Control of plastid inheritance by environmental and genetic factors

Chung, Kin Pan; Gonzalez-Duran, Enrique; Ruf, Stephanie; Endries, Pierre; Bock, Ralph

doi:10.1038/s41477-022-01323-7

Control of plastid inheritance by environmental and genetic factors

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Published: 16 January 2023

Volume 9, pages 68–80, (2023)
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Control of plastid inheritance by environmental and genetic factors

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Abstract

The genomes of cytoplasmic organelles (mitochondria and plastids) are maternally inherited in most eukaryotes, thus excluding organellar genomes from the benefits of sexual reproduction and recombination. The mechanisms underlying maternal inheritance are largely unknown. Here we demonstrate that two independently acting mechanisms ensure maternal inheritance of the plastid (chloroplast) genome. Conducting large-scale genetic screens for paternal plastid transmission, we discovered that mild chilling stress during male gametogenesis leads to increased entry of paternal plastids into sperm cells and strongly increased paternal plastid transmission. We further show that the inheritance of paternal plastid genomes is controlled by the activity of a genome-degrading exonuclease during pollen maturation. Our data reveal that (1) maternal inheritance breaks down under specific environmental conditions, (2) an organelle exclusion mechanism and a genome degradation mechanism act in concert to prevent paternal transmission of plastid genes and (3) plastid inheritance is determined by complex gene–environment interactions.

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Main

Cytoplasmic genomes are maternally inherited in most eukaryotes^1,2. It is generally believed that the uniparental inheritance of organelles and their genomes makes them asexually reproducing genetic systems^3,4,5. Lack of sexual recombination is expected to lead to the eventual mutational meltdown of organellar genomes, a phenomenon widely known as Muller’s ratchet^6,7,8. This is due to the accumulation of deleterious mutations that cannot be separated from (only rarely occurring) beneficial mutations and can be considered as a case of ‘genetic hitchhiking’⁹. While there must be evolutionary forces that explain the strong prevalence of uniparental inheritance, there must also be compensatory mechanisms that allow organellar genomes to escape mutational meltdown.

In plants, the two organellar genomes (plastids and mitochondria) have lower mutation rates than nuclear genomes^{10,1) were analysed in Model 3 (n_rep.total = 13 harvests, ~2.65 million seedlings; Extended Data Tables 1 and 2). Black horizontal bars show mean rates per genotype/treatment combination. Rates per experimental group were estimated (coloured horizontal lines) with CI95s (coloured boxes). Dashed lines depict the basal plastid paternal transmission (grey) and the theoretical maximum (black). Effect estimates were tested by simultaneous two-tailed Wald z-tests: dpd1 genotype (P = 6.22 × 10⁻³⁵), chilling treatment (P = 2.20 × 10⁻¹²⁶) and the interaction between both factors (P = 3.17 × 10⁻¹⁰) were significant. ***P < 0.001, α = 0.05. f, Visualization of paternal plastid transmission by spectinomycin selection (Experiments 2 and 3; Table 1). Blue arrowheads indicate green sectors (paternal plastids). Insets show magnified examples.}

Full size image

Consistent with previous findings reported in A. thaliana, the tobacco dpd1 mutant does not display a noticeable vegetative growth phenotype. Plant growth, reproductive transition and floral development in the dpd1 mutant were comparable to wild-type plants (Extended Data Fig. 5a). However, a reduction in pollen viability was observed in the dpd1 mutant (Extended Data Fig. 5b,c). Confocal laser-scanning microscopy and quantitative PCR analysis revealed retention of plastid DNA in mature pollen grains of the dpd1 mutant (Fig. 3c,d and Extended Data Fig. 5d). Large-scale inheritance assays showed that the dpd1 mutant displayed greatly elevated levels of paternal plastid transmission into the progeny (Fig. 3e,f and Table 1), thus identifying the rate of plastid genome degradation as a genetic factor determining the mode of plastid inheritance.

Synergistic gene–environment control of plastid transmission

Next, we wanted to examine whether the combined action of genes and environment confers even higher levels of biparental plastid inheritance. To this end, we comparatively assessed the effects of chilling stress on the transmission frequency of paternal plastids in the wild-type and the dpd1 genetic backgrounds. Indeed, pollen development at low temperature in the dpd1 mutant resulted in an approximately tenfold increase in paternal plastid transmission compared with the effect of the dpd1 genotype alone or that of the environment alone (Table 1 and Fig. 3e,f). Paternal plastid transmission reached frequencies between 2.2% and 3.2% (in two independent experiments; Fig. 3e and Table 1). These data show that genes and environment act synergistically in the control of plastid inheritance, and their combined action can result in substantial levels of biparental inheritance (Fig. 3e). It is also noteworthy that the environmental factor low temperature and the genetic factor dpd1 are not entirely independent in their effects on plastid inheritance, as revealed by the negative interaction between them (Fig. 3e).

Paternally transmitted plastids readily enter the germline

Biparental inheritance events in which the inherited paternal plastids are present in the shoot apical meristem (Extended Data Fig. 2b,c and Fig. 4a,b) are particularly relevant in that from there, they enter the germline (upon transition of the apical meristem into a floral meristem) and become heritable. Therefore, large-scale screens were conducted to quantitatively assess the entry of paternal plastids into shoot and root apical meristems in the wild-type and the dpd1 genetic backgrounds under ambient temperature or chilling stress. Presence of paternal plastids was evidenced by GFP fluorescence in the meristem (Extended Data Fig. 2b,c) and continued plant growth in the presence of spectinomycin (Fig. 4a). The presence of paternal plastids in the shoot apical meristem was observed at high frequency, in almost half of the biparental transmission events in dpd1 under chilling stress (cf. Table 1 and Fig. 4b).

**Fig. 4: Transmission of paternal plastids into apical meristems and their inheritance in the next generation.**

To determine whether the inherited paternal plastid genomes can be stably maintained across generations, plants with paternal plastids present in the shoot apical meristem were grown in the greenhouse and self-pollinated for seed production. Uniform resistance of the progeny to spectinomycin (Fig. 4c) demonstrated homoplasmy of the (paternally acquired) plastid genome, which is maternally inherited into the next generation in the absence of stress. Together, these data show that paternally inherited plastids readily enter the germline and are passed on to the next generation, suggesting that the discovered phenomenon has evolutionary significance for adaptation⁴³ and, potentially, speciation⁴⁴.

Discussion

In the course of this work, we have discovered two distinct factors that jointly determine plastid inheritance in plants. While the environmental factor temperature affects plastid distribution during the asymmetric cell division occurring in PMI, the genetic factor DPD1 affects organellar genome degradation during male gametogenesis (Fig. 5).

**Fig. 5: Mechanistic model of uniparental versus biparental plastid inheritance.**

Surprisingly, the possibility that plastid inheritance is influenced by the environment has not been considered previously. Here we discovered that maternal inheritance breaks down when male gametogenesis occurs at low temperature. We identified the underlying mechanism by showing that chilling promotes inclusion of paternal plastids in the GC. Chilling stress has been shown to result in destabilization of the cytoskeleton during gametogenesis⁴⁵. The occurrence of altered cell division patterns in PMI upon plant exposure to low temperatures (Extended Data Fig. 3b) suggests that compromised cytoskeletal function is responsible for incomplete plastid exclusion from the GC.

The second elimination mechanism uncovered by our work acts at the level of organellar genome stability. The exonuclease DPD1 degrades organellar genomes in maturing pollen, thus providing a fail-safe mechanism that genetically inactivates those plastids that escaped the organelle exclusion mechanism operating in PMI. Consequently, when both mechanisms (plastid exclusion and genome degradation) are compromised, substantial levels of biparental inheritance ensue (Figs. 3e and 5). Together, our findings suggest that plastid inheritance is a complex trait that is affected by both genetic and environmental factors.

Although most organisms inherit their plastids and/or mitochondria maternally, biparental inheritance has evolved multiple times independently in both animals and plants⁴⁶. It will be interesting to investigate species that exhibit biparental inheritance of plastids and/or mitochondria, and determine the expression patterns of genes for cytoskeletal proteins (especially those that have been implicated in organelle movement^47,48,49) and the activity of organelle-targeted nucleases during male gametogenesis. DPD1 homologues are present in both angiosperms and gymnosperms⁴², and the cytoskeletal components are also generally highly conserved. These candidate genes and their expression patterns could also explain the variation in the rates of biparental plastid inheritance in natural populations²⁰, and the switches in inheritance modes seen over evolutionary timescales².

Given the high conservation of the male gametogenesis programme in seed plants, we speculate that the mechanisms identified in our work are likely to be relevant to all seed plant species. However, it is important to note that the mechanisms reported here are primarily operating during PMI. It seems possible that maternal factors and/or post-fertilization mechanisms also contribute to the control of cytoplasmic inheritance in plants.

Our work also provides simple methods to alter organellar inheritance patterns. In the light of our findings, shifts to biparental inheritance can be achieved either temporarily (by applying chilling stress) or permanently (by genetic interventions, for example, DPD1 gene editing). Changing the mode of inheritance of organelles has important applications in plant breeding. In most crops, both plastids and mitochondria are maternally inherited, making it notoriously difficult to separate the influence of the plastid from that of the mitochondrial genome on important phenotypic traits such as growth, yield and stress tolerance^23,43,50,51. Induction of biparental transmission will allow separation of plastid from mitochondrial effects by taking advantage of the random segregation and sorting of the two organelle types in the progeny.

Finally, our finding that mild chilling stress triggers substantial rates of biparental plastid inheritance has far-reaching consequences for our understanding of organellar genome evolution. Low temperatures (10 °C) are ubiquitous in nature, hence our findings provide a possibility for organelles to participate frequently in sexual reproduction. Assuming that our findings reported here extend to mitochondrial inheritance, the paternally transmitted mitochondria would be able to fuse with their maternal counterparts, leading to genome recombination^52,53. This could efficiently counteract Muller’s ratchet by generating new variation in mitochondrial genomes and facilitating the combination of favourable mutations.

In contrast to plant mitochondria, fusion and genome recombination in plastids appear to be rare^17,18,19. However, temperature-induced biparental inheritance also provides a mechanism for plastids to spread through pollen between species and populations, a phenomenon known as chloroplast capture^22,54. Interestingly, accumulating evidence suggests that the plastid genome plays roles in tolerance to low temperatures^55,56, thus potentially providing a direct link between chilling-induced biparental plastid inheritance, selection and adaptation.

In summary, our data reported here demonstrate that plastid inheritance is controlled at two distinct steps in male gametogenesis and determined by synergistic gene–environment interaction. Our finding that uniparental inheritance of plastids breaks down under conditions of mild environmental stress indicates that organellar genomes experience episodes of biparental inheritance, and casts considerable doubt on the long-held tenet that organelles are asexual genetic systems.

Methods

Plant material and growth conditions

Tobacco plants (Nicotiana tabacum cv. Petit Havana) were grown in standard greenhouse conditions at ~300 μE m⁻² s⁻¹ light intensity under a 16 h light/8 h dark regime (day temperature: ~25 °C, night temperature: ~20 °C). Transplastomic lines (with a wild-type nuclear background; WT^ptGFP) used as pollen donor harbour a gfp expression cassette and the selectable marker aadA encoding an enzyme conferring spectinomycin resistance³¹. Transplastomic lines (WT^ptDsRed) used for confocal microscopic studies harbour a DsRed expression cassette (driven by the ribosomal RNA operon promoter and a chimeric 5′ untranslated region from the chloroplast psbA and clpP genes) and the spectinomycin resistance marker aadA⁵⁷. Seed germination and plant cultivation in vitro were performed on agar-solidified synthetic medium containing 3% (w/v) sucrose⁵⁸. Abiotic stress treatments were performed in controlled-environment chambers.

Crossing experiments and screening for paternal plastid transmission

Homoplasmic transplastomic plants were raised under standard greenhouse conditions and transferred to the designated abiotic stress condition in controlled-environment chambers upon onset of flower development (Fig. 1a). To ensure that male gametophyte development occurred under stress, flower buds ≥10 mm in length were removed upon transfer (Extended Data Fig. 1). High light stress was applied by shifting plants to 1,000 μE m⁻² s⁻¹ light intensity. Drought stress was applied by withholding water until visible loss of turgor and wilting occurred. Heat stress was applied by shifting plants to 35 °C, and chilling stress was performed by shifting plants to 10 °C. Upon pollen maturation, large-scale crosses were conducted by manual pollination³¹, using pollen from transplastomic plants (WT^ptGFP or T₁ dpd1^ptGFP) grown in standard greenhouse conditions or under abiotic stress. Greenhouse-grown plants with wild-type plastids were used as maternal recipients: male-sterile Nt-nms plants in Experiment 1³¹, and emasculated wild-type plants in Experiments 2 and 3.

The resulting progeny was assayed for paternal plastid transmission by germinating seeds on synthetic medium in the presence of spectinomycin (500 µg ml⁻¹), at a density of approximately 500 seedlings per 180-mm-diameter Petri dish or 120 mm square plate. Seedling phenotypes were analysed by visual screening under a stereomicroscope (Zeiss), followed by inspection of detected green (antibiotic-resistant) sectors by UV and confocal microscopy to test for GFP fluorescence. Paternal plastid inheritance (PPI) lines were transferred to soil, grown to maturity in the greenhouse and selfed for seed production. Seeds were assayed for stable paternal plastid inheritance by germination on agar-solidified synthetic medium with spectinomycin (500 µg ml⁻¹).

Tissue culture and plant regeneration

To regenerate plants that contain paternal plastids, green (spectinomycin-resistant) sectors were excised from cotyledons or primary leaves and regenerated on agar-solidified plant regeneration medium⁵⁸ containing 3% (w/v) sucrose, 0.1 mg ml⁻¹ 1-naphtaleneacetic acid (NAA), 1.0 mg ml⁻¹ 6-benzylaminopurine (BAP) and 500 µg ml⁻¹ spectinomycin. To eliminate spontaneous spectinomycin-resistant mutants, tissue samples were exposed to double selection on medium containing spectinomycin and streptomycin (500 µg ml⁻¹ each). Spontaneous spectinomycin-resistant cells bleach out on this medium, whereas cells with aadA-expressing transgenic plastids remain green and continue to grow^59,60. To obtain seeds, regenerated homoplasmic lines with paternal plastids were rooted and propagated on synthetic medium in the presence of spectinomycin (500 µg ml⁻¹).

Light microscopy, UV microscopy and confocal laser-scanning microscopy

Green sectors potentially harbouring paternal plastids were identified in germinating seedlings by visual inspection and/or light microscopy using a stereomicroscope (Stemmi 2000-C; Zeiss). GFP fluorescence was detected with an MZ FLIII fluorescence stereomicroscope (Leica) using filters GFP2 (excitation filter: BP 480/40 nm, barrier filter: LP 510 nm) and GFP3 (excitation filter: BP 470/40 nm, barrier filter: BP 525/50 nm). Subcellular localization of GFP fluorescence was determined by UV microscopy with an Axioskop 2 (Zeiss; excitation filter: BP 450/90 nm, barrier filter: BP 515/65 nm), or by confocal laser-scanning microscopy (TCS SP8; Leica) using an argon laser for excitation (at 488 nm), a 500–510 nm filter for detection of GFP fluorescence and a 610–700 nm filter for detection of chlorophyll fluorescence. Subcellular localization of DsRed fluorescence was determined by confocal laser-scanning microscopy (TCS SP8; Leica) using a diode-pumped solid-state laser for excitation (at 561 nm), and a 575–605 nm filter for detection of DsRed fluorescence.

Aniline blue staining

To visualize the transient CCW, EBP grains were extracted from flower buds with size of 15–18 mm (plants grown at 20–25 °C) or 32–35 mm (plants grown at 10 °C). The CCW was stained by decolorized aniline blue solution (0.1% (w/v) aniline blue (Sigma) in 0.1 M K₃PO₄ (pH 11.0)) at room temperature for 10 min. Stained samples were imaged by confocal laser-scanning microscopy (TCS SP8; Leica) using an argon laser for excitation at 405 nm, and a 494–544 nm filter for detection of fluorescence emission. Z-stack imaging was performed with a Leica TCS SP8 using the Galvo flow system and optimized settings with a step size of ~0.34 µm.

SYTO11 staining of in vitro germinating pollen tubes

To visualize the nuclear DNA of the generative cell within pollen tubes, two to three anthers were detached from flowers at anthesis and collected in a 2 ml Eppendorf tube. Mature pollen grains were released from the stamen by vortexing. Subsequently, the stamens were discarded and 100 µl pollen germination buffer (1.6 mM boric acid, 3.0 mM calcium nitrate, 1.0 mM potassium nitrate, 0.8 mM magnesium sulfate, 10% sucrose, pH 7.4) supplemented with 1 µM SYTO11 stain (Thermo Fisher) were added to the pollen. Incubation was done in the dark at room temperature for 120–180 min. Stained pollen tubes were imaged by confocal laser-scanning microscopy (TCS SP8; Leica) using a 488 nm argon laser for excitation and a 500–550 nm filter for detection of the SYTO11 signal.

Pollen viability assay

To determine the effect of temperature on pollen viability, anthers were detached from flowers at anthesis and collected in a 5 ml Eppendorf tube. Pollen grains were harvested by vortexing with pollen harvesting buffer (100 mM Na₃PO₄ (pH 7.0) and 1 mM Na₂EDTA (pH 8.0)), followed by centrifugation at 1,000 × g for 5 min. The pollen pellet was then resuspended and stained in pollen viability solution (100 mM Na₃PO₄ (pH 7.0), 1 mM Na₂EDTA (pH 8.0), 10 μM propidium iodide (Abcam) and 20 μg ml⁻¹ fluorescein diacetate Sigma) at room temperature for 30 min. Stained samples were imaged by confocal laser-scanning microscopy (TCS SP8; Leica). Fluorescein diacetate was excited with a 488 nm argon laser and the emission signal was collected at 500–550 nm. Propidium iodide was excited with a 561 nm argon laser and its emission signal was collected at 600–650 nm.

DAPI staining

To visualize nuclear and organellar DNA, uninucleate microspores (UNM) and mature pollen grains (MPG) were extracted from flower buds with sizes of approximately 10 mm (for UNM) or from flowers at anthesis (for MPG). DNA was stained with 4,6-diamidino-2-phenylindole (DAPI) staining solution (100 mM Na₃PO₄ (pH 7.0), 0.1% (v/v) Triton X-100, 1 mM Na₂EDTA (pH 8.0) and 1 μg ml⁻¹ DAPI) at room temperature for 30 min. Stained samples were imaged by confocal laser-scanning microscopy (TCS SP8; Leica) using a 405 nm laser diode for excitation and a 430–495 nm filter for detection of fluorescence emission. For the pollen squash method, stained pollen grains were placed on a glass slide and gently squashed by tap** on the cover slip⁴¹. Squashed pollen were imaged by confocal microscopy (Leica TCS SP8), with z-stack imaging using the Galvo flow system and optimized settings with a step size of ~0.30 µm.

NtDPD1 gene identification and sgRNA design for genome editing

The protein sequence of DPD1 from Arabidopsis thaliana (AT5G26940) was used as query in a search against the genomic scaffolds in the available draft genome of N. tabacum⁶¹. Two homologues were identified and named NtDPD1S and NtDPD1T (scaffolds Nitab4.5_0002715 and Nitab4.5_0014337, respectively). The transcript structures of both loci were deduced from the sequences present in the associated transcriptome (Nitab4.5_0002715g0070.1 and Nitab4.5_0014337g0020.1). Both NtDPD1 genomic sequences were divided into 2 kb fragments and used as input for sgRNA generation using CRISPOR⁶². A pair of sgRNAs was chosen for NtDPD1 gene editing (sgRNA L1: 5′-GTCCAGACATTAGTCAATCC...-3′; sgRNA R1: 5′- GATGCCCTTCAATAAGAGAA...-3′, with nucleotides 2–20 corresponding to the target sequence). The targeted sequences are fully conserved in both NtDPD1 loci.

Construction of a plant transformation vector for genome editing of NtDPD1

Plasmid pEG001 was assembled for constitutive expression of cas9 in plant cells. pEG001 is a derivative of plasmid pJF1031 described previously⁶³. The backbone of pJF1031 was excised by digestion with the restriction enzymes SpeI and XbaI and purification of the ~14.5 kb fragment obtained, resulting in the removal of the promoter driving cas9. The ~1.5 kb promoter of the HPL gene of A. thaliana (hydroxyperoxide lyase, AT4G15440) was amplified from the pORE E2 plasmid⁶⁴ using primers oEG126 and oEG127 (Extended Data Table 4), and integrated into the pJF1031 backbone through a Gibson assembly reaction⁶⁵ (New England Biolabs), resulting in plasmid pEG001. For gene editing at the NtDPD1 loci, vector pEG037 was constructed. Primers oEG315 and oEG320 (Extended Data Table 4) were used to amplify a fragment of plasmid pJF1046⁶³ containing part of an sgRNA scaffold, the U6 promoter and the U6 terminator. The primers introduce the targeting sequences of sgRNA L or R, respectively, as well as BsaI sites at both ends of the fragment. The PCR product was cloned into pEG001 through a simultaneous BsaI restriction and ligation reaction⁶⁶. The resulting binary vector pEG037 for mutagenesis of NtDPD1 contains a hygromycin resistance marker for selection in planta and a kanamycin marker for selection in bacteria. Sequences of all oligonucleotides used in this study are provided in Extended Data Table 4.

Generation of a quadruple dpd1 knockout line

Transplastomic plants with a wild-type nuclear background (WT^ptGFP) were supertransformed with Agrobacterium tumefaciens strain GV2260 harbouring vector pEG037 using the leaf disc infiltration method. Selected hygromycin-resistant plant lines were maintained in medium containing hygromycin (15 mg l⁻¹) and cefotaxime (250 mg l⁻¹) for continued selection of transgenic plant cells and elimination of residual Agrobacterium cells, respectively. Extended Data Fig. 3 illustrates the PCR-based genoty** (Reactions 1–3) of transgenic lines and the screening procedure that led to the isolation of the double dpd1 mutant (tetra-allelic). Due to the allotetraploidy of the N. tabacum genome, the screening process was tailored to identify plants with four knockout DPD1 homeoalleles in the diploid state (that is, two S and two T alleles; Fig. 4a). PCR and sequencing were employed to identify plants with somatic Cas9 activity and desirable knockout mutations in dpd1 (Extended Data Fig. 4). Additional regeneration steps in tissue culture helped with purification of desired mutation events and reduction of the allele complexity arising from constitutive Cas9 expression.

Polymorphisms between the S and T loci allowed classification of the amplified sequences as S or T homeoalleles after sequencing (Fig. 3a,b and Extended Data Fig. 4). Reaction 1 (Extended Data Fig. 4b) generates a short product only when a large deletion was generated between targeting sites L and R. Presence of this product implies that the plant line possesses an active Cas9, and at least one mutant allele can confidently be identified by this PCR. The primers used in Reaction 2 (Extended Data Fig. 4b) flank the site targeted by sgRNA L. This reaction was performed to identify the other three mutant alleles by direct Sanger sequencing of amplified PCR products from ~320 T₀ samples analysed. Occurrence of small InDels often caused Reaction 2 to produce electropherograms showing several overlap** peak profiles. Whenever profiles of up to three overlap** sequences were obtained, deconvolution of sequence strings was attempted by visual inspection. At the L site, Cas9 frequently caused small InDels (from −4 to +1 bp) and produced recognizable shifted peak patterns compared to the known S and T wild-type sequences, which could be deconvoluted even when three sequences were overlap**. After three rounds of PCR screening and two rounds of regeneration in tissue culture, a double dpd1 mutant plant line was isolated (for simplicity, referred to as dpd1). Sanger sequencing confirmed that dpd1 possesses four distinct mutant alleles at the L site (Fig. 3a,b and Extended Data Fig. 4). One of the sequences was a large deletion, and the three other sequences were frameshift mutations.

The T₀ dpd1 plant was transferred to the greenhouse where T₁ seeds were obtained by selfing. Presence of the mutations at the L site was confirmed by genoty** T₁ seedlings (performing Reactions 1 and 2; Extended Data Fig. 4b). Allele segregation in the T₁ generation facilitated the individual sequencing of the four alleles after PCR and gel purification, as Reaction 2 yields products of different size for the S and T loci. Mutations at the sgRNA R site were analysed in T₁ seedlings by Reaction 3 (Extended Data Fig. 4b), applying visual deconvolution when necessary. The linkage between mutations in the L and R target sites was established by observing co-segregation of genotyped alleles in the T₁ generation (23 seedlings analysed). No other alleles were identified in the final dpd1 line, suggesting that the four mutant alleles detected in the T₀ generation had become fixed.

Isolation of nucleic acids and PCR

For DNA gel blot analysis, total plant DNA was isolated using a cetyltrimethylammoniumbromide (CTAB)-based protocol⁶⁷. Extracted DNA samples were digested with the restriction enzymes XhoI and EcoRV, separated by gel electrophoresis in 0.8% agarose gels and blotted onto Hybond N nylon membranes (GE Healthcare) using standard protocols. For hybridization, α[³²P]dCTP-labelled probes were generated by random priming (Multiprime DNA labelling kit, GE Healthcare). A PCR product covering part of the 16S rRNA gene (amplified by primers P16Srrn-F and P16Srrn-R, and purified by agarose gel electrophoresis) was used as probe for RFLP analysis. Hybridizations were carried out at 65–68 °C in rapid hybridization buffer (GE Healthcare) following the manufacturer’s instructions.

For genoty** reactions, genomic DNA was extracted from leaf tissue with the Extract-N-Amp kit (Sigma-Aldrich). One µl was used as template for PCR. For cloning and for the initial PCR-based screening of dpd1 mutations, Phusion DNA polymerase (Thermo Fisher) was used. In all other PCR reactions, DreamTaq DNA polymerase (Thermo Fisher) was used. PCR products were column-purified before sequencing (NucleoSpin Gel and PCR Clean-up, Macherey-Nagel).

Quantification of plastid DNA in pollen

Total DNA (comprising both nuclear and organellar DNA) of enriched pollen fractions was prepared using a published protocol⁴¹. Tobacco pollen grains were collected by rubbing dehiscent anthers from three opened flowers (WT^ptGFP and dpd1^ptGFP plants, respectively) onto the wall of a 1.5 ml Eppendorf tube, followed by addition of 100 µl of distilled water for resuspension. Due to the procedure used for pollen harvest, some level of contamination by other anther cell types cannot be avoided, hence the samples should be considered as enriched pollen preparations (rather than pure pollen fractions). The enriched pollen suspension was incubated at 95 °C for 5 min, centrifuged at 16,000 x g for 5 min and then subjected to real-time PCR to determine the relative amounts of nuclear and plastid DNA. 18S rDNA (nuclear gene) and aadA (transgene present in the plastid genome of WT^ptGFP and dpd1^ptGFP) were used as proxies for nuclear and plastid DNA abundance, respectively. Primers oCK68 and oCK69 were used for 18S rDNA amplification, and primers oKPC579 and oKPC580 for aadA amplification (Extended Data Table 4). Quantitative real-time PCR was conducted using the LightCycler 480 Real-Time PCR system and LightCycler SYBR Green reaction mixtures (Roche). The relative amount of plastid DNA in the enriched pollen fractions was determined by the abundance of the aadA amplicon normalized to the abundance of the 18S rDNA. Relative quantification was performed using the ΔΔCt method⁶⁸.

Modelling and statistics

The quantitative effects of genotype and stress treatments on paternal plastid transmission, plastid inclusion in the generative cell and pollen survival (after chilling stress, or when comparing WT^ptGFP and dpd1^ptGFP mutant) were estimated using generalized linear models. Models were constructed with the maximum-likelihood method in R v3.5.3 (https://www.R-project.org/). Genotype, treatment and experiment (stage of visualization in pollen; independent experiments to determine plastid transmission) were proposed a priori as explanatory variables. Five models were selected for data analysis (Models 1–5).

The datasets for plastid inclusion in the generative cell (for Model 2), pollen survival under chilling stress (Model 4) and pollen survival in the wild-type and the dpd1 genotypes (Model 5) were modelled as proportions of binary outcomes using the binomial distribution. The total amount of pollen grains analysed per unit of replication was provided in the ‘weights’ argument of glm(), as recommended⁶⁹. The two paternal plastid transmission datasets (one representing Experiment 1, the other including Experiments 1, 2 and 3) were modelled using the negative binomial distribution (for Models 1 and 3), after evidence of overdispersion was found in preliminary Poisson versions of these models. The unit of replication in Experiments 1, 2 and 3 is the ‘harvest’: a batch of seeds obtained from a set of maternal recipients (5–30 plants) fertilized by pollen donors from a single treatment (3–8 plants). For modelling of proportions and rates, the count of seedlings with sectors per unit of replication was set as the response variable, while the total amount of seedlings analysed was provided as offset. ‘log’ was used as the link function for all models.

For each dataset, a starting model was generated that includes effect parameters for proposed explanatory variables and interactions. All effect parameters were modelled relative to specific levels of the explanatory variable (‘contrast to treatment’). Models were selected according to minimal AIC (Akaike’s Information Criterion), as they are considered the most parsimonious⁷⁰: a version corrected for small sample sizes (AICc) was used as recommended previously⁷¹. From the starting and following models, parameters were removed sequentially and only if they led to a reduction in AICc. Overdispersion of the preliminary Poisson models was checked by performing a likelihood ratio test (LRT): this change resulted in a large reduction in AICc. The final models with minimal AICc (one per dataset) were designated Models 1–5. Goodness of fit was tested for these models using LRTs comparing (1) each selected model against a ‘saturated’ model, where the test provides a general measure of whether the model fits the data well, or (2) the selected model against a competing model to measure relative goodness of fit. Models 1–5 were confirmed to fit as well as the saturated model. Parameter estimates in the selected models were evaluated with simultaneous Wald’s z-tests (two-tailed). No corrections for multiple comparisons were applied for z-tests.

Negative binomial models were constructed using the glm.nb() function of MASS package v7.3.54⁷². Parameter values, standard errors and Wald z-statistics with P values were retrieved using the insight package v0.2.0 (https://easystats.github.io/insight/), or directly from the glm() and glm.nb() outputs (Extended Data Table 2). Parameters, errors and CI95s (95% confidence intervals) were transformed for the models to be log-linear in base 10. AICc were obtained through the MuMIn package v1.43.6 (https://CRAN.R-project.org/package=MuMIn). Wald CI95s for parameter estimates were calculated using the confint.default() function in the base package. Means fitted by the model were obtained from the glm() output, and CI95s for the estimated means were obtained by using the ciTools package v0.6.1 (https://CRAN.R-project.org/package=ciTools). LRTs for overdispersion were calculated using odTest() from the pscl package v1.5.5 (https://github.com/atahk/pscl/), and the remaining LRTs were performed using the base stats package. R was accessed through R Studio v2022.07.2 Build 576.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Data supporting the findings of this work are available within the paper and its Supplementary Information files. Sequences from Arabidopsis (AT5G26940.1 and AT4G15440) are available through TAIR (https://www.arabidopsis.org/). Genomic sequences from Nicotiana (scaffolds Nitab4.5_0002715 and Nitab4.5_0014337) and transcripts (Nitab4.5_0002715g0070.1 and Nitab4.5_0014337g0020.1) are available at Sol Genomics Network (https://solgenomics.net/). Source data are provided with this paper.

Code availability

The R code used for assembling the statistical models is available on GitHub at https://github.com/egonzalezduran/modelsplastidinheritance.

References

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Acknowledgements

We thank the following MPI-MP team members: C. Hasse, S. Braune, R. Martens, S. Otto, C. Schmidt, S. Grebe, J. Bartetzko, N. Wulff, G. Olar and J. Abaya for help with crosses and seed assays; X. Kroop and A. Schadach for help with tissue culture and hybridization experiments; C. Krämer for providing oligonucleotides for qPCR analysis; F. Kragler for expert help and advice on confocal microscopy; and the MPI-MP GreenTeam for help with plant cultivation and nuclear transformation. This research was financed by the Max Planck Society.

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Open access funding provided by Max Planck Society.

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These authors contributed equally: Kin Pan Chung, Enrique Gonzalez-Duran and Stephanie Ruf.

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Max-Planck-Institut für Molekulare Pflanzenphysiologie, Potsdam-Golm, Germany
Kin Pan Chung, Enrique Gonzalez-Duran, Stephanie Ruf & Ralph Bock
Universität Hamburg, Institut für Pflanzenwissenschaften und Mikrobiologie, Hamburg, Germany
Pierre Endries

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Kin Pan Chung
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Enrique Gonzalez-Duran
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Stephanie Ruf
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Pierre Endries
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Contributions

K.P.C., S.R., E.G.-D. and R.B. designed the research. K.P.C., S.R., E.G.-D. and P.E. performed the experiments. All authors contributed to data analysis and interpretation. R.B. wrote the manuscript, with input from K.P.C., S.R. and E.G.-D. All co-authors commented on the manuscript draft.

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Correspondence to Ralph Bock.

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Extended data

Extended Data Fig. 1 Stress application to develo** microspores.

Abiotic stress was applied at early stages of development of the male gametophyte. Flower buds ≥10 mm in length were removed from WT^ptGFP plants prior to plant transfer to controlled environment chambers for stress application. White arrowheads indicate the developmental window of the flower buds at the time of plant transfer to the stressful condition. Pollen development in these buds (<10 mm long) largely occurs under stress. Scale bar, 10 mm.

Extended Data Fig. 2 Detection of paternal plastid transmission in the absence of selection.

a, Seeds harvested from crossing experiments were germinated in medium without antibiotics. Scale bar, 10 mm. b, Stereomicroscopic images of a seedling containing paternally inherited plastids grown in antibiotic-free medium. The microscopic bright field picture (left), the GFP fluorescence image (middle) and the chlorophyll fluorescence image (right) are shown. Arrowheads 1 and 2 point to two sectors containing paternal plastid as evidenced by GFP fluorescence. Scale bar, 1 mm. c, Visualization of paternal plastids by their antibiotic resistance. The seedling shown in b was transferred to spectinomycin-containing medium that causes bleaching of cells that contain only maternal plastids. Photographs were taken 7 days (left; scale bar, 1 mm), 20 days (middle; scale bar, 1 mm) and 48 days (right; scale bar, 10 mm) after transfer. d, Frequency of paternal plastid transmission observed in the absence of spectinomycin selection.

Extended Data Fig. 3 Effect of chilling stress on floral and pollen development in Nicotiana tabacum.

a, Flowers developed under standard greenhouse conditions (left, 20–25 °C) compared to flowers developed under chilling stress (right, 10 °C). b, Confocal laser-scanning microscopy images of early binucleate pollen (EBP) harvested from WT^ptDsRed plants grown at 10 °C. The callose cell wall separating the generative cell from the vegetative cell was stained with aniline blue to visualize the division pattern. In addition to the normal cell division pattern (left, asymmetric), various defects in cell division (middle and right) are observed in pollen developed at 10 °C. This experiment was repeated four times and representative images are shown. Scale bar, 10 μm. c, Confocal microscopy images of mature pollen grains (MPGs) from wild-type tobacco plants grown under standard greenhouse conditions (left, 20–25 °C) or under chilling stress (right, 10 °C). MPGs were stained with fluorescein diacetate (FDA) and propidium iodide (PI) to determine pollen viability. Viable pollen is evidenced by green fluorescence, because FDA is converted to fluorescein by cytoplasmic esterase activity. Dead pollen displays only PI fluorescence inside the grain, due to absence of FDA derived fluorescence and permeability to PI. This experiment was repeated three times and representative images are shown. Scale bar, 100 μm. d, Quantification of pollen viability. The proportions of viable pollen per biological replicate (imaging session) were modelled using the binomial distribution (Model 4 constructed with n_rep.total = 6, 3 sessions per group, ~3100 pollen grains analysed; see Extended Data Tables 1 and 2). Shaded coloured boxes show the 95% confidence interval (CI95) of the mean estimates. β_chilling represents the effect of the chilling treatment on pollen survival (P = 1.64 × 10⁻¹⁷⁶), expressed as log₁₀ of the fold change (Extended Data Table 2). The parameter estimate was tested by a two-tailed Wald z-test. ***, P < 0.001; α = 0.05.

Extended Data Fig. 4 Generation of dpd1 mutants in the allotetraploid species Nicotiana tabacum.

a, Primer (oEG) binding sites in the NtDPD1 genomic sequence. Boxes represent exons. TSS is the transcription start site according to transcriptome data. Blue and orange areas indicate the protein-coding sequences in the two diploid subgenomes (S: Nicotiana sylvestris; T: Nicotiana tomentosiformis). The conserved exonuclease domain is represented by the darker color. The sequences targeted by CRISPR/Cas9-based genome editing (sgRNA-binding sites; sgRNA L: ‘left’ sgRNA; sgRNA R: ‘right’ sgRNA) are indicated by red vertical bars and black arrowheads. Start and stop codons are also shown. b, PCR reactions used for genoty** of the DPD1 loci. Red marks represent the mutations produced by Cas9 cleavage at the L and R target sites. Large deletions (ΔΔ) are the result of double cleavage and loss of the fragment between the L and R sites. c, Structure of the T-DNA in transformation vector pEG037 used for mutagenesis of NtDPD1. RB and LB are the T-DNA borders. P_U6 and T_U6 are promoter and terminator, respectively, of the U6 snRNA gene from Arabidopsis thaliana. The sgRNAs R and L are shown as red boxes. P_HPL is the promoter from the HPL gene of A. thaliana (AT4G15440). zCas9 is a cas9 gene version that was codon-optimized for Zea mays. T_RbcS, Rubisco small subunit terminator; P_35S, double CaMV 35S promoter; hpt, hygromycin phosphotransferase gene conferring resistance to hygromycin in planta. d, Schematic overview of the generation of dpd1 mutants. After Agrobacterium-mediated transformation with vector pEG037, PCR-based genoty** and additional regeneration cycles in tissue culture were conducted. Somatic expression of Cas9 from the HPL promoter potentially causes different mutations in different parts of the plants. Plants with putative knock-out alleles of DPD1 were identified by PCR genoty** and DNA sequencing, and the additional regeneration rounds were performed to reduce the mosaicism in the mutant lines by obtaining clonal regenerants from single cells. The procedure led to the isolation of plant lines in which all cells contain the same set of mutated DPD1 alleles (dpd1 mutant).

Extended Data Fig. 5 Phenotypic analysis of the tobacco dpd1 mutant.

a,WT^ptGFP and dpd1^ptGFP plants were grown in the greenhouse. Pictures were taken from 6-week-old plants, 9-week-old plants and fully opened flowers. b, Confocal microscopy images of pollen grains from WT^ptGFP and dpd1^ptGFP plants grown under standard greenhouse conditions. Pollen grains were stained with fluorescein diacetate (FDA) and propidium iodide (PI) to determine pollen viability. This experiment was repeated three times and representative images are shown. Scale bar, 100 μm. c, Quantification of pollen viability. The proportions of viable pollen per biological replicate (imaging session) were modelled using the binomial distribution (Model 5; constructed with n_rep.total = 6; 2,228 pollen grains analysed; see Extended Data Tables 1 and 2). Each replicate consisted of a single harvest of pollen from 2–3 flowers of a single plant, which were stained and imaged in the same session (delimited by dashed lines). During data analysis, it was found that there was significant variation between replicates (represented by replicate parameters with P < 0.05 in Extended Data Table 2). To better represent this variation, the graph shows the proportions of viable pollen in the 5–9 image fields (coloured circles) photographed in each replicate. The mean estimates per genotype are indicated with a black horizontal bar. β_dpd1 represents the effect of the dpd1 genotype on pollen survival (P = 3.00 × 10⁻⁹) compared to the wild type at normal temperature conditions (20–25 °C), expressed as log₁₀ of the fold change (Extended Data Table 2). The parameter estimate was tested by a two-tailed Wald z-test. ***, P < 0.001; α = 0.05. d, Confocal images of WT^ptGFP and dpd1^ptGFP mature pollen stained with DAPI. Images were taken after applying the pollen squash method to release the mature pollen from the anthers. Z-stack overlaid images with maximized projection are displayed (total width of z-slices of WT^ptGFP and dpd1^ptGFP is 2.15 μm and 1.78 μm, respectively). Arrowheads indicate the vegetative nucleus (VN) and the generative nucleus (GN). Arrows indicate organellar DNA released from squashed mature dpd1^ptGFP pollen. This experiment was repeated two times and representative images are shown. Scale bar, 10 μm.

Extended Data Table 1 Specification of generalized linear models for analysis of paternal plastid transmission and plastid inclusion

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Extended Data Table 2 Parameter estimates obtained from modeling

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Extended Data Table 3 Proportions of plastid inclusion in the generative cell across imaging experiments

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Extended Data Table 4 Synthetic oligonucleotides used in this study

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Supplementary information

Reporting Summary

Supplementary Video 1

Three-dimensional confocal laser-scanning imaging of early binucleate pollen develo** under standard greenhouse conditions. Early binucleate pollen was harvested from transplastomic WT^ptDsRed plants grown under standard conditions, followed by staining with aniline blue. The callose cell wall (grey) separating the vegetative cell and the newly formed generative cell was visualized by z-stack confocal imaging with optimized settings and a step size of ~0.34 µm (width of the slices: 8.5 µm). No plastids (magenta) are observed within the generative cell, in line with efficient exclusion of paternal plastids from the generative cell during male gametogenesis. This experiment was repeated four times and a representative video is shown.

Supplementary Video 2

Three-dimensional confocal imaging of early binucleate pollen develo** under chilling stress. Early binucleate pollen were harvested from transplastomic WT^ptDsRed plants grown under chilling stress (10 °C), followed by staining with aniline blue. The callose cell wall (grey) separating the vegetative cell and the newly formed generative cell was visualized by z-stack confocal imaging with optimized settings and a step size of ~0.34 µm (width of the slices: 11 µm). A paternal plastid (magenta) is observed within the generative cell, indicating that the plastid exclusion mechanism during male gametogenesis is compromised at low temperature. This experiment was repeated four times and a representative video is shown.

Supplementary Video 3

Time-lapse confocal imaging of early pollen tube development during in vitro germination under chilling stress. Mature pollen grains were harvested from transplastomic WT^ptDsRed plants grown under chilling stress (10 °C), followed by pollen germination in vitro and staining with SYTO11, a nucleic acid-binding dye. The generative cell nucleus (GCN) within the growing pollen tube is visualized by green SYTO11 fluorescence. Paternal plastids (arrowhead, magenta) are closely associated with the GCN and enclosed by the generative cell membrane. Consequently, these plastids display similar movement dynamics as the GCN. By contrast, plastids outside of the generative cell (asterisk) display a much more rapid movement due to intense cytoplasmic streaming in the growing tube. This experiment was repeated ten times and a representative video is shown.

Source data

Source Data Fig. 1

Unprocessed agarose gel and southern blot for assembly of Fig. 1e. Left: agarose gel with markers. Right: samples of the gel transferred to the blot.

Source Data Fig. 2

Ct values and calculations for relative quantification of plastid DNA.

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Chung, K.P., Gonzalez-Duran, E., Ruf, S. et al. Control of plastid inheritance by environmental and genetic factors. Nat. Plants 9, 68–80 (2023). https://doi.org/10.1038/s41477-022-01323-7

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Received: 21 July 2022
Accepted: 26 November 2022
Published: 16 January 2023
Issue Date: January 2023
DOI: https://doi.org/10.1038/s41477-022-01323-7
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Control of plastid inheritance by environmental and genetic factors

Abstract

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Main

Synergistic gene–environment control of plastid transmission

Paternally transmitted plastids readily enter the germline

Discussion

Methods

Plant material and growth conditions

Crossing experiments and screening for paternal plastid transmission

Tissue culture and plant regeneration

Light microscopy, UV microscopy and confocal laser-scanning microscopy

Aniline blue staining

SYTO11 staining of in vitro germinating pollen tubes

Pollen viability assay

DAPI staining

NtDPD1 gene identification and sgRNA design for genome editing

Construction of a plant transformation vector for genome editing of NtDPD1

Generation of a quadruple dpd1 knockout line

Isolation of nucleic acids and PCR

Quantification of plastid DNA in pollen

Modelling and statistics

Reporting summary

Data availability

Code availability

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