# NEW INSIGHTS INTO MECHANISMS OF EPIGENETIC MODIFIERS IN PLANT GROWTH AND DEVELOPMENT

EDITED BY : Ming Luo, Gabino Ríos, Tomasz Jacek Sarnowski, Shoudong Zhang, Marc Libault, Nitin Mantri and Jean-Benoit Charron PUBLISHED IN : Frontiers in Plant Science

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ISSN 1664-8714 ISBN 978-2-88963-466-8 DOI 10.3389/978-2-88963-466-8

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# NEW INSIGHTS INTO MECHANISMS OF EPIGENETIC MODIFIERS IN PLANT GROWTH AND DEVELOPMENT

Topic Editors:

Ming Luo, South China Botanical Garden, Chinese Academy of Sciences, China Gabino Ríos, Instituto Valenciano de Investigaciones Agrarias, Spain Tomasz Jacek Sarnowski, Institute of Biochemistry and Biophysics (PAN), Poland Shoudong Zhang, The Chinese University of Hong Kong, China Marc Libault, University of Nebraska-Lincoln, United States Nitin Mantri, RMIT University, Australia Jean-Benoit Charron, McGill University, Canada

Citation: Luo, M., Ríos, G., Sarnowski, T. J., Zhang, S., Libault, M., Mantri, N., Charron, J.-B., eds. (2020). New Insights into Mechanisms of Epigenetic Modifiers in Plant Growth and Development. Lausanne: Frontiers Media SA. doi: 10.3389/978-2-88963-466-8

# Table of Contents

	- Yunmin Xu, Lu Zhang and Gang Wu

Shenxiu Du, Liang Chen, Liangfa Ge and Wei Huang

*81 Functional Coordination of the Chromatin-Remodeling Factor AtINO80 and the Histone Chaperones NRP1/2 in Inflorescence Meristem and Root Apical Meristem*

Huijia Kang, Jing Ma, Di Wu, Wen-Hui Shen and Yan Zhu

Stéphane Maury, Mamadou Dia Sow, Anne-Laure Le Gac, Julien Genitoni, Clément Lafon-Placette and Iva Mozgova

*121 Chromatin Evolution-Key Innovations Underpinning Morphological Complexity*

Mohsen Hajheidari, Csaba Koncz and Marcel Bucher

Taras Pasternak and Denes Dudits

*150 Dynamic Changes in Genome-Wide Histone3 Lysine27 Trimethylation and Gene Expression of Soybean Roots in Response to Salt Stress* Lei Sun, Guangshu Song, Weijun Guo, Weixuan Wang, Hongkun Zhao, Tingting Gao, Qingxue Lv, Xue Yang, Fan Xu, Yingshan Dong and Li Pu

# Editorial: New Insights Into Mechanisms of Epigenetic Modifiers in Plant Growth and Development

Ming Luo1,2\*, Gabino Ríos 3 , Tomasz Jacek Sarnowski 4 , Shoudong Zhang5 , Nitin Mantri 6 , Jean-Benoit Charron7 and Marc Libault 8\*

1 Key Laboratory of South China Agricultural Plant Molecular Analysis and Genetic Improvement, Guangdong Provincial Key Laboratory of Applied Botany, South China Botanical Garden, Chinese Academy of Sciences, Guangzhou, China, 2 Center of Economic Botany, Core Botanical Gardens, Chinese Academy of Sciences, Guangzhou, China, 3 Department of Citriculture and Plant Production, Instituto Valenciano de Investigaciones Agrarias (IVIA), Valencia, Spain, 4 Department of Protein Biosynthesis, Institute of Biochemistry and Biophysics Polish Academy of Sciences, Warsaw, Poland, 5 Centre for Soybean Research, State Key Laboratory of Agrobiotechnology and School of Life Sciences, The Chinese University of Hong Kong, Shatin, Hong Kong, 6 School of Science, The Pangenomics Group, RMIT University, Melbourne, VIC, Australia, 7 Department of Plant Science, McGill University, Sainte-Anne-de-Bellevue, QC, Canada, 8 Department of Agronomy and Horticulture, Center for Plant Science Innovation, University of Nebraska, Lincoln, NE, United States

Keywords: epigenetic regulation, plant development, histone modification, chromatin modification, DNA methylation, small RNAs

#### Edited and reviewed by:

Jin-Gui Chen, Oak Ridge National Laboratory (DOE), United States

#### \*Correspondence:

Ming Luo<EMAIL_ADDRESS>Marc Libault<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science

Received: 30 October 2019 Accepted: 25 November 2019 Published: 10 January 2020

#### Citation:

Luo M, Ríos G, Sarnowski TJ, Zhang S, Mantri N, Charron J-B and Libault M (2020) Editorial: New Insights Into Mechanisms of Epigenetic Modifiers in Plant Growth and Development. Front. Plant Sci. 10:1661. doi: 10.3389/fpls.2019.01661 Editorial on the Research Topic

New Insights Into Mechanisms of Epigenetic Modifiers in Plant Growth and Development

In eukaryotic cells, chromatin, a highly dynamic nucleoprotein complex, plays a critical role in controlling gene expression notably by regulating the interaction between transcription factors and regulatory elements. The structure of the chromatin is determined by epigenetic mechanisms, including DNA methylation, histone modifications, and chromatin remodeling. A growing body of evidence indicates that epigenetic regulations are involved in plant adaptation to environmental stresses, and in plant development, including flowering control, fruit and root development, as well as seed maturation and germination. Furthermore, epigenetic mechanisms have the potential to stabilize cell identity and maintain tissue organization. Hence, epigenetic diversity is now emerging as a new source of phenotypic variation to improve adaptation to changing environment and ensure yield and quality of crops. The 14 articles published in this Research Topic highlight recent progresses, opinions, and reviews to advance our knowledge in the role of the epigenome on controlling plant development, plant response to environmental stresses, and plant evolution. For instance, gene duplication and chromatin remodeling contribute to increase the morphological and cellular complexity of plants during their evolution according to Hajheidari et al.

Chromatin modifications, including DNA methylation and histone modifications, are critical in regulating gene transcription, and thus may reprogram cell differentiation and development (Inácio et al.; Zhang et al.; Hajheidari et al.). For instance, Inácio et al. immunolocalized various epigenetic marks and correlated epigenomic changes with transcriptional regulation when studying cork formation and quality in cork oak, a genuinely forest-specific process. Furthermore, changes in the acetylation levels of the lysine 9 of the histone H3 (H3K9) and lysine 5 of the histone H4 (H4K5) were found associated with the heat stress-dependent inhibition of lateral root formation in maize (Zhang et al.). Interestingly, whereas a global increase in histone acetylation was observed in response to heat stress, H3K9 and H4K5 acetylation decreased significantly in the promoter region of the haem oxygenase-1 (ZmHO-1) and giberellic acid–stimula ted like-1 (ZmGSL-1) genes, two inhibitors of lateral root forma tion (Zhang et al.).

Plant cells have the capability to dedifferentiate in totipotent cells, a prerequisite to asexual embryogenesis. Recent papers support a role of histone deacetylation and DNA methylation in cellular reprogramming leading to callus formation and asexual embryogenesis through the regulation of key developmental genes such as Wuschel (Pasternak and Dudits). In addition to somatic embryogenesis, the epigenome also controls the juvenile-to-adult developmental transition notably by modula ting the expression of regulatory genes. Indeed, in Arabidopsis plants, this transition is regulated by miR156/157 and its ta rget-squamosa promoter binding protein-like gene (Xu et al.). Other epigenetic changes controlling the juvenile-to-adult developmental transition include DNA methylation, and histone modification (Xu et al.). Ultimately, these chemical cha nges lead to a remodeling of the chromatin. The SWI/SNF chromatin remodeling complexes play a central role in this biological process by controlling phytohormone biosynthesis, the establishment and maintenance of meristems, organ development, and floral transition (Ojolo et al.; Maury et al.). Supporting the central role of chromatin remodeling and histone modifications in controlling development of plant, Kang et al. studied the role of the chromatin-remodeling factor inositol auxotrophy 80 and the histone chaperones nap1-related protein 1 and 2 in modulating auxin fluxes and the activity of the inflorescence and root apical meristems. Another interesting study highlights the impact of the epigenome in controlling tra nscriptional initiation. The single-stranded DNA-binding protein whirly1 promoted the acetylation of H3K9 and repressed the trimethylation of H3K4 to enhance the recruitment of the RNA polymerase II on the wrky53 promoter (Huang et al.).

Epigenetic alterations also control the response of plants to environmental stresses including light perception and various abiotic stresses (e.g. salinity, drought, UV-B radiation, temperature, and heavy metal toxicity). As described by Lee et al. the circadian regulation of two proteins of the Sin3-histone deacetylase complex, encoded by SAP30 function-related 1 (AFR1) and AFR2 genes, is critical for the proper regulation of Arabidopsis circadian rhythm. These two proteins directly bind to the circadaian clock associated 1 (CCA1) and pseudo-response regulator 9 (PRR9) promoters in order to locally deacetylate the histone H3 and negatively affect their expression. This is just a first level of the epigenetic regulation of the Arabidopsis circadia n clock. Indeed, Hung et al. described a more complex tra nscriptional regulation of the circadian clock: the recruitment of the lysine-specific demethylase 1 (LSD1)-like 1/2 (LDL1/2) and histone deacetylase 6 (HDA6) proteins by circadian clock associated 1 (CCA1)/late elongated hypocotyl (LHY) is needed to repress the expression of timing of cab expression 1 (TOC1). Acting as a negative feedback regulatory loop, TOC1 also intera cts with LDL1/2 and HDA6 proteins to repress the expression of CCA1/LHY. A broader picture of the role of the epigenome on the plant circadian clock is provided in the Du et al. review paper.

Environmental stresses also induce the formation of stressresponding agents such as nitric oxide. In soybean, Sun et al. revealed that the de novo deposition of trimethylated histone H3 lysine 27 residue in the promoter and coding sequence of plant genes is needed to repress their transcription in response to salt stress. Mechanistically, Ageeva-Kieferle et al. described in their review the role of nitric oxide as inhibitors of histone deacetylase through the S-nitrosation of selected cysteine residues. Nitric oxide also regulates the epigenome by controlling the expression of genes encoding DNA and histone methyltransferases and demethylases. Taken together, nitric oxide is a chemical agent controlling plant gene activity in response to environmental stresses notably by regulating the activity of various histone acetyltransferases, deacetylases, methyltransferases and demethylases, and DNA methyl transferases and demethylases.

# CONCLUDING REMARKS

This special topic clearly highlights the central role of the epigenome in the regulation of gene expression that influences many plant biological processes such as plant development and plant response to environmental stresses. A deeper analysis of the chromatin remodeling and transcription related mechanisms will be needed to better understand the epigenetic regulation of gene expression. Single cell -omic technologies such as single cell RNA-seq and ATAC-seq will enable further discoveries by capturing the transcriptome and epigenome for each cell composing a complex organ. While single cell RNA-seq was recently applied on Arabidopsis root protoplasts, there is a need to develop plant single cell ATAC-seq technology to gain a more complete picture of the plant cell epigenome.

# AUTHOR CONTRIBUTIONS

All authors listed have made a substantial, direct and intellectual contribution to the work, and approved it for publication.

# FUNDING

The work was supported by Youth Innovation Promotion Association, Chinese Academy of Sciences (2017399), Guangdong Natural Science Foundation (2018A030313350), the Strategic Priority Research Program of the Chinese Academy of Sciences (XDA13020603), the National Science Centre, Poland UMO-2014/13/B/NZ2/01187, Natural Sciences and Engineering Research Council of Canada (06679), the Chinese University of Hong Kong direct grant for research (#4053383), the National Science Foundation (awards #1854326 and #1339194), the Spanish Ministry of Economy, Industry and CompetitivenessINIA-FEDER (RTA2017-00011-C03-01) and by the Nebraska Research Initiative core facility research grant.

Conflict of Interest: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2020 Luo, Ríos, Sarnowski, Zhang, Mantri, Charron and Libault. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Epigenetic Regulation of Juvenile-to-Adult Transition in Plants

#### Yunmin Xu† , Lu Zhang† and Gang Wu\*

State Key Laboratory of Subtropical Silviculture, School of Agriculture and Food Sciences, Zhejiang Agriculture and Forestry University, Hangzhou, China

Epigenetic regulation is referred to as changes in gene function that do not involve changes in the DNA sequence, it is usually accomplished by DNA methylation, histone modifications (repressive marks such as H3K9me, H3K27me, H2Aub, or active marks such as H3K4me, H3K36me, H3Ac), and chromatin remodeling (nucleosome composition, occupancy, and location). In plants, the shoot apex produces different lateral organs during development to give rise to distinguishable phases of a juvenile, an adult and a reproductive phase after embryogenesis. The juvenile-to-adult transition is a key developmental event in plant life cycle, and it is regulated by a decrease in the expression of a conserved microRNA-miR156/157, and a corresponding increase in the expression of its target genes encoding a set of plant specific SQUAMOSA PROMOTER BINDING PROTEIN-LIKE (SPL) proteins. Recent work has revealed that the miR156/157-SPL pathway is the master regulator of juvenile-to-adult transition in plants, and genes in this pathway are subjected to epigenetic regulation, such as DNA methylation, histone modifications, and chromatin remodeling. In this review, we summarized the recent progress in understanding the epigenetic regulation of the miR156/157-SPL pathway during juvenile-to-adult transition and bring forward some perspectives of future research in this field.

Keywords: epigenetic regulation, miR156, SPL, juvenile-to-adult transition, plants

# INTRODUCTION

Unlike mammals, in which organ formation is completed during embryonic development, plants produce new organs from self-sustaining stem cell populations known as meristems in different developmental processes. In plants, post-embryonic development can be divided into a juvenile vegetative phase, an adult vegetative phase and a reproductive phase, and each developmental phase is marked by changes in a series of distinct phase-specific traits (Poethig, 1990; Kerstetter and Poethig, 1998). The transition from the juvenile vegetative phase to the adult vegetative phase was referred to as the juvenile-to-adult transition or vegetative phase change.

In Arabidopsis, the juvenile-to-adult transition is characterized by the formation of leaf abaxial trichomes, an increase in leaf length/width ratio and serration, and a decrease in cell size (Telfer et al., 1997; Tsukaya et al., 2000; Usami et al., 2009). Genetic and molecular analyses demonstrated that the conserved miRNA-miR156/157 and its target genes-SQUAMOSA PROMOTER BINDING PROTEIN-LIKE (SPL) genes act sequentially with miR172, another miRNA that targets a class of AP2-like transcription factors (TFs), to regulate juvenile-to-adult transition in plants (Wu and Poethig, 2006; Wu et al., 2009; He et al., 2018). miR156/157 is highly expressed in juvenile phase and its abundance declines gradually, while its target SPL genes increases during shoot development.

#### Edited by:

Ming Luo, South China Botanical Garden (CAS), China

#### Reviewed by:

Abdelali Hannoufa, Agriculture and Agri-Food Canada (AAFC), Canada Rosario Muleo, Università degli Studi della Tuscia, Italy

#### \*Correspondence:

Gang Wu<EMAIL_ADDRESS>†These authors have contributed

#### Specialty section:

equally to this work.

This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science

Received: 27 April 2018 Accepted: 28 June 2018 Published: 17 July 2018

#### Citation:

Xu Y, Zhang L and Wu G (2018) Epigenetic Regulation of Juvenile-to-Adult Transition in Plants. Front. Plant Sci. 9:1048. doi: 10.3389/fpls.2018.01048

**8**

miR156/157 negatively regulates SPL gene expression through transcript cleavage or translational inhibition. SPLs were also responsive to photoperiodic induction and exhibited an miR156/157-independent expression pattern (Schmid et al., 2003; Jung et al., 2012). Therefore, the outcome of SPL levels finetuned by both miR156/157 and exogeneous cues orchestrates the timing of juvenile-to-adult transition (Huijser and Schmid, 2011; Poethig, 2013).

The Arabidopsis genome encodes eight miR156 genes (MIR156A∼H) and four miR157 genes (MIR156A∼D), and those genes function redundantly. The mir156a mir156c double mutant exhibited a similar phenotype to the 35S::MIMICRY156 transgenic plants with significantly reduced levels of miR156, which indicates that MIR156A and MIR156C are the two main loci contributing to the level of miR156 and have dominant roles in vegetative phase change within the miR156 family in Arabidopsis (Yang L. et al., 2013; Yu et al., 2013). miR157 functions redundantly with miR156, but has a much smaller effect on shoot morphology and SPL gene expression than miR156 (He et al., 2018). miR156/157 targets 10 out of 16 different SPL genes in Arabidopsis. Based on the amino acid sequence of the SBP domain, the miR156/157-targeted SPL genes can be classified into five clades, SPL3/SPL4/SPL5, SPL9/SPL15, SPL2/SPL10/SPL11, SPL6, and SPL13A/B (Xie et al., 2006; Riese et al., 2007; Preston and Hileman, 2013). Genetic and functional analysis of the role of SPL genes in vegetative phase change indicated that SPL2/SPL9/SPL10/SPL11/SPL13/SPL15, but not SPL3/4/5/6, contribute to the juvenile-to-adult transition with SPL9/SPL13/SPL15 being more important for juvenile-to-adult transition than SPL2/SPL10/SPL11 (Xu et al., 2016a).

As the master regulator of the juvenile-to-adult transition, miR156/157-SPL pathway has been shown to be subjected to transcriptional and post-transcriptional regulation. Those include the transcriptional regulation of pri-MIR156/157 and SPLs genes, the regulation of miR156/157 biogenesis, and posttranscriptional regulation of SPL genes (**Figure 1**). Here, we review our current understanding of epigenetic regulation of the miR156/157-SPL pathway and the roles of corresponding players in juvenile-to-adult transition in plants.

# DNA METHYLATION

DNA methylation [5-Methylcytosine (5mC)] is a hallmark of epigenetic gene silencing in both plants and mammals (Feng et al., 2010; Law and Jacobsen, 2010). DNA methylation is found at CG or non-CG sites including CHH and CHG (H represents A, T, or C) in plants in contrast to CG sites only in mammals (Henderson and Jacobsen, 2007; Cokus et al., 2008). In plants, CG methylation is carried out by DNA METHYLTRANSFERASE 1 (MET1), whereas DOMAINS-REARRANGED METHYLTRANSFERASEs (DRM) and CHROMOMETHYLASE 3 (CMT3) are responsible for the non-CG methylation (Law and Jacobsen, 2010).

The first indication of DNA methylation plays a role in phases of shoot development comes from the work done by Brink. In the 1950s, Brink noticed the similarity between phase change in plants and changes in cell states in non-plant organisms, he proposed that phases of shoot development might be regulated by reversible changes in chromatin based on his research on paramutation in maize (Brink, 1962). Subsequent work on Spm transposable elements (Banks and Fedoroff, 1989) and the Robertson's Mutator (Mu) element (Martienssen et al., 1990) suggest that DNA methylation may be the underlying mechanism for maintaining phases of shoot development in plants. Recent work in peach also demonstrated that levels of nuclear DNA methylation was higher in adult meristems than that in juvenile and juvenile-like meristems (Bitonti et al., 2002), and an increase in DNA methylation during development seems widespread in plants (Fraga et al., 2002; Ruiz-García et al., 2005). In Arabidopsis, the triple DNA methyltransferase mutant drm1 drm2 cmt3 exhibited a developmental retardation phenotype (Cao and Jacobsen, 2002), indicating that DNA methylation is important for normal growth and development in plants. However, genome-wide DNA methylation analysis of 5-weekold Columbia wild type, met1 and drm1 drm2 cmt3 triple mutant (Zhang et al., 2006), and 25-day-old Columbia wild type (Zilberman et al., 2007) indicated that only the coding sequence of the SPL10 gene contains non-CG methylation. These results suggest that genes upstream or downstream of the miR156/157-SPL pathway, instead of miR156/157 or SPL genes, might be regulated by DNA methylation. Therefore, phenotypic characterization of vegetative phase change phenotype of mutants of DNA methyltransferases (MET1, DRM, and DNMT2) or demethylation enzymes (ROS1, DME, DML2, and DML3), as well as bisulfite sequencing of MIR156/157 and SPLs loci, will facilitate to uncover the role of DNA methylation in regulation of miR156/157-SPL pathway and juvenile-to-adult transition in plants.

# HISTONE MODIFICATION

Histone modification at specific lysine sites functions as transcription repressive marks such as H3K9me, H3K27me, H2Aub, etc., or active marks such as H3K4me, H3K36me, H3Ac, etc., this modification is catalyzed by Polycomb group (PcG) protein complexes and Trithorax group (TrxG) protein complexes, respectively (Pien and Grossniklaus, 2007; Köhler and Hennig, 2010; Grossniklaus and Paro, 2014; Kingston and Tamkun, 2014). PcG complexes are repressors of gene transcription, and function in multi-subunit complexes, such as Polycomb Repressor Complex 1 (PRC1) or Polycomb Repressor Complex 2 (PRC2) (Grossniklaus and Paro, 2014).

# PRC2 AND H3K27me3 MODIFICATION

PRC2 is a highly conserved and well-characterized PcG complex, and it represses target gene expression by trimethylating histone H3 at lysine 27 (H3K27me3) through the E(z) SET domain (Köhler and Hennig, 2010; Grossniklaus and Paro, 2014). In the

and SPL loci, respectively. Triangle indicates gradual increase or decrease in the epigenetic modification levels of MIR156 loci.

no obvious roles in juvenile-to-adult transition, miR156/157 targeted SPL genes are largely devoid of the H3K27me3 mark. These results imply that the PRC2 complex promotes SPL gene transcription indirectly by repressing the transcription of

During juvenile-to-adult transition in Arabidopsis, the decrease in the transcription of MIR156A and MIR156C loci is associated with an increase in the binding of the PRC2 complex to these two loci, causing an increase in the H3K27me3 mark in their promoter and transcribed regions as well as a decrease in the H3K27ac mark in the region immediately after transcription start sites (TSS) (Xu et al., 2016b,c). Loss-of-function mutant of SWN, but not the loss-of-function mutant of CLF, exhibited an obvious delayed juvenile-to-adult transition phenotype (Xu et al., 2016b,c). H3K27me3 was completely lost in clf swn double

fpls-09-01048 July 20, 2018 Time: 14:48 # 3

FIGURE 1 | Epigenetic regulation of juvenile-to-adult transition in plants. Active and repressive epigenetic regulators were marked in red and black at MIR156/157

mutant and it eventually dedifferentiated into a callus-like tissue, making it impossible to determine the phenotype of juvenile-to-adult transition (Xu et al., 2016b). Therefore, the question of if SWN and CLF functions redundantly in vegetative phase change remains unknown. However, the H3K27me3 mark at MIR156A/MIR156C loci was significantly reduced in clf mutants, but that in swn mutant remains controversial, which indicates that SWN and CLF may function redundantly to repress MIR156A/MIR156C by catalyzing H3K27me3 (Xu et al., 2016b,c).

#### PRC1 AND HISTONE UBIQUITINATION

PRC1 is thought to recognize the H3K27me3 mark to confer stable transcriptional repression (Lund and van Lohuizen, 2004). PRC1 is more dissimilar between Arabidopsis and animals, but it has related functions. In Arabidopsis, the function of PRC1 can be histone 2A mono-ubiquitination (H2Aub) dependent or independent. H2Aub dependent group requires the E3 ubiquitin ligase activity of Arabidopsis B lymphoma Moloney murine leukemia virus insertion region1 homolog 1A (AtBMI1A)/B/C or AtRING1A/B, while H2Aub independent group requires the activity of the EMBRYONIC FLOWER 1 (EMF1) (Yang C. et al., 2013; Calonje, 2014). BMI1-PRC1 and RING1-PRC1 are required for the repression of seed maturation program after germination, whereas EMF1-PRC1 is required for floral repression (Moon et al., 2003; Calonje et al., 2008; Chen et al., 2010).

PRC1 has been shown to be involved in juvenile-to-adult transition in Arabidopsis. BMI1-PRC1 maintains the repression of miR156 and accelerates juvenile-to-adult transition (Picó et al., 2015). The levels of MIR156A and MIR156C were upregulated in atbmi1a/b mutant and the juvenile phase was prolonged with the H2Aub and H3K27me3 marks being decreased in the TSS region of MIR156A and MIR156C (Picó et al., 2015).

RING1-PRC1 and EMF1-PRC1 function to repress SPLs to delay juvenile-to-adult transition (Li et al., 2017). In ring 1a ring 1b double mutant, the H2Aub mark was obviously decreased in the promoter and coding region of SPL3, SPL9 and SPL10, causing upregulation of these genes to accelerate the appearance of adult traits (Li et al., 2017). Therefore, PRC1 variants function in vegetative phase change mainly by targeting different MIR156/157 loci or SPL genes in the miR156/157-SPL pathway, and they have opposing roles in this process. However, how PRC1 variants recognize distinct targets still remains unclear, and more work is required to explore the mechanism of how PRC1 works.

# ATXR7 AND H3K4me3 MODIFICATION

The Arabidopsis genome encodes three H3K4 methyltransferase, namely ARABIDOPSIS TRITHORAX1 (ATX1), ATX2, and ATXR7 (Avramova, 2009). ATX1 and ATX7 are members of the Trithorax family, and ATXR7 is the only member of the SET1 subfamily in Arabidopsis (Tamada et al., 2009). atxr7-1, but not atx1-1, atx2-1, or atx1 atx2 double mutant, exhibits a precocious juvenile-to-adult transition phenotype. Chromatin immunoprecipitation (ChIP) analyses indicated that ATXR7 binds to a region adjacent to the TSS of MIR156A and deposits the H3K4me3 mark to activate MIR156A transcription (Xu et al., 2018).

# HAG1 AND HISTONE ACETYLATION

Histone acetylation is generally considered as an active epigenetic mark, which is a balanced process regulated by histone acetyltransferases (HAG1) and histone deacetylases (HDA1, HAD6). Spt–Ada–Gcn5–acetyltransferase-like histone acetyltransferase complex (SAGA-like complex) is conserved in mammals, plants, files and yeast, and General Control Nonrepressed 5 (GCN5) functions as the catalytic component for this complex (Turner, 2000).

In Arabidopsis, loss-of-function mutants in HAG1 (the Arabidopsis homolog of GCN5), hag1-6 and hag1-7, exhibited a significantly delayed juvenile-to-adult transition phenotype (Kim et al., 2015). In hag1-6 mutant, transcripts of MIR156 loci and mature miR156 remained stable; however, those of SPL3, SPL4, SPL5, SPL9, SPL11, SPL13, SPL15, and SPL8 were greatly reduced, suggesting that the regulation of SPLs by HAG1 is independent of miR156. ChIP results showed HAG1 was bound to the promoters and transcribed regions of SPL3 and SPL9 directly, leading to histone acetylation at the H3K9, H3K14, and H3K27 sites in these genes (Kim et al., 2015). HAG1-mediated H3 acetylation (H3Ac) of SPL9 is also responsive to light signals, which indicates that HAG1-mediated H3Ac of SPL9 might function as a sensor of environmental conditions to modulate the developmental process in plants (Kim et al., 2015).

# CHROMATIN REMODELING

Chromatin remodeling includes changes in nucleosome composition, nucleosome occupancy, nucleosome location, and the accessibility of the DNA to other transcriptional regulators.

# SWR1-C AND H2A.Z HISTONE VARIANT

ATP-dependent SWR1 chromatin remodeling complex (SWR1- C) functions in exchanging the histone H2A-H2B dimer with the H2A.Z-H2B dimer, and then produces nucleosome variant (Mizuguchi et al., 2004; Luk et al., 2010). In Arabidopsis, mutations in the SWR1-C subunit coding genes (ARP6, SEF, and PIE1) and H2A.Z coding genes (HTA8, HTA9, and HTA11) exhibited a similar pleiotropic phenotype, which indicates that the primary function of SWR1-C is to deposit H2A.Z (Mizuguchi et al., 2004; Wu et al., 2005). However, the mechanism of H2A.Z modification by SWR1-C to regulate different target gene expression is distinguishable in that H2A.Z can change the nucleosome occupancy to destabilize nucleosomes or to increase nucleosome stability and/or to function with H3K4me3 mark together (Martin-Trillo et al., 2006; Kumar and Wigge, 2010; Choi et al., 2013).

In arp6 and hat9/hat11 mutants, MIR156A/MIR156C transcripts were reduced and juvenile-to-adult transition was accelerated (Choi et al., 2016; Xu et al., 2018). ChIP with H2A.Z antibody showed that H2A.Z was enriched at the first 500 nucleotides after TSS of MIR156A/MIR156C, and the level of H2A.Z was significantly reduced in arp6 mutant. However, H2A.Z level does not change significantly during juvenile-to-adult transition, suggesting that H2A.Z and SWR1-C contribute to maintaining the expression of MIR156A/MIR156C early in shoot development, but do not regulate the timing of juvenile-to-adult transition (Xu et al., 2018). MIR156A transcript was reduced in arp6 mutant due to higher nucleosome occupancy in its promoter region (Choi et al., 2016); however, it was suggested that H2A.Z increases the expression of MIR156A/MIR156C by promoting the deposition of H3K4me3 rather than by decreasing nucleosome occupancy in the MIR156A promoter region (Xu et al., 2018).

#### ATP-DEPENDENT CHROMATIN REMODELING PROTEIN

BRAHMA (BRM) is the ATPase subunit of the most widely studied SWI2/SNF2 chromatin remodeling protein complex. It uses the energy derived from ATP hydrolysis to change the histone octamer-DNA interaction (Saha et al., 2006; Clapier and Cairns, 2009). BRM regulates MIR156A transcription by directly binding to the promoter region and maintaining low occupancy of the −2 and +1 nucleosomes proximal to the TSS. brm mutants exhibit an accelerated juvenile-to-adult transition phenotype by reducing the transcription of MIR156A (Xu et al., 2016c). BRM also antagonizes the function of SWN in the PRC2 complex to remove H3K27me3 repressive mark in MIR156A (Xu et al., 2016c).

PICKLE (PKL) is a CHD3 ATP-dependent nucleosome remodeling protein, which is physically associated with the nucleosome remodeling and deacetylation complex (Perruc et al., 2007; Zhang et al., 2008; Ho et al., 2013). PKL is bound to the TSS adjacent region of MIR156A/MIR156C to promote the juvenile-to-adult transition by repressing the transcription of MIR156A/MIR156C. In pkl mutants, MIR156A/MIR156C transcripts were elevated due to the reduction in nucleosome occupancy at the +1 position, an increase in the H3K27ac mark, and a corresponding decrease in the H3K27me3 mark in the promoter and transcribed region (Xu et al., 2016b).

 Recent studies have revealed that DNA methylation, histone modification, chromatin remodeling play important roles in regulating the expression of some components in the miR156/157-SPL pathway. However, there are still some critical questions remain to be solved as illustrated in **Figure 2**.

#### HOW ARE EPIGENETIC REGULATORS RECRUITED TO THE MIR156/157 AND/OR SPLs LOCI?

MIR156/157 and/or SPL loci are subjected to epigenetic regulation to modulate juvenile-to-adult transition in plants. However, these epigenetic regulators, by their own, have no DNA binding specificity. Therefore, a central question is how these epigenetic regulators are recruited to their target genes.

PRC2-mediated H3K27me3 is a conserved epigenetic modification between plants and the animal kingdom (Mozgova and Hennig, 2015; Xiao and Wagner, 2015). Recent genomic study in Arabidopsis showed that PRC2 components bind to specific DNA motifs called Polycomb response elements (PREs) by interacting with specific TFs (Xiao et al., 2017). Interestingly, six top enriched motifs (CTCC, CCG, G-box, GA repeat, ACrich, and Telobox motifs) out of 170 computationally defined PREs were present at the MIR156A locus (Xiao et al., 2017). The GA repeat and Telobox motifs were present adjacent to the TSS region of MIR156A and MIR156C loci together, these motifs are the potential binding sites for class I BPC and C1-2iD TFs, respectively. This information will be helpful to identity TFs through which the PRC2 complex interacts to be recruited to the MIR156A/MIR156C loci during juvenile-to-adult transition.

#### EPIGENETIC MODIFICATION OF MIR156/157 AND/OR SPL LOCI BY STRESS?

Plants are sessile organisms and they are forced to adapt to the changing environment. The miR156/157-SPL pathway functions as the master regulator of juvenile-to-adult transition and flowering (Wang et al., 2009; Wu et al., 2009). Therefore, plants evolved a precise mechanism to adapt to the environment by shortening or prolonging the juvenile phase or changing the flowering time. Under salt or drought stress conditions, miR156 was induced to maintain plants in the juvenile phase for a relatively longer time; when they were returned to favorable conditions, miR156 was suppressed to accelerate the developmental transition (Cui et al., 2014). Under UV-B radiation conditions, the PRC2-mediated H3K27me3 modification in the MIR156A/MIR156C loci was decreased, and the corresponding up-regulation of miR156 delayed juvenile-to-adult transition (Dotto et al., 2018). Other studies also indicate that the expression of miR156 is responsive to ambient temperature (Stief et al., 2014), phosphate starvation (Hsieh et al., 2009), CO2 treatment (May et al., 2013), suggesting a tight interaction between juvenile-to-adult transition and environment through the miR156/157-SPL pathway.

Epigenetic modification is a reversible mark, which can be removed or deposited to target genes to affect their expression in response to changing environment. It will be of great interest

to learn how epigenetic modification patterns of MIR156/157 and/or SPL loci change in response to external cues, especially to environment stresses, as well as how this changing environment affects the juvenile-to-adult transition.

#### REVERSIBLE EPIGENETIC REGULATION OF miR156/157 RESETTING?

During juvenile-to-adult transition, miR156/157 transcription was reduced or silenced gradually to ensure the plant to enter the adult phase and flower. This is achieved by disposing of active epigenetic marks such as H3K4me3, H3K27ac and depositing some repressive epigenetic marks such as H3K27me3 to miR156 loci. Interestingly, this silencing process needs to be reset to an active state in each generation as miR156/157 is de-repressed again to be highly expressed in the pro-embryo stage (Nodine and Bartel, 2010) after flowering.

A similar example of Off-Resetting pattern in plant lifecycle is the regulation of FLOWERING LOCUS C (FLC). FLC is silenced by depositing H3K27me3 mark under winter cold treatment, and the silenced state was maintained in the mature pollen grains and the egg cells (De Lucia et al., 2008; Sheldon et al., 2008). In pro-embryo stage, FLC is activated by depositing active epigenetic marks such as H3K4me3, H3K36me3, and disposing of repressive marks such as H3K27me3. LEAFY COTYLEDON1 (LEC1), a seed-specific pioneer TF (Tao et al., 2017), and EARLY FLOWERING 6 (ELF6), a H3K27me3 demethylase (Crevillén et al., 2014), were shown to play critical roles in FLC reactivation.

As for MIR156/157, it is still unknown where and when the de novo re-activation occurs. Moreover, whether the resetting of miR156/157 depends on a reversible epigenetic regulation still remains elusive. Further study of when, where and how miR156/157 Off-Reset pattern is initiated during plant life cycle will be an important future task.

# AUTHOR CONTRIBUTIONS

YX and LZ wrote the article, and GW revised it.

# FUNDING

This work was supported by the National Natural Science Foundation of China (Grant Nos. 31770209 and 31700249), a start-up fund from the Zhejiang Agricultural & Forestry University (Grant No. 2013FR084), and a fund from the Education Department of Zhejiang Province (Grant No. Y201635908).

#### REFERENCES

fpls-09-01048 July 20, 2018 Time: 14:48 # 7

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Xu, Zhang and Wu. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

fpls-09-01048 July 20, 2018 Time: 14:48 # 8

# Regulation of Plant Growth and Development: A Review From a Chromatin Remodeling Perspective

Simon P. Ojolo1,2† , Shijiang Cao1,3† , S. V. G. N. Priyadarshani1,2, Weimin Li1,4 , Maokai Yan1,5, Mohammad Aslam1,6, Heming Zhao1 and Yuan Qin1 \*

1 Fujian Provincial Key Laboratory of Haixia Applied Plant Systems Biology, Key Laboratory of Genetics, Breeding and Multiple Utilization of Crops, Ministry of Education, State Key Laboratory of Ecological Pest Control for Fujian and Taiwan Crops, Fujian Agriculture and Forestry University, Fuzhou, China, 2 College of Crop Science, Fujian Agriculture and Forestry University, Fuzhou, China, 3 College of Forestry, Fujian Agriculture and Forestry University, Fuzhou, China, 4 College of Resources and Environment, Fujian Agriculture and Forestry University, Fuzhou, China, 5 College of Life Sciences, Fujian Agriculture and Forestry University, Fuzhou, China, 6 College of Plant Protection, Fujian Agriculture and Forestry University, Fuzhou, China

#### Edited by:

Ming Luo, South China Botanical Garden (CAS), China

#### Reviewed by:

Chenlong Li, Agriculture and Agri-Food Canada (AAFC), Canada Xigang Liu, Institute of Genetics and Developmental Biology (CAS), China

#### \*Correspondence:

Yuan Qin<EMAIL_ADDRESS><EMAIL_ADDRESS>
†These authors have contributed equally to this work

#### Specialty section:

This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science

Received: 11 March 2018 Accepted: 03 August 2018 Published: 22 August 2018

#### Citation:

Ojolo SP, Cao S, Priyadarshani SVGN, Li W, Yan M, Aslam M, Zhao H and Qin Y (2018) Regulation of Plant Growth and Development: A Review From a Chromatin Remodeling Perspective. Front. Plant Sci. 9:1232. doi: 10.3389/fpls.2018.01232 In eukaryotes, genetic material is packaged into a dynamic but stable nucleoprotein structure called chromatin. Post-translational modification of chromatin domains affects the expression of underlying genes and subsequently the identity of cells by conveying epigenetic information from mother to daughter cells. SWI/SNF chromatin remodelers are ATP-dependent complexes that modulate core histone protein polypeptides, incorporate variant histone species and modify nucleotides in DNA strands within the nucleosome. The present review discusses the SWI/SNF chromatin remodeler family, its classification and recent advancements. We also address the involvement of SWI/SNF remodelers in regulating vital plant growth and development processes such as meristem establishment and maintenance, cell differentiation, organ initiation, flower morphogenesis and flowering time regulation. Moreover, the role of chromatin remodelers in key phytohormone signaling pathways is also reviewed. The information provided in this review may prompt further debate and investigations aimed at understanding plant-specific epigenetic regulation mediated by chromatin remodeling under continuously varying plant growth conditions and global climate change.

Keywords: chromatin remodeling, SWI/SNF complexes, histones, gene regulation, meristem, hormone signaling, plant development, flowering

# INTRODUCTION

The packaging of genetic material into nucleosomes is a distinctive evolutionary feature of eukaryotic cells. Nucleosomes are repetitive units consisting of 147 bp DNA strands tightly wrapped around a central octamer comprising one histone heterotetramer (H3/H4) and two histone heterodimers (H2A/H2B). Histones are highly conserved proteins found in all eukaryotes as core nucleosome units (Suzuki and Bird, 2008). Together, the histone octamer and DNA resemble "beads" on a string and are referred to as chromatin (Luger et al., 1997). This packaging ensures that long DNA strands are tightly condensed by supercoiling to precisely fit

into the nucleus. Such compaction of nucleosomes in chromatin hinders DNA accessibility to important regulatory proteins essential for various nuclear processes. However, cells have evolved in response to this impediment through chromatin remodeling (Groth et al., 2007).

Gene regulatory mechanisms in eukaryotic cells (such as transcription, DNA repair and replication) act upon chromatin structure as a substrate. These activities induce cellular changes and regulate gene expression in a number of biological processes, including genome stability, recombination, developmental reprogramming and response to extracellular signals (Feng et al., 2010; Soria et al., 2012; Zhu et al., 2013). The regulatory precision of these fundamental genome processes relies on the high fidelity of chromatin remodeling mechanisms that permit temporal access to – or blocking of – vital DNA sequences, such as gene promoters. Histone modifications, nucleosome remodeling and DNA methylation have been shown to regulate chromatin structure and gene expression (Henikoff and Shilatifard, 2011).

There are two main players regulating chromatin dynamics: (1) chromatin remodelers that alter DNA-histone interactions by energy harnessed through ATP hydrolysis and (2) nucleosome-modifying enzymes that modulate DNA and histone residues by specifically adding or removing covalent modifications (Jerzmanowski, 2007). Organisms rely on gene expression regulation to achieve normal cell differentiation, organogenesis, growth and development. Moreover, gene expression regulation is temporally and spatially coordinated via crosstalk between chromatin remodeling complexes (CRCs) and the transcription machinery (Deem et al., 2012). In plants, such precise control of gene expression mediated by chromatin modifications in response to endogenous and environmental stimuli is fundamental for proper development and reproductive success (Sarnowski et al., 2005; Bezhani et al., 2007; Han et al., 2012; Archacki et al., 2013; Efroni et al., 2013; Sarnowska et al., 2013; Qin et al., 2014; Vercruyssen et al., 2014).

Plant development can be divided into the embryonic and postembryonic phases. The embryonic development phase includes the establishment of the seedling apical-basal axis, the shoot apical meristem (SAM) and the root apical meristem (RAM). The subsequent establishment of the leaf, stem and flower meristems occurs in the postembryonic phase. In this review, we discuss recent investigations underpinning the involvement of SWI/SNF chromatin remodeling ATPases in regulating key plant growth and development processes, including meristem establishment and maintenance, cell differentiation, organ initiation, flower morphogenesis and flowering time regulation. We also discuss the role of chromatin remodelers in the plant response to key phytohormone signals. This overview may be useful for framing the current knowledge gaps, thereby stimulating further debate and research aimed at a comprehensive understanding of epigenetic regulatory mechanisms in plant development. This in turn may guide molecularbased plant improvement techniques for desirable agronomic traits.

# CLASSIFICATION OF EUKARYOTIC CHROMATIN REMODELING COMPLEXES

Chromatin remodeling complexes are evolutionarily conserved multi-unit protein complexes that regulate chromatin structure by altering nucleosome composition and interactions (Narlikar et al., 2002). A common feature of all purified CRCs is that they harbor an ATPase/helicase of the SWITCHING DEFECTIVE2/SUCROSE NON-FERMENTING2 (SWI2/SNF2) family as their catalytic core. The SWI/SNF family is part of superfamily2 (SF2), a large family of helicases and translocases, and is named after the first identified CRC (Peterson et al., 1994). SWI/SNF CRCs utilize the energy derived from ATP hydrolysis to control accessibility to vital DNA sequences by influencing nucleosome structure and position and determining the kind of variant histone subspecies to be incorporated (Eisen et al., 1995; Clapier and Cairns, 2009).

A number of Arabidopsis SWI/SNF complex subunits have been identified based on sequence similarity with metazoan subunits. These include four SWI2/SNF2 ATPases (BRAHMA [BRM], SPLAYED [SYD], MINU1/CHR12 and MINU2/CHR23) (Knizewski et al., 2008; Han et al., 2015); four SWI3 proteins (SWI3A to SWI3D); two SWI/SNF ASSOCIATED PROTEINS 73 (SWP73A/CHC2 and SWP73B/CHC1, also called BRAHMA ASSOCIATED FACTOR 60 (BAF60) in humans); two ACTIN RELATED PROTEINS belonging to SWI/SNF complexes (ARP4 and ARP7); and the BUSHY (BSH) protein (Meagher et al., 2005; Jerzmanowski, 2007; Kwon and Wagner, 2007; Sang et al., 2012). Eukaryotic SWI2/SNF2 family chromatin remodelers can be categorized into four classes/subfamilies based on phylogenetic analysis, and all four of these subfamilies are represented in plants (Farrona et al., 2008).

#### SWI/SNF Subfamily Remodelers

The SWI/SNF subfamily of remodelers initially purified from Saccharomyces cerevisiae is probably the most comprehensively studied. SWI/SNF subfamily CRCs have a well-established role in gene expression regulation. The SWI/SNF subfamily remodelers contain 8–14 subunits, and the catalytic ATPases of most SWI/SNF subfamily remodelers are composed of a helicase-SANT domain, a post-HSA domain and a C-terminal bromodomain. While the fungal SWI/SNF subfamily ATPases consist of a pair of actin-related proteins (ARPs) (Cairns et al., 1998), higher orthologs of SWI/SNF subfamily complexes contain a dimer of an ARP and actin (Lessard et al., 2007). The mode of action of SWI/SNF complexes has been characterized as sliding and/or ejecting nucleosomes at many target loci, but they reportedly have no vital role in the assembly of eukaryotic chromatin structure (Clapier and Cairns, 2009). It was recently reported that the Arabidopsis SWI/SNF complex expedites its role in activating and repressing target gene expression by binding to both promoters and terminators and that it regulates the expression of both promoter-centered genes and non-coding RNAs (Archacki et al., 2017).

# Imitation Switch (ISWI) Subfamily Remodelers

The ISWI subfamily remodelers have 2–4 subunits. Eukaryotic ISWI remodeler complexes typically have 1 or 2 distinct catalytic subunits and specialized attendant proteins that give rise to various domains. These domains include plant homeodomains, bromodomains, DNA-binding histone fold motifs and additional DNA-binding motifs (Clapier and Cairns, 2009). ISWI complexes were initially purified from Drosophila melanogaster. They include the nucleosome remodeling factor (dNURF), chromatin accessibility complex (dCHRAC) and ATP-utilizing chromatin assembly and remodeling factor (dACF) (Corona and Tamkun, 2004). SANT-like ISWI ATPases contain a SANT domain adjacent to a SLIDE domain in their C-terminus, which acts as a binding site for unmodified histone tails and DNA. The diversity resulting from attendant subunits is exhibited in the modes of action of the ISWI family complexes. Both ACF and CHRAC have been shown to promote chromatin assembly and transcription repression through the optimization of nucleosome spacing, whereas NURF assists RNA polymerase II (RNAPII) activation by randomizing nucleosome spacing (Corona and Tamkun, 2004).

# Chromodomain Helicase DNA-Binding (CHD) Subfamily Remodelers

The CHD subfamily remodelers were first purified from Xenopus laevis and comprise 1–10 subunits. The N-terminus of their catalytic subunit consists of two chromodomains arranged tandemly, with the composition varying from monomeric in lower eukaryotes to multimeric in vertebrates (Marfella and Imbalzano, 2007). CHD subfamily remodelers vary in their structure, composition and function, due in part to the diversity of their chromodomains. One of the members of the CHD subfamily in vertebrates, the nucleosome remodeling and deacetylase (Mi-2/NuRD) complex, has repressive roles due to its inclusion of histone deacetylases (HDAC1/2) and methyl CpG-binding domain (MBD) proteins. In contrast, other CHD subfamily remodelers have been reported to promote transcription by either ejecting or sliding nucleosomes along the DNA double strand (Denslow and Wade, 2007).

# Inositol Requiring 80 (INO80) Subfamily Remodelers

The INO80 subfamily is the most recently characterized chromatin remodeler subfamily. Many INO80 subunit homologs have been identified in yeast, fruit fly, mammals and plants, making it the most conserved subfamily. The presence of a split ATPase subunit distinguishes INO80 complexes from the SWI/SNF, CHD and ISWI subfamilies (Morrison and Shen, 2009). In S. cerevisiae, the INO80 subfamily is represented by two complexes, INO80 and Swi2/snf2-related 1 (SWR1); in mammals, it is represented by three complexes, INO80, Snf2-related CBP activator protein (SRCAP) and p400. D. melanogaster has INO80 and p400 complexes (Morrison and Shen, 2009; Bao and Shen, 2011). S. cerevisiae INO80 and SWR1 ATPases are approximately 1.2–1.5 MDa in mass and are reported to contain 15 and 14 subunits, respectively (Shen et al., 2000; Krogan et al., 2003; Mizuguchi et al., 2004). In general, the INO80 subfamily has a relatively conserved composition of individual complexes and a high degree of homology in the ATPase subunit. Studies of the S. cerevisiae SWR1 complex indicate the importance of a spacer (split) region in the ATPase subunit, as deletion of this region leads to dissociation of a number of subunits from the complex, including the RuvB-like Rvb1 and Rvb2 subunits (Wu et al., 2005). The INO80 subfamily is responsible for various functions in eukaryotic cells including transcriptional activation and DNA double-strand break (DSB) repair (Ebbert et al., 1999).

#### INVOLVEMENT OF SWI/SNF ATPases IN MERISTEM ESTABLISHMENT AND MAINTENANCE

Plant development is characterized by the presence of stem cells with the capacity to self-renew and transform into tissuespecific founder cells. The stem cells of the RAM and SAM have been widely studied in plants and are found at the root and shoot tips, respectively (Shen and Xu, 2009). Stem cell homeostasis and identity in both plants and mammals are associated with the activity of SWI/SNF chromatin remodeling ATPases (Ori et al., 2000; Kwon et al., 2005; Ho et al., 2009; Aichinger et al., 2011). Perturbations in the Polycomb group (PcG) repressive complex, histone acetylation or chromatin assembly may lead to improper RAM or SAM development (Phelps-Durr et al., 2005; Schubert et al., 2006; Barrero et al., 2007; Xu and Shen, 2008; Kornet and Scheres, 2009).

SYD promotes SAM maintenance by binding to the promoter region of its target gene WUSCHEL (WUS) and activating WUS expression in the organizing center (OC) region of the SAM (**Figure 1**). Consistent with this function, Arabidopsis syd mutant plants exhibit reduced WUS transcript levels accompanied by abnormal SAM development (Kwon et al., 2005).

The CHR12 and CHR23 ATPases have redundant roles in plant development processes including meristem initiation and maintenance. Plants with strong double mutant chr12chr23 combinations show embryonic lethality, endosperm defects and failure to initiate stem cell populations in both roots and shoots, whereas plants with weak double mutant chr12chr23 combinations are small viable plants with noticeable defects in RAM and SAM maintenance (Sang et al., 2012).

A study by Wu et al. (2015) demonstrated a role of chromatin in local developmental progressions in the root and shoot. In the SAM, cells with high auxin levels initiate proteolysis of Aux/IAA leading to the dissociation of the corepressor complex TOPLESS-HISTONE DEACETYLASE 19

(TPL-HDA19). This enables the recruitment of SYD and BRM to direct the acquisition of flower primordium founder cell fate (Wu et al., 2015). In the RAM, a small group of mitotically inactive cells known as the quiescent center (QC) maintains the root stem cells. BRM was found to act in the PLETHORA (PLT) pathway to maintain the root stem cell niche by altering the expression of PIN-FORMED (PIN) genes responsible for auxin distribution in the RAM, as illustrated in **Figure 1**. brm mutants exhibit faulty root stem cell niche maintenance, reduced meristem activity and retarded root growth (Yang et al., 2015).

To attain continuous plant growth, cells derived from pluripotent stem cells must undergo asymmetric cell division and expansion. This allows for the generation of lateral organs and the maintenance of stem cell populations for additional growth (Jarillo et al., 2009). The role of BRM in ensuring asymmetric cell division and cell fate determination in both plant and animal stem cells was discussed extensively in a recent review (Pillitteri et al., 2016).

Our current knowledge of mechanisms involving chromatinmediated regulation of plant stem cell initiation and maintenance remains limited to the few Arabidopsis SWI/SNF complexes discussed above. However, emerging evidence suggests complex interconnections among many players including key hormone signaling pathways. This underscores the ability of plants to activate or arrest lateral organ founder cell formation in response to both internal developmental signals and external biotic and abiotic stress signals. These attributes may eventually be harnessed or targeted for crop improvement applications.

#### CHROMATIN REMODELING IN CELL DIFFERENTIATION, ORGAN INITIATION AND DEVELOPMENT

Stem cells in the SAM initiate organ founder cells that later differentiate into aboveground organ-specific tissues such as leaves and stems (Sinha, 1999; Laux, 2003). Plant roots are formed from a reservoir of undifferentiated cells in the RAM called the root stem cells. Mature plant organs usually maintain relatively undifferentiated cells as a fallback for hormonal stimuli or mechanical injury, helping to generate new tissues and organs through cellular reprogramming (Ikeuchi et al., 2015). Cellular differentiation results from global changes in gene expression patterns and these changes involve many transcription regulators and are epigenetically mediated by chromatin remodeling (Bruex et al., 2012; Taylor-Teeples et al., 2015).

ANGUSTIFOLIA3 (AN3)/GRF-INTERACTING FACTOR1 (GIF1), a member of the GIF family of transcriptional coactivators, plays a key role in Arabidopsis shoot development (Kim and Kende, 2004), cotyledon identity establishment during

embryogenesis (Kanei et al., 2012) and leaf size increase resulting from increased cell number (Horiguchi et al., 2005; Lee et al., 2009). Transcriptional coactivators often work together with DNA-binding transcription factors to promote transcription either by recruiting chromatin remodelers or stimulating general complex formation around RNA polymerase II (Pol II). Genetic interaction between AN3 and the Arabidopsis SWI/SNF chromatin remodeling complex BRM (**Figure 2**) suggests that AN3 recruits SWI/SNF complexes to promote cell division during leaf development (Vercruyssen et al., 2014). BRM is recruited to its target gene loci via association with the plant-specific H3K27 demethylase RELATIVE OF EARLY FLOWERING 6 (REF6). Through its zinc-finger (ZnF) domains, REF6 recognizes genomic loci harboring a CTCTGYTY motif (Li et al., 2016). A forward genetic screen in Arabidopsis indicated that BRM and SWINGER (SWN) (a key component of Polycomb Group Repressive Complex 2 in plants) antagonistically control vegetative phase change through the temporal expression of miR156 at the nucleosome level. Specifically, the accelerated vegetative phase change of brm mutants was accompanied by reduced miR156 expression and increased levels of H3K27me3 at the MIR156A locus (Xu et al., 2016).

SWP73 subunits of Arabidopsis SWI/SNF CRCs function in various plant development pathways including the regulation of leaf and flower development (Vercruyssen et al., 2014; Sacharowski et al., 2015). For example, SWP73B directly binds to the promoters of the KANADI1/3, YABBY2/3/5 and ASYMMETRIC LEAVES1/2 genes involved in leaf development (**Figure 2**) (Sacharowski et al., 2015). RNA interference (RNAi) silencing of SWP73B leads to dwarfism in Arabidopsis, evidencing its role in plant development (Crane and Gelvin, 2007).

A study of the chromatin remodeler protein ZmCHB101, which is the core subunit of maize SWI3, revealed its key roles in normal maize growth and development. ZmCHB101 controls the transcriptional reprogramming of a set of genes involved in gene expression regulation, photosynthesis, metabolic regulation and stress response. The RNAi maize lines generated in the study exhibited improper tassel and cob development and abaxially curling leaves caused by increased bulliform cell numbers (Yu et al., 2016).

Chlorophyll biosynthesis is a critical mark of the transition from heterotrophic to autotrophic growth in plants. Physical interaction between BRM and the transcription factor PHYTOCHROME-INTERACTING FACTOR 1 (PIF1) was shown to regulate chlorophyll biosynthesis in Arabidopsis (**Figure 2**). When exposed to light, dark-grown brm plants exhibit higher greening rates, reduced protochlorophyllide accumulation and lower levels of reactive oxygen species (ROS) compared to wild-type plants; there is also increased expression of NADPH:protochlorophyllide oxidoreductase A (PORA), PORB and PORC, enzymes which accelerate a key step in chlorophyll biosynthesis (Zhang et al., 2017).

The importance of chromatin remodelers for proper plant growth progression and development is exemplified by their involvement in asymmetric cell differentiation and in initiating organ founder cells. Proper plant growth also involves precise developmental phase transitions, and the antagonism between SWI/SNF ATPases and PcG repressive complex components is crucial for these shifts.

#### CHROMATIN REMODELER-MEDIATED FLOWERING AND FLOWER MORPHOGENESIS

The precise regulation of the transition from vegetative growth to flowering is paramount for plant reproductive success. This process is characterized by the transition of the vegetative

SAM into an inflorescence meristem (IM) and the initiation of floral meristems (FMs) (Sablowski, 2007; Kaufmann et al., 2010). While SAM maintenance ensures indeterminate plant growth, the determinate nature of FMs determines reproductive success, seed development and the yield of agricultural crops (Liu et al., 2011). Complex regulatory networks of transcription factors and chromatin remodelers guide flowering time and flower development while integrating both internal and external signals (Wils and Kaufmann, 2017). These networks comprise photoperiod, vernalization and thermo-sensory pathways for sensing long days, cold winter and ambient temperature, respectively, together with pathways responsive to internal factors, such as the age pathway and the gibberellins (GA) signaling pathway (He, 2012).

In Arabidopsis, a repressor complex that consists of the two MADS box transcription factors MADS AFFECTING FLOWERING 4/5 (MAF4/5), FLOWERING LOCUS C (FLC) and SHORT VEGETATIVE PHASE (SVP) serves as a negative regulator of flowering time. The components directly repress the expression of the floral pathway integrators FLOWERING LOCUS T (FT) and SUPPRESSOR OF OVEREXPRESSION OF CONSTANS 1 (SOC1) (Lee et al., 2007; Li et al., 2008; He, 2012). FRIGIDA (FRI) promotes higher FLC levels that inhibit flowering (Michaels and Amasino, 2001) through the recruitment of multiple active chromatin modifications at the FLC locus (Choi et al., 2011). Vernalization overrides the FRI-mediated activation of FLC expression and thereby enables flowering (Kim et al., 2009; Crevillenn and Dean, ´ 2011).

The core subunit components of the Arabidopsis SWR1 chromatin remodeling complex (including PHOTOPERIOD-INDEPENDENT EARLY FLOWERING 1 (PIE1), ACTIN RELATED PROTEIN 6 (ARP6) and SWR1 COMPLEX 6 (SWC6)/SERRATED LEAVES AND EARLY FLOWERING (SEF)) have been shown to play important roles in regulating the proper growth and development of most plant organs. Importantly, SWR1 controls plant development by generating a balance between microRNAs and target mRNAs at the transcriptional level (Choi et al., 2016). It was recently demonstrated that SWR1 regulates gene expression by establishing lowly accessible and highly accessible nucleosome structure at the first nucleosome upstream and downstream of the transcription start site (TSS), respectively (Dai et al., 2017). SWR1 subunit loss-offunction mutants exhibit pleiotropic phenotypes including early flowering, serrated leaves, frequent absence of inflorescence internodes, bushy growth and flowers with altered organ number and size (Choi et al., 2005; Deal et al., 2005; Martin-Trillo et al., 2006; March-Diaz et al., 2007; Qin et al., 2014). H2A.Z deposition around the FLC TSS by SWR1 is necessary for FRI-meditated activation of FLC (**Figure 3**). One line of evidence is that functional disruption of SWR1 prevents H2A.Z deposition at FLC chromatin, suppressing its expression and leading to early flowering (Choi et al., 2007; Deal et al., 2007). Additionally, PIE1 mutations were earlier shown to substantially reduce FLC transcript levels, with a concomitant conversion of the winter-annual habit to the rapid flowering summer-annual habit (Noh and Amasino, 2003). A recent study revealed an Arabidopsis homolog of the yeast SANT domain protein Swc4/Eaf2 as a novel SWR1-C subunit. SWC4 is a DNA-binding protein that contributes to the recruitment SWR1-C through specific recognition of AT-rich DNA segments in chromatin regions of target genes to deposit H2A.Z. Further, knock-out and knockdown studies showed that SWC4 is essential for both embryo viability and the control of postembryonic processes, including flowering time, by repressing transcription of a number of genes including the floral integrator FT and key transcription factors (Gomez-Zambrano et al., 2018).

The AtINO80 complex is required for somatic homologous recombination (HR) and expression regulation of FLC and MAF4/5 by facilitating the enrichment of H2A.Z at their ends. Independent Atino80-5 and Atino80-6 mutant alleles display similar pleiotropic phenotypes of small plant size, reduced organ size and late flowering. These observations provide a link between plant responses to environmental and developmental signals and epigenetic mechanisms involved in plant chromatin stability (Zhang et al., 2015).

The SWP73B (BAF60) subunit of the Arabidopsis SWI/SNF complex has also been implicated in flowering time control through its involvement in chromatin loop formation at the FLC locus (Jegu et al., 2014). SWP73A has been shown to be confined to flowering time modulation under short day conditions, with functional overlap between SWP73A and SWP73B during embryogenesis (Sacharowski et al., 2015).

SVP, another key flowering repressor, is highly expressed during the vegetative phase to promote growth (Hartmann et al., 2000) but is down-regulated during the floral transition by the autonomous and GA signaling pathways (Li et al., 2008). FT and SOC1 are subsequently activated to promote flowering (**Figure 3**). One study proposed that BRM controls flowering time in Arabidopsis by directly activating SVP expression (Li et al., 2015). Genome-wide analysis of H3K27me3, a histone mark associated with gene repression, in brm mutant seedlings revealed increased H3K27me3 deposition at several genes including SVP, indicating an antagonism between BRM and PcG repressor proteins (Li et al., 2015).

Another key flowering time regulator in Arabidopsis activated by CONSTANS (CO) under long days to induce flowering is the FT-encoded florigen protein (FT) (Corbesier et al., 2007). FT moves from the phloem to the SAM and forms a complex with the bZIP transcription factor FD to activate the expression of the FM identity genes LEAFY (LFY) and APETALA1 (AP1). This activation leads to the formation of the floral primordium, which sequentially generates three types of lateral floral organs (sepals, petals and stamens) (Zik and Irish, 2003; Abe et al., 2005; Wigge et al., 2005). LFY recruits SYD and BRM ATPases to induce the expression of the floral homeotic regulator AGAMOUS (AG) by removing H3K27me3 marks on its second intron (**Figure 3**) (Wu et al., 2012). The SWR1 complex is also involved in FT expression regulation, with functional disruption of SWR1 leading to temperatureinsensitive FT activation and early flowering. An ambient temperature increase from 17 to 27◦C causes eviction of the

H2A.Z nucleosomes, which enables FT transcription by Pol II

(Kumar and Wigge, 2010). Studies of the CHD3 chromatin remodeler PICKLE (PKL) have highlighted its role in plant reproductive development by promoting crosstalk between the sporophytic and gametophytic generations. Loss of PKL in the maternal sporophyte leads to improper development of the Arabidopsis female gametophyte, integument and pollen tube, accompanied by delayed ovule and embryo development (Carter et al., 2016).

It was recently shown that during megasporogenesis, somatic cells cooperatively use SWR1 to restrict female reproductive founder cell specification to a single cell in the ovule primordia by incorporating H2A.Z at a particular WRKY28 nucleosome and promoting expression of the gene (Zhao et al., 2018). The WRKY28 transcription factor is exclusively expressed in hypodermal somatic cells surrounding the megasporocyte and represses those cells from differentiating into functional megasporocytes (Zhao et al., 2018). Other separate findings demonstrated that both BRM and SWR1 are involved in determining inflorescence architecture in response to developmental cues (Zhao et al., 2015; Cai et al., 2017).

The above findings reveal mechanisms involving the recruitment of SWI/SNF ATPases to specific flower development regulatory pathways at an appropriate time. They also emphasize the centrality of SWI/SNF complexes in shaping plant growth and ensuring the continuity of plant viability through generations by precise spatiotemporal control of the relevant developmental processes. Nevertheless, these conclusions about important regulatory mechanisms are generally based on findings from model plants such as Arabidopsis. Thus, the understanding of such regulatory mechanisms in agriculturally important crop species remains inadequate and will require further investigation.

#### CHROMATIN REMODELERS IN PHYTOHORMONE SIGNALING PATHWAYS

Plant hormones are critical for the proper regulation of plant growth and development processes including seed germination, vegetative and reproductive growth and abiotic stress response. The biosynthesis and degradation of plant hormones are also

tightly controlled through the transcriptional regulation of target genes. SWI/SNF ATPases are involved in plant hormone responses through physical interaction with hormone signaling pathway components and transcriptional regulators of genes involved in hormone biosynthesis and perception (Sarnowska et al., 2016).

Abscisic acid (ABA) is an important phytohormone that promotes seed dormancy and arrests growth in postgermination embryos under water stress conditions. In the absence of stress stimuli, BRM was initially implicated in repressing the activity of the ABA pathway, with adult brm mutants exhibiting increased drought tolerance (Han et al., 2012). It was later revealed that important ABA signaling pathway components physically interact with BRM, leading to post-translational phosphorylation/dephosphorylation of BRM. For example, phosphorylation of BRM by SnRK2 kinases leads to its inhibition, while PP2CA-mediated dephosphorylation restores the ability of BRM to repress the ABA response (**Figure 4**). Moreover, the phosphomimetic BRM mutant shows hypersensitivity to ABA (Peirats-Llobet et al., 2016).

BRM has also been shown to play a direct role in the positive regulation of GA biosynthesis by binding to chromatin near the GA3ox1 promoter, thereby activating the gene (**Figure 4**). Arabidopsis brm null mutants exhibit a significant decrease in active GA levels (Archacki et al., 2013). Furthermore, transcriptional profiling revealed that most genes involved in the GA and auxin signaling pathways are affected in both syd and brm null mutants (Bezhani et al., 2007). Previous studies showed that plants over-expressing CHR12 had enhanced growth arrest when exposed to drought or heat stress (Mlynarova et al., 2007). In addition, SYD was linked to ethylene (ET)- and jasmonic acid (JA)-dependent gene regulation (Walley et al., 2008), and the SWI3B subunit was found to interact with a negative regulator of ABA signaling, the PP2C-type phosphatase HAB1 (Saez et al., 2008).

During development, plants transition from the embryonic stage to the seedling stage. Previous studies have characterized the joint effects of PKL and GA in repressing embryonic traits during plant development transitions (Zhang et al., 2008; Zhang et al., 2012). Specifically, PKL expression represses the embryonic state by controlling a significant number of GA-responsive genes. pkl seedlings exhibit a semi-dwarf phenotype similar to that of GA-response mutants and are able to express embryonic traits under GA-biosynthesis inhibition. In contrast, treatment of pkl mutants with GA greatly decreases their characteristic pickleroot phenotype (Henderson et al., 2004). Both PKL and BRM

CRCs have been proposed to act as positive regulators of the GA pathway possibly through distinct mechanisms. Unlike brm mutants, pkl mutant plants have an increased abundance of active GA (Archacki et al., 2013). Recently, PKL was found to positively regulate most GA-mediated developmental processes including promoting vegetative growth (hypocotyl, leaf, and inflorescence stem elongation) and phase transitions (i.e., the juvenile-toadult and vegetative-to-reproductive transitions) (Park et al., 2017).

Excess cytokinin (CK) production inhibits primary root elongation and weakens lateral root development (Kuderova et al., 2008). SWP73B is one of the accessory subunits of Arabidopsis SWI/SNF complexes shown to positively regulate root development and cell cycle progression in the root meristem by suppressing CK biosynthesis (**Figure 4**). SWP73B negatively regulates the CK biosynthesis genes ADENOSINE PHOSPHATE-ISOPENTENYLTRANSFERASE (IPT3) and (IPT7) and the CKregulated cell cycle inhibitor KIP-RELATED PROTEIN7 (KRP7) by hindering the deposition of active histone marks on their gene bodies (Jegu et al., 2015).

In aggregate, these findings indicate that both phytohormone biosynthesis and degradation in response to extracellular and intracellular cues partly depend on SWI/SNF chromatin remodeling ATPases. The findings also suggest the role of SWI/SNF ATPases as a hub in plant perception and response to phytohormone signaling as well as hormone crosstalk through direct physical interactions with hormone signaling pathway components. The outcomes of these interactions influence the plant response to abiotic stress, enabling plant adaptability under changing climates.

#### CONCLUDING REMARKS AND FUTURE PERSPECTIVES

Chromatin remodeling has been extensively studied in the context of various regulatory and developmental processes in a number of eukaryotic organisms, including humans, mice, yeast, fruit fly and Arabidopsis. Phylogenetic analysis suggests that the chromatin remodeling machinery and their modes of operation are evolutionarily conserved across eukaryotes. However, SWI/SNF subunit loss-of-function studies have uncovered varied organism-specific phenotypes. These phenotypes also depend on the strength of a given mutation, leading to phenotypic differences among single and double mutants of SWI/SNF subunit genes. The functional redundancy of some SWI/SNF complex subunits also necessitates further in-depth analyses of their interaction patterns in different plant development pathways.

As discussed above, a number of plant regulatory and developmental transitions are controlled by chromatin modifiers through complex pathways in response to both endogenous and environmental factors. These precisely orchestrated mechanisms are heritable, underscoring their importance for plant survival in changing habitats. Moreover, the dynamics of these epigenetic controls go hand in hand with changes in plant growth conditions. Positive attributes that could potentially be integrated into plant breeding schemes are of interest to many plant scientists and breeders insofar as they may yield varieties that can withstand adverse weather conditions, early flowering or various biotic stresses and maximize nutrient utilization for improved yield. High-throughput technologies together with the vast genomic data presently available in public repositories should facilitate comparative and functional studies aimed at plant improvement. Future studies in other species of interest (e.g., rice and other crops) will likely build upon the findings on ATP-dependent CRCs in Arabidopsis and more recently in maize. Plant-specific molecular and genetic approaches should also characterize regulatory differences between developmental stages and under different growth conditions for a more comprehensive understanding of the involvement of SWI/SNF chromatin modifiers in plant growth and development.

# GENERAL OUTLOOK

The following research questions and topics are of particular interest in terms of how CRCs may be involved in various processes either directly or indirectly related to plant development.

# AUTHOR CONTRIBUTIONS

SO and SC conceived the idea, wrote the manuscript, and drafted the figures. SP designed and organized figures and revised the

manuscript. WL, MY, MA, and HZ revised the manuscript and organized the figures. YQ conceived the idea and revised and organized the manuscript.

#### FUNDING

This work was supported by NSFC<PHONE_NUMBER>4, U1605212, and 31470284) and a Newton Advanced Fellowship to YQ.

#### REFERENCES

#### ACKNOWLEDGMENTS

We extend our apologies to all our colleagues whose work could not be reviewed here owing to space limitations. We are indebted to Fujian Agriculture and Forestry University for its financial support and to members of the Qin Lab (Haixia Institute of Science and Technology, Centre for Genomics and Biotechnology) for their motivation and suggestions for this manuscript.

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2018 Ojolo, Cao, Priyadarshani, Li, Yan, Aslam, Zhao and Qin. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Cork Oak Young and Traumatic Periderms Show PCD Typical Chromatin Patterns but Different Chromatin-Modifying Genes Expression

#### Vera Inácio1 \*, Madalena T. Martins1 , José Graça2 and Leonor Morais-Cecílio1

1 Linking Landscape, Environment, Agriculture and Food (LEAF), Institute of Agronomy, University of Lisbon, Lisbon, Portugal, 2 Forest Research Center (CEF), Institute of Agronomy, University of Lisbon, Lisbon, Portugal

Plants are subjected to adverse conditions being outer protective tissues fundamental to their survival. Tree stems are enveloped by a periderm made of cork cells, resulting from the activity of the meristem phellogen. DNA methylation and histone modifications have important roles in the regulation of plant cell differentiation. However, studies on its involvement in cork differentiation are scarce despite periderm importance. Cork oak periderm development was used as a model to study the formation and differentiation of secondary protective tissues, and their behavior after traumatic wounding (traumatic periderm). Nuclei structural changes, dynamics of DNA methylation, and posttranslational histone modifications were assessed in young and traumatic periderms, after cork harvesting. Lenticular phellogen producing atypical non-suberized cells that disaggregate and form pores was also studied, due to high impact for cork industrial uses. Immunolocalization of active and repressive marks, transcription analysis of the corresponding genes, and correlations between gene expression and cork porosity were investigated. During young periderm development, a reduction in nuclei area along with high levels of DNA methylation occurred throughout epidermis disruption. As cork cells became more differentiated, whole nuclei progressive chromatin condensation with accumulation in the nuclear periphery and increasing DNA methylation was observed. Lenticular cells nuclei were highly fragmented with faint 5-mC labeling. Phellogen nuclei were less methylated than in cork cells, and in lenticular phellogen were even lower. No significant differences were detected in H3K4me3 and H3K18ac signals between cork cells layers, although an increase in H3K4me3 signals was found from the phellogen to cork cells. Distinct gene expression patterns in young and traumatic periderms suggest that cork differentiation might be under specific silencing regulatory pathways. Significant correlations were found between QsMET1, QsMET2, and QsSUVH4 gene expression and cork porosity. This work evidences that DNA methylation and histone modifications play a role in cork differentiation and epidermis induced tension-stress. It also provides the first insights into chromatin dynamics during cork and lenticular cells differentiation pointing to a distinct type of remodeling associated with cell death.

Keywords: CORK, phellogen, lenticels, lenticular phellogen, 5-mC, H3K4Me3, H3K9me2, H3K18Ac

#### Edited by:

Gabino Ríos, Instituto Valenciano de Investigaciones Agrarias, Spain

#### Reviewed by:

Shaojun Xie, Purdue University, United States Rodrigo J. Hasbun, Universidad de Concepción, Chile

> \*Correspondence: Vera Inácio<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science

Received: 04 May 2018 Accepted: 25 July 2018 Published: 27 August 2018

#### Citation:

Inácio V, Martins MT, Graça J and Morais-Cecílio L (2018) Cork Oak Young and Traumatic Periderms Show PCD Typical Chromatin Patterns but Different Chromatin-Modifying Genes Expression. Front. Plant Sci. 9:1194. doi: 10.3389/fpls.2018.01194

#### INTRODUCTION

fpls-09-01194 August 27, 2018 Time: 16:15 # 2

Plants are exposed to adverse environmental conditions like desiccation, freezing, heat injury, mechanical traumas, and disease. Enveloping protective tissues are thus fundamental to plants survival. Epidermis, the protecting tissue in primary tissues, is replaced by the periderm during stems and roots secondary growth (Evert, 2006). An exceptionally thick periderm is found in the cork oak (Quercus suber L.), the commercial cork, which due to several valuable properties, like imperviousness to liquids and insulation, is used for a wide number of important industrial applications (Pereira, 2007). Cork is the result of phellogen (cork cambium) meristematic activity followed by a particular differentiation process, involving cork cells expansion, cell walls suberization and deposition of waxes, ending with cell death and complete emptiness of the cells (Natividade, 1950; Pereira, 2007). In cork oak stems, the phellogen arises in the first year of growth in the subepidermal cell layer (Graça and Pereira, 2004) and continuously produces cork cells throughout the tree's lifespan accumulating a thick periderm very rapidly. Cork is allowed to be firstly harvested when the stem perimeter reaches the legal size (Oliveira and Costa, 2012). The separation of cork is obtained by the physical rupture of phellogen cells, leading to its death. A new traumatic phellogen is formed after cork extraction by a process of meristematic activation within the exposed non-conducting phloem (Fortes et al., 2004).  This process is thereafter cyclically repeated allowing the sustainable exploration of corkoak trees for more than 200 years. The cork produced by traumatic phellogens (amadia cork) has the best characteristics for industrial transformation, as opposed to the first cork divided by the original phellogen. However, even this cork can have widely variable characteristics, presumably due to both environmental and genetic factors, expressed as its industrial "quality." Cork quality is defined by the cork tissue thickness and homogeneity (Silva et al., 2005). The cumulative yearly layers of cork cells are locally crossed at certain points by lenticular channels, named cork pores. These channels are formed by the activity of particular regions of the phellogen, the lenticular phellogen, and are thought to permit gas diffusion between the inward living tissues, and the external environment. Cork porosity, meaning the number, dimension, and distribution of lenticular channels is widely variable in corks from different trees (Graça and Pereira, 2004). Corks with high levels of porosity strongly depreciate its industrial and economic value.

DNA methylation, post translational modifications of histones (HPTMs) and RNA-directed DNA methylation (RdDM) are hallmarks in modifying the functional state of chromatin, and together with nucleosome remodeling can alter the nuclear architecture during plant cell differentiation [reviewed in (Pikaard and Scheid, 2014; Ikeuchi et al., 2015; Takatsuka and Umeda, 2015; Latrasse et al., 2016)]. Plant genomes are methylated in CG, CHG, and CHH contexts which requires the activity of specific DNA methyltransferases (DNMTs) DNA METHYLTRANSFERASE 1 (MET1) maintains CG methylation; CHROMOMETHYLASE 3 (CMT3) maintains non-CG methylation in a self-reinforcing loop between histone H3K9 methylation and DNA methylation by requiring KRYPTONITE (KYP/SUVH4), SUVH5, and SUVH6, a H3K9 methyltransferases; and DOMAINS REARRANGED METHYLTRANSFERASES 1 (DRM1) and 2 (DRM2) responsible for CHH de novo methylation through the RdDM pathway [reviewed in (Pikaard and Scheid, 2014)]. CG methylation can result in gene silencing when found in promoter regions or be correlated with moderately high transcription when present within gene body (Zhang et al., 2006; Lister et al., 2008). Non-CG methylation is associated with the transcriptional silencing of transposable elements (TEs) (Cokus et al., 2008). Histone PTMs are also important components of chromatin-level control of gene activity contributing to define distinct chromatin states that modulate the access of transcription machinery to DNA [reviewed in (Kouzarides, 2007)]. Among these modifications, the dimethylation of histone H3 at lysine 9 (H3K9me2) is a highly conserved repressive mark found in heterochromatic regions with a particular role in the silencing of TEs and other repetitive DNA (Bernatavichute et al., 2008). Different HPTMs may be associated with transcriptionally active chromatin such as the trimethylation of histone H3 at lysine 4 (H3K4me3) which is a hallmark of transcription initiation (Roudier et al., 2011; Sequeira-Mendes et al., 2014) specifically accomplished by ATXR3 (Berr et al., 2010; Guo et al., 2010) and ATX3 (Chen et al., 2017). Another modification related to active chromatin is the acetylation of histone H3 at lysine 18 (H3K18ac) mainly found in regions surrounding the transcription start site and associated with transcription enhancers (Wang et al., 2008).

The regulation of specific plant developmental processes involving DNA methylation and HPTMs is well known [reviewed in (Pikaard and Scheid, 2014; Ikeuchi et al., 2015; Takatsuka and Umeda, 2015; Latrasse et al., 2016)]. Few studies were conducted in cork oak such as in pollen nuclei (Ribeiro et al., 2009), revealing an unexpected pattern of marks associated either with gene silencing or activation. Also, DNA methylation levels were correlated with tissue maturity during embryogenesis (Rodríguez-Sanz et al., 2014; Pérez et al., 2015), and with differences in cork cellular characteristics (cork quality traits) which could be directly related to original (Inácio et al., 2017) or traumatic phellogen activity (Ramos et al., 2013; Inácio et al., 2017). Notwithstanding the crucial role played by the cork as a protective tissue and its highly valued product, studies on the role of these modifications in cork formation and differentiation are still scarce.

In this work, we used cork oak periderm as a model to seek for the first insights into the formation and differentiation of secondary protective tissues at the chromatin level. We studied the chromatin organization and nuclei structural changes during cork cells differentiation together with the dynamics of DNA methylation and HPTMs in young and traumatic periderms, formed after cork extraction wounding. The relative expression of the corresponding chromatin-modifying genes was compared through qRT-PCR in both periderms. In addition, relationships between gene expression and the most relevant cork quality traits were investigated.

#### MATERIALS AND METHODS

fpls-09-01194 August 27, 2018 Time: 16:15 # 3

#### Sampling

Young periderms were collected from sprigs harvested in randomly chosen cork oak adult trees located in Tapada da Ajuda field, Lisbon: herbaceous 'just burst' sprigs (hereafter referred as herbaceous); around one-year-old sprigs slightly lignified (hereafter referred as one-year old); and three-year-old sprigs heavily lignified (hereafter referred as three-year old). This was performed either by harvesting whole sprigs or by peeling off the periderm tissues.

Traumatic periderms in the form of cork planks were extracted from selected cork-oak trees during the harvesting season. The planks were harvested at breast height (at 1.30 m height from soil) from two adult trees located at Tapada da Ajuda field, Lisbon and seven adult trees located at a cork oak stand (montado) in Herdade dos Leitões, Montargil, Portugal. Trees were selected based on their cork quality parameters previously characterized (Inácio et al., 2017). All tissues were harvested during the period of more intense phellogen activity between July and September 2016. Samples of phellogen with contiguous differentiating tissue were collected by scraping the inner surface of cork planks and stored in liquid nitrogen until further use. Small pieces were also cut from the inner surface of the same cork planks and fixed for histologic and cytogenomic analysis. Neighboring cork planks from the same trees were collected for quality traits assessment [described elsewhere (Inácio et al., 2017)].

#### Fixation and Sectioning

Immediately after collection, detached periderms were fixed in 4% paraformaldehyde in 1× PBS (phosphate buffered saline: 137 mM NaCl; 0.27 mM KCl; 1 mM phosphate buffer, pH 7.4) under vacuum followed by overnight incubation in fresh fixative at 4◦C. Periderms were then dehydrated with a graded ethanol series (50, 70, 85, 95, and 100%), cleared with histoclear (VWR Chemicals), and embedded in paraffin (VWR Chemicals). Tissue sections of 7 µm were made using a microtome and mounted in slides previously coated with poly-L-lysine (1 mg/mL, Sigma-Aldrich, Spain).

To preserve the 3D structure of the nuclei, thick sections of whole sprigs with one and three years old were made using the technique described by Conde et al. (2012).

Traumatic periderm and herbaceous sprigs were fixed in FAA (formaldehyde 37%, acetic acid, ethanol 50%, 1:1:18) under vacuum followed by overnight incubation in fresh fixative at 4 ◦C, and dehydrated with a graded ethanol series (70, 96, and 100%) before embedding in glycol methacrylate – GMA (resinbased product – Technovit R 7100) following the manufacturer's instructions. Semi-thin sections (2 µm thick) were made using glass knifes on a piramitome LKB Bromma 11800.

#### Anatomy and Histology Studies

To identify the structures present in the herbaceous and oneyear-old sprigs, these were stained with toluidine blue O (1%), and observed on a Leitz Biomed microscope (Leica Microsystems, Germany) under bright field and photographed with an AxioVision color camera (Carl Zeiss, Germany).

The traumatic periderms were stained with a combination of berberine 0.1% and crystal violet 0.5% to identify the cell walls composition. Preparations were observed in an Axio Imager.Z1 epifluorescence microscope (Carl Zeiss, Germany), and images were acquired with an AxioVision HRm camera (Carl Zeiss, Germany) using the Zeiss filter set 49 (445/50 nm).

The detection of autofluorescence in herbaceous, one-year-old sprigs and traumatic periderms observed with an Axio Imager.Z1 epifluorescence microscope (Carl Zeiss, Germany) using the Zeiss filter sets 49 (445/50 nm), 44 (530/50 nm), and 14 (590 nm) allowed the identification of tissues in the immunodetection experiments.

#### Immunolocalization of 5-Methylcytosine and Posttranslational Histone Modifications

Immunodetections of 5-methylcytosine and HPTMs in one and three-year-old sprigs were performed according to (Ribeiro et al., 2009) with some modifications. The antibodies used were chosen since they have been extensively tested in several studies in both plants and animals (Bianco-Miotto et al., 2010; Carvalho et al., 2010; She and Baroux, 2015; Groth et al., 2016) and their specificity has been verified (Nettersheim et al., 2013). Briefly, cell walls were partially digested with 2% cellulose (Sigma-Aldrich, Spain) in 1x PBS and hydrolysable tannins digested with 2 U/ml of tannase (Thermo Fisher Scientific, Waltham, MA, United States). The antigens were further retrieved using the microwave technique at full power for 8 min (Nic-Can et al., 2013). After cooling down, the sections were incubated with 5% BSA (bovine serum albumin, Sigma-Aldrich, Spain) to block non-specific binding, before the incubation with primary antibodies: anti-5mC (1:100 dilution, Abcam AB10805, Cambridge, United Kingdom), anti-H3K9me2 (1:5 dilution, Abcam AB1220, Cambridge, United Kingdom), anti-H3K4me3 (1:50 dilution, Abcam AB8580, Cambridge, United Kingdom), and anti-H3K18ac (1:100 dilution, Abcam AB1191, Cambridge, United Kingdom). Goat polyclonal secondary antibody to mouse or rabbit IgG – H&L conjugated to Alexa Fluor R 488 (Abcam AB150113, AB150077, Cambridge, United Kingdom) were added in a 1:100 dilution accordingly. The slides were mounted in VECTASHIELD Mounting Medium with DAPI (Vector Laboratories, United Kingdom). Thick sections were examined on a Leica SP5 confocal coupled to a Leica DMI6000 (Leica, Germany) using a 63× 1.4 NA Oil immersion objective and HyD detectors in Standard Mode. Laser lines 405 and 488 nm where then used to excite DAPI and Alexa Fluor R 488 fluorochromes with spectral detection adjusted for each. Z-stacks with 0.5 µm were acquired to allow for the identification of entire nuclei.

Immunolocalization of 5-methylcytosine (5-mC) on semithin sections of herbaceous sprigs and traumatic periderms was carried out also according to (Ribeiro et al., 2009) but with a prior permeabilization with 1× PBS containing 0.5% Triton X-100. Sections were incubated with mouse monoclonal to 5-mC

(Abcam AB10805, Cambridge, United Kingdom), followed by incubation with anti-mouse CyTM3-labeled secondary antibody (Sigma-Aldrich C2181, Spain). The slides were mounted in VECTASHIELD Mounting Medium with DAPI (Vector Laboratories, United Kingdom) and examined with Zeiss Axio Imager.Z1 epifluorescence microscope (Carl Zeiss, Germany) with a 63× 1.25 NA Oil immersion objective. Images were acquired with AxioVision HRm camera (Carl Zeiss, Germany) using Zeiss filter sets 49 (445/50 nm) and 14 (590 nm).

#### Fluorescence Intensity Quantification

To achieve precise and reliable comparisons between signals observed at distinct cell differentiation stages, confocal analysis was performed using the same laser excitation and sample emission capture settings (El-Tantawy et al., 2014). The same procedure was applied to epifluorescence analysis on herbaceous sprigs and traumatic periderms preparations. Measurements of the fluorescent signal intensity and nuclei area were performed on the different tissues and/or cork cell layers using Fiji (Schindelin et al., 2019). Projections of maximum fluorescence images from confocal Z-series were obtained and used to quantify fluorescence (Conde et al., 2012). The contour of each nucleus was manually outlined, and the fluorescence intensity was measured (sum of the fluorescence on each pixel within the outlined area). Fluorescence intensity in a non-labeled region was used to normalize all quantifications. The nucleus area was used to normalize the fluorescence intensity to avoid an artificial positive correlation between fluorescence intensity and nuclei size. The ratio fluorescence intensity/nucleus area reflects the amount of DNA methylation or histone modifications. Semithin sections of GMA (2 µm) and paraffin (7 µm) led to the segmentation of each nucleus, thus, for statistical analysis, these were grouped in classes according to its area (<15[, [15;25[, [>25 µm2 ). Data are presented in standard boxplots as minimum, first quartile (bottom of box), median, third quartile (top of box), and maximum with the actual spread of individual observations represented by jittered dots. Differences in the ratio fluorescent intensity/nucleus area and in the nuclei area between tissues and/or cork cell layers were tested through Student's t-test and one-way ANOVA followed by Tukey's multiple comparison test, at a 5% significance level. For non-normally distributed data, Kruskal-Wallis non-parametric and Dunn's Multiple Correction post hoc tests were used. All statistical tests were performed using GraphPad Prism V5.0 software (GraphPad©, San Diego, CA, United States).

#### RNA Extraction

Total RNA was extracted from detached periderms and traumatic periderms with the SpectrumTM Plant Total RNA kit (Sigma-Aldrich, Spain) according to manufacturer's instructions except for some minor modifications: the isolation buffer was supplemented with one volume of Plant RNA Isolation Aid (Thermo Fisher Scientific, Waltham, MA, United States) per unit mass of fresh tissue. Total RNA integrity was assessed by 1% (w/v) agarose gel electrophoresis. mRNA was isolated from total RNA using the Dynabeads mRNA Purification Kit (Thermo Fisher Scientific, Waltham, MA, United States) following the manufacturer's directions. cDNA was synthesized from 45 ng of mRNA using oligo(dT)18 in a 20 µL-reaction volume using RevertAid H Minus Reverse transcriptase (Thermo Scientific, Waltham, MA, United States) according to the manufacturer's protocol. cDNA was stored at −20◦C until further use.

#### Putative Quercus suber Histone Methyltransferases Characterization

The cork oak QsDNMTs (QsCMT3, QsDRM2, QsMET1, and QsMET2), and QsSWC4 evaluated by qRT-PCR in this study were previously characterized (Ramos et al., 2011).

The putative cork oak sequences homologous of characterized histone methyltransferases (HMTs – QsSUVH4, QsATXR3, and QsATX3) were obtained by performing a BLAST at cork oak database1 (Pereira-leal et al., 2014), using the Arabidopsis protein sequences as a query. Complete sequences were retrieved from the cork oak genome version 1.0 (Ramos et al., 2018), except for QsATXR3. Protein structure was analyzed using NCBI-CD2 (Marchler-Bauer et al., 2016) and SMART searches3 (Letunic and Bork, 2017). The potential cork oak QsHMTs orthologous proteins in other angiosperms (**Supplementary Table 1**) were obtained by performing a BLAST at the NCBI database4 (Altschul et al., 1997). All sequences were aligned with MUSCLE5 (Edgar, 2004) and the alignment was trimmed with GBLOCKS6 (Dereeper et al., 2008). QsHMTs were used to perform a phylogenetic analysis with orthologous sequences. Phylogeny analysis was obtained with MEGA 7 software (Kumar et al., 2016), using the maximum likelihood method and a bootstrap of 1000.

#### Primer Selection and qRT-PCR Analysis

Seven target genes – QsCMT3, QsDRM2, QsMET1, QsMET2, QsSWC4, QsSUVH4, QsATXR3, and QsATX3 – and four housekeeping genes – ACT (actin), CACs (clathrin adaptor complexes medium subunit family protein), GAPDH (glyceraldehyde 3-phosphate dehydrogenase), EF-1α (elongation factor 1-alfa) – were evaluated in this study. Primers were designed using Primer Premier 5.0 software (PREMIER Biosoft International, Palo Alto, CA, United States) except for QsDRM2 gene and housekeeping genes that was chosen from a previous gene expression studies in cork oak (Marum et al., 2012; Ramos et al., 2013). qRT-PCR experiments were performed in all tissues except for QsSUVH4 which were only performed in three-yearold sprigs and traumatic periderms. Gene description, NCBI nucleotide and protein sequences accession numbers, primer sequences, and amplicon size are described in **Supplementary Table 2**.

The real-time qPCR was performed in 96 well white reaction plates (Bio-Rad, Hercules, CA, United States), using an IQ5 Real Time PCR (Bio-Rad, Hercules, CA, United States) with at least

1http://corkoakdb.org

2https://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi

3http://smart.embl.de

4https://blast.ncbi.nlm.nih.gov/Blast.cgi

5https://www.ebi.ac.uk/Tools/msa/muscle/

6http://phylogeny.lirmm.fr/phylo\_cgi/one\_task.cgi?task\_type=gblocks

six individuals and three technical replicates. All cDNA samples were diluted 20-fold and were amplified in triplicate in two independent PCR runs. The reaction mixture was composed of 1 µL diluted cDNA, 0.5 µM of each gene-specific primer and 5 µL master mix (SsoFast EvaGreen Supermix, Bio-Rad, Hercules, CA, United States). The following program was applied: initial polymerase activation, 95◦C, 3 min; then 40 cycles at 94◦C for 10 s (denaturation), 61◦C (except for ATXR3 which was 55◦C) for 20 s (annealing), 72◦C 15 s (extension), followed by a melting curve analysis to confirm the correct amplification of target gene fragments and the lack of primer dimmers. No template controls were also included in triplicate for each primer pair. Amplification efficiencies of all genes were estimated with the LinRegPCR quantitative PCR data analysis program (Ruijter et al., 2009) using the raw fluorescence data as input. According to NormFinder algorithm (Andersen et al., 2004) the GAPDH and ACT genes were chosen as the most stable ones to be used as references. The target genes relative expression ratio was calculated based on amplification efficiencies and expressed in comparison to the geometric mean of reference genes according to (Pfaffl, 2001).

Statistical analysis was performed by clustering data from each group of tissues (young periderms – one and three-years-old – and traumatic periderms) and evaluating statistical differences between them as well as between genes, through Student's t-tests and one-way ANOVA followed by Tukey's multiple comparison tests, at a 5% significance level.

Also, relationships between DNMTs, HMTs, and QsSCW4 relative gene expression and cork quality traits previously assessed (Inácio et al., 2017) were analyzed by Pearson's correlations using 'rcorr' function from R environment. The studied traits were porosity coefficient, and pores area, length, and roundness (**Supplementary Table 3**), described in detail elsewhere (Inácio et al., 2017).

#### RESULTS

#### Cork Cells Keep Their Nuclei Up to the Last Phases of the Differentiating Process

In the herbaceous sprigs immediately after burst, only primary growth was detected. From the outside to the inside, epidermis with pluricellular trichomes, cortical parenchyma, primary phloem and xylem, and pith were identified. Periclinal divisions were detected in outermost cortex cell layer establishing the precursor of the phellogen (**Figure 1A**). Different tissues from the outermost to the innermost region of the one-year-old sprigs were clearly recognized: residues of the epidermis with trichomes, few cork cell layers, phellogen, phelloderm, cortex (**Figure 1B**), secondary and primary phloem, secondary and primary xylem, and pith. In the detached periderms the protection tissue is well individualized but the phellogen cell layer was not present since it was torn during the removal. In three-year-old sprigs, due to the increase in thickness of the stem, resulting from the underlying phellogen activity, the epidermis was no longer detectable, and increasing cork cell layers were detected (**Figure 1C**). Cork cell layers exhibited intense autofluorescence when excited with ultraviolet light, indicating the deposition of suberin in their cell walls, which favored the identification of the phellogen, a cell layer without fluorescence right below cork in whole sprigs.

The traumatic periderms, cork planks identical to commercial cork, had the thickness resulting from the nine years growth cycle. In the inner surface of cork planks, the tear zone, i.e., the region where the cork planks were detached from the tree, the cells were disrupted with several contiguous layers of differentiating cork cells already showing suberized walls from the early stages of the differentiation process (**Figure 1D**). Cork living cells showing entire nuclei (DAPI positive) with cytoplasmic content were observed in three-year-old sprigs and traumatic periderms several layers beyond the phellogen and the tear zone (**Figure 1E**). It was impossible to count the total number of living cork cell layers in traumatic periderms, since these were heavily corrugated. The cork cell layers will be referred hereafter as c1, c2, c3, c4, and c5 from phellogen to the outermost layers according to its age, being c1 the cork cell layer most recently formed and c5 the latter differentiation stages.

#### Chromatin Condenses, and Nuclei Area Decreases as Differentiation Proceeds in Cork Cells

Alterations in nuclei structure were noticed in differentiating cork cells in both young periderms (one and three-year-old sprigs) and traumatic periderms. Decreases in nuclei area from the phellogen to the cork layers at later stages of differentiation were detected ranging from 2.1-fold in one-year-old sprigs (p < 0.001, Tukey's multiple comparison test, **Figure 2A**) to 4.2-fold in traumatic periderm samples (p < 0.001, Tukey's multiple comparison test, **Figure 2B**). These changes were corroborated by the higher nuclei number with less than 15 µm2 found in cork cells at later differentiating stages, particularly in traumatic periderm where no nuclei higher than 15 µm2 were found (**Supplementary Figure 1**). The reduction in area was accompanied by drastic changes at the chromatin level. Chromatin progressively condensed as cork cells became more differentiated, and the condensed chromatin preferentially localized at the nuclear periphery.

Nuclei area in epidermis also suffered changes in morphology by decreasing its area from entire to disrupted stage (from herbaceous to one-year-old sprigs) (**Figure 2C**, p < 0.001, Tukey's multiple comparison test), as denoted by the absence of nuclei with areas higher than 25 µm2 in the latter (**Figure 2D**).

#### Nuclear Fragmentation Is Present in Lenticels of Older Periderms

In younger periderms from one-year-old sprigs, small areas with intense meristematic activity denoted the lenticular phellogen. This meristem produces huge number of cells known as filling tissue forming the lenticels. The high number of cell layers produced by the lenticular phellogen propels the epidermis upwards causing its fracture (**Figure 3A**). In three-year-old

FIGURE 1 | Anatomical analysis of cross-sections of young sprigs (herbaceous, and one and three-year-old sprigs) and traumatic periderms. (A) Toluidine Blue O staining of the herbaceous sprigs showing the epidermis with pluricellular trichomes, cortical parenchyma, and the first periclinal divisions originating the precursor of the phellogen. (B) Autofluorescence detection under UV light of cork cells and residues of the epidermis with trichomes in cross-sections of one-year-old sprigs. Right below the cork cells is the phellogen, and the underlying tissues phelloderm and cortical parenchyma. (C) Autofluorescence detection under UV light of several cork cell layers resulting from the phellogen activity in cross-sections of three-year-old sprigs. (D) Berberine/crystal violet staining of traumatic periderms observed under UV light. The cells from the tear zone presented disrupted cellulosic reddish walls (arrows). The contiguous layer of cork cells already shows suberized walls at early stages of differentiation (arrowhead), and highly suberized walls at later stages of differentiation (intense green). (E) Cells at later stages of cork differentiation showing entire nuclei. DNA was counterstained with DAPI. Bar = 10 µm. Ep, epidermis; Tr, pluricellular trichomes; Co, cortex; Ph, phellogen; C, cork cells; Phd, Phelloderm; Tz, tear zone; N, nucleus.

sprigs, the lenticels were larger due to continuous lenticular phellogen activity during sprig development (**Figure 3B**). The wall of the cells produced by lenticular phellogen emitted much less autofluorescence than cork cells, indicative of a low suberin content, and a distinct cell wall composition. Contrastingly to cork cells, nuclei from older lenticels (from three-year-old sprigs) became misshapen (**Figure 3C**), and highly fragmented as cells differentiate, with portions of chromatin protruding from a central less condensed chromatin mass to the outermost regions of the lenticels (**Figure 3D**).

### DNA Methylation and Posttranslational Histone Modifications Are Highly Dynamic During Cork Cells Differentiation

To assess the nuclear distribution of well-known markers of different chromatin functional states, immunolocalization of 5-mC and H3K9m2 (repressive marks), and H3K4me3 and H3K19ac (active marks) were performed. These studies revealed differences in the intensity and distribution patterns at distinct tissues and differentiation stages.

In very young herbaceous sprigs the nuclei showed discrete 5 mC signals distributed in small spots all over the chromatin both in cortex and epidermis (**Figure 4A**) with similar intensity level (p > 0.05, Unpaired t-test with Welch's correction, **Figure 4B**). In one-year-old sprigs, 5-mC signals were dispersed throughout the nucleus in all cork cell layers (**Figure 4C**) and significant higher intensity levels were detected in all cork cells when compared with phellogen, except for c2 (p < 0.01 for Ph vs. c1, p < 0.05 for Ph vs. c2, Dunn's multiple comparison test, **Figure 4D**). In fact, a 2.5-fold increase from phellogen to the more differentiated cork cell layer (c3) was noticed. Simultaneously, high levels of DNA methylation were found in epidermal nuclei in cells that remained alive (**Figure 4C**, arrow). In three-year-old sprigs,

every individual measurement is represented by jittered dots.

5-mC signals were dispersed throughout the nucleus in cork cells, however, the highest intensity was observed in nuclear periphery (**Figure 4E**). There was a significant increment in 5 mC intensity from c2 to c5 at the latter stages of differentiation (p < 0.05, Dunn's multiple comparison test, **Figure 4F**). In traumatic periderm, the tear zone, which locates at the immediate vicinity of the phellogen, showed faint 5-mC fluorescence signals distributed as very small spots all over the chromatin that became more intense at nuclear periphery in the more differentiated cork cells (**Figure 4G**). In nuclei with areas less than 15 µm2 , a remarkable 6.4-fold increase in the 5-mC level from the tear zone to cork cells at later differentiation stages (c4) was detected (p < 0.01, Dunn's multiple comparison test, **Figure 4H**).

The immunodetection of 5-mC in lenticels exposed differences in fluorescence signal between lenticel filling cells compared to the underlying lenticular phellogen, and neighboring phellogen and cork cells (**Figure 5A**). Although 5-mC signals were distributed throughout the nuclei within lenticels (**Figure 5B**), the quantification of changes in 5 mC levels along lenticular cell layers was unfeasible since nuclei were highly fragmented. Nevertheless, the less intensity in 5-mC signals in lenticular filling tissue compared with the surrounding tissues was quite evident (**Figure 5A**). Furthermore, lenticular phellogen exhibited significantly lower levels of 5-mC than the contiguous cork-forming phellogen (p < 0.0001, Unpaired t-test with Welch's correction, **Figure 5C**).

portions of chromatin protruding from a central less condensed chromatin mass. DNA was counterstained with DAPI. Bars in (A,B) = 20 µm and in (C–E) = 10 µm.

Regarding the dimethylation of histone H3 at lysine 9 (H3K9me2) in one-year-old sprigs, two condensed chromatin knobs in the periphery of the nucleolus could be observed (**Figure 6**). The H3K9me2 signals were not quantified due to the reduced number of labeled nuclei.

Ep, epidermis; Lph, lenticular phellogen; C, cork cells; Lc, lenticular cells.

The trimethylation of histone H3 at lysine 4 (H3K4me3) signals were distributed throughout the nuclei both in phellogen and differentiating cork cell layers in one and three-year-old sprigs (**Figures 7A,B**). In one-year-old sprigs, a 2.4 and 2.7-fold enrichment in the level of H3K4me3 was found from the phellogen to the two contiguous cork cell layers, respectively (p < 0.001, Dunn's multiple comparison test, **Figure 7C**), while in three-year-old an average of 2.6-fold increase was noticed from phellogen to each of the differentiating cork cell layers (p < 0.001, Dunn's multiple comparison test), with no differences between them (**Figure 7D**).

The nuclear distribution pattern of acetylation of histone H3 at lysine 18 (H3K18ac) revealed nuclei equally and thoroughly labeled in all cork cell layers in one-year-old sprigs (**Figure 8A**) and in all cork cell layers and in phellogen in three-year old sprigs (**Figure 8B**), respectively. No significant differences were detected in the intensity levels (p > 0.05 for both sprigs, Tukey's multiple comparison test, **Figures 8C,D**).

# Quercus suber HMTs Proteins Showed All Expected Domains

The QsDNMTs and QsSWC4 studied in this work were previously characterized (Ramos et al., 2013). The three QsHMTs analyzed revealed that QsSUVH4 encodes a putative complete protein comprising all four domains found in SUVH4 proteins: SRA-YDG, Pre-SET, SET, and post-SET from N to C terminal; QsATXR3 encodes a putative partial protein lacking the N-terminal, but with the SET domain of ATXR3 proteins detected at the C terminal; QsATX3 encodes a putative complete protein, containing all domains described for ATX3 proteins: PWWP, plant homeodomains (PHD) finger, SET, and post-SET from N to C terminal (**Supplementary Figure 2**). All domains seemed conserved in the angiosperms used in this study (see **Supplementary Table 1** for list of angiosperms). Each QsHMT grouped with their orthologous sequences establishing three well individualized groups (**Supplementary Figure 3**), revealing a high degree of conservation.

# HMTs Differential Gene Expression Was Found Between Young and Traumatic Periderms

The relative expression of several QsDNMTs, QsHMTs, and QsSWC4 was compared in young and traumatic periderms through qRT-PCR.

A high accumulation of transcripts was found for QsDRM2, QsMET1, QsATXR3, and QsATX3 genes in both young and traumatic periderms. Although variability was found between individuals, QsDNMTs, QsHMTs, and QsSWC4 showed a tendency to be expressed at lower levels in traumatic than in young periderms, except for QsSUVH4 and QsATXR3 (**Figure 9**). QsSUVH4 was amongst the most expressed genes in traumatic periderms, but significantly down-regulated in the young ones (p = 0.03, Mann Whitney test). In the latter, QsSUVH4 was significantly down-regulated when compared with QsCMT3 (p = 0.03, Unpaired t-test). The expression of QsMET1 was positively and significantly correlated with porosity coefficient (r≈0.95, p = 0.003, 'rcorr' R function), while QsMET2 and QsSUVH4 expressions showed negative and significant correlations with porosity-related traits (pore length and roundness), one of the most relevant defects found in cork of (r≈0.78, p = 0.04, 'rcorr' R function; **Supplementary Table 4**).

differences. DNA was counterstained with DAPI. Bars in (A,C,E) = 10 µm and in (G) = 2 µm. Ph, phellogen; Ep, epidermis; Co, cortex; Tz, tear zone.

#### DISCUSSION

In this work, we used for the first time cork oak periderm as a model to study the formation and differentiation of secondary plant protective tissues and their behavior after traumatic wounding. Periderms are composed of three different types of tissues: the phellogen, a single cell layer secondary meristem with high cell cycling activity; the cork, a multicell layer tissue interrupted locally by lenticular channels; and the phelloderm, a single cell layer composed of living parenchymatous cells (Evert, 2006). An integrated approach that combines the immunodetection of several chromatinmodifying marks, and the expression of genes required for the imposition of these modifications enabled a detailed view of the dynamics of active and repressive chromatin marks in nuclei of periderm cells. Also, the establishment of correlations between gene expression and one of the most relevant cork quality traits was achieved. Although periderm formation and development has been extensively studied at the chemical and molecular level [(Miguel et al., 2015; Wunderling et al., 2018) and reviewed in (Franke et al., 2012; Graça et al., 2015;

with DAPI. Bars = 10 µm. Lph, lenticular phellogen; Ph, phellogen; Lc, lenticular cells; C, cork cells; Co, cortex.

Vishwanath et al., 2015)], our results at the chromatin level brings novelty and adds significant value to the comprehension of its ontogeny.

### Drastic Changes in Chromatin Structure Occurs During Cork Cells Differentiation

The periderm formation in cork oak involves several processes, including cork cells expansion, cell wall suberization (Graça and Pereira, 2004; Pereira, 2007), and likely programmed cell death (PCD) as a terminal differentiation step, resulting in several layers of dead cork cells with empty lumens, with a key function in insulation and protection. During cork cells differentiation, a repressive nuclear compartment was defined by a high level of DNA methylation at the nuclear periphery. Methylation at the nuclear periphery is usually associated with transposable elements (TEs) and repressed genes location (Bi et al., 2017). This suggests a striking reallocation of chromatin within these nuclei in oak meristematic cells since TEs are known to have an interspersed arrangement in potential gene-rich regions, accompanied by a dispersed 5-mC pattern all over the nuclei

(Alves et al., 2012). Moreover, different types of nuclei can present different chromatin organizations, as detected by several epigenetic marks patterns in cork oak pollen nuclei (Ribeiro et al., 2009), corroborating a strong chromatin remodeling associated with development and differentiation processes in cork oak. Similar to cork cells differentiation are the changes in nuclear morphology and chromatin organization found amongst the most typical structural features in cells undergoing PCD [reviewed in (Van Hautegem et al., 2014; Latrasse et al., 2016)]. These alterations accompanied by increased levels and a change in the distribution of DNA methylation in differentiating cork cells has also been observed in tapetum cells PCD correlating with an up-regulation of MET1 (Solís et al., 2014), responsible for CpG methylation maintenance in cycling cells (Huang et al., 2010). Indeed, QsMET1 was amongst the genes with higher levels of relative expression in the cork tissue contradicting, however, previous results in corks with different qualities (Ramos et al., 2013) where was argued that QsMET2 might be substituting QsMET1 function to maintain the CpG methylation during phellogen activity. In our work, although at lower levels than QsMET1 this gene was amongst the genes with the highest expression. DNMT2 or MET2 are known to have weak or no DNA methyltransferase activity although still present in all eukaryotes (Ponger and Li, 2005). For these reasons it has been suggested that, at least in some species, it may have alternative roles (Vieira et al., 2017). In addition, it has been shown that AtDNMT2 interacts with AtHD2s, a unique plant-specific type of histone deacetylase family (Song et al., 2010). Moreover, increase in HD2 expression has been detected during fruit senescence (Kuang et al., 2011), indicating a possible role of DNMT2 in plant cell death programs through the association with histone deacetylases. Therefore, the preferential expression in tissues undergoing PCD, and the other functions might contribute to its high expression. Another important process in meristematic derivative cells is the de novo methylation mainly accomplished by DRM1 and DMR2 through the RdDM pathway [reviewed in (Pikaard and Scheid, 2014)]. In differentiating cork cells where DNA methylation is increasing it was not surprising to find the highest levels of gene expression for QsDRM2, according to previous results (Ramos et al., 2013). Different methyltransferases act together with chromatin remodeling complexes in an intricate interplay to modify chromatin structure and regulate transcription. CMT3 is required for CHG methylation maintenance, preferring hemimethylated CHG sites (Du et al., 2012), having an active role in TEs silencing (Tompa et al., 2002; Kato et al., 2003; Lippman et al., 2003). Indeed, in traumatic periderms, which are composed of phellogen and contiguous differentiating cork cells, around 20% of methylated CCG loci were found (Inácio et al., 2017), although a global DNA methylation view is compromised since the methylation in all other contexts could not be assessed. Considering the emergence of the repressive chromatin domain in these nuclei, and CMT3 function in silencing TEs, its expression in a tissue with intense meristematic activity and in its derivatives, is not surprising.

Epigenetic modifications are read by several protein complexes. DMAP1 is the human homologous of the Arabidopsis SWC4, which is a subunit of the chromatin remodeling complex SWR1C, and also a component of the NuA4 histone acetyltransferase complex, both involved in transcriptional regulation (Bieluszewski et al., 2015) by interacting with the promoters of target genes (Mozgová et al., 2015). SWC4 which has nucleosome acetyltransferase H4 activity (Doyon et al., 2004; Bieluszewski et al., 2015), is involved in several cellular processes such as, the regulation of mitotic cell cycle progression (Shin et al., 2010), DNA repair (Lee et al., 2010), and has been considered essential for plant development (Mozgová et al., 2015). The human DMAP1 is also a co-regulator that stimulates global maintenance of DNA methylation by co-working with MET1 (Rountree et al., 2000) at sites of double strand break repair (Negishi et al., 2009; Shin et al., 2010). The diverse functions of the DMAP1/SWC4 protein, as well as its involvement in two chromatin remodeling complexes associated with gene expression, should account for the expression detected in cork tissues during its differentiation, as already seen in traumatic corks (Ramos et al., 2013).

Like in cork cells, epidermis nuclei decrease in area whereas DNA methylation level increases from herbaceous to one-year-old sprigs, until this cell layer gradually disappear. During the enlargement of stems and roots due to secondary growth, the one-layered epidermis is replaced by the periderm, which thereafter assures the protective role (Evert, 2006). This replacement is caused by internal mechanical pressure driving to epidermis rupture. Thus, epidermis of woody species is a shortlived tissue that undergoes PCD likely as a response to mechanical stress signals, as the changes in chromatin structure observed are