specialized plastids trigger tissue-specific signaling for ... · kamaldeep s. virdi, b christian...

672 Plant Physiology®, October 2018, Vol. 178, pp. 672–683, www.plantphysiol.org © 2018 American Society of Plant Biologists. All Rights Reserved.

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Plastids comprise a defining feature of the plant cell, present-day outcomes of an endosymbiotic event that distinguishes the plant kingdom. Arising through fission rather than de novo biogenesis during the development of a plant, plastids undergo specialized differentiations. The transition from proplastid to chloroplast defines tissues that will be photosynthetic within a plant, with amyloplasts occurring in nonphotosynthetic tissues (Enami et al., 2011), chromoplasts within reproduc-tive and senescing organs (Barsan et al., 2012), lipid- containing plastids during flower and seed development (Rottet et al., 2015), and so on. These transitions involve proteome and metabolic changes that, in most cases, are not yet fully defined. Integration of mutational, metabolic, and infrastructural data for plastid devel-opment shows plastid biogenesis to be surprisingly complex (Charuvi et al., 2012; Pogson et al., 2015; van Wijk and Kessler, 2017).

Evidence has accumulated for a class of plastids that is specialized for stress sensing and acclimation. For example, plastids within the epidermis that are ap-proximately 30% the size of a mesophyll chloroplast have been associated with osmotic stress response (Veley et al., 2012) and coordination of auxin induction for stem growth under shade conditions (Procko et al., 2016). Similarly sized plastids within the vascular pa-renchyma have been associated with ABA biosynthe-sis and high-light and drought responses (Endo et al., 2008; Galvez-Valdivieso et al., 2009). These two groups of plastids appear to share at least a portion of their proteomes distinct from mesophyll chloroplasts (Virdi et al., 2016), suggesting that more detailed pro-teome investigations would provide insight into these specializations.

Studies of vascular and epidermal plastids identify MSH1, a plant-specific mitochondrial protein (Abdelnoor et al., 2003; Xu et al., 2011) that is also associated with this specialized plastid type. Spatiotemporal regulation ensures that MSH1 functions in epidermis, vascular parenchyma, and reproductive cells; the GFP-tagged protein localizes to small plastids within these tissues (Virdi et al., 2016). Steady-state MSH1 transcript lev-els decline in plants under abiotic stress (Shedge et al., 2010; Xu et al., 2011), and RNA interference (RNAi) suppression of MSH1 triggers constitutive and wide- ranging stress and developmental pathways in a plant grown under normal conditions (Xu et al., 2012; Virdi et al., 2016). Detailed analysis of msh1 T-DNA mu-tants shows dramatically altered expression of genes controlling abiotic and biotic stress, oxidative and re-dox, phytohormone response, and circadian rhythm

Specialized Plastids Trigger Tissue-Specific Signaling for Systemic Stress Response in Plants1[OPEN]

Jesús Beltrán,a,b Yashitola Wamboldt,b Robersy Sanchez,a Evan W. LaBrant,b Hardik Kundariya,a Kamaldeep S. Virdi,b Christian Elowsky,b and Sally A. Mackenziea,2,3

aDepartments of Biology and Plant Science, The Pennsylvania State University, University Park, Pennsylvania 16802bDepartment of Agronomy and Horticulture, University of Nebraska, Lincoln, Nebraska 68588ORCID IDs: 0000-0002-8095-4757 (J.B.); 0000-0002-5239-1459 (Y.W.); 0000-0002-5246-1453 (R.S.); 0000-0003-0742-9363 (E.W.L.); 0000-0002-4065-3888 (H.K.); 0000-0003-2077-5607 (S.A.M.)

Plastids comprise a complex set of organelles in plants that can undergo distinctive patterns of differentiation and redifferentia-tion during their lifespan. Plastids localized to the epidermis and vascular parenchyma are distinctive in size, structural features, and functions. These plastids are termed “sensory” plastids, and here we show their proteome to be distinct from chloroplasts, with specialized stress-associated features. The distinctive sensory plastid proteome in Arabidopsis (Arabidopsis thaliana) derives from spatiotemporal regulation of nuclear genes encoding plastid-targeted proteins. Perturbation caused by depletion of the sensory plastid-specific protein MutS HOMOLOG1 conditioned local, programmed changes in gene networks controlling chro-matin, stress-related phytohormone, and circadian clock behavior and producing a global, systemic stress response in the plant. We posit that the sensory plastid participates in sensing environmental stress, integrating this sensory function with epigenetic and gene expression circuitry to condition heritable stress memory.

1This work was supported by the Bill and Melinda Gates Founda-tion (OPP1088661).

2Author for contact: [email protected] author.The author responsible for distribution of materials integral to

the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Sally A. Mackenzie ([email protected]).

S.A.M. conceptualized the research; S.A.M, K.S.V., and J.B. de-signed the experiments; J.B., Y.W., R.S., E.W.L., H.K., K.S.V., and C.E. conducted the experiments; S.A.M. and J.B. wrote the manuscript.

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pathways (Shao et al., 2017). The MSH1-RNAi or msh1 mutant phenotype is also characterized by a range in phenotype intensity, plant variegation, variable growth rate, delayed maturity and flowering, altered circa-dian rhythm, altered nonphotoactive redox behavior, and abiotic stress tolerance (Shedge et al., 2010; Xu et al., 2012). Earlier hemicomplementation experiments showed that this integrated and pleiotropic response to MSH1 suppression emanates from the plastid (Xu et al., 2011). These observations imply that the “sensory” plastid, in which MSH1 resides, differentiates in a spa-tially and temporally defined manner and influences a wide range of integrated stress-response and devel-opmental networks.

The global stress phenotype that derives from MSH1 RNAi knockdown is characterized by genome-wide DNA methylome repatterning (Virdi et al., 2015), im-plying an epigenomic component to the phenotype. Transgene-null progeny from these lines can retain the msh1 phenotype, more uniform and slightly attenuated, as heritable “memory” lines (Xu et al., 2012) that display altered methylation (X. Yang, R. Sanchez, H. Kundariya, Y. Wamboldt, J. Barreras, and S. Mackenzie, unpub-lished data). These data provide nascent evidence of epigenetic changes, induced by this unusual plastid type, that are programmed and presumably adaptive for the plant.

Based on observations by our group and others, we postulate that epidermal cells, in direct contact with the external environment, and vascular parenchyma cells, which are able to transmit signals systemically, harbor a plastid type that is specialized for environ-mental sensing. Here, we investigate this putative “sensory plastid” in Arabidopsis (Arabidopsis thaliana), its proteome and stress-responsive behavior relative to neighboring mesophyll chloroplasts, and show the sensory plastid and chloroplast proteomes to be over-lapping but distinct. Many stress-related proteins ear-lier assigned to the chloroplast appear, in fact, to be localized to sensory plastids. Investigation of the trans-latome within sensory plastid-containing cells, follow-ing MSH1 disruption, provides evidence of calcium signaling, redox, chromatin remodeling, and circadian clock network changes that, together, may account for the strikingly pleiotropic developmental phenotypes and the plant’s capacity for transgenerational stress memory.

RESULTS

Sensory Plastids and Chloroplasts Can Be Differentially Sorted from Arabidopsis Floral Stem Tissues

Previous work shows that MSH1 expression within the epidermis and vascular parenchyma results in MSH1 localization to the nucleoid and thylakoid mem-brane of the sensory plastid (Virdi et al., 2016). PPD3, a putative stress protein related to PsbP, also localizes

to sensory plastids and may physically interact with MSH1 (Virdi et al., 2016), providing evidence of dis-tinctive sensory plastid proteome features. We predicted that differential accumulation of proteins in sensory plastid versus chloroplast populations would reveal distinct functional information if the organelles could be physically separated. Taking advantage of functional complementation of the msh1 mutant with a native promoter::MSH1::GFP construction (Virdi et al., 2016), we adapted fluorescence-activated cell sorting (FACS)-based procedures to separate Arabidopsis sensory plastids from mesophyll chloroplasts (Fig. 1A).

Complemented lines stably expressing the MSH1:: GFP fusion (Fig. 1B) were used to extract total plas-tids from floral stems (Fig. 1, C and D), where MSH1- expressing tissues are enriched (Virdi et al., 2016). Sensory plastids and mesophyll chloroplasts were fractionated by FACS, discriminating based on positive GFP signal in sensory plastids and separable levels of autofluorescence in the two fractions (Supplemental Figs. S1, A and B). Multiple sorting replicates recovered sufficient amounts of the two populations for down-stream experiments (Supplemental Fig. S2C). Postsort-ing evaluation of organelle populations indicated a sufficiently high level of purity (∼90%) for separation of sensory plastids (Supplemental Fig. S2D).

Sorted plastids were inspected for integrity using laser-scanning confocal microscopy for differential GFP fluorescence. Punctate GFP signal could be seen within nucleoids of sensory plastids (Fig. 1E) and was not detectable in mesophyll chloroplasts (Fig. 1F). Ob-served organelle size differences were consistent with previous in planta observations (Virdi et al., 2016). Elec-tron microscopy imaging (Fig. 1, G and H) showed dif-ferences in internal structural features. Sorted sensory plastids had a planar area of 3.98 ± 0.72 µm2 (Fig. 1I), relative to chloroplasts at 11.76 ± 0.67 µm2, with granal stacking appearing less complex in sensory plastids and plastoglobuli counts lower (Fig. 1J). These data served as confirmation for the successful isolation of a distinct plastid type. Quantitative measurement of chlorophyll fluorescence per equal counts in each plastid population (Fig. 2A) indicated that, following normalization for plastid size, the chlorophyll level of sensory plastids was approximately 66% that of chlo-roplasts (Fig. 2B).

The Chloroplast Proteome Is Predominantly Organized for Photosynthesis, Whereas the Sensory Plastid Proteome May Be Optimized for Stress Response

Sorted plastid populations were subjected to mass spectrometry-based proteomics using nano liquid chromatography-tandem mass spectrometry (nanoLC-MS/MS). Since our objective was to elucidate possible differences between sensory plastid and chloroplast samples and because protein numbers in proteomic experiments can vary significantly from study to study, we filtered and retained only those hits predicted to be chloroplast localized or present in either the PPDB

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(Sun et al., 2009) or the AT_CHLORO (Bruley et al., 2012) databases to enhance stringency (Supplemental Fig. S2, A and B). Protein presence/absence was as-sessed if identification could be established at greater than 99% probability for at least two identified pep-tides (see “Materials and Methods”). Surprisingly, sen-sory plastids exhibited significantly more hits overall (833) than mesophyll chloroplasts (304), while 246 hits were shared by both populations (Fig. 2C; Supplemen-tal Data Set 1). Given these observations, we tested whether we could reconstitute, by combining sensory plastid and chloroplast data sets, the functional (Gene Ontology [GO], biological function) ranking for the list of plastid proteins within the PPDB database. As shown in Figure 2D, a combined sensory plastid and chloroplast assembly shows compelling similarity to the available chloroplast proteome database for GO categories, adding confidence in our proteome results.

Some differences between the reconstituted sample and the publicly available proteome data were antic-ipated because of discrepancies between databases (PPDB versus AT_CHLORO); these factors were taken into account within the reference list to filter the

proteome data. Although our study is foundational for the model plant Arabidopsis, for maize (Zea mays), the PPDB database also reports 173 fewer hits for me-sophyll chloroplast than for the bundle sheath only. While it is difficult to assess the purity of plastid prepa-rations used in all previous studies, it is possible that Arabidopsis chloroplast databases contain, to varying degrees, a mixture of hits coming from heterogeneous plastid populations. Another way to validate our results involved intersection of the sensory plastid proteome data set with single-gene studies that investigated tis-sue-specific expression of plastid-localized proteins within the vascular tissue, as in the case of reticulata mutants. For example, PHOSPHOENOLPYRUVATE/PHOSPHATE TRANSLOCATOR1 (PPT1 or CUE1), NAD(P)H-THIOREDOXIN REDUCTASE, CARBAMOYL PHOSPHATE SYNTHETASE α, CARBAMOYL PHOS-PHATE SYNTHETASE β, and At3g08640, encoding an alpha-tandem protein, are all vascular parenchyma specific or vascular enriched (Lundquist et al., 2014) and were captured in our sensory plastid proteome data set but not in the chloroplast data set (Supplemen-tal Table S1).

Figure 1. Sorting of sensory plastids and chloroplasts. A, A chart of the experimental procedure. Sensory plastids were tagged with MSH1::GFP expressed under the control of the MSH1 native promoter. FACS was used to separate GFP-tagged sensory plastids from mesophyll chloroplasts. Mass spectrometry allowed proteome analysis of the individual fractions. B, Plants ex-pressing an MSH1::GFP fusion in mutant msh1 background (DUAL) were used to extract plastids. C and D, Cross sections of floral stems showing GFP-associated sensory plastids confined to vascular parenchyma and epidermal cells (C; scale bar, 50 µm) and a nontransgenic plant for comparison (D; scale bar, 50 µm). E and F, Confocal laser-scanning image of sorted sensory plastids (E; scale bar, 5 µm) and chloroplasts (F; scale bar, 5 µm). Electron microscopy image of sorted sensory plastid (G; scale bar, 2 µm) and chloroplast (H; scale bar, 2 µm). I, Planar area of sensory plastids and chloroplasts. Values are means ± sd (n = 4). J, Plastoglobule count in sensory plastids and chloroplasts. Values are means ± sd (n = 4).

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We interrogated the functional categorization of the derived sensory plastid and chloroplast proteomes to find considerable differences between the two data sets. Not surprisingly, the main functional category in mesophyll chloroplasts was “photosynthesis” (Fig. 2E). However, functional categories for “oxidation-reduction process,” “response to cytokinin,” and “response to cadmium ion” ranked above “photosynthesis” in sen-sory plastids (Fig. 2F). Proteins shared by both sensory and chloroplast populations included proteins for the photosynthetic machinery, including proteins related to PSI and PSII assembly and components of electron transport important for photosynthesis (Supplemental

Table S2). These results imply that the sensory plastid may be photosynthetically competent to some extent. Moreover, a previous study showed that MSH1 asso-ciates and copurifies with electron transport compo-nents in the sensory plastid (Virdi et al., 2016).

To better understand the proteome differences found, we performed GO enrichment analysis for protein hits that were present exclusively in each plastid popula-tion. Some of the components that were present within the chloroplast but seemingly reduced or absent from sensory plastid fell into categories of “transport” or “photosynthesis” (Supplemental Fig. S3A; Supplemen-tal Data Set 1), but these may contribute to optimization

Figure 2. Features distinguishing sensory plastids and chloroplasts. A, Chlorophyll mean fluorescence intensity (MFI) in sensory plastids and chloroplasts. B, Normalized chlorophyll MFI per planar area (chloroplast MFI as 100%). C, Distribution of proteins identified in sensory plastids and chloroplasts. D, GO enrichment analysis of combined data sets obtained in this study (“sen-sory + chloroplast”) in comparison to the plastid data set curated in PPDB. E, GO terms (% of counts) in the total proteome of chloroplasts. F, GO terms (% of counts) in the total proteome of sensory plastids. Fuchsia color in E and F denotes the ranking for the category “photosynthesis” in each plastid population.



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rather than essential factors. In contrast, categories within the sensory plastid-specific data set were mainly stress-related, including “oxidation-reduction process,” “response to cadmium ion,” “cytokinin,” “cold,” “salt,” “oxidative stresses,” and “defense response to bacte-rium” (Supplemental Fig. S3B). Additionally, biosyn-thetic pathways for fatty acid and amino acid synthesis were enriched in the sensory plastid data set, empha-sizing biosynthetic capacity of these organelles. In fact, nearly the entire shikimate pathway (from PEP to cho-rismate) was localized to the sensory plastids. Notably, nearly all the “reticulate” mutants for plastid-localized proteins in the vasculature or bundle sheath cells are affected in plastid primary metabolism (Lundquist et al., 2014). These data, collectively, indicate that me-sophyll chloroplasts and “sensory” plastids represent functionally distinct plastid populations, with sensory plastids displaying features of specialization for en-hanced stress response and metabolic functions.

Sensory Plastid-Containing Cells Undergo Precise Changes in Gene Expression with Depletion of MSH1

GFP tagging of the MSH1 sensory plastid-specific protein permitted in planta investigation of its spatio-temporal expression patterns. We explored the pattern of expression of the Arabidopsis MSH1 gene under its native promoter in transgenic tobacco (Nicotiana tabacum). In tobacco epidermis, MSH1 localized to small plastids in the trichome stalk but was not detected in the trichome head (Fig. 3, A and B). These small plas-tids showed evidence of autofluorescence, in contrast to those in Arabidopsis, where trichome plastids show little or none (Fig. 3, C and D). The trichome head in tobacco harbored larger autofluorescent chloroplasts, which have been shown by others to undergo active photosynthesis and express a trichome head-specific Rubisco (Laterre et al., 2017). From our observations, the small-sized, MSH1-expressing plastids in the stalk are presumed to be sensory plastids. Noteworthy is the observation in tobacco, as in Arabidopsis, that plastid differentiation is accompanied by differential expres-sion of MSH1 (Fig. 3E); these observations in tobacco emphasize the precise “on-off” promoter behavior re-quired for sensory plastid differentiation across contig-uous tissues (Fig. 3F).

Continuity of sensory plastid behavior between Arabidopsis and tobacco is reiterated by the observa-tion that RNAi suppression of MSH1 elicits markedly similar programmed changes in plant growth pattern and epigenetic memory in six different plant species (Xu et al., 2012). These observations prompted us to investigate the local cellular changes that occur in re-sponse to sensory plastid perturbation. Since MSH1 is down-regulated by environmental stresses and its de-pletion elicits a strong transcriptomic stress response, we used the msh1 phenomenon as a first proxy to un-derstand sensory plastid functions. Capitalizing on the specificity of the MSH1 promoter to drive expression of a FLAG-tagged ribosome protein, together with

affinity-purification and RNA-sequencing (RNA-seq; translatome), we investigated sensory plastid cell- specific responses in plants depleted of MSH1 by the translating ribosome affinity purification (TRAP)-seq method (Supplemental Fig. S4A; Reynoso et al., 2015). The experiments involved introduction of large-subunit ribosomal protein L18 fusion with N-terminal FLAG epitope tag under the control of the MSH1 promoter, introduced to both the msh1 mutant and Col-0 wild type (Fig. 4A; Supplemental Fig. S4B).

Principal component analysis of derived trans-latome data clustered the mutant samples distinctly from wild type, providing evidence of a differential

Figure 3. Sensory plastids in tobacco. A, Trichome of transgenic tobacco line expressing MSH1::GFP in sensory plastids. Inset is an enlargement of sensory plastids in the trichome stalk taken from an independent experiment in the region indicated by the white arrow. B, Trichome of nontransgenic tobacco. C, Trichome of stable transgenic Arabidopsis line expressing MSH1::GFP in sensory plastids. Inset is a zoom image of sensory plastids from the region indicated by the white arrow. D, Trichome of nontransgenic Arabidopsis. Inset is a zoom im-age of sensory plastids. E, Epidermal sensory plastids are positive for MSH1::GFP. F, On and off modulation of MSH1 in different tissues of tobacco and Arabidopsis. VP, Vascular parenchyma; M, mesophyll; E, epidermis; T, trichome. All scale bars are 25 µm.


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gene expression response following MSH1 depletion (Fig. 4B). Cell-specific translatome profiles captured evidence of differential translation for 3,124 genes in MSH1-depleted cells, whereas total transcriptome pro-filing from the same tissue was able to distinguish only 300 genes with differential expression. Similar differ-ential enrichment effects have been reported for other systems (Mustroph et al., 2009; Endo et al., 2014), perhaps due to ribosome enrichment prior to RNA ex-traction in translatome procedures. Thus, this study with pooled floral stem samples and previous studies with selected phenotypes and rosettes tissues (Shao et al., 2017) were unable to capture the entire translatome gene response via transcriptome analysis (Supplemen-tal Fig. S5C; Supplemental Data Set 2). Differentially expressed gene number in the msh1 mutant can vary markedly depending on the generation and phenotype intensity (Shao et al., 2017). Yet, GO ontology analysis of the transcriptome in this study (Supplemental Data Set 3) confirms similar global stress response to earlier reports (Shao et al., 2017), with categories altered for response to cold, wounding, oxidative stress, response to hormones (jasmonic acid, auxin, gibberellin, and ABA) and circadian rhythm (Fig. 4C).

For translatome analysis, we used the full list of differentially translated genes to perform GO enrich-ment (Fig. 4C), network enrichment analysis test and network-based enrichment analysis (Fig. 4D). KEGG networks that were identified included MAPK signal-ing, circadian rhythm, Arg and Pro metabolism, meta-bolic pathways, biosynthesis of secondary metabolites, SNARE interactions in vascular transport, autophagy, phagosome, plant-pathogen interaction, and plant hor-mone signal transduction. GO enrichment analysis identified several related categories (Supplemental Data Set 3), the most prominent involving redox, stress

response and signaling, chromatin remodeling, and cellular growth behavior. Within the “response to oxi-dative stress” category were genes for peroxidases, glutathione transferases, and thioredoxins, including ASCORBATE PEROXIDASE1, scavenging hydrogen peroxide in plant cells, and GLUTATHIONE PEROX-IDASE8, for suppression of oxidative damage in the nucleus and cytosol. In cell redox homeostasis, thi-oredoxins were highly enriched, including THIORE-DOXIN F-TYPE1 in the short-term activation of carbon metabolism necessary for growth under short-day con-ditions (Naranjo et al., 2016). These observations are indicative of ROS changes in the cell.

Direct-measure oxido-reduction changes were earlier correlated with sensory plastid perturbation in floral stems of msh1 mutants (Virdi et al., 2016) and in msh1 hemicomplementation tests (Xu et al., 2012; Virdi et al., 2016). Oxidative stress can elicit plastid signaling events accompanied by calcium activation of calmodulins to trigger signaling cascades and influence nuclear stress response (Stael et al., 2012; Guo et al., 2016). We detected sensory cell-specific gene expression changes in calcium- mediated signaling pathways, including numerous calmodulins and rapid alkalization factor-like signal elements. Rapid alkalization factor elements play a role in cellular signaling under stress in Arabidopsis (Atkinson et al., 2013). MAPK phosphatase2 (AT3G06110), a positive regulator of the cellular response to oxi-dant challenge (Jin and Ellis, 2007), and WRKY DNA- binding protein 25 (AT2G30250), involved in abiotic stress response (Jiang and Deyholos, 2009), were also altered in the translatome. These data support the par-ticipation of ROS- and calcium-mediated stress signal-ing in sensory plastid-containing cells.

MSH1 depletion was associated with altered expres-sion of members of the circadian clock mechanism

Figure 4. Cell-specific translatome profil-ing. A, Representative plants expressing a FLAG::RPL-18 fusion in both the wild type (WT; Col-0 RPL-18) and the msh1 mutant (msh1 RPL-18) backgrounds. B, Principal component analysis plot for translatome (IP) and transcriptome (total) reads. C, Prominent GO categories for biological function in the translatome and the transcriptome (this study) and total transcriptome (T; total plant data set from Shao et al., 2017). D, Network enrichment analysis of KEGG-derived pathways for the sensory plastid-associated translatome output.



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(LHY, GI, FT, SPA1, PRR9, CCA1, TCP11, and PRR5), several of which were similarly prominent in msh1- associated epigenetic memory response (X. Yang, R. Sanchez, H. Kundariya, Y. Wamboldt, J. Barreras, and S. Mackenzie, unpublished data). This intersection with earlier independent studies of the msh1 phenomenon suggests that MSH1 depletion primes a programmed signaling response pattern within sensory plastid- containing cells, where it is precisely localized. The identified networks provide a model for the highly pleiotropic msh1 phenotypes that emerge.

The most significant category identified by GO enrichment analysis was “nucleosome assembly” and its related categories of “chromatin silencing,” “pro-duction of siRNA,” and “mRNA splicing, via spliceo-some” (Fig. 5, A–C and E). These functional categories may link to the stress-related, genome-wide methyla-tion changes that have been reported in msh1 mutants (Virdi et al., 2015) and with subsequent transgenera-tional memory (X. Yang, R. Sanchez, H. Kundariya, Y. Wamboldt, J. Barreras, and S. Mackenzie, unpublished data). Numerous histone genes were up-regulated in response to MSH1 depletion, including histones HTB1, HTB11, HTB4, HTB4, HTB9, HTB2, HIS4, a “winged- helix DNA-binding transcription factor family protein” (H1.1), and a high-mobility group A (HMGA) protein. H1.1 is a linker histone important for chromatin dy-namics (Kotliński et al., 2017). There were also three HISTONE3-RELATED (HTR) genes, HTR4 (AT4g40030), HTR5, and HTR8, that encode H3.3, required for depo-sition or maintenance of DNA methylation over gene bodies (Wollmann et al., 2017). Specifically, H3.3 knock-down results in loss of methylation over gene bodies, and its enrichment near gene transcription start sites is positively associated with transcription (Stroud et al., 2012; Wollmann et al., 2012). SUVR5, which binds DNA to repress the expression of a subset of stimulus- response genes (Caro et al., 2012), was reduced in ex-pression in msh1, perhaps factoring into the observed unleashing of several stimuli responders. ARGO-NAUTE4, involved in siRNA-mediated gene silencing (Zilberman et al., 2004), was also downregulated (Fig. 5, C and D). The expression levels of other members of the methylation and siRNA production machinery were altered following MSH1 depletion, including MET1, DME, and RDR2 (Fig. 5D). These data provide a unique profile of epigenomic reprogramming, and its cell-specific induction, when aligned with the marked changes in genome-wide methylation that are documented in msh1 (Virdi et al., 2015) and its heritable memory (X. Yang, R. Sanchez, H. Kundariya, Y. Wamboldt, J. Barreras, and S. Mackenzie, unpublished data).

To better understand how sensory plastid-containing cells respond to induced stress, we divided the differ-entially translated genes into up-regulated and down- regulated subgroups, revealing a much larger num-ber of transcription factors to be activated (120) than down-regulated (38), with basic helix-loop-helix, C2H2, AP2-EREBP, and MYB families most prominent (Fig. 6A).

Globally, nonredundant GO category analysis showed that MSH1-depleted cells up-regulate pathways for chromatin remodeling and stress responses, including nucleosome assembly, chromatin assembly or disas-sembly, heterochromatin organization, response to hormones, and response to oxidative stress and other stresses like cold, salt, drought, and biotic factors. Meanwhile, down-regulated networks involved pre-dominantly the regulation of cell shape, cell wall or-ganization, and secondary metabolism (Supplemental Fig. S5, A and B), consistent with the reduced plant growth phenotype of msh1 that counterbalances en-hanced stress response (Xu et al., 2011, 2012). The ob-served changes in gene expression networks within sensory plastid-containing cells were accompanied by changes in stress-related small molecules. Visualization of ROS species (H2O2 and O2

-) in msh1 mutants showed a higher concentration proximal to vascular tissues (Fig. 6, B and C), which likely accounts for generally higher measurements of ROS in MSH1-RNAi lines (Xu et al., 2012), and with oxidative stress responses ob-served in gene expression analyses.

DISCUSSION

The sensory plastid is distinguished from mesophyll chloroplast not only by size and morphology but proteome features. We found significant overlap for proteins related to PSI and PSII assembly, as well as components of the electron transport chain important for photosynthesis, but also markedly distinctive fea-tures for stress-related and metabolic pathway com-ponents. Therefore, sensory plastids may be uniquely equipped to trigger stress and developmental response by the plant. Studies of the reticulata mutant pheno-types, characterized by changes in vascular plastid be-havior (Kinsman and Pyke, 1998), show that mesophyll chloroplast functions are influenced by, and responsive to, vascular plastid signals (Lundquist et al., 2014). The reticulata phenotype is recognized by leaf patterning with distinctive venation due to altered signaling be-tween vascular plastids and mesophyll chloroplasts. The phenotype can be conditioned by several distinct loci, revealing vascular plastid-specific proteins that serve to validate our own data for sensory plastid-specific pro-teome components. The reticulata phenomenon invites an intriguing model for systemic stress signal transmis-sion in plant systems, initiating with vascular tissue- localized signals that emanate to alter chloroplast behavior (Lundquist et al., 2014). A component of this signaling appears to involve phosphoenolpyruvate import to sensory plastids, which can be complemented by exogenous application of aromatic amino acids (Staehr et al., 2014). While both “sensory” plastids and mesophyll chloroplast data sets contain the category for response to oxidative stress (Fig. 2, E and F), our study lends support to localized plastid-derived stress responses.


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It is not clear whether the epidermal and vascular organelles are a singular population in proteome com-position or comprise distinct organelle types. Analysis of the tobacco trichome showed that differentiation of sensory plastid relative to chloroplast is precise and not defined by meristematic origin. The epidermal plastid contains thylakoid grana that are already vis-ible in L1 undifferentiated meristem cells (Charuvi et al., 2012). This is the case for plastids within the L3 layer that derives the vascular tissues as well. Earlier studies have suggested that epidermal plastids are nonphotosynthetic (Charuvi et al., 2012), implying that assembly and granal stacking of the thylakoid mem-brane reflects a functional electron transport chain that may lack other components of the photosynthetic apparatus. However, as shown in this study, purified Arabidopsis epidermal and vascular plastid fractions autofluoresce and contain chlorophyll. Our analysis of sensory plastid proteome data did not reveal key com-ponents of photosynthesis to be lacking, suggesting

that sensory plastids may contain a full complement of the photosynthetic apparatus. Consequently, it is possible that sensory plastids are photosynthetically competent but are additionally equipped with stress and metabolic signaling mechanisms that specialize their functions.

Our study showed the chloroplast to be less complex in proteome than previously estimated (http://ppdb.tc.cornell.edu/). This observation implies that some plastid-associated stress response features may have been misattributed in earlier studies to the chloroplast due to difficulties in fractionation of the two organelle types. It is possible that retrograde signaling circuits may differ across tissue types and that crop engineer-ing efforts could be refined to target the correct plastid type, therefore avoiding undesired effects.

Observations detailed here and in earlier studies are consistent with the hypothesis that the MSH1 system, presumably one of several that appear to emanate from the sensory plastid, plays a role in plant adaptation.

Figure 5. Cell-specific influence on chromatin remodeling, DNA methylation, and calcium signaling gene expression in msh1 T-DNA mutants based on translatome data. Panels show gene expression changes in nucleosome assembly (A), chromatin si-lencing (B), production of siRNA (C), methylation machinery (D), mRNA splicing (E), and calcium-mediated signaling (F). Gene expression changes are measured by log2 fold-change (log2FC).



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Localization within the epidermis, vascular parenchyma, and bundle sheath cells likely serves to accommodate environmental sensing and signaling. MSH1 disruption is sufficient to trigger global stress response and devel-opmental changes in plants, an effect that requires two generations to fully amplify (Shao et al., 2017). This global stress genomic posture, once achieved, includes extreme methylation repatterning that, in subsequent generations, evokes heritable nongenetic memory (Xu et al., 2012; Virdi et al., 2015; X. Yang, R. Sanchez, H. Kundariya, Y. Wamboldt, J. Barreras, and S. Mackenzie, unpublished data).

To understand the msh1 global stress behavior and derived memory, we characterized translatome fea-tures within cells that contain sensory plastids under wild-type and msh1 conditions. Pathways identified within the translatome are presumed to represent at least some of the initial responses to MSH1 depletion. Networks for calcium signaling, oxidative stress, nu-cleosome and chromatin remodeling, and circadian clock were prominent sensory plastid-proximal re-sponses. These outcomes appear strikingly resonant with studies of msh1 memory, the state that persists with MSH1 restoration following RNAi knockdown (Xu et al., 2012; X. Yang, R. Sanchez, H. Kundariya, Y. Wamboldt, J. Barreras, and S. Mackenzie, unpublished data). The memory phenotype involves DNA methy-lome repatterning, with altered expression of circadian clock, ABA, and oxidative stress responses (X. Yang, R. Sanchez, H. Kundariya, Y. Wamboldt, J. Barreras, and S. Mackenzie, unpublished data). The vascular circadian network is known to exert robust regulatory dominance over neighboring mesophyll cells, with transcriptional and physiological consequences (Endo et al., 2014).

The msh1 T-DNA-null mutant produces a significant range in phenotype intensity and contains a much broader, more variable, and more extreme effect on gene expression networks (Shao et al., 2017) than we see in the memory phenotype. We postulate that cell-specific sensory plastid effects serve to trigger a system-wide plant response, with the intensity and range of pheno-type displayed by each msh1 mutant plant conditioned by downstream factors, and with a central core of the phenomenon retained in heritable memory.

Aspects of the MSH1 system appear to intersect with findings by other groups. Perturbation of plastids re-siding within epidermal and vascular parenchyma cells alters ABA- (Endo et al., 2008; Gorecka et al., 2014) and auxin-related (Procko et al., 2016) plant stress be-haviors. Salt (Veley et al., 2012) and drought (Endo et al., 2008) stress pathways are triggered within these plastids, as is high-light response (Galvez-Valdivieso et al., 2009). Selection for second-site suppressors of the REPRESSOR OF SILENCING1 (ROS1) locus, which participates in cytosine demethylation (Shen et al., 2009), identifies the CHLOROPHYLL A/B BINDING PROTEIN UNDEREXPRESSED1 (CUE1) gene, which encodes a phosphoenolpyruvate translocator on the envelope membrane of vascular sensory plastids. This observation appears to link vascular plastid functions and epigenetic changes within the nucleus, influenced by availability of aromatic amino acids (Shen et al., 2009).

The reports by other groups provide evidence that perturbation of sensory plastids results in ABA-mediated stress responses in the plant, as well as nuclear epi-genetic changes. The msh1 mutant displays enhanced tolerance not only to drought (Virdi et al., 2016), but to heat (Shedge et al., 2010) and high light (Xu et al., 2011) as well. MSH1 suppression alters expression of the ABA pathway and methylation repatterning of genes involved in the regulation of ABA (Virdi et al., 2015; X. Yang, R. Sanchez, H. Kundariya, Y. Wamboldt, J. Barreras, and S. Mackenzie, unpublished data).

Collectively, these studies support a model of sensory plastids as distinctly differentiated organelles that func-tion in environmental sensing (Fig. 7). Translatome features in sensory plastid cells under perturbation suggest the involvement of ROS, calcium, and MAPK in this process, components of plastid stress signaling (Stael et al., 2012; Guo et al., 2016). Under natural con-ditions, MSH1 undergoes transcriptional suppression in response to stress and, through its depletion, elicits strongly pleiotropic plant responses (Xu et al., 2011, 2012). It will be interesting to learn whether this type of stress responsiveness is observed for other sensory plastid-specific protein genes. The MSH1 protein is presumed to have been mitochondrially localized early in its evolution (Abdelnoor et al., 2006), subsequently expanded in targeting to the plastid (Xu et al., 2011) and acquiring gene promoter features supporting spa-tiotemporal and stress-responsive regulation (Virdi et al., 2016). These components of the gene’s evolu-tion appear fundamental to its neofunctionalization

Figure 6. Phenotypes mediated in msh1 by sensory plastid perturba-tion. A, Graph of transcription factor changes. B, Differential accumu-lation of H2O2 in floral stem transversal cuts from msh1 and wild-type (WT) control by nitro-tetrazolium blue chloride (NBT) staining. Arrows indicate areas with differential staining relative to the wild type. C, Accumulation of O2

− in transversal cuts of floral stems from msh1 and wild-type control by 3,3′-diaminobenzidine (DAB) staining. Arrows in-dicate areas with differential staining relative to the wild type.


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for triggering programmed plant stress and heritable memory behaviors, and this evolutionary scheme may prove instructive in modeling the origins of other sen-sory plastid-specific proteins.

MATERIALS AND METHODS

Plant Materials

Arabidopsis (Arabidopsis thaliana) plants were grown in a controlled- environment walk-in chamber at 12 h day length and 22°C. The Col-0 line used in this study for transformation and control has been DNA sequenced and confirmed to represent the reference genome (Shao et al., 2016). Col-0 msh1 line complemented with MSH1P::MSH1::GFP was derived previously (Xu et al., 2011). The msh1 T-DNA mutant (SAIL-877-F01) was obtained from The

Arabidopsis Stock Center (http://www.arabidopsis.org/). Col-0 transgenic plants expressing MSH1P::FLAG::RPL18 were generated in both msh1 homo-zygous mutant (SAIL-877-F01) and wild-type backgrounds with the cloned MSH1 sequence described by Xu et al. (2011) and with transformation by the floral-dip method (Clough and Bent, 1998). Positive transgenic lines were screened for FLAG tag by immunoblotting methods. Stable tobacco transgenic lines were generated by introducing the MSH1P::SSU-MSH1::GFP fusion construct (Xu et al., 2011) to leaf discs and regenerating transgenic plantlets as described (Virdi et al., 2016). Tobacco plants were grown under standard greenhouse conditions, and tissues were excised for confocal microscopy, with nontransgenic isolines as control.

Plastid Extraction

Crude plastid preparations from floral stems were carried out in extraction buffer (20 mm Tricine-NaOH, pH 8.4, 300 mm sorbitol, 10 mm KCl, 10 mm EDTA, 0.25% [w/v] bovine serum albumin, 4.5 mm Na-ascorbate, and 5 mm l-Cys). Samples were centrifuged at 2,000g for 5 min at 4°C and supernatants recovered and centrifuged at 4,000g for 10 min at 4°C. Plastid pellets were resuspended in wash buffer (20 mm HEPES, 300 mm sorbitol, 10 mm KCl, 2.5 mm EDTA, and 5 mm MgCl2), recovered by centrifugation at 5,000g for 10 min at 4°C and resuspended again in 5 mL of wash buffer for sorting.

FACS

Plastids were sorted on a BD FACS Aria II system. GFP and chlorophyll autofluorescence were detected with a 488-nm blue laser. The GFP signal was detected with a 505LP and 530/30BP filter combination and chlorophyll aut-ofluorescence detected with a 670LP and 685/35BP filter combination. The sample flow rate was between 9,000 and 12,000 events/s in buffer containing 20 mm HEPES, 300 mm sorbitol, 10 mm KCl, 2.5 mm EDTA, and 5 mm MgCl2. The experiment was performed twice. For each experiment, four independent sortings were combined. Each independent sorting required the floral stems of 36 plants.

Protein Extraction and Mass Spectrometry

Plastids were lysed by addition of RapiGest, a cleavable, mass spectrometry- friendly detergent (Waters). The samples were then reduced with 5 mm DTT at 80°C, cooled, and alkylated with 3× molar excess of iodoacetamide. The io-doacetamide was quenched with additional DTT before digestion with 500 ng trypsin overnight at 37°C. The detergent was cleaved by the addition of triflu-oroacetic acid and incubation at 37°C. Insoluble components of the detergent were carefully removed; samples were partially dried in a Savant SpeedVac concentrator SVC100H (SpectraLab Scientific) and then dissolved into mass spectrometry sample loading buffer (2.5% [v/v] acetonitrile and 0.1% [v/v] formic acid) according to yield. Samples were spun at 21,000g for 10 min be-fore equal amounts were analyzed by nanoLC-MS/MS using a 2-h gradient on a 0.075 mm × 250 mm C18 Waters CSH column feeding into a Q-Exactive HF mass spectrometer.

Proteomics

All MS/MS samples were analyzed with Mascot (Matrix Science; version 2.5.1). Mascot was set to search NCBInr_20160821database (Arabidopsis thaliana, 94,081 entries), and cRAP_20150130 database (117 entries) assuming the diges-tion enzyme trypsin. Mascot was searched with a fragment ion mass tolerance of 0.060 D and a parent ion tolerance of 10.0 PPM. Deamidation of Asn and Gln, oxidation of Met, and carbamidomethyl of Cys were specified in Mascot as variable modifications.

Scaffold (version Scaffold_4.7.2; Proteome Software) was used to validate MS/MS-based peptide and protein identifications. Peptide identifications were accepted if they could be established at greater than 80% probability by the Peptide Prophet algorithm (Keller et al., 2002) with Scaffold delta-mass correction. Protein identifications were accepted if they could be established at greater than 99% probability and contained at least two identified pep-tides. Protein probabilities were assigned by the Protein Prophet algorithm (Nesvizhskii et al., 2003). Proteins that contained similar peptides and could not be differentiated based on MS/MS analysis alone were grouped to sat-isfy the principles of parsimony. Proteins sharing significant peptide evi-dence were grouped into clusters. Peptide hits were annotated using UniProt

Figure 7. Proposed model for plastid-specific modulation of chromatin remodeling and stress response pathways mediated by MSH1. We pos-tulate that experimental or stress-associated depletion of MSH1 from the thylakoid/nucleoid site induces ROS/redox changes and release of calcium from the plastid, binding calmodulins (CaMs) to activate calmodulin-binding protein kinases (CBKs) and protein kinase C to ac-tivate MAPKs via phosphorylation. Chromatin remodeling events may unleash a localized stress response, involving changes in translation of stress-related transcription factors, ABA regulators, and circadian clock components. A subset of methylome changes that accompany this pro-cess display transgenerational heritability for the msh1 memory effect. Emboldened gene classes reflect observed gene expression changes in the sensory plastid translatome data set.



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(http://www.uniprot.org), and TAIR ID numbers were retrieved and retained only if present in two independent plastid sample replicates. To increase stringency, hits were filtered and retained only if predicted to be chloroplast- localized or present in either the PPDB (Sun et al., 2009) and/or the At_CHLORO (Bruley et al., 2012) databases. For functional categorization, we used unique representative ID numbers rather than gene families.

Confocal Microscopy

Confocal laser-scanning microcopy of sorted plastids and plant tissues was performed on a Nikon A1 confocal laser-scanning microscope mounted on the Nikon Eclipse 90 upright compound microscope. Images were acquired using a Nikon NIS Elements version 4.20. GFP (excitation, 488 nm; emission, 500–550 nm) and chlorophyll autofluorescence (excitation, 640.6 nm; emission, 663–738 nm). Alternatively, a Zeiss LSM 510 laser-scanning microscope was used with similar excitation/emission parameters with the corresponding Zen software.

Transmission Electron Microscopy

FACS-sorted plastids were fixed in 2.5% (v/v) glutaraldehyde in 0.05 m sodium cacodylate (pH 7.4), and transmission electron microscopy was car-ried out with a Hitachi H7500-I microscope and standard procedures at the University of NE Center for Biotechnology Microscopy Facility. Area measure-ments of plastid photographs were made with the ImageJ2 program (Rueden et al., 2017).

Western-Blot Analysis

Plant tissues were ground in liquid nitrogen and total proteins extracted in lysis buffer (50 mm sodium phosphate buffer, pH 7.0, 10 mm EDTA, 1% [v/v] Triton, 0.1% [w/v] SDS, 1× protease inhibitor, and β-mercaptoethanol) for 75 min at 4°C. Lysates were centrifuged at 20,000g for 1 h and protein samples were separated using the NuPAGE system (Invitrogen). Proteins were trans-ferred to nitrocellulose membranes (Optitran; Whatman) for incubation in 1× PBS blocking buffer (137 mm NaCl, 2.7 mm KCl, 8 mm Na2HPO4, and 2 mm KH2PO4), 3% (w/v) bovine serum albumin, and 1% (v/v) Tween 20 for 1 h at room temperature, then for 1 h at room temperature with anti-FLAG antibody. After washing, membranes were incubated with the respective secondary an-tibody, and immunodetection performed with the Alkaline Phosphatase sys-tem (Roche Diagnostics).

TRAP

Polysomal mRNA from MSH1-specific cells was obtained by ribosome pu-rification using the TRAP method essentially as reported by Reynoso et al. (2015). In brief, floral stem tissues were ground in liquid nitrogen and stored at −80°C. One hundred milliliters lysis buffer (0.2 m Tris, pH 9.0, 0.2 m KCl, 0.025 m EGTA, 0.035 m MgCl2, 1% [v/v] detergent mix [20% (w/v) poly-oxyethylene (23) lauryl ether, 20% (v/v) Triton X-100, 20% (v/v) octylphenyl- polyethylene glycol, 20% (v/v) polyoxyethylene sorbitan monolaurate 20], 1% (w/v) DOC, 1% (v/v) PTE, 5 mm DTT, 1 mm PMSF, 50 µg/mL cyclohexim-ide, 50 µg/mL chloramphenicol) were added to 50 mL of ground tissue. Then, 1 to 2 mL of anti-FLAG agarose beads (EZview Red ANTI-FLAG M2 Affin-ity Gel) was added per sample and incubated overnight at 4°C with gentle rotation. Samples were washed three times with wash buffer (0.2 m Tris, pH 9.0, 0.2 m KCl, 0.025 m EGTA, 0.035 m MgCl2, 5 mm DTT, 1 mm PMSF, 50 μg/mL cycloheximide, and 50 μg/mL chloramphenicol). Elution of samples was performed with wash buffer containing 24 µL FLAG peptide, 2 µL RNase inhibitor, and 2 µL RNase A inhibitor. Final RNA extraction was performed with the standard Trizol method (Invitrogen) by adding 1 µL of glycogen.

RNA-Seq and Analysis

Transcriptome and translatome for msh1 samples were sequenced by BGI-Tech on the HiSeq 4000 analyzer (Illumina) with paired-end 100-bp option for read length. Alignments were performed using RUM 2.0.4 (−min-length 70 parameters; Grant et al., 2011), retaining only uniquely mapped reads. The read count data were generated from the SAM files with QoRTs software pack-age at −minMAPQ 25 flag (chloroplast and mitochondria reads were dropped; Hartley and Mullikin, 2015). DESeq2 (Love et al., 2014) was used for gene

count normalization and to identify differentially expressed genes (FDR < 0.05, |log2FC| > 0.5). GO enrichment analyses were performed with the Da-tabase for Annotation, Visualization and Integrated Discovery (DAVID) v6.8, applying the Benjamini-Hochberg method for multiple testing adjustment of P values. Custom R scripts were used to generate heat maps. Enrichment-Browser R package was applied for Network Based Enrichment Analysis, and the R package “neat” 705 version 1.1.1 was used for Network Enrichment Analysis Test.

Accession Numbers

Sequence data from this article have been submitted to the GenBank data libraries under accession number GSE114199.

Supplemental Data

The following supplemental materials are available.

Supplemental Figure S1. Assessment of the plastid sorting technique.

Supplemental Figure S2. Comparative proteomes of sensory plastids and chloroplasts relative to public database information.

Supplemental Figure S3. Proteome distinctions between the two plastid types.

Supplemental Figure S4. Translatome profiling.

Supplemental Figure S5. Most significant GO biological processes associ-ated with the cell-specific translatome response in msh1.

Supplemental Table S1. List of reticulata-associated genes (Lundquist et al., 2014) found in the “sensory” plastid proteome and absent in chlo-roplast proteome.

Supplemental Data Set 1. List of proteome outputs for chloroplast and sensory plastid.

Supplemental Data Set 2. msh1 translatome and msh1 transcriptome.

Supplemental Data Set 3. GO categories for translatome and transcriptome.

ACKNOWLEDGMENTS

We thank Jose R. Barreras for bioinformatics assistance, Dirk Anderson for assistance in flow cytometry, Michael Naldrett from the proteomics and metabolomics facility of the Center for Biotechnology at University of Nebraska- Lincoln, and Drs. Xiaodong Yang and Vikas Shedge for helpful discussions. We also thank Sarah Pfaff for help with PCR screening.

Received July 5, 2018; accepted August 13, 2018; published August 22, 2018.

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