708
ml Corex tube]. The gradients were centrifuged at 10,000 g for 10 min. Intact
709
chloroplasts were recovered from the gradient’s interphase and washed in 200 ml
710
ChloroWash. After repeating the washing steps in smaller volumes, the isolated
711
chloroplasts were re-suspended in a suitable buffer for further use.
712
713
DNA isolation
714
Total DNA for Illumina sequencing was isolated from rosette leaves using a phenol-
715
based DNA isolation protocol developed for Oenothera (Ågren et al., 2015). In
716
contrast to the previously described protocol, we used 100 µl of a 10 mg/mL RNAse
717
A solution (50 U/mg; Roche Diagnostics GmbH, Manheim, Germany). For PCR
718
analyses, we used total DNA obtained with the DNeasy Plant Mini Kit (Qiagen,
719
Hilden, Germany; Rauwolf et al., 2008). For certain Oenothera strains, especially
720
when older material was used, DNA was additionally purified by an anion-exchange
721
column (Nucleobond AX 20; Macherey-Nagel, Düren, Germany). This procedure
722
was found to be very effective in removing remaining phenolic components,
723
mucilage and/or polysaccharides that potentially inhibit enzymatic reactions from
724
Oenothera DNA isolations in general.
22
725
Total high molecular weight DNA for PacBio sequencing was obtained from
726
etiolated seedlings. To this end, seeds were sown on a wet sterile filter paper in Petri
727
dishes and supplemented with 0.05% (v/v) of Plant Preservative Mixture (Plant Cell
728
Technology Store, Washington, DC, USA) to prevent fungal contamination.
729
Subsequently, seeds were incubated at 27°C in the light. Immediately after
730
germination (when root tips became visible), Petri dishes were wrapped with
731
aluminium foil and after three days, material was harvested and frozen in liquid
732
nitrogen. 100 mg of seedlings were ground in liquid nitrogen, subsequently
733
supplemented with 1550 µL CTAB buffer [130 mM Tris/HCl (pH 8.0), 1.8 M NaCl,
734
1.3% (w/v) PVP-40, 2.6% (w/v) CTAB] and 250 µL 1-thioglycerol. Throughout the
735
procedure, samples were mixed by gentle inversion. The aqueous DNA solution was
736
pipetted with a cut tip. After adding 100 µL of a 10 mg/mL RNase A solution (see
737
above), samples were incubated for 30 min at 60°C. Afterwards, 100 µL of
738
proteinase K (10 mg/mL; Sigma-Aldrich, Steinheim, Gemany) was added and
739
samples were incubated at the same conditions as described above. To remove cell
740
debris, samples were centrifuged at 20,000 g for 5 min at room temperature. The
741
supernatant was treated with 2 mL phenol/chloroform/isoamyl alcohol (25:25:1, pH
742
7.5) and 2 mL chloroform/isoamyl alcohol (24:1). DNA was precipitated with 1/10
743
volume of 5 M NH4-acetate and 1 volume of isopropanol. Further purification was
744
achieved using QIAGEN Genomic-tip 20/G columns (Qiagen, Hilden, Germany) and
745
(optionally) with a “High Salt Phenol Chloroform Cleanup” recommended by Pacific
746
Biosciences (Menlo Park, CA, USA). Deviating from the Pacific Biosciences protocol,
747
we omitted EDTA from our buffers. As judged from pulsed-field gel electrophoresis,
748
our procedure yields DNA fragments between 20 kb and 200 kb, with ODs of ~1.8
749
(A260/A280) and ~2.1 (A260/A230). Prior to library preparation, the DNA was subjected
750
to an additional AMPure PB purification step (Pacific Biosciences, Menlo Park, CA,
751
USA).
752
To obtain ptDNA for 454 sequencing, isolated chloroplasts were re-
753
suspended in 400 µL of TENTS buffer [100 mM Tris/HCl (pH 8.0), 50 mM EDTA, 0.5
754
M NaCl, 0.2% (v/v) Triton X-100, 1% (w/v) SDS] and incubated at 60˚C for 15 min.
755
Next, the lysate was supplemented with 100 µL RNase A (10 mg/mL, see above)
756
and incubated for 1.5 h at 37°C. Subsequently, 100 µL of 10 mg/mL proteinase K
757
(see above) solution was added and samples were placed overnight at room
758
temperature. After extraction with phenol/chloroform/isoamyl alcohol (25:24:1, pH
23
759
7.5) and with chloroform/isoamylalcohol (24:1), the DNA was precipitated at -20°C
760
overnight as described above.
761
762
RNA isolation
763
Total RNA for RNA gel blot analyses was isolated from Oenothera leaves using the
764
Spectrum Plant Total RNA Kit (Sigma, Steinheim, Germany). Columns and buffers
765
used were included by the manufacturer. To cope with mucilage and high amounts
766
of phenolic components in the tissue, minor modifications were introduced to the
767
provided protocol: Up to 25 mg of frozen tissue was ground in liquid nitrogen.
768
Material was lysed in 750 μL of freshly prepared Lysis Solution (50 μL β-
769
mercaptoethanol for 1 mL of supplied Lysis Solution). The lysate was incubated at
770
room temperature for 5 min and filtrated on a Filtration Column. Centrifugation time
771
was extended up to 5 minutes. 750 μL of Binding Solution was mixed with the
772
clarified lysate, mixed gently through pipetting and centrifuged on a Binding Column.
773
An additional elution step was performed to increase of RNA yields.
774
775
Detection of RNA via RNA gel blot
776
1.5 - 2.0 µg of total RNA was separated on 1% agarose gels (w/v) under denaturing
777
condition. RNA was transferred to a nylon membrane (Hybond–XL, GE Healthcare,
778
Buckinghamshire, UK) by capillary transfer using 10x SSC buffer (1x SSC: 0.015 M
779
sodium citrate, 0.15 M NaCl). The RNA was covalently linked to the membrane using
780
a UV cross linker (Vilber Lourmat, Ebertharfzell, Germany). Radiolabelled probes
781
were used for strand-specific detection of transcripts in RNA gel blot hybridization.
782
For this purpose transcription of gene specific PCR products was performed with the
783
MAXIscript T7 Kit (Ambion, Darmstadt, Germany) using radiolabelled
784
(Hartman Analytic GmbH, Braunschweig, Germany). Gene specific PCR products for
785
ycf3, petB and psaC were obtained employing the following primers in a PCR with
786
total Oenothera DNA as a template: ycf3ex2_north_fr: 5’-ggatgtcggctcagtcag-3’,
787
ycf3ex2_north_rv:
788
petBex2_north_fr:
789
taatacgactcactatagggaatacagctagaaccacgcctgtg-3’,
790
gtcggatattcattctcgtatctggagc-3’,
791
taatacgactcactatagggagttgctcgttccaggcctc-3’,
792
taatacgactcactatagggacccatacttcgagttgtttcatgc-3’,
32
P dUTP
5’-taatacgactcactataggggtaagaacgggtttcgttctagtg-3’,
5’-gtctcgagattcaggcgattgc-3’,
petBex2_north_rv:
5’-
petBint_north_fr:
5’-
petBint_north_rv:
24
psaC_north_fr:
and
psaC_north_rv:
5’5’5’-
793
atgtcgcattcagtaaagatttatgatacctg-3’. RNA gel blots were performed according to a
794
standard protocol. Nylon membrane with RNA was incubated in Church buffer [0.5 M
795
NaH2PO4, 7% SDS (w/v), 1 mM EDTA; pH 7.2] for one hour. After this time
796
radioactive probes were transferred to hybridization tube and incubated overnight in
797
65°C. Next day three washing steps were performed: once in 1x SSC supplemented
798
with 0.2% SDS (w/v) for 20 min and twice in 0.5x SSC supplemented with 0.2% SDS
799
(w/v) for 20 min each. Visualization of radioactive signals was performed at different
800
time points, depending on the strengths of the mRNA signal.
801
802
DNA sequencing
803
Sanger sequencing was performed with PCR products using standard methods (at
804
Eurofins MWG Operon, Ebersberg, Germany). Primers were obtained from the same
805
source.
806
Illumina paired-end sequencing (375 bp insert size) was performed as
807
described previously (Ågren et al., 2015). To improve sequence quality at very long
808
oligo(N) stretches and to avoid a bias in the Illumina libraries against certain (TA)n
809
rich sequences in some regions of the Oenothera chloroplast genome, “PCR free”
810
Illumina libraries were employed at a later stage of the project. In most cases, about
811
10% to 30% of the sequenced DNA was found to be ptDNA. The observed variance
812
results from the developmental stage of the leaf tissue used and/or the particular
813
genotype of the line (cf. Rauwolf et al., 2010). From these data, we could deduce
814
that roughly 5 million total DNA 100 bp Illumina paired-end reads are sufficient to
815
sequence a chloroplast genome of Oenothera at 500x - 1500x coverage. In this
816
context, it is relevant to mention that the mutant tissue (obtained from random
817
sorting-out of mutant and nursery plastomes) used in this work for Illumina
818
sequencing (see above), was confirmed to be fully homoplasmic for the mutant
819
chloroplast genome.
820
Prior to 454 sequencing, 2 - 5 μg of isolated ptDNA were pre-amplified to 20
821
µg using the GenomiPhi HY DNA Amplification Kit (GE Healthcare, Chalfont St
822
Giles, UK). Single-end read libraries were sequenced on a Roche/GS FLX Titanium
823
platform to an approximate coverage of 20x per chloroplast genome. 454
824
sequencing was performed at Eurofins MWG Operon, Ebersberg, Germany. To
825
correct sequence uncertainties that are frequent in 454 datasets and a potential bias
25
826
in the template DNA introduced by the rolling cycle amplification with the GenomiPhi
827
Kit, all chloroplast genomes obtained from 454 data were re-sequenced by Illumina.
828
PacBio sequences were obtained from a 7 kb cut-off insert library using P4-
829
C2 chemistry. Size selection, library preparation and sequencing were performed at
830
the Max Planck-Genome-Centre in Cologne, Germany. PacBio sequencing is part of
831
the Oenothera genome project and will be communicated elsewhere.
832
833
Plastome assembly
834
In total, the complete sequences of 65 chloroplast genomes were determined in this
835
study, many of them through multiple methods. For a summary of the different
836
sequencing and assembly methods used, see Supplemental Tables 3 and 5. De
837
novo assemblies of the chloroplast genomes and reference-guided mutation
838
mapping were performed using the standard parameters of SeqMan NGen v.12.1.0
839
or earlier (DNASTAR, Madison, WI, USA) and/or CLC Genomic Workbench v.7.5.1
840
or earlier (CLC bio, Aarhus, Denmark). First we determined whether Illumina libraries
841
obtained from total DNA contain plastome sequences translocated to the nuclear
842
genome (NUPTs; Kleine et al., 2009). Translocated sequences, potentially
843
containing polymorphisms compared to the true cytoplasmic ptDNA, might interfere
844
with the assembly/mutation identification. Therefore, we independently de novo
845
assembled 454 reads from ptDNA of isolated chloroplasts and Illumina reads from
846
total DNA. For this, we used three wild-type lines and eleven chloroplast mutants.
847
Both approaches (Illumina and 454 of purified ptDNA from the same materials) gave
848
comparable results and NUPTs could not be detected with certainty in the Illumina
849
sequences. Hence, the remaining chloroplast genomes were solely obtained from
850
Illumina reads of total DNA. However, both assemblers and sequencing techniques
851
did not yield satisfactory results in the de novo assemblies of the notoriously
852
repetitive regions of the Oenothera plastome, such as upstream and within accD and
853
ycf1 (tic214), within ycf2 and in the rrn16-trnI-GAU spacer. In these regions,
854
individual repetitive elements can span more than the read length and/or the insert
855
size of a paired-end library. In the wild-type reference plastomes, those gaps were
856
closed by Sanger sequencing and even consistent assemblies were routinely verified
857
with the same technique. In the johansen Standard line, these regions were
858
additionally verified by single PacBio reads, fully spanning the repetitive regions. All
859
repetitive regions discussed in this work are supported by Sanger reads. Chloroplast
26
860
mutations, if not identified from a de novo assembly, were identified by a reference-
861
guided assembly using
862
Misassemblies, indicating larger indels, were identified by paired-end mapping.
863
Smaller mutations could be directly identified in the reference-guided assembly. The
864
notoriously repetitive regions as well as some of the mutant alleles, especially those
865
harbouring larger indels, were verified at random by Sanger sequencing.
the
chloroplast genome
of
the
reference
strain.
866
867
Sequence annotation
868
Wild-type chloroplast genomes were annotated using GeSeq v.0.2.1, GenBank
869
submission files were generated by GenBank 2 Sequin (https://chlorobox.mpimp-
870
golm.mpg.de). These software tools were developed for this project to annotate and
871
process organelle sequences (manuscripts in preparation). The existing annotation
872
of the reference Oenothera chloroplast genome from O. elata ssp. hookeri strain
873
johansen Standard (AJ271079) was used as a reference. Also, in the course of this
874
work, the annotation of our previously sequenced Oenothera plastomes (Greiner et
875
al., 2008a) was revised, and the exon/intron borders of protein-coding genes are
876
now verified by cDNA sequences. Further, some names of genes and gene products
877
were updated (see AJ271079.4, EU262887.2, EU262889.2, EU262890.2 and
878
EU262891.2). For Illumina transcriptome sequencing, total RNA extractions,
879
sequencing and assemblies, see Johnson et al. (2012) and Hollister et al. (2015).
880
881
Repeat analysis
882
Longer repetitive elements were identified manually or using the EMBOSS suite
883
(Rice et al., 2000). Larger imperfect palindromes were identified with the standard
884
parameters of mfold (Zuker, 2003). For macro repeat analyses incorporated into
885
Figure 2 refer to Greiner et al. 2008a. Since we found that repetitive elements of no
886
larger than 3 bp are able to induce replication slippage in the chloroplast, a
887
sequence inspection by eye was necessary to identify these elements. For this
888
reason, the sequence context 20 bp upstream and downstream of a micro indel (<
889
10 bp) was inspected for (perfect/imperfect/nested) micro (<10 bp) tandem, direct,
890
palindrome and inverted repeats or combinations thereof.
891
892
Global sequence analyses and visualization of the Oenothera plastome
27
893
For
global
sequence
analyses,
including
statistical
analysis,
GC
content
894
determination, repeat analyses and search for sequence motif abundance, the
895
reference Oenothera chloroplast genome of the johansen Standard strain (O. elata
896
ssp. hookeri) was used (AJ271079.4). Especially coding regions of the Oenothera
897
plastome are largely identical between Oenothera species (Greiner et al., 2008a).
898
Chloroplast genome maps were drawn by OrganellarGenomeDRAW (Lose et al.
899
2007) and edited in CorelDarw X5 (Corel Corporation, Ottawa, ON, Canada).
900
901
Statistical analyses
902
χ2 tests were performed using GraphPad (http://www.graphpad.com) with Yates'
903
correction. Two-tailed p-value equals are given.
904
905
Accession Numbers:
906
Sequence data from this article can be found in the EMBL/GenBank data libraries under
907
accession numbers AJ271079.4, EU262887.2, EU262889.2 - EU262891.2, and KT881169.2
908
- KT881177.1.
909
910
Supplemental Data
911
Supplemental Figure 1. Sequence context of the detected mutations at oligo(N) stretches.
912
Supplemental Figure 2. Sequence context of the detected micro tandem duplications.
913
Supplemental Figure 3. Sequence context of the detected micro deletions.
914
Supplemental Figure 4. Sequence context of the detected macro tandem duplications.
915
Supplemental Figure 5. Sequence context of the detected macro tandem duplication and
916
deletion.
917
Supplemental Figure 6. Sequence context of the detected macro deletions.
918
919
Supplemental Table 1. Origin of the mutant lines in the Oenothera plastome mutant
920
collection.
921
Supplemental Table 2. Oenothea wild type, nuclear genome mutants and chloroplast
922
substitution lines used this work.
923
Supplemental Table 3. Summary on the wild type Oenothera chloroplast genomes
924
determined de novo or re-sequenced in this work.
925
Supplemental Table 4. Major phenotypic classes of the Oenothera plastome mutants.
926
Supplemental Table 5. Sequencing and assembly methods of the full chloroplast genomes
927
determined in this study.
28
928
929
Supplemental Data Set 1. Summary of the sequence context of the identified mutations in
930
the chloroplast mutant collection.
931
Supplemental Data Set 2. Macroscopic phenotypes of deletion mutants in the chloroplast
932
genome.
933
934
Acknowledgments
935
Dr. Wilfried Stubbe is acknowledged for compiling and previously maintaining
936
Oenothera plastome mutant collection, and Dr. Reinhold G. Herrmann, Dr. Herbert
937
Hopf and Elisabeth Gerick for maintaining the material after the retirement of Dr.
938
Wilfried Stubbe. Thanks go to Dr. Dirk Walther for advice in statistical analysis, Drs.
939
Jörg Meurer and Serena Schwenkert for sharing information on the macroscopic
940
phenotypes of psbI and psbK knock-out plants. We wish to thank Dr. Barbara B.
941
Sears for providing her mutant material, which served as control samples. She
942
obtained her material earlier from Dr. Wilfried Stubbe’s collection. Dr. Barbara B.
943
Sears and two anonymous reviewers are acknowledged for fruitful discussion and
944
helpful comments on the manuscript. Transcriptome sequencing was made possible
945
through Dr. Gane Ka-Shu Wong, the 1kp initiative (1kp.org) and BGI-Shenzhen. The
946
work was supported by grants from the Deutsche Forschungsgemeinschaft (DFG;
947
GR 4193/1-1) to S.G. and R.B. (BO 1482/17-1), and by the Max Planck Society.
948
M.T.J.J. is supported by an NSERC Discovery Grant and an Early Researcher
949
Award from the Ontario Government. S.I.W. is funded by a Natural Sciences and
950
Engineering Research Council of Canada (NSERC) grant.
951
952
Author Contributions
953
A.M., J.S., L.Y.-R., E.U.-J., A.Z., T.P., J.S., and S.G. performed experiments. All
954
authors analyzed and discussed the data. S.G. designed the study and wrote the
955
manuscript. R.B. participated in writing.
956
29
957
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958
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37
Table 1. Phenotypes, major defects and mutated loci of the mutant lines in the Oenothera plastome mutant collection
Mutant1)
Defect in
Gene or
locus
M1/M4
I-alpha/
I-delta
PS II
psbD
M2
I-beta
Cytb6f
petD
Leaf phenotype2)
Young
Medium
4
3-4
4
M2
I-beta
M3/M5
I-gamma/
I-epsilon
-
PS II
3
Old
3
2-3
5 bp duplication
(CTTGG), + 18 to + 22
psbD, truncated PsbD,
alternative start codon [?]
1
old leaves often with
small green spots
4 bp duplication (ATTT),
+ 998 to + 1001 petD,
truncated PetD
66 bp deletion
(AATTT…TATGG), - 258
to - 323 ycf1 (tic214)3)
1
older leaves with
enhanced greening
close to vascular
tissue
2 bp duplication (GT), +
82 to + 83 psbA,
truncated PsbA
4
M6
I-zeta
PS I
psaA
1-2
1
1
-
M7
I-eta
PS I
psaA
2
1-2
1
-
M8
I-theta
PS I
psaB
2
1-2
1
-
M9
I-iota
ATP
synthase
atpB
(1)
(1-4)
(4)
38
Mutation and predicted
molecular
consequence
very old leaves with
narrow whitish
zones, older leaves
sometimes darker
green
SSC-IRA
junction
psbA
Description
mottled, young
leaves white, old
leaves nearly green
5 bp duplication
(GTTTT), + 1350 to +
1354 psbA, truncated
PsaA
4 bp duplication (GATT),
+ 1088 to + 1090 psaA,
truncated PsaA
32 bp duplication
(TTCCA…ATTAG), +
2132 to + 2163 psaB,
truncated PsaB
1 bp insertion (A), + 58
atpB, truncated AtpB [?]
Table 1. (continued)
Mutant1)
Defect in
Gene or
locus
Leaf phenotype2)
Young
Medium
Old
Description
M10/M15
I-kappa/
I-omikron
PS I
psaA
1-2
1-2
1-2
-
M11
I-lambda
PS II
psbA
3-4
1-2
1
-
M12
I-my
Cytb6f
ccsA
4
3-4
1
old leaves often with
small green spots
M13
I-ny
PS I
psaB
2
1-2
1
-
M14
I-xi
PS II
psbD
4
3-4
1
-
M16
I-pi
PS II
psbA
3-4
2-3
1
-
M17
I-rho
PS II
psbA
3-4
2
1
greening declining
from leaf base to tip
4
4
4
-
M17
I-rho
-
upstream
accD
M184)
I-sigma
RuBisCO
rbcL
39
Mutation and predicted
molecular
consequence
70 bp duplication
(ACGAT…CCCGT), +
1437 to + 1506 psaA,
truncated PsaA
4 bp deletion (GCTT), +
859 to + 862 psbA,
truncated PsbA
31 bp duplication
(GAATT…TTAAA), + 540
to + 570 ccsA; truncated
CcsA
4 bp deletion (AGAT), +
399 to +402 psaB,
truncated PsaB
5 bp deletion (CAGCT), +
530 to + 534 psbD,
truncated PsbD
5 bp deletion (TCTCT), +
208 to + 212 psbA,
truncated PsbA
5 bp deletion (ACAAC), +
797 to + 801 psbA,
truncated PsbA
30 bp deletion
(CCTTT…CTTTA), -30 to
-1 accD3)
5 bp deletion (TTAAC),
+808 to +812 rbcL,
truncated RbcL
Table 1. (continued)
Mutant1)
M19
M20
Defect in
I-tau
translation
I-ypsilon
ATP
synthase
Gene or
locus
rps7
Description
Mutation and predicted
molecular
consequence
periodical
bleaching/regreening (virescent),
viable on soil at
homoplasmic state,
retarded in cell
growth
4 bp duplication (ATTT),
+ 462 to + 465 rps7, last
2 aa of Rps7 modified
and 2 aa added
Leaf phenotype2)
Young
(1-4)
Medium
(1-4)
Old
(1-4)
atpB
(1)
(1)
(1)
mottled, but spots
very rare
M20
I-ypsilon
-
ycf1 (tic214)
M21
I-phi
Cytb6f
ccsA
3-4
2
2
-
M22
I-chi
PS I
psaA
1-2
1
1
-
40
34 bp duplication
(GATAC…AATTA), +
461 to + 494 atpB,
truncated AtpB
21 bp deletion
(AAAAG…ACGTC),
+2404 to + 2424 ycf1 (tic
214), 21 bp duplication
(TCAAA…CGAAA),
+2486 to + 2406 ycf1
(tic214), 3 aa exchanges
in the variable part of
Ycf1 (Tic214)3)
4 bp duplication (ATTT),
+ 10 to + 13 ccsA,
truncated CcsA
5 bp duplication
(CCGCT), + 1420 to +
1424 psaA, truncated
PsaA
Table 1. (continued)
Mutant1)
Defect in
Gene or
locus
Leaf phenotype2)
Young
Medium
Old
Description
M23
I-psi
PS I
ycf3 (intron)
1-2
2
2
-
M24
I-omega
PS I
psaA
1-2
1
1
-
M25
I-do
Cytb6f
ccsA
3
2
2
-
M26/M27
I-re/I-mi
Cytb6f
petA
2-3
1-2
1-2
-
M28
II-alpha
M29
III-epsilon
M304)
M31
PS I
5)
ycf4
3
2-3
1
PS I
ycf3
2
1
1
II-gamma
PS II
psbE
4
2-3
1-2
II-delta
-
ndhE-psaC
spacer
4
4
4
41
old leaves of field
grown plants with
characteristic
yellowish/whitish tip
old leaves often with
small green spots
no change during
development
Mutation and predicted
molecular
consequence
5 bp duplication
(ATTGA), + 316 to + 320
ycf3 (intron), splicing
deficiency
4 bp duplication (TCTT),
+ 2226 to + 2229 psaA,
changed/extended PsaA
6 bp deletion (ACAATA),
+ 298 to + 303 ccsA,
deletion of 2 aa in CcsA
1 bp insertion (G), + 667
petA, truncated PetA
4 bp duplication (AAAA),
+ 442 to + 445 ycf4;
truncated Ycf4
29 bp duplication
(ACAAG…GCTTT), +
996 to + 1024 ycf3,
truncated Ycf3
5 bp duplication
(ATTAT), + 42 to + 46
psbE, truncated PsbE
24 bp deletion
(TATAT…AATCA), - 60
to - 83 psaC, transcript
processing affected
Table 1. (continued)
Mutant1)
M32
M33
II-epsilon
II-zeta
Defect in
Gene or
locus
PS I
psaA
PS II
psbB
Leaf phenotype2)
Description
Young
Medium
Old
2
1
1
-
1
old leaves with
enhanced greening
close to vascular
tissue
4
2-3
M34
II-eta
Cytb6f
petB
4
3
1
old leaves often with
small green spots
M354)
II-theta
Cytb6f
petB
(intron)
4
3
1
old leaves often with
small green spots
M36/M47
II-iota/
IV-delta
PS I
psaA
2
2
2
-
M37
II-kappa
PS I
ycf3 (intron)
old leaves in parts
darker than younger
leaves, viable on soil
at homoplasmic
state
M37
II-kappa
-
ndhF-rpl32
spacer
M38
II-lambda
Cytb6f
petA
3-4
3-4
4
4
3-4
2-3
42
-
Mutation and predicted
molecular
consequence
4 bp duplication (TGAG),
+ 124 to + 127 psaA,
truncated PsaA
4 bp duplication (CTTT),
+ 117 to + 120 psbB,
truncated PsbB
5 bp duplication
(ATTAC), + 1346 to +
1350 petB, truncated
PetB, transcript
processing/stability
affected
2 bp deletion (AC), + 430
to + 431 petB (intron),
splicing deficiency
5 bp duplication
(TTAAC), + 667 to + 671
psaA, truncated PsaA
transversion G/T, + 1438
ycf3 (intron), splicing
deficiency
single bp
insertion/deletion, results
in a transversion (C/T), 511 rpl323)
39 bp deletion
(GGTTT...CCGAG), +
337 to + 375 petA, 13 aa
deleted in PetA
Table 1. (continued)
Mutant1)
M39
II-my
Defect in
Gene or
locus
Cytb6f
petA
M39
II-my
-
ndhG-ndhI
spacer
M40
II-ny
PS I
ycf4
M40
II-ny
PS II
psbB
M417)
III-alpha
PS II
psbD
M42
M43
III-beta
III-gamma
PS I
PS I
M43
III-gamma
-
M44
III-delta
M44
III-delta
PS I
ATP
synthase
psaB
Leaf phenotype2)
Description
Young
Medium
Old
3-4
3-4
1-2
-
2
1-2
1
-
4
3-4
1
-
1-3
1-3
1-3
bleaches under
higher light
conditions,
sometimes
inhomogeneous
psaA
1-28)
1-2
1
-
1
1
1
-
trnQ-UUGaccD
spacer
psaB
atpB
43
Mutation and predicted
molecular
consequence
5 bp duplication
(TAGCC), + 352 to + 356
petA, truncated PetA
single bp deletion (A), 310 ndhG3)
mutation of M28 (IIalpha)
2 bp deletion (CT), + 791
to + 792 psbB, truncated
PsbB6)
6 bp duplication
(TATATT), + 540 to +
545 psbD, 2 additional aa
in PsbD
5 bp duplication
(ATAAA), + 1246 to +
1250 psaB, truncated
PsaB
8 bp duplication
(AATTTAGT), + 1540 to
+ 1547 psaA, truncated
PsaA
12 bp deletion
(CTCGGCTGATGA), 689 to - 678 accD3)
mutation of M42 (III-beta)
1 bp deletion (A), + 57
atpB, truncated AtpB6)
Table 1. (continued)
Mutant1)
Gene or
locus
Leaf phenotype2)
Young
Medium
Old
Description
M45
IV-alpha
PS I
psaA
3
1-2
1
-
M464)
IV-beta
RuBisCO
rbcL
4
4
3-4
-
M48
IV-epsilon
PS II
psbA
3
2-3
1-2
greening declining
from leaf base to tip
V-alpha
RuBisCO9)
(tentative)
rbcL9)
(tentative)
M49
M50
1222
1223
1224
Defect in
V-delta
PS I
4
4
4
grayish green
phenotype, older
leaves in parts
darker
ycf3
M50
V-delta
-
accD
M51
I-fa
Cytb6f
ccsA
4
3
1
-
3
2-3
1-2
-
1)
Mutation and predicted
molecular
consequence
5 bp duplication
(GTACT), + 2173 to +
2177 psaA, truncated
PsaA
transversion (G/C), + 337
rbcL, single aa exchange
8 bp deletion
(AGTGGGAA), + 389 to
+ 396 psbA, truncated
PsbA
6 bp deletion (GATGAA),
+ 750 to + 755 rbcL,
deletion of 2 aa in RbcL
1,364 bp deletion in ycf3
affecting 11 bp of the 3'end of exon 1, the full
intron 1, the full exon 2,
and 403 bp of the 5'-end
of intron 2
153 bp deletion +208 to +
360 accD, deletion of 51
aa in the variable part of
AccD3)
5 bp duplication
(TTAAC), + 702 to + 705
ccsA, truncated CcsA
Mutants are designated according to their basic plastome genotype (I – V) and Greek letters indicating the sequence of mutant
isolation (with alpha representing the first mutant isolated). In the case of plastome I, for which more mutants have been isolated than
44
1225
1226
1227
1228
1229
1230
1231
1232
1233
1234
1235
1236
1237
1238
1239
1240
1241
1242
1243
1244
1245
1246
1247
1248
1249
1250
1251
1252
letters exist in the Greek alphabet, additional mutants are designated by the seven tone syllables. For details, see Kutzelnigg and
Stubbe (1974), and Stubbe (1989).
2)
1 = nearly white, 2 = yellowish cream-colored, 3 = light green, 4 = nearly green
3)
Potentially phenotypically neutral second site mutation. For details, see text.
4)
Mutations in the mutants M18 (I-sigma), M30 (II-gamma), M35 (II-theta) and M46 (IV-beta) were determined previously and could
be verified in this study. For summary and references, see Greiner (2012).
5)
This mutant was originally obtained under the name II-beta, but re-designated in this work. For details, see Supplemental Table 1.
6)
Second site mutation isolated based on a phenotypic change in the primary mutant. For details, see text.
7)
With respect to its reference plastome (deserens de Vries; GenBank accession number: KT881172.1), the mutant M41 (III-alpha)
carries a 6 bp deletion (TAATCA) in the rbcL-atpB intergenic spacer, - 113 to - 118 of rbcL. However, this mutation must have arisen
in the wild type reference after the isolation of M41 (III-alpha). Hence, it is not treated as second site mutation in M41 (III-alpha). For
details, see text.
8)
The phenotype of the mutant M43 (III-gamma) was described as “lighter green (3)” in young tissue by W. Stubbe. However, in our
hands the mutant appeared to be nearly white to yellowish (1-2) at most developmental stages. It is possible that this phenotypic
change is due to a second site mutation upstream of accD which was not present when W. Stubbe initially assessed the phenotypes.
For details, see text.
9)
The corresponding wild-type strain (argillicola Douthat 4b) of the mutant M50 (V-alpha) is no longer present in our genetic stocks.
For mutant allele identification, the two available O. argillicola plastomes (EU262887.2 and KT881177.1) were used. See Stubbe and
Herrmann (1982), for assignment of this mutant to a physiological defect in RuBisCO.
45
1253
1254
Figure Legends
1255
1256
Figure 1. Representative phenotypes in the Oenothera chloroplast mutant collection.
1257
For details, see text and Table 1. (A) I-ny, variegated individual with completed sorting-
1258
out giving raise to homoplasmic white (mutated) and green (nursery plastome) sectors.
1259
(B) Developmental series of homoplasmic IV-epsilon leaves, showing age-dependent
1260
bleaching. (C) Age-dependent greening and mottled pattern of leaves of mutant I-iota.
1261
From left to right: homoplasmic young nearly white leaf, homoplasmic medium-age
1262
mottled leaves and older green leaf. Note that the oldest leaf displays variegation. As a
1263
result of a completed sorting-out, the leaf blade is homoplasmic for the nearly re-
1264
greened I-iota plastome. The margin is homoplasmic for the wild type chloroplast
1265
(indicated by the arrow). (D) I-tau, growing photoautotrophically in soil as homoplasmic
1266
plant. It displays a reversible bleaching (virescent) phenotype, alternatingly forming
1267
bleached and green leaves. The homoplasmic state of all mutants was confirmed by
1268
Illumina deep sequencing. For molecular causes of the mutations, see Table 1. Scale
1269
bars: 4 cm for (A), (B) and (D); leaf sizes in (C) from left to right: ~5 cm, 10 cm, 12 cm
1270
and 20 cm.
1271
1272
Figure 2. Distribution of the mutated alleles of the plastome mutant collection across
1273
the Oenothera plastome. The physical map of the plastome is shown and the origins of
1274
replication (oriA and oriB), the GC content (grey inner circle) and the macro repeats (red
1275
inner circle) are indicated. Blue circles denote mutations conferring a mutant phenotype;
1276
orange circles denote second site mutations which may be phenotypically neutral.
1277
Green circles refer to phenotypically neutral mutations in the closely related O.
1278
glazioviana and O. elata strains. Note clustering of these mutations in the highly
1279
repetitive regions around accD and ycf1 (tic214). Numbers refer to the number of
1280
individual mutations identified for a gene or locus. Class I genes are indicated in color,
1281
Class II genes and Class III genes in grey or white. For details on the definition of the
1282
gene classes, see text. For macro repeat identification, see Greiner et al. (2008a), for
1283
location of the origins of replication, see Chiu and Sears (1992).
46
1284
1285
Figure 3. Distribution of mutations in the mutational hotspots psaA, psbA and ccsA.
1286
Note that solely duplications could be identified in psaA. Also note the uneven
1287
distribution of mutations within the genes. For molecular causes of the mutations and
1288
associated repetitive elements, see Table 1, Supplemental Data Set 1, and
1289
Supplemental Figures 2 - 4.
1290
1291
Figure 4: Defects in mRNA maturation due to mutations in introns, a coding region, or
1292
an intergenic region in the chloroplast mutants I-psi, II-kappa, II-eta, II-theta and II-delta
1293
(A). Schematic overview of the ycf3, psaC and petB regions of the plastome. Positions
1294
of the mutations are marked with stars, hybridization probes are indicated as green
1295
bars. (B) RNA gel blot analyses of mRNA accumulation and RNA processing patterns.
1296
Upper row, left and middle panel: Splicing defects in ycf3 as found in the mutants I-psi
1297
and II-kappa, respectively. In I-psi, a 5 bp duplication in intron 1 leads to loss of splicing
1298
of this intron, correlating with a severe bleaching phenotype (Table 1). In II-kappa, a
1299
transversion in intron 2 results in a reduced splicing activity. Note that some mature
1300
transcript (0.7 kb) is still produced, consistent with the homoplasmic mutant being viable
1301
upon cultivation in soil (Table 1). Upper row, right panel: Processing defect in the psaC
1302
operon of the mutant II-delta. The mutant harbors a 24-bp deletion in the psaC-ndhE
1303
intergenic spacer. Whereas a precursor transcript of about 8.0 kb overaccumulates in
1304
the mutant, accumulation of processed mRNA species is strongly reduced. Some
1305
mature monocistronic mRNA (0.6 kb) is produced in the mutant, consistent with its mild
1306
phenotype (Table 1). Lower row: Splicing and processing defects in transcripts from the
1307
psbB operon (containing the petB gene) in the mutants II-eta and II-theta. A 2-bp
1308
deletion is present in the petB intron in II-theta. In II-eta, a 5 bp duplication is present in
1309
the second exon of petB. Left panel: hybridization to a petB exon probe, right panel:
1310
intron probe. In II-theta, a petB splicing defect is clearly visible. Transcripts
1311
overaccumulating in the mutant (i.e., the 2.2 kb and 1.5 kb RNA species) likely
1312
represent the unspliced petB/petD dicistron and the unspliced petB, respectively. The
1313
putative mature transcript (0.8 kb) does not accumulate in the mutant, in agreement with
1314
its severe phenotype (Table 1). Interestingly, in II-eta, the mutation in exon 2 leads to
47
1315
loss of the spliced psbH/petB/petD tricistron (1.7 kb in the WT; Westhoff and Herrmann,
1316
1988), and instead to accumulation of a 1.4 kb transcript likely containing the spliced
1317
petB/petD dicistron. The mature transcript (0.8 kb) seems to accumulate which,
1318
however, carries a frameshift. This leads to a largely identical phenotype as II-theta
1319
(Table 1). For details on the mutations, see Table 1. 1 and 2 represent biological
1320
replicates of all plant lines analyzed here. To confirm equal loading, the ethidium
1321
bromide-stained agarose gel prior to blotting is also shown.
1322
1323
Figure 5. Polymerase slippage during DNA replication at repetitive elements leading to
1324
duplication or deletions. Arrows indicate repeats and pairing homology. (A) Slippage
1325
conferred by misalignment at tandem or direct repeats. If slippage occurs at the nascent
1326
(upper) strand, replication leads to duplication. Insertions are a result of slippage at the
1327
leading (lower) strand. (B) Slippage conferred by a hairpin structure at palindrome or
1328
inverted repeats. Duplications are a result of a hairpin at the nascent (upper) strand. A
1329
hairpin at the leading (lower) strand confers a deletion.
1330
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Spontaneous chloroplast mutants mostly occur by replication slippage and show a biased pattern in
the plastome of Oenothera
Amid Massouh, Julia Schubert, Liliya Yaneva-Roder, Elena S. Ulbricht-Jones, Arkadiusz Zupok, Marc T.
J. Johnson, Stephen Wright, Tommaso Pellizzer, Johanna Sobanski, Ralph Bock and Stephan Greiner
Plant Cell; originally published online April 6, 2016;
DOI 10.1105/tpc.15.00879
This information is current as of July 31, 2017
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