- Email: [email protected]
25 Years of Cell Cycle Research: What's Ahead?
Opinion
25 Years of Cell Cycle Research: What's Ahead? Crisanto Gutierrez1,* We have reached 25 years since the first molecular approaches to plant cell cycle. Fortunately, we have witnessed an enormous advance in this field that has benefited from using complementary approaches including molecular, cellular, genetic and genomic resources. These studies have also branched and demonstrated the functional relevance of cell cycle regulators for virtually every aspect of plant life. The question is – where are we heading? I review here the latest developments in the field and briefly elaborate on how new technological advances should contribute to novel approaches that will benefit the plant cell cycle field. Understanding how the cell division cycle is integrated at the organismal level is perhaps one of the major challenges.
Trends Research carried out mainly over the past 25 years has led to a detailed understanding of plant cell cycle at the physiological, cellular, molecular, genetic, and genomic levels. Many aspects of plant life, including formative cell divisions during organogenesis, the exit to differentiation, chromatin and epigenetics, metabolism and stem cell potential, circadian clock, and innate immunity, among others, impinge or depend directly on cell cycle regulators.
A Timeline of Plant Cell Cycle Research Cell cycle research witnessed a turning point in 1987 when a human homolog of the S. cerevisiae p34cdc2, a cyclin-dependent kinase (CDK) crucial for cell cycle progression, was isolated by a functional complementation screen [1]. This study, together with others, was worth a Nobel Prize in 2001, and demonstrated that a key cell cycle regulator was conserved in very distantly related eukaryotes. It also triggered a new interest of plant researchers to investigate if plant cells could use CDK–cyclin complexes as part of the cell cycle progression control. Thus, 25 years ago Peter John and colleagues, including Melanie Lee (who identified the human p34cdc2 protein), were the first to (i) identify a p34 protein in plant extracts (including Chlamydomonas reinhardtii and Arabidopsis thaliana) with high immunological similarities to the yeast and human p34cdc2 protein, and (ii) show that its phosphorylation status changed during the cell cycle [2]. The kinase activity of plant p34cdc2 complexes as well as the cloning of the p34 encoding gene in pea [3] and Arabidopsis [4] were reported soon afterwards. An immediate challenge to fully confirm the presence of such complexes in plants was to identify the cyclin (CYC) moieties. The effort of several laboratories succeeded in the identification and cloning of various cyclins, later identified as A- or B-type cyclins [5–7], highly similar to the yeast proteins. Plant cell cycle research has passed through several distinct stages marked by successive breakthrough discoveries and technological advances (Figure 1). Thus, to be fair, interest in plant cell cycle bloomed much earlier, 50 years ago. The initial efforts were focused on cytological and physiological approaches to mitotic activity and kinetics [8–10], environmental effects [11,12], and S-phase progression [13], to mention a few representative examples. Later, the 1990s were marked with advances that revolutionized cell cycle research because they initiated molecular and cell biology studies with two main series of findings (Figure 1). One, already mentioned above, that allowed the cloning of several CDK and cyclin genes, most likely involved in the G2/M transition. Another, the identification of key cell cycle regulators that were not present in yeast, such as D-type cyclins [14,15] and the retinoblastoma protein, a homolog of the human tumor-suppressor [16–18]. Together with other reports, they provided the framework to begin to understand that the plant cell cycle machinery, with some unique features, was much more like that in animal cells than in yeast. The beginning of the 21st century was marked by the
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Future challenges aim at obtaining an integrative understanding of cell proliferation in the whole plant. Recent advances in next-generation sequencing and single-cell omics, sophisticated imaging, and cell typespecific genome editing are among the novel approaches that should contribute to solve major questions on cell cycle research in model and crop species.
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Centro de Biologia Molecular Severo Ochoa, Consejo Superior de Investigaciones Científicas (CSIC), Universidad Autónoma de Madrid (UAM), Nicolas Cabrera 1, 28049 Madrid, Spain
*Correspondence: [email protected] (C. Gutierrez).
http://dx.doi.org/10.1016/j.tplants.2016.06.007 © 2016 Elsevier Ltd. All rights reserved.
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Figure 1. Timeline of Plant Cell Cycle Research.
explosion of several ‘omic’ approaches (Figure 1), greatly facilitated by the availability of the full genome sequence of Arabidopsis, the first multicellular organism, together with Drosophila, to be sequenced [19]. It is obvious that all the different experimental approaches are still fruitfully combined and will contribute to provide further insights into the plant cell cycle regulatory networks in individual cells. Where are we moving, and what are the challenges ahead? In the following paragraphs I will expand upon these ideas, highlighting some of the various lines that may drive plant cell cycle research in the coming years.
Formative Cell Divisions and Cell Fate Cell proliferation is an integral part of morphogenesis. Formative cell divisions, also known as asymmetric cell divisions (ACDs), are one widespread example of a specific type of cell division that impact on morphogenesis. Typically, ACD produces two daughter cells that will take uniquely different fates, breaking the paradigm that cell division produces two identical cells. They occur in different locations across the plant body to generate new cell types and/or expand organogenesis. It was originally thought that developmental signals would trigger a signaling cascade that ultimately impinges on cell division as a downstream process. Indeed, recent evidence shows that various components of the cell cycle machinery are direct and immediate targets of developmental regulators that control ACD (Figure 2). The case of cyclin CYCD6;1 illustrates such coordination. CYCD6;1 is required for ACD in the root apical meristem and, specifically, triggers division of the cortex-endodermis initial, a process that is inhibited by the RETINOBLASTOMA-RELATED1/SCARECROW/SHORTROOT (RBR1/ SCR/SHR) module [20,21]. CYCD6;1-containing CDKA and CDKB1 complexes phosphorylate RBR1, releasing the inhibition of ACD [22,23]. It is not known whether these complexes act sequentially or in an alternative manner. Although it is known that CYCD6;1 availability is controlled by an E3 ubiquitin ligase [24], learning about how CYCD6;1 expression and stability are controlled in different locations will be enlightening. The daughter cells resulting from the asymmetrical division of stem cells in the root apical meristem undergo several division cycles to constitute the root meristem (also called transit amplifying compartment) before exit to differentiation. The plant-specific GROWTH-REGULATING FACTORS (GRFS) are expressed in proliferating cells in the meristem, but are excluded from stem cells where they are repressed by miR396 [25]. Importantly, GRFs repress PLETHORA (PLT) genes in the stem cell derivatives and are required to prevent periclinal formative divisions.
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Figure 2. Summary of the Physiological Processes that Impinge Directly on Cell Cycle Regulators.
Further understanding on how this module regulates downstream targets and whether they are cell cycle regulators is of primary relevance (see also below). Crucial to the establishment of an asymmetry in the daughter cells is not only the signaling cascade channeled through cell cycle regulators but also the control of the cell division plane. Choosing between symmetrical/asymmetrical and/or periclinal/anticlinal divisions is not trivial because it involves a dramatic change in the internal organization of the cytoskeleton relative to that of the mother cell. Aurora (AUR) kinases are likely to be key players in such decisions, in particular ACDs, because they are involved in the orientation of the cell division plane in various stages of plant development [26,27]. Strikingly, AUR proteins have putative RBR1 binding sites [28], which leads us to speculate on a possible link between AUR-controlled ACD and other processes impinging on ACD, although direct evidence is still lacking. This possible link also reveals the complexity of ACD control depending on the phosphorylation status of RBR1 during the cell cycle, the unknown set of AUR targets, and their availability. The coordination of cell proliferation and cell fate acquisition is a fundamental aspect of patterning. Remarkably, several cell cycle regulators have been identified as direct targets of developmentally regulated master genes (Figure 2), such as (i) FOURLIPS (FLP/MYB88) that controls guard mother cell division in the stomatal lineage by regulating the expression of CYCA2;3, CDKA;1, and CDKB1 [29], and (ii) ACR4 (ARABIDOPSIS CRINKLY4), that is necessary for restricting cell division at early stages of lateral root development [30], and which also controls columella stem cells division [31] and giant cell identity in sepals [32]. Interestingly, acr4 mutants show hyperproliferation and, concomitantly, reduced endoreplication [32]. Whether ACR4 and other master regulators of cell specification directly target other cell cycle genes, as well as on endoreplication genes, to control their activity remains to be studied.
Division and Differentiation Balance Organogenesis relies on two intimately coordinated processes: the production of a sufficient number of cells and progression of selected groups through distinct differentiation pathways to generate the specific cell types of a particular organ (Figure 2). An excess of the division program (or a delayed entry into differentiation) could lead to hyperplasia and dysfunctional organs. Likewise, premature exit from the cell cycle to differentiation (or a reduction in proliferative
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potential) generally leads to hypoplasia through exhaustion of the pool of dividing cells. The postembryonic nature of plant organogenesis, together with its responsiveness not only to internal (developmental cues, hormonal signals) but also to external (environmental challenges) inputs, represents an additional layer of regulation that confers plasticity to plant organogenesis. The root meristem has provided insightful information supporting the notion that some cell cycle regulators are at the crossroads of the balance between cell division and differentiation [33]. There are several examples illustrating that RBR1 is a crucial hub for integrate hormonal signals that not only control organ size but also determine the division potential of various cell types within the root. Thus, activation of the AUXIN RESPONSE FACTOR19 (ARF19), a factor that promotes differentiation of root meristem cells, requires RBR1 and ARABIDOPSIS RESPONSE REGULATOR12 (ARR12), a cytokinin-dependent transcription factor [34]. Another example is the division of quiescent center (QC) cells, which is extremely rare but it occurs in response to various stimuli. This is strictly controlled by the balance between the ETHYLENE RESPONSE FACTOR115 (ERF115)-mediated activation [35] and the RBR1dependent repression [36]. ERF115 expression depends on BRAVO and the brassinosteroid signaling [37] and CELL CYCLE SWITCH52 (CCS52A2), an activator of the anaphasepromoting complex 35, whereas RBR1 and its interaction with SCR depend on the activity of CYCD3;3 and WUSCHEL-RELATED HOMEOBOX5 (WOX5) [38]. The involvement of RBR1 in the division and differentiation decisions seems to be general in many, perhaps all, cell types. This has been demonstrated for the interaction of RBR1 with the transcription factor FAMA, which prevents overproliferation of stomatal precursors [39]. Whether these RBR1dependent processes are regulated by the same set of CDK–cyclin-dependent phosphorylation events that affect cell cycle progression is completely unknown. Nevertheless, the possible involvement of CDKA;1 is likely because the effects of a loss of CDKA activity can be restored, at least partially, by reducing RBR1 function [23,40]. In turn, whether the RBR1 functions are cell cycle-dependent (in the sense of whether cells respond to maintain a proper division/differentiation balance at any cell cycle stage) is also an open question [41]. In fact, RBR1 is a substrate for different kinases, including S6K, and RBR1 function depends on its phosphorylation state at various residues [42,43]. Some developmental regulators target other cell cycle components. One example is TCP4 (a member of the TEOSINTE BRANCHED1–CYCLIN-PROLIFERATING CELL FACTOR family), a strong repressor of cell proliferation that arrests cells in G2–M. TCP4 plays a dual role as an activator of the inhibitor of CDK–cyclin, also known as KRP1 (KIP-RELATED PROTEIN1) and of miR396b [44]. Therefore, it will not be surprising to identify novel molecular links between regulators of organ development and the cell cycle machinery.
Cell Cycle Progression and Chromatin Accumulating evidence support that important chromatin-associated events are directly linked to cell cycle transitions (Figure 2) [41]. We can envisage, at least, two major questions that need to be addressed in the future to understand the links between chromatin dynamics during the cell cycle and transcriptional regulation and genome replication. Regarding the first, a recent study reported on the identification of the DREAM (DP/RB/E2F/ MuvB) complex, a crucial regulatory complex active in G2 [45], and assigned functions different from those of the RBR–E2F–DP complex acting in G1 (Figure 3). Delineating how CDK–cyclin activity modulates the function of DREAM in G2 is a future challenge. Access of these multiprotein complexes to their target genes is facilitated not only by transcription factors but also directly by changes in chromatin organization. Thus, the activity of nucleosome remodeling complexes and histone deacetylases await detailed analysis and identification (see also below). Interestingly, RBR likely interacts with SWI/SNF (SWITCH/SUCROSE NON FERMENTABLE)
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Figure 3. Coupling of Chromatin Modifications to Cell Cycle Progression. Cell cycle progression is coordinated with chromatin-associated modifications. These are mainly acetylation (ac) and methylation (me) of histones H3 and H4, methylation of cytosine residues (mC), and incorporation of the canonical histone H3.1. The cell cycle is defined by the interphase (G1, S, G2) and mitotic (P, prophase; M, metaphase; A, anaphase; T, telophase) phases. Note that in most cases the causal relationship between specific chromatin marks and cell cycle transitions has not been demonstrated. Abbreviations: DREAM, DP/RB/E2F/MuvB complex; E2F, transcription factor E2F; MSA, M-specific activator; ORI, origin of replication; RBR, RETINOBLASTOMA-RELATED.
complexes, although only circumstantial evidence is available supporting the regulation of cell cycle gene expression [46]. Genome replication initiates at multiple sites called DNA replication origins (ORIs). All potential sites are bound by pre-replication complexes, but only a subset of them are activated in each Sphase. ORI specification does not depend on a DNA consensus sequence but seems to be directly influenced by the chromatin landscape [47]. However, the chromatin features that define ORIs are poorly known (Figure 3) [48]. Advances in this field will require the availability of protocols and experimental strategies that allow the study of ORI function in whole plants because this would greatly facilitate the use of appropriate mutants and genetic approaches to address directly the relevance of chromatin marks on ORI specification and function. DNA replication timing appears to be also intimately linked to gene expression patterns [49].
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Therefore, a further step ahead would be the study of S-phase initiation and progression in the context of different cell types defined by their specific transcriptional programs.
Cell Division Potential and Chromatin Polycomb repressor complexes (PRC2) are responsible for depositing H3K27me3 at various stages of plant development [50,51] and are known to regulate the expression of many developmental and tissue-specific genes. The post-embryonic nature of plant organogenesis requires the continuous production of cells to form new organs. Thus, given the coupling of cell proliferation potential, cell fate decisions, and the regulation of developmental phase transitions, it would be interesting to explore whether and how Polycomb activity relates to cell cycle progression and, as discussed above, with cell specification (Figure 3). Further evidence in the same direction comes from phenotypic analysis of PRC2 mutants at the cellular level. These fail to retain the differentiated root hair state, and generate calli composed of undifferentiated cells, but, surprisingly, root hair nuclei develop the endoreplication cycle and eventually reinitiate mitotic division [52]. Altered levels of the PRC2 target genes are responsible for this phenotype, although a functional link to modifications of cell cycle machinery has not been yet determined. In another context, misexpression of MUTE and SPEECHLESS leads to a severe loss of H3K27me3 marks, strongly suggesting a link with PRC2 function [39]. Incidentally, this overproliferation phenotype is reminiscent of RBR1 loss of function in leaves that leads to hyperplasia [53], opening the question of whether this phenotype is solely dependent on the alteration of cell cycle progression or is partially dependent on the coupling to appropriate H3K27me3 deposition. Repression of the floral regulator FLC (FLOWERING LOCUS C), a paradigmatic Polycombregulated gene, is linked to dividing cells [54,55], indirectly suggesting that DNA replication is a prerequisite for stable maintenance of FLC repression. This finding underscores the observed coupling between PRC2 activity and DNA replication through the participation of DNA polymerases. The molecular mechanisms are not fully delineated but it is clear that mutations in the INCURVATA2 (ICU2) gene, encoding the catalytic subunit of DNA polymerase /, lead to defects in FLC repression and stable maintenance of H3K27me3 level, although PRC2 recruitment and H3K27me3 deposition are normal [56,57]. ESD7 (EARLY IN SHORT DAYS7), which encodes the catalytic subunit of DNA polymerase e, interacts with ICU2 and is also required for correct repression of flowering through epigenetic mechanisms [58]. Finally, mutations in DNA polymerase d also affect flowering but through a different mechanism because the H3K4me3 level of the SEPALLATA3 (SEP3) gene is increased and, as a consequence of its ectopic expression, FLOWERING LOCUS T (FT) transcription is promoted [59]. These data reveal that the three major replicative DNA polymerases that act during S-phase are crucial for genome replication and maintenance of epigenetic marks in genes involved in plant development (Figures 2,3). Furthermore, another component involved in replication fork progression, topoisomerase 1/ (TOP1/), also plays a role in PRC2-mediated repression of WUS, as well as other targets [60]. Whether other genes are also affected and the detailed molecular interactions responsible for linking S-phase, DNA replication fork progression, and the maintenance of chromatin modifications are totally unexplored areas. It is worth noting that recent data obtained in animal cells could illuminate the path to follow [61]. One major component of chromatin organization is the set of histone (canonical and variants) proteins that form each nucleosome. In addition to the canonical forms (H2A, H2B, H3, H4) there are multiple variants (H2A.Z, H2A.W, and H3.3, among others) that play distinct roles in chromatin organization and function [62]. Because in every S-phase only canonical histones are assembled onto newly synthesized DNA (Figure 3), there must be mechanisms coordinating histone incorporation and variant exchange with the division or differentiation status of the cell. Initial results in this direction have been recently obtained by direct determination of the balance
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between canonical H3.1 and variant H3.3 histones. It has been found that cells with a high proliferation rate within the root meristem possess a high H3.1:H3.3 ratio, while cells undergoing their last cell cycle before entering a differentiation pathway show a low ratio owing to a massive eviction of H3.1 in G2 [63]. Moreover, the last cell cycle in the proliferation domain of the root apical meristem is characterized by a longer G2 phase that most likely favors H3.1 eviction. Thus, histone dynamics could be used as a read-out of proliferation status. This is likely to be of general applicability because the H3.1:H3.3 pattern is observed in various cell types and also in endoreplicating cells [63]. Interestingly, chromatin-remodeling factors interact with GRFs [64] that affect the proliferation potential of stem cell derivatives in the root [25]. Therefore, it is tempting to speculate that different transcription factors, for example GRFs and PLT, which impinge on the proliferation potential, may act coordinately with the H3.1 deposition and eviction processes, although direct evidence of such coupling awaits further experimentation. In this context, regeneration is a unique feature of plant explants with potential biotechnological applications because it allows the generation of somatic embryos or new organs, primarily roots and shoots. Reactivation of post-mitotic cells by dedifferentiation and entering into a cell division program is a prerequisite. Ideally, manipulating the activity of cell cycle activators (e.g., cyclins, E2F–DP) and/or repressors (e.g., RBR1, KRPs) could provide a useful framework for improving regeneration efficiency. Other cellular factors, such as chromatin organization, should perhaps be considered as additional players [65].
The Endocycle Many proliferating cell types in diverse plant species arrest the cell cycle and enter the endocycle. This is a special cell cycle in which the genome is duplicated once or several times (endoreplication) without chromosome segregation, leading to polyploid cells. The endocycle program is frequently developmentally and environmentally regulated to initiate various differentiation pathways [66]. The molecular mechanisms controlling the initiation of endoreplication have been delineated and the basic requirements are known [67]. However, the maintenance and arrest of the endocycle program are much less well understood and deserve special attention (Figure 2). Furthermore, many cellular factors participate in both the cell cycle and the endocycle, for example CDK–cyclins, CDK inhibitors, and DNA replication proteins, but their availability and regulation seem to be different in the two cases. Whether the DNA replication origins active during the cell cycle possess the same chromosomal features as those used during the endocycle is not presently known. One interesting hypothesis to test is that the 3D organization of the nucleus and the existence of chromatin domains are relevant for the endocycle process. This is conceivable based on observations of chromatin dynamics in diploid guard cells and polyploid pavement cells in the epidermis of Arabidopsis, revealed using a GFP-mediated chromatin-tagging system, which demonstrated a larger nuclear territory in endoreplicated cells [68].
DNA Damage Response A large amount of DNA modifications occur on a daily basis as a result of both normal metabolism and the effect of environmental challenges to which plants are particularly exposed. These DNA alterations are very harmful to genomic integrity. All cells have evolved mechanisms to cope with DNA damage, and plants are no exception [69]. DNA damage tolerance pathways are frequently linked to modulation of the activity of cell cycle regulators. One of the primary responses is the phosphorylation of SOG1 that transcriptionally regulates the expression of hundreds of DNA damage-response genes, many of which are direct regulators of both the cell cycle and endoreplication [70,71]. The mechanistic details of the SOG1-mediated cell cycle arrest and its coupling with endocycle onset are still far from being fully understood. Another component of the DNA damage response is WEE1 that participates in the intra-S checkpoint [69,72]. In cases where genome damage cannot be repaired, a cell death pathway is triggered,
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but plants have evolved mechanisms to prevent loss of all proliferating stem cells as a result of extensive DNA damage. This relies on QC cells that are more resistant to DNA damage than other cells, and hence they provide a back-up mechanism to restore a stem cell population that can resume organ growth. Importantly, stem cells are actually hypersensitive to DNA damage, which likely contributes to preventing the accumulation of mutations in the stem cell derivatives [73]. Future studies should provide an integrative view of how plants tolerate DNA damage at the organismal level.
Concluding Remarks The examples discussed in previous paragraphs serve to highlight the relevance of cell cycle control as a hub integrating many aspects of plant physiology (Figure 1), although they do not constitute a comprehensive list. In fact, there are other cases that suggest even more unanticipated functional interactions, as illustrated in the following reports: (i) the impact of metabolic balance on stem cell activity directly via E2F and S-phase genes [74]; (ii) the role of reactive oxygen species on the proliferation and differentiation status in the root apical meristem [75] and during lateral root development [76], the coupling of cell cycle to the circadian clock [77]; and (iii) the altered innate immunity response of mutants in the SIM–SMR and cyclin D genes [78,79]. Further studies will be necessary in these and several other directions to fully understand in molecular terms the functional relevance of cell cycle regulators as targets of a plethora of cellular processes (see Outstanding Questions). Cell cycle research, as many other fields, has always benefited from technological advances that allowed exploring questions from a different perspective or at a different level of resolution. One such example is the refinement of image acquisition and analysis techniques that now allow the reconstruction of whole organs in 3D and their analysis over time, adding a new dimension (4D) [80]. These approaches could be combined with appropriate cell cycle markers to analyze in detail cell proliferation dynamics during organogenesis, thus obtaining information at the singlecell level of how they interact with each other. Moreover, the use of computational modeling can allow us to explore how cell proliferation dynamics interacts with patterning rules and developmental cues to establish cellular homeostasis during development [81,82]. Advanced imaging at single-cell resolution can be also potentially very informative, as illustrated by studies of the replication of the specialized structures occurring at telomeres. Single-cell telomere identification and length quantification has demonstrated that different cell lineages, including stem cells, have characteristic telomere lengths [83]. These data show that telomere dynamics during S-phase is intimately coupled to cell proliferation potential and meristem activity. The question remains as to whether other cell cycle features are also cell type-specific. Again a combination of appropriate cell cycle markers with single-cell imaging could be highly informative. Advances in single-cell technologies also have great potential in genomics and proteomics. Thus, single-cell RNA-seq has been applied successfully to plant cells [84,85]. This is not the case yet for chromatin immunoprecipitation (ChIP)-seq, which will be necessary for studies of cell cycle-dependent transcription factor binding and epigenetic studies. Therefore, it is conceivable that different versions of single-cell approaches will be crucial to define molecular events associated with cell cycle progression, arrest, and exit to differentiation, as well as to dedifferentiation (or trans-differentiation) during organ regeneration and wound repair. Because these approaches have an inherently high signal-to-noise ratio [86,87], an important demand for robust bioinformatics tools is increasing. Information derived from single-cell approaches needs to be transferred at the organ level, and ideally to the organismal level [88]. In this context, different growth conditions and adaptations
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Outstanding Questions Do we need to develop more powerful resources to extract novel information relevant for plant cell cycle research? Will we be able to integrate cell cycle regulation with other aspects of plant life? Can synthetic biology take advantage of new integrated knowledge on the cell cycle to generate better-performing plants that contribute to a better life?
have an effect on morphogenesis, for example changes in lateral root density, which are related to specific cell proliferation and differentiation patterns. Therefore, a potential source of valuable information that has been relatively underexploited in cell cycle studies is natural variation [89], and this could facilitate the identification of novel roles of cell cycle regulators in an integrative manner. The analysis of gene function and the acquisition of novel traits is now facilitated by advances in genome editing using CRISPR/Cas9 technology [90,91]. This technology has not been applied yet to cell cycle research on a routine basis but may represent a significant step ahead because, in many cases, functional redundancy is a major obstacle in cell cycle research. In any case, tailored and cell type-specific genome editing would be of great relevance given the essential nature of many cell cycle regulators. Furthermore, because the genome editing technologies are becoming available for crop species [92], it is likely that cell cycle knowledge obtained primarily in Arabidopsis could be useful for plants of commercial interest. Reevaluation with new eyes, concepts, and tools of the role of cell cycle regulators beyond their function as part of the machinery to produce new cells could be extremely enlightening to understand plant growth and development in an integrated manner. Cell cycle regulatory factors are direct targets that control ‘where’ these cells need to be produced (spatial control), ‘when’ during development (temporal control), and ‘what’ type of division is required (symmetrical or asymmetrical, periclinal or anticlinal). The famous statement of Dobzansky comes to mind: ‘Nothing in biology makes sense except in the light of evolution’ [93]. Let me paraphrase it: ‘Nothing in development and growth makes sense except in the light of cell proliferation and differentiation’. Thus, exciting times are ahead because studies in these and other promising directions should lead to discoveries that will widen considerably our perspective in the field beyond the cellular level into the organismal level. Acknowledgments I would like to thank all members of the laboratory for helpful discussions and comments, Z. Vergara for help with the figures, and E. Martinez-Salas for suggestions. This work has been supported by grants BFU2012-34821 and BFU2013-50098 (MINECO) and BFU2015-68396 (MINECO/FEDER), and by an institutional grant from Fundacion Ramon Areces to the Centro de Biologia Molecular ‘Severo Ochoa’.
References 1. Lee, M.G. and Nurse, P. (1987) Complementation used to clone a human homologue of the fission yeast cell cycle control gene cdc2. Nature 327, 31–35 2. John, P.C. et al. (1989) A homolog of the cell cycle control protein p34cdc2 participates in the division cycle of Chlamydomonas, and a similar protein is detectable in higher plants and remote taxa. Plant Cell 1, 1185–1193 3. Feiler, H.S. and Jacobs, T.W. (1990) Cell division in higher plants: a cdc2 gene, its 34-kDa product, and histone H1 kinase activity in pea. Proc. Natl. Acad. Sci. U.S.A. 87, 5397–5401 4. Ferreira, P.C. et al. (1991) The Arabidopsis functional homolog of the p34cdc2 protein kinase. Plant Cell 3, 531–540 5. Hata, S. et al. (1991) Isolation and characterization of cDNA clones for plant cyclins. EMBO J. 10, 2681–2688 6. Hemerly, A. et al. (1992) Genes regulating the plant cell cycle: isolation of a mitotic-like cyclin from Arabidopsis thaliana. Proc. Natl. Acad. Sci. U.S.A. 89, 3295–3299 7. Hirt, H. et al. (1992) Alfalfa cyclins: differential expression during the cell cycle and in plant organs. Plant Cell 4, 1531–1538 8. Van’T Hof, J. and Wilson, G.B. (1962) Studies on the control of mitotic activity: the effect of respiratory inhibitors on mitotic cycle time in the root meristem of Pisum sativum. Chromosoma 13, 39–46 9. Van’t Hof, J. (1965) Cell population kinetics of excised roots of Pisum sativum. J. Cell Biol. 27, 179–189
10. Gimenez-Martin, G. et al. (1965) A new method of labeling cells. J. Cell Biol. 26, 305–309 11. Van’t Hof, J. and Sparrow, A.H. (1965) Radiation effects on the growth rate and cell population kinetics of actively growing and dormant roots of Tradescantia paludosa. J. Cell Biol. 26, 187–199 12. Lopez-Saez, J.F. et al. (1966) Duration of the cell division cycle and its dependence on temperature. Z. Zellforsch. Mikrosk. Anat. 75, 591–600 13. Bryant, J.A. and Wildon, D.C. (1971) Cytoplasmic deoxyribonucleic acid in roots of Pisum sativum. Biochem. J. 121, 5P 14. Soni, R. et al. (1995) A family of cyclin D homologs from plants differentially controlled by growth regulators and containing the conserved retinoblastoma protein interaction motif. Plant Cell 7, 85–103 15. Dahl, M. et al. (1995) The D-type alfalfa cyclin gene cycMs4 complements G1 cyclin-deficient yeast and is induced in the G1 phase of the cell cycle. Plant Cell 7, 1847–1857 16. Xie, Q. et al. (1995) Identification and analysis of a retinoblastoma binding motif in the replication protein of a plant DNA virus: requirement for efficient viral DNA replication. EMBO J. 14, 4073–4082 17. Grafi, G. et al. (1996) A maize cDNA encoding a member of the retinoblastoma protein family: involvement in endoreduplication. Proc. Natl. Acad. Sci. U.S.A. 93, 8962–8967
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18. Xie, Q. et al. (1996) Plant cells contain a novel member of the retinoblastoma family of growth regulatory proteins. EMBO J. 15, 4900–4908 19. AGI (2000) Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408, 796–815 20. Wildwater, M. et al. (2005) The RETINOBLASTOMA-RELATED gene regulates stem cell maintenance in Arabidopsis roots. Cell 123, 1337–1349 21. Sozzani, R. et al. (2010) Spatiotemporal regulation of cell-cycle genes by SHORTROOT links patterning and growth. Nature 466, 128–132 22. Cruz-Ramirez, A. et al. (2012) A bistable circuit involving SCARECROW-RETINOBLASTOMA integrates cues to inform asymmetric stem cell division. Cell 150, 1002–1015 23. Weimer, A.K. et al. (2012) Retinoblastoma related1 regulates asymmetric cell divisions in Arabidopsis. Plant Cell 24, 4083–4095 24. Kinoshita, A. et al. (2015) A plant U-box protein, PUB4, regulates asymmetric cell division and cell proliferation in the root meristem. Development 142, 444–453 25. Rodriguez, R.E. et al. (2015) MicroRNA miR396 regulates the switch between stem cells and transit-amplifying cells in Arabidopsis roots. Plant Cell 27, 3354–3366 26. Van Damme, D. et al. (2011) Arabidopsis alpha Aurora kinases function in formative cell division plane orientation. Plant Cell 23, 4013–4024 27. Weimer, A.K. et al. (2016) Aurora kinases throughout plant development. Trends Plant Sci. 21, 69–79 28. Petrovska, B. et al. (2012) Plant Aurora kinases play a role in maintenance of primary meristems and control of endoreduplication. New Phytol. 193, 590–604 29. Yang, K. et al. (2014) Requirement for A-type cyclin-dependent kinase and cyclins for the terminal division in the stomatal lineage of Arabidopsis. J. Exp. Bot. 65, 2449–2461 30. De Smet, I. et al. (2008) Receptor-like kinase ACR4 restricts formative cell divisions in the Arabidopsis root. Science 322, 594–597 31. Yue, K. et al. (2016) PP2A-3 interacts with ACR4 and regulates formative cell division in the Arabidopsis root. Proc. Natl. Acad. Sci. U.S.A. 113, 1447–1452
44. Schommer, C. et al. (2014) Repression of cell proliferation by miR319-regulated TCP4. Mol. Plant 7, 1533–1544 45. Kobayashi, K. et al. (2015) Transcriptional repression by MYB3R proteins regulates plant organ growth. EMBO J. 34, 1992–2007 46. Jegu, T. et al. (2015) A SWI/SNF chromatin remodelling protein controls cytokinin production through the regulation of chromatin architecture. PLoS One 10, e0138276 47. Sequeira-Mendes, J. and Gutierrez, C. (2015) Links between genome replication and chromatin landscapes. Plant J. 83, 38–51 48. Costas, C. et al. (2011) Genome-wide mapping of Arabidopsis origins of DNA replication and their associated epigenetic marks. Nat. Struct. Mol. Biol. 18, 395–400 49. Bass, H.W. et al. (2015) Defining multiple, distinct, and shared spatiotemporal patterns of DNA replication and endoreduplication from 3D image analysis of developing maize (Zea mays L.) root tip nuclei. Plant Mol. Biol. 89, 339–351 50. Kuwabara, A. and Gruissem, W. (2014) Arabidopsis retinoblastoma-related and Polycomb group proteins: cooperation during plant cell differentiation and development. J. Exp. Bot. 65, 2667– 2676 51. Xiao, J. and Wagner, D. (2015) Polycomb repression in the regulation of growth and development in Arabidopsis. Curr. Opin. Plant Biol. 23, 15–24 52. Ikeuchi, M. et al. (2015) PRC2 represses dedifferentiation of mature somatic cells in Arabidopsis. Nat. Plants 1, 15089 53. Desvoyes, B. et al. (2006) Cell type-specific role of the retinoblastoma/E2F pathway during Arabidopsis leaf development. Plant Physiol. 140, 67–80 54. Finnegan, E.J. and Dennis, E.S. (2007) Vernalization-induced trimethylation of histone H3 lysine 27 at FLC is not maintained in mitotically quiescent cells. Curr. Biol. 17, 1978–1983 55. Berry, S. et al. (2015) Local chromatin environment of a Polycomb target gene instructs its own epigenetic inheritance. Elife 4, e07205 56. Barrero, J.M. et al. (2007) INCURVATA2 encodes the catalytic subunit of DNA polymerase alpha and interacts with genes involved in chromatin-mediated cellular memory in Arabidopsis thaliana. Plant Cell 19, 2822–2838
32. Roeder, A.H. et al. (2012) Cell cycle regulates cell type in the Arabidopsis sepal. Development 139, 4416–4427
57. Hyun, Y. et al. (2013) The catalytic subunit of Arabidopsis DNA polymerase alpha ensures stable maintenance of histone modification. Development 140, 156–166
33. Harashima, H. and Sugimoto, K. (2016) Integration of developmental and environmental signals into cell proliferation and differentiation through RETINOBLASTOMA-RELATED 1. Curr. Opin. Plant Biol. 29, 95–103
58. del Olmo, I. et al. (2010) EARLY IN SHORT DAYS 7 (ESD7) encodes the catalytic subunit of DNA polymerase epsilon and is required for flowering repression through a mechanism involving epigenetic gene silencing. Plant J. 61, 623–636
34. Perilli, S. et al. (2013) RETINOBLASTOMA-RELATED protein stimulates cell differentiation in the Arabidopsis root meristem by interacting with cytokinin signaling. Plant Cell 25, 4469–4478
59. Iglesias, F.M. et al. (2015) The Arabidopsis DNA polymerase delta has a role in the deposition of transcriptionally active epigenetic marks, development and flowering. PLoS Genet. 11, e1004975
35. Heyman, J. et al. (2013) ERF115 controls root quiescent center cell division and stem cell replenishment. Science 342, 860–863 36. Cruz-Ramirez, A. et al. (2013) A SCARECROW-RETINOBLASTOMA protein network controls protective quiescence in the Arabidopsis root stem cell organizer. PLoS Biol. 11, e1001724 37. Vilarrasa-Blasi, J. et al. (2014) Regulation of plant stem cell quiescence by a brassinosteroid signaling module. Dev. Cell 30, 36–47 38. Forzani, C. et al. (2014) WOX5 suppresses CYCLIN D activity to establish quiescence at the center of the root stem cell niche. Curr. Biol. 24, 1939–1944 39. Matos, J.L. et al. (2014) Irreversible fate commitment in the Arabidopsis stomatal lineage requires a FAMA and RETINOBLASTOMA-RELATED module. Elife 3, e03271 40. Nowack, M.K. et al. (2012) Genetic framework of cyclin-dependent kinase function in Arabidopsis. Dev. Cell 22, 1030–1040
60. Liu, X. et al. (2014) DNA topoisomerase I affects polycomb group protein-mediated epigenetic regulation and plant development by altering nucleosome distribution in Arabidopsis. Plant Cell 26, 2803–2817 61. Alabert, C. et al. (2015) Two distinct modes for propagation of histone PTMs across the cell cycle. Genes Dev. 29, 585–590 62. Talbert, P.B. and Henikoff, S. (2013) Phylogeny as the basis for naming histones. Trends Genet. 29, 499–500 63. Otero, S. et al. (2016) Histone H3 dynamics uncovers domains with distinct proliferation potential in the Arabidopsis root. Plant Cell http://dx.doi.org/10.1105/tpc.15.01003 64. Debernardi, J.M. et al. (2014) Post-transcriptional control of GRF transcription factors by microRNA miR396 and GIF co-activator affects leaf size and longevity. Plant J. 79, 413–426
41. Desvoyes, B. et al. (2014) Looking at plant cell cycle from the chromatin window. Front. Plant Sci. 5, 369
65. Ikeuchi, M. et al. (2015) Control of plant cell differentiation by histone modification and DNA methylation. Curr. Opin. Plant Biol. 28, 60–67
42. Henriques, R. et al. (2010) Arabidopsis S6 kinase mutants display chromosome instability and altered RBR1-E2F pathway activity. EMBO J. 29, 2979–2993
66. Polyn, S. et al. (2015) Cell cycle entry, maintenance, and exit during plant development. Curr. Opin. Plant Biol. 23, 1–7
43. Magyar, Z. et al. (2012) Arabidopsis E2FA stimulates proliferation and endocycle separately through RBR-bound and RBR-free complexes. EMBO J. 31, 1480–1493
832
Trends in Plant Science, October 2016, Vol. 21, No. 10
67. Edgar, B.A. et al. (2014) Endocycles: a recurrent evolutionary innovation for post-mitotic cell growth. Nat. Rev. Mol. Cell Biol. 15, 197–210
68. Kato, N. and Lam, E. (2003) Chromatin of endoreduplicated pavement cells has greater range of movement than that of diploid guard cells in Arabidopsis thaliana. J. Cell Sci. 116, 2195–2201 69. Hu, Z. et al. (2016) Mechanisms used by plants to cope with DNA damage. Ann. Rev. Plant Biol. 67, 439–462 70. Yoshiyama, K. et al. (2009) Suppressor of gamma response 1 (SOG1) encodes a putative transcription factor governing multiple responses to DNA damage. Proc. Natl. Acad. Sci. U.S.A. 106, 12843–12848 71. Yoshiyama, K.O. et al. (2013) ATM-mediated phosphorylation of SOG1 is essential for the DNA damage response in Arabidopsis. EMBO Rep. 14, 817–822 72. Cools, T. et al. (2011) The Arabidopsis thaliana checkpoint kinase WEE1 protects against premature vascular differentiation during replication stress. Plant Cell 23, 1435–1448 73. Fulcher, N. and Sablowski, R. (2009) Hypersensitivity to DNA damage in plant stem cell niches. Proc. Natl. Acad. Sci. U.S.A. 106, 20984–20988 74. Xiong, Y. et al. (2013) Glucose–TOR signalling reprograms the transcriptome and activates meristems. Nature 496, 181–186 75. Tsukagoshi, H. et al. (2010) Transcriptional regulation of ROS controls transition from proliferation to differentiation in the root. Cell 143, 606–616 76. Manzano, C. et al. (2014) The emerging role of reactive oxygen species signaling during lateral root development. Plant Physiol. 165, 1105–1119 77. Miyagishima, S.Y. et al. (2014) Translation-independent circadian control of the cell cycle in a unicellular photosynthetic eukaryote. Nat. Commun. 5, 3807 78. Hamdoun, S. et al. (2016) Differential roles of two homologous cyclin-dependent kinase inhibitor genes in regulating cell cycle and innate immunity in Arabidopsis. Plant Physiol. 170, 515–527 79. Eichmann, R. and Schafer, P. (2015) Growth versus immunity – a redirection of the cell cycle? Curr. Opin. Plant Biol. 26, 106–112
80. Bassel, G.W. and Smith, R.S. (2016) Quantifying morphogenesis in plants in 4D. Curr. Opin. Plant Biol. 29, 87–94 81. Ortiz-Gutierrez, E. et al. (2015) A dynamic gene regulatory network model that recovers the cyclic behavior of Arabidopsis thaliana cell cycle. PLoS Comput. Biol. 11, e1004486 82. von Wangenheim, D. et al. (2016) Rules and self-organizing properties of post-embryonic plant organ cell division patterns. Curr. Biol. 26, 439–449 83. Gonzalez-Garcia, M.P. et al. (2015) Single-cell telomere-length quantification couples telomere length to meristem activity and stem cell development in Arabidopsis. Cell Rep. 11, 977–989 84. Efroni, I. and Birnbaum, K.D. (2016) The potential of single-cell profiling in plants. Genome Biol. 17, 65 85. Efroni, I. et al. (2015) Quantification of cell identity from single-cell gene expression profiles. Genome Biol. 16, 9 86. Buettner, F. et al. (2015) Computational analysis of cell-to-cell heterogeneity in single-cell RNA-sequencing data reveals hidden subpopulations of cells. Nat. Biotechnol. 33, 155–160 87. McDavid, A. et al. (2016) The contribution of cell cycle to heterogeneity in single-cell RNA-seq data. Nat. Biotechnol. 34, 591–593 88. Chaiwanon, J. et al. (2016) Information integration and communication in plant growth regulation. Cell 164, 1257–1268 89. Zhu, W. et al. (2015) Natural variation identifies ICARUS1, a universal gene required for cell proliferation and growth at high temperatures in Arabidopsis thaliana. PLoS Genet. 11, e1005085 90. Ding, Y. et al. (2016) Recent advances in genome editing using CRISPR/Cas9. Front. Plant Sci. 7, 703 91. Puchta, H. (2016) Genome engineering using CRISPR/Cas: getting more versatile and more precise at the same time. Genome Biol. 17, 51 92. Khatodia, S. et al. (2015) The CRISPR/Cas genome-editing tool: application in improvement of crops. Front. Plant Sci. 7, 506 93. Dobzhansky, T. (1973) Nothing in biology makes sense except in the light of evolution. Am. Biol. Teacher 35, 125–129
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25 Years of Cell Cycle Research: What's Ahead? Crisanto Gutierrez1,* We have reached 25 years since the first molecular approaches to plant cell cycle. Fortunately, we have witnessed an enormous advance in this field that has benefited from using complementary approaches including molecular, cellular, genetic and genomic resources. These studies have also branched and demonstrated the functional relevance of cell cycle regulators for virtually every aspect of plant life. The question is – where are we heading? I review here the latest developments in the field and briefly elaborate on how new technological advances should contribute to novel approaches that will benefit the plant cell cycle field. Understanding how the cell division cycle is integrated at the organismal level is perhaps one of the major challenges.
Trends Research carried out mainly over the past 25 years has led to a detailed understanding of plant cell cycle at the physiological, cellular, molecular, genetic, and genomic levels. Many aspects of plant life, including formative cell divisions during organogenesis, the exit to differentiation, chromatin and epigenetics, metabolism and stem cell potential, circadian clock, and innate immunity, among others, impinge or depend directly on cell cycle regulators.
A Timeline of Plant Cell Cycle Research Cell cycle research witnessed a turning point in 1987 when a human homolog of the S. cerevisiae p34cdc2, a cyclin-dependent kinase (CDK) crucial for cell cycle progression, was isolated by a functional complementation screen [1]. This study, together with others, was worth a Nobel Prize in 2001, and demonstrated that a key cell cycle regulator was conserved in very distantly related eukaryotes. It also triggered a new interest of plant researchers to investigate if plant cells could use CDK–cyclin complexes as part of the cell cycle progression control. Thus, 25 years ago Peter John and colleagues, including Melanie Lee (who identified the human p34cdc2 protein), were the first to (i) identify a p34 protein in plant extracts (including Chlamydomonas reinhardtii and Arabidopsis thaliana) with high immunological similarities to the yeast and human p34cdc2 protein, and (ii) show that its phosphorylation status changed during the cell cycle [2]. The kinase activity of plant p34cdc2 complexes as well as the cloning of the p34 encoding gene in pea [3] and Arabidopsis [4] were reported soon afterwards. An immediate challenge to fully confirm the presence of such complexes in plants was to identify the cyclin (CYC) moieties. The effort of several laboratories succeeded in the identification and cloning of various cyclins, later identified as A- or B-type cyclins [5–7], highly similar to the yeast proteins. Plant cell cycle research has passed through several distinct stages marked by successive breakthrough discoveries and technological advances (Figure 1). Thus, to be fair, interest in plant cell cycle bloomed much earlier, 50 years ago. The initial efforts were focused on cytological and physiological approaches to mitotic activity and kinetics [8–10], environmental effects [11,12], and S-phase progression [13], to mention a few representative examples. Later, the 1990s were marked with advances that revolutionized cell cycle research because they initiated molecular and cell biology studies with two main series of findings (Figure 1). One, already mentioned above, that allowed the cloning of several CDK and cyclin genes, most likely involved in the G2/M transition. Another, the identification of key cell cycle regulators that were not present in yeast, such as D-type cyclins [14,15] and the retinoblastoma protein, a homolog of the human tumor-suppressor [16–18]. Together with other reports, they provided the framework to begin to understand that the plant cell cycle machinery, with some unique features, was much more like that in animal cells than in yeast. The beginning of the 21st century was marked by the
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Future challenges aim at obtaining an integrative understanding of cell proliferation in the whole plant. Recent advances in next-generation sequencing and single-cell omics, sophisticated imaging, and cell typespecific genome editing are among the novel approaches that should contribute to solve major questions on cell cycle research in model and crop species.
1
Centro de Biologia Molecular Severo Ochoa, Consejo Superior de Investigaciones Científicas (CSIC), Universidad Autónoma de Madrid (UAM), Nicolas Cabrera 1, 28049 Madrid, Spain
*Correspondence: [email protected] (C. Gutierrez).
http://dx.doi.org/10.1016/j.tplants.2016.06.007 © 2016 Elsevier Ltd. All rights reserved.
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Figure 1. Timeline of Plant Cell Cycle Research.
explosion of several ‘omic’ approaches (Figure 1), greatly facilitated by the availability of the full genome sequence of Arabidopsis, the first multicellular organism, together with Drosophila, to be sequenced [19]. It is obvious that all the different experimental approaches are still fruitfully combined and will contribute to provide further insights into the plant cell cycle regulatory networks in individual cells. Where are we moving, and what are the challenges ahead? In the following paragraphs I will expand upon these ideas, highlighting some of the various lines that may drive plant cell cycle research in the coming years.
Formative Cell Divisions and Cell Fate Cell proliferation is an integral part of morphogenesis. Formative cell divisions, also known as asymmetric cell divisions (ACDs), are one widespread example of a specific type of cell division that impact on morphogenesis. Typically, ACD produces two daughter cells that will take uniquely different fates, breaking the paradigm that cell division produces two identical cells. They occur in different locations across the plant body to generate new cell types and/or expand organogenesis. It was originally thought that developmental signals would trigger a signaling cascade that ultimately impinges on cell division as a downstream process. Indeed, recent evidence shows that various components of the cell cycle machinery are direct and immediate targets of developmental regulators that control ACD (Figure 2). The case of cyclin CYCD6;1 illustrates such coordination. CYCD6;1 is required for ACD in the root apical meristem and, specifically, triggers division of the cortex-endodermis initial, a process that is inhibited by the RETINOBLASTOMA-RELATED1/SCARECROW/SHORTROOT (RBR1/ SCR/SHR) module [20,21]. CYCD6;1-containing CDKA and CDKB1 complexes phosphorylate RBR1, releasing the inhibition of ACD [22,23]. It is not known whether these complexes act sequentially or in an alternative manner. Although it is known that CYCD6;1 availability is controlled by an E3 ubiquitin ligase [24], learning about how CYCD6;1 expression and stability are controlled in different locations will be enlightening. The daughter cells resulting from the asymmetrical division of stem cells in the root apical meristem undergo several division cycles to constitute the root meristem (also called transit amplifying compartment) before exit to differentiation. The plant-specific GROWTH-REGULATING FACTORS (GRFS) are expressed in proliferating cells in the meristem, but are excluded from stem cells where they are repressed by miR396 [25]. Importantly, GRFs repress PLETHORA (PLT) genes in the stem cell derivatives and are required to prevent periclinal formative divisions.
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Formave cell divisions Cell fate
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Figure 2. Summary of the Physiological Processes that Impinge Directly on Cell Cycle Regulators.
Further understanding on how this module regulates downstream targets and whether they are cell cycle regulators is of primary relevance (see also below). Crucial to the establishment of an asymmetry in the daughter cells is not only the signaling cascade channeled through cell cycle regulators but also the control of the cell division plane. Choosing between symmetrical/asymmetrical and/or periclinal/anticlinal divisions is not trivial because it involves a dramatic change in the internal organization of the cytoskeleton relative to that of the mother cell. Aurora (AUR) kinases are likely to be key players in such decisions, in particular ACDs, because they are involved in the orientation of the cell division plane in various stages of plant development [26,27]. Strikingly, AUR proteins have putative RBR1 binding sites [28], which leads us to speculate on a possible link between AUR-controlled ACD and other processes impinging on ACD, although direct evidence is still lacking. This possible link also reveals the complexity of ACD control depending on the phosphorylation status of RBR1 during the cell cycle, the unknown set of AUR targets, and their availability. The coordination of cell proliferation and cell fate acquisition is a fundamental aspect of patterning. Remarkably, several cell cycle regulators have been identified as direct targets of developmentally regulated master genes (Figure 2), such as (i) FOURLIPS (FLP/MYB88) that controls guard mother cell division in the stomatal lineage by regulating the expression of CYCA2;3, CDKA;1, and CDKB1 [29], and (ii) ACR4 (ARABIDOPSIS CRINKLY4), that is necessary for restricting cell division at early stages of lateral root development [30], and which also controls columella stem cells division [31] and giant cell identity in sepals [32]. Interestingly, acr4 mutants show hyperproliferation and, concomitantly, reduced endoreplication [32]. Whether ACR4 and other master regulators of cell specification directly target other cell cycle genes, as well as on endoreplication genes, to control their activity remains to be studied.
Division and Differentiation Balance Organogenesis relies on two intimately coordinated processes: the production of a sufficient number of cells and progression of selected groups through distinct differentiation pathways to generate the specific cell types of a particular organ (Figure 2). An excess of the division program (or a delayed entry into differentiation) could lead to hyperplasia and dysfunctional organs. Likewise, premature exit from the cell cycle to differentiation (or a reduction in proliferative
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potential) generally leads to hypoplasia through exhaustion of the pool of dividing cells. The postembryonic nature of plant organogenesis, together with its responsiveness not only to internal (developmental cues, hormonal signals) but also to external (environmental challenges) inputs, represents an additional layer of regulation that confers plasticity to plant organogenesis. The root meristem has provided insightful information supporting the notion that some cell cycle regulators are at the crossroads of the balance between cell division and differentiation [33]. There are several examples illustrating that RBR1 is a crucial hub for integrate hormonal signals that not only control organ size but also determine the division potential of various cell types within the root. Thus, activation of the AUXIN RESPONSE FACTOR19 (ARF19), a factor that promotes differentiation of root meristem cells, requires RBR1 and ARABIDOPSIS RESPONSE REGULATOR12 (ARR12), a cytokinin-dependent transcription factor [34]. Another example is the division of quiescent center (QC) cells, which is extremely rare but it occurs in response to various stimuli. This is strictly controlled by the balance between the ETHYLENE RESPONSE FACTOR115 (ERF115)-mediated activation [35] and the RBR1dependent repression [36]. ERF115 expression depends on BRAVO and the brassinosteroid signaling [37] and CELL CYCLE SWITCH52 (CCS52A2), an activator of the anaphasepromoting complex 35, whereas RBR1 and its interaction with SCR depend on the activity of CYCD3;3 and WUSCHEL-RELATED HOMEOBOX5 (WOX5) [38]. The involvement of RBR1 in the division and differentiation decisions seems to be general in many, perhaps all, cell types. This has been demonstrated for the interaction of RBR1 with the transcription factor FAMA, which prevents overproliferation of stomatal precursors [39]. Whether these RBR1dependent processes are regulated by the same set of CDK–cyclin-dependent phosphorylation events that affect cell cycle progression is completely unknown. Nevertheless, the possible involvement of CDKA;1 is likely because the effects of a loss of CDKA activity can be restored, at least partially, by reducing RBR1 function [23,40]. In turn, whether the RBR1 functions are cell cycle-dependent (in the sense of whether cells respond to maintain a proper division/differentiation balance at any cell cycle stage) is also an open question [41]. In fact, RBR1 is a substrate for different kinases, including S6K, and RBR1 function depends on its phosphorylation state at various residues [42,43]. Some developmental regulators target other cell cycle components. One example is TCP4 (a member of the TEOSINTE BRANCHED1–CYCLIN-PROLIFERATING CELL FACTOR family), a strong repressor of cell proliferation that arrests cells in G2–M. TCP4 plays a dual role as an activator of the inhibitor of CDK–cyclin, also known as KRP1 (KIP-RELATED PROTEIN1) and of miR396b [44]. Therefore, it will not be surprising to identify novel molecular links between regulators of organ development and the cell cycle machinery.
Cell Cycle Progression and Chromatin Accumulating evidence support that important chromatin-associated events are directly linked to cell cycle transitions (Figure 2) [41]. We can envisage, at least, two major questions that need to be addressed in the future to understand the links between chromatin dynamics during the cell cycle and transcriptional regulation and genome replication. Regarding the first, a recent study reported on the identification of the DREAM (DP/RB/E2F/ MuvB) complex, a crucial regulatory complex active in G2 [45], and assigned functions different from those of the RBR–E2F–DP complex acting in G1 (Figure 3). Delineating how CDK–cyclin activity modulates the function of DREAM in G2 is a future challenge. Access of these multiprotein complexes to their target genes is facilitated not only by transcription factors but also directly by changes in chromatin organization. Thus, the activity of nucleosome remodeling complexes and histone deacetylases await detailed analysis and identification (see also below). Interestingly, RBR likely interacts with SWI/SNF (SWITCH/SUCROSE NON FERMENTABLE)
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Figure 3. Coupling of Chromatin Modifications to Cell Cycle Progression. Cell cycle progression is coordinated with chromatin-associated modifications. These are mainly acetylation (ac) and methylation (me) of histones H3 and H4, methylation of cytosine residues (mC), and incorporation of the canonical histone H3.1. The cell cycle is defined by the interphase (G1, S, G2) and mitotic (P, prophase; M, metaphase; A, anaphase; T, telophase) phases. Note that in most cases the causal relationship between specific chromatin marks and cell cycle transitions has not been demonstrated. Abbreviations: DREAM, DP/RB/E2F/MuvB complex; E2F, transcription factor E2F; MSA, M-specific activator; ORI, origin of replication; RBR, RETINOBLASTOMA-RELATED.
complexes, although only circumstantial evidence is available supporting the regulation of cell cycle gene expression [46]. Genome replication initiates at multiple sites called DNA replication origins (ORIs). All potential sites are bound by pre-replication complexes, but only a subset of them are activated in each Sphase. ORI specification does not depend on a DNA consensus sequence but seems to be directly influenced by the chromatin landscape [47]. However, the chromatin features that define ORIs are poorly known (Figure 3) [48]. Advances in this field will require the availability of protocols and experimental strategies that allow the study of ORI function in whole plants because this would greatly facilitate the use of appropriate mutants and genetic approaches to address directly the relevance of chromatin marks on ORI specification and function. DNA replication timing appears to be also intimately linked to gene expression patterns [49].
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Therefore, a further step ahead would be the study of S-phase initiation and progression in the context of different cell types defined by their specific transcriptional programs.
Cell Division Potential and Chromatin Polycomb repressor complexes (PRC2) are responsible for depositing H3K27me3 at various stages of plant development [50,51] and are known to regulate the expression of many developmental and tissue-specific genes. The post-embryonic nature of plant organogenesis requires the continuous production of cells to form new organs. Thus, given the coupling of cell proliferation potential, cell fate decisions, and the regulation of developmental phase transitions, it would be interesting to explore whether and how Polycomb activity relates to cell cycle progression and, as discussed above, with cell specification (Figure 3). Further evidence in the same direction comes from phenotypic analysis of PRC2 mutants at the cellular level. These fail to retain the differentiated root hair state, and generate calli composed of undifferentiated cells, but, surprisingly, root hair nuclei develop the endoreplication cycle and eventually reinitiate mitotic division [52]. Altered levels of the PRC2 target genes are responsible for this phenotype, although a functional link to modifications of cell cycle machinery has not been yet determined. In another context, misexpression of MUTE and SPEECHLESS leads to a severe loss of H3K27me3 marks, strongly suggesting a link with PRC2 function [39]. Incidentally, this overproliferation phenotype is reminiscent of RBR1 loss of function in leaves that leads to hyperplasia [53], opening the question of whether this phenotype is solely dependent on the alteration of cell cycle progression or is partially dependent on the coupling to appropriate H3K27me3 deposition. Repression of the floral regulator FLC (FLOWERING LOCUS C), a paradigmatic Polycombregulated gene, is linked to dividing cells [54,55], indirectly suggesting that DNA replication is a prerequisite for stable maintenance of FLC repression. This finding underscores the observed coupling between PRC2 activity and DNA replication through the participation of DNA polymerases. The molecular mechanisms are not fully delineated but it is clear that mutations in the INCURVATA2 (ICU2) gene, encoding the catalytic subunit of DNA polymerase /, lead to defects in FLC repression and stable maintenance of H3K27me3 level, although PRC2 recruitment and H3K27me3 deposition are normal [56,57]. ESD7 (EARLY IN SHORT DAYS7), which encodes the catalytic subunit of DNA polymerase e, interacts with ICU2 and is also required for correct repression of flowering through epigenetic mechanisms [58]. Finally, mutations in DNA polymerase d also affect flowering but through a different mechanism because the H3K4me3 level of the SEPALLATA3 (SEP3) gene is increased and, as a consequence of its ectopic expression, FLOWERING LOCUS T (FT) transcription is promoted [59]. These data reveal that the three major replicative DNA polymerases that act during S-phase are crucial for genome replication and maintenance of epigenetic marks in genes involved in plant development (Figures 2,3). Furthermore, another component involved in replication fork progression, topoisomerase 1/ (TOP1/), also plays a role in PRC2-mediated repression of WUS, as well as other targets [60]. Whether other genes are also affected and the detailed molecular interactions responsible for linking S-phase, DNA replication fork progression, and the maintenance of chromatin modifications are totally unexplored areas. It is worth noting that recent data obtained in animal cells could illuminate the path to follow [61]. One major component of chromatin organization is the set of histone (canonical and variants) proteins that form each nucleosome. In addition to the canonical forms (H2A, H2B, H3, H4) there are multiple variants (H2A.Z, H2A.W, and H3.3, among others) that play distinct roles in chromatin organization and function [62]. Because in every S-phase only canonical histones are assembled onto newly synthesized DNA (Figure 3), there must be mechanisms coordinating histone incorporation and variant exchange with the division or differentiation status of the cell. Initial results in this direction have been recently obtained by direct determination of the balance
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between canonical H3.1 and variant H3.3 histones. It has been found that cells with a high proliferation rate within the root meristem possess a high H3.1:H3.3 ratio, while cells undergoing their last cell cycle before entering a differentiation pathway show a low ratio owing to a massive eviction of H3.1 in G2 [63]. Moreover, the last cell cycle in the proliferation domain of the root apical meristem is characterized by a longer G2 phase that most likely favors H3.1 eviction. Thus, histone dynamics could be used as a read-out of proliferation status. This is likely to be of general applicability because the H3.1:H3.3 pattern is observed in various cell types and also in endoreplicating cells [63]. Interestingly, chromatin-remodeling factors interact with GRFs [64] that affect the proliferation potential of stem cell derivatives in the root [25]. Therefore, it is tempting to speculate that different transcription factors, for example GRFs and PLT, which impinge on the proliferation potential, may act coordinately with the H3.1 deposition and eviction processes, although direct evidence of such coupling awaits further experimentation. In this context, regeneration is a unique feature of plant explants with potential biotechnological applications because it allows the generation of somatic embryos or new organs, primarily roots and shoots. Reactivation of post-mitotic cells by dedifferentiation and entering into a cell division program is a prerequisite. Ideally, manipulating the activity of cell cycle activators (e.g., cyclins, E2F–DP) and/or repressors (e.g., RBR1, KRPs) could provide a useful framework for improving regeneration efficiency. Other cellular factors, such as chromatin organization, should perhaps be considered as additional players [65].
The Endocycle Many proliferating cell types in diverse plant species arrest the cell cycle and enter the endocycle. This is a special cell cycle in which the genome is duplicated once or several times (endoreplication) without chromosome segregation, leading to polyploid cells. The endocycle program is frequently developmentally and environmentally regulated to initiate various differentiation pathways [66]. The molecular mechanisms controlling the initiation of endoreplication have been delineated and the basic requirements are known [67]. However, the maintenance and arrest of the endocycle program are much less well understood and deserve special attention (Figure 2). Furthermore, many cellular factors participate in both the cell cycle and the endocycle, for example CDK–cyclins, CDK inhibitors, and DNA replication proteins, but their availability and regulation seem to be different in the two cases. Whether the DNA replication origins active during the cell cycle possess the same chromosomal features as those used during the endocycle is not presently known. One interesting hypothesis to test is that the 3D organization of the nucleus and the existence of chromatin domains are relevant for the endocycle process. This is conceivable based on observations of chromatin dynamics in diploid guard cells and polyploid pavement cells in the epidermis of Arabidopsis, revealed using a GFP-mediated chromatin-tagging system, which demonstrated a larger nuclear territory in endoreplicated cells [68].
DNA Damage Response A large amount of DNA modifications occur on a daily basis as a result of both normal metabolism and the effect of environmental challenges to which plants are particularly exposed. These DNA alterations are very harmful to genomic integrity. All cells have evolved mechanisms to cope with DNA damage, and plants are no exception [69]. DNA damage tolerance pathways are frequently linked to modulation of the activity of cell cycle regulators. One of the primary responses is the phosphorylation of SOG1 that transcriptionally regulates the expression of hundreds of DNA damage-response genes, many of which are direct regulators of both the cell cycle and endoreplication [70,71]. The mechanistic details of the SOG1-mediated cell cycle arrest and its coupling with endocycle onset are still far from being fully understood. Another component of the DNA damage response is WEE1 that participates in the intra-S checkpoint [69,72]. In cases where genome damage cannot be repaired, a cell death pathway is triggered,
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but plants have evolved mechanisms to prevent loss of all proliferating stem cells as a result of extensive DNA damage. This relies on QC cells that are more resistant to DNA damage than other cells, and hence they provide a back-up mechanism to restore a stem cell population that can resume organ growth. Importantly, stem cells are actually hypersensitive to DNA damage, which likely contributes to preventing the accumulation of mutations in the stem cell derivatives [73]. Future studies should provide an integrative view of how plants tolerate DNA damage at the organismal level.
Concluding Remarks The examples discussed in previous paragraphs serve to highlight the relevance of cell cycle control as a hub integrating many aspects of plant physiology (Figure 1), although they do not constitute a comprehensive list. In fact, there are other cases that suggest even more unanticipated functional interactions, as illustrated in the following reports: (i) the impact of metabolic balance on stem cell activity directly via E2F and S-phase genes [74]; (ii) the role of reactive oxygen species on the proliferation and differentiation status in the root apical meristem [75] and during lateral root development [76], the coupling of cell cycle to the circadian clock [77]; and (iii) the altered innate immunity response of mutants in the SIM–SMR and cyclin D genes [78,79]. Further studies will be necessary in these and several other directions to fully understand in molecular terms the functional relevance of cell cycle regulators as targets of a plethora of cellular processes (see Outstanding Questions). Cell cycle research, as many other fields, has always benefited from technological advances that allowed exploring questions from a different perspective or at a different level of resolution. One such example is the refinement of image acquisition and analysis techniques that now allow the reconstruction of whole organs in 3D and their analysis over time, adding a new dimension (4D) [80]. These approaches could be combined with appropriate cell cycle markers to analyze in detail cell proliferation dynamics during organogenesis, thus obtaining information at the singlecell level of how they interact with each other. Moreover, the use of computational modeling can allow us to explore how cell proliferation dynamics interacts with patterning rules and developmental cues to establish cellular homeostasis during development [81,82]. Advanced imaging at single-cell resolution can be also potentially very informative, as illustrated by studies of the replication of the specialized structures occurring at telomeres. Single-cell telomere identification and length quantification has demonstrated that different cell lineages, including stem cells, have characteristic telomere lengths [83]. These data show that telomere dynamics during S-phase is intimately coupled to cell proliferation potential and meristem activity. The question remains as to whether other cell cycle features are also cell type-specific. Again a combination of appropriate cell cycle markers with single-cell imaging could be highly informative. Advances in single-cell technologies also have great potential in genomics and proteomics. Thus, single-cell RNA-seq has been applied successfully to plant cells [84,85]. This is not the case yet for chromatin immunoprecipitation (ChIP)-seq, which will be necessary for studies of cell cycle-dependent transcription factor binding and epigenetic studies. Therefore, it is conceivable that different versions of single-cell approaches will be crucial to define molecular events associated with cell cycle progression, arrest, and exit to differentiation, as well as to dedifferentiation (or trans-differentiation) during organ regeneration and wound repair. Because these approaches have an inherently high signal-to-noise ratio [86,87], an important demand for robust bioinformatics tools is increasing. Information derived from single-cell approaches needs to be transferred at the organ level, and ideally to the organismal level [88]. In this context, different growth conditions and adaptations
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Outstanding Questions Do we need to develop more powerful resources to extract novel information relevant for plant cell cycle research? Will we be able to integrate cell cycle regulation with other aspects of plant life? Can synthetic biology take advantage of new integrated knowledge on the cell cycle to generate better-performing plants that contribute to a better life?
have an effect on morphogenesis, for example changes in lateral root density, which are related to specific cell proliferation and differentiation patterns. Therefore, a potential source of valuable information that has been relatively underexploited in cell cycle studies is natural variation [89], and this could facilitate the identification of novel roles of cell cycle regulators in an integrative manner. The analysis of gene function and the acquisition of novel traits is now facilitated by advances in genome editing using CRISPR/Cas9 technology [90,91]. This technology has not been applied yet to cell cycle research on a routine basis but may represent a significant step ahead because, in many cases, functional redundancy is a major obstacle in cell cycle research. In any case, tailored and cell type-specific genome editing would be of great relevance given the essential nature of many cell cycle regulators. Furthermore, because the genome editing technologies are becoming available for crop species [92], it is likely that cell cycle knowledge obtained primarily in Arabidopsis could be useful for plants of commercial interest. Reevaluation with new eyes, concepts, and tools of the role of cell cycle regulators beyond their function as part of the machinery to produce new cells could be extremely enlightening to understand plant growth and development in an integrated manner. Cell cycle regulatory factors are direct targets that control ‘where’ these cells need to be produced (spatial control), ‘when’ during development (temporal control), and ‘what’ type of division is required (symmetrical or asymmetrical, periclinal or anticlinal). The famous statement of Dobzansky comes to mind: ‘Nothing in biology makes sense except in the light of evolution’ [93]. Let me paraphrase it: ‘Nothing in development and growth makes sense except in the light of cell proliferation and differentiation’. Thus, exciting times are ahead because studies in these and other promising directions should lead to discoveries that will widen considerably our perspective in the field beyond the cellular level into the organismal level. Acknowledgments I would like to thank all members of the laboratory for helpful discussions and comments, Z. Vergara for help with the figures, and E. Martinez-Salas for suggestions. This work has been supported by grants BFU2012-34821 and BFU2013-50098 (MINECO) and BFU2015-68396 (MINECO/FEDER), and by an institutional grant from Fundacion Ramon Areces to the Centro de Biologia Molecular ‘Severo Ochoa’.
References 1. Lee, M.G. and Nurse, P. (1987) Complementation used to clone a human homologue of the fission yeast cell cycle control gene cdc2. Nature 327, 31–35 2. John, P.C. et al. (1989) A homolog of the cell cycle control protein p34cdc2 participates in the division cycle of Chlamydomonas, and a similar protein is detectable in higher plants and remote taxa. Plant Cell 1, 1185–1193 3. Feiler, H.S. and Jacobs, T.W. (1990) Cell division in higher plants: a cdc2 gene, its 34-kDa product, and histone H1 kinase activity in pea. Proc. Natl. Acad. Sci. U.S.A. 87, 5397–5401 4. Ferreira, P.C. et al. (1991) The Arabidopsis functional homolog of the p34cdc2 protein kinase. Plant Cell 3, 531–540 5. Hata, S. et al. (1991) Isolation and characterization of cDNA clones for plant cyclins. EMBO J. 10, 2681–2688 6. Hemerly, A. et al. (1992) Genes regulating the plant cell cycle: isolation of a mitotic-like cyclin from Arabidopsis thaliana. Proc. Natl. Acad. Sci. U.S.A. 89, 3295–3299 7. Hirt, H. et al. (1992) Alfalfa cyclins: differential expression during the cell cycle and in plant organs. Plant Cell 4, 1531–1538 8. Van’T Hof, J. and Wilson, G.B. (1962) Studies on the control of mitotic activity: the effect of respiratory inhibitors on mitotic cycle time in the root meristem of Pisum sativum. Chromosoma 13, 39–46 9. Van’t Hof, J. (1965) Cell population kinetics of excised roots of Pisum sativum. J. Cell Biol. 27, 179–189
10. Gimenez-Martin, G. et al. (1965) A new method of labeling cells. J. Cell Biol. 26, 305–309 11. Van’t Hof, J. and Sparrow, A.H. (1965) Radiation effects on the growth rate and cell population kinetics of actively growing and dormant roots of Tradescantia paludosa. J. Cell Biol. 26, 187–199 12. Lopez-Saez, J.F. et al. (1966) Duration of the cell division cycle and its dependence on temperature. Z. Zellforsch. Mikrosk. Anat. 75, 591–600 13. Bryant, J.A. and Wildon, D.C. (1971) Cytoplasmic deoxyribonucleic acid in roots of Pisum sativum. Biochem. J. 121, 5P 14. Soni, R. et al. (1995) A family of cyclin D homologs from plants differentially controlled by growth regulators and containing the conserved retinoblastoma protein interaction motif. Plant Cell 7, 85–103 15. Dahl, M. et al. (1995) The D-type alfalfa cyclin gene cycMs4 complements G1 cyclin-deficient yeast and is induced in the G1 phase of the cell cycle. Plant Cell 7, 1847–1857 16. Xie, Q. et al. (1995) Identification and analysis of a retinoblastoma binding motif in the replication protein of a plant DNA virus: requirement for efficient viral DNA replication. EMBO J. 14, 4073–4082 17. Grafi, G. et al. (1996) A maize cDNA encoding a member of the retinoblastoma protein family: involvement in endoreduplication. Proc. Natl. Acad. Sci. U.S.A. 93, 8962–8967
Trends in Plant Science, October 2016, Vol. 21, No. 10
831
18. Xie, Q. et al. (1996) Plant cells contain a novel member of the retinoblastoma family of growth regulatory proteins. EMBO J. 15, 4900–4908 19. AGI (2000) Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408, 796–815 20. Wildwater, M. et al. (2005) The RETINOBLASTOMA-RELATED gene regulates stem cell maintenance in Arabidopsis roots. Cell 123, 1337–1349 21. Sozzani, R. et al. (2010) Spatiotemporal regulation of cell-cycle genes by SHORTROOT links patterning and growth. Nature 466, 128–132 22. Cruz-Ramirez, A. et al. (2012) A bistable circuit involving SCARECROW-RETINOBLASTOMA integrates cues to inform asymmetric stem cell division. Cell 150, 1002–1015 23. Weimer, A.K. et al. (2012) Retinoblastoma related1 regulates asymmetric cell divisions in Arabidopsis. Plant Cell 24, 4083–4095 24. Kinoshita, A. et al. (2015) A plant U-box protein, PUB4, regulates asymmetric cell division and cell proliferation in the root meristem. Development 142, 444–453 25. Rodriguez, R.E. et al. (2015) MicroRNA miR396 regulates the switch between stem cells and transit-amplifying cells in Arabidopsis roots. Plant Cell 27, 3354–3366 26. Van Damme, D. et al. (2011) Arabidopsis alpha Aurora kinases function in formative cell division plane orientation. Plant Cell 23, 4013–4024 27. Weimer, A.K. et al. (2016) Aurora kinases throughout plant development. Trends Plant Sci. 21, 69–79 28. Petrovska, B. et al. (2012) Plant Aurora kinases play a role in maintenance of primary meristems and control of endoreduplication. New Phytol. 193, 590–604 29. Yang, K. et al. (2014) Requirement for A-type cyclin-dependent kinase and cyclins for the terminal division in the stomatal lineage of Arabidopsis. J. Exp. Bot. 65, 2449–2461 30. De Smet, I. et al. (2008) Receptor-like kinase ACR4 restricts formative cell divisions in the Arabidopsis root. Science 322, 594–597 31. Yue, K. et al. (2016) PP2A-3 interacts with ACR4 and regulates formative cell division in the Arabidopsis root. Proc. Natl. Acad. Sci. U.S.A. 113, 1447–1452
44. Schommer, C. et al. (2014) Repression of cell proliferation by miR319-regulated TCP4. Mol. Plant 7, 1533–1544 45. Kobayashi, K. et al. (2015) Transcriptional repression by MYB3R proteins regulates plant organ growth. EMBO J. 34, 1992–2007 46. Jegu, T. et al. (2015) A SWI/SNF chromatin remodelling protein controls cytokinin production through the regulation of chromatin architecture. PLoS One 10, e0138276 47. Sequeira-Mendes, J. and Gutierrez, C. (2015) Links between genome replication and chromatin landscapes. Plant J. 83, 38–51 48. Costas, C. et al. (2011) Genome-wide mapping of Arabidopsis origins of DNA replication and their associated epigenetic marks. Nat. Struct. Mol. Biol. 18, 395–400 49. Bass, H.W. et al. (2015) Defining multiple, distinct, and shared spatiotemporal patterns of DNA replication and endoreduplication from 3D image analysis of developing maize (Zea mays L.) root tip nuclei. Plant Mol. Biol. 89, 339–351 50. Kuwabara, A. and Gruissem, W. (2014) Arabidopsis retinoblastoma-related and Polycomb group proteins: cooperation during plant cell differentiation and development. J. Exp. Bot. 65, 2667– 2676 51. Xiao, J. and Wagner, D. (2015) Polycomb repression in the regulation of growth and development in Arabidopsis. Curr. Opin. Plant Biol. 23, 15–24 52. Ikeuchi, M. et al. (2015) PRC2 represses dedifferentiation of mature somatic cells in Arabidopsis. Nat. Plants 1, 15089 53. Desvoyes, B. et al. (2006) Cell type-specific role of the retinoblastoma/E2F pathway during Arabidopsis leaf development. Plant Physiol. 140, 67–80 54. Finnegan, E.J. and Dennis, E.S. (2007) Vernalization-induced trimethylation of histone H3 lysine 27 at FLC is not maintained in mitotically quiescent cells. Curr. Biol. 17, 1978–1983 55. Berry, S. et al. (2015) Local chromatin environment of a Polycomb target gene instructs its own epigenetic inheritance. Elife 4, e07205 56. Barrero, J.M. et al. (2007) INCURVATA2 encodes the catalytic subunit of DNA polymerase alpha and interacts with genes involved in chromatin-mediated cellular memory in Arabidopsis thaliana. Plant Cell 19, 2822–2838
32. Roeder, A.H. et al. (2012) Cell cycle regulates cell type in the Arabidopsis sepal. Development 139, 4416–4427
57. Hyun, Y. et al. (2013) The catalytic subunit of Arabidopsis DNA polymerase alpha ensures stable maintenance of histone modification. Development 140, 156–166
33. Harashima, H. and Sugimoto, K. (2016) Integration of developmental and environmental signals into cell proliferation and differentiation through RETINOBLASTOMA-RELATED 1. Curr. Opin. Plant Biol. 29, 95–103
58. del Olmo, I. et al. (2010) EARLY IN SHORT DAYS 7 (ESD7) encodes the catalytic subunit of DNA polymerase epsilon and is required for flowering repression through a mechanism involving epigenetic gene silencing. Plant J. 61, 623–636
34. Perilli, S. et al. (2013) RETINOBLASTOMA-RELATED protein stimulates cell differentiation in the Arabidopsis root meristem by interacting with cytokinin signaling. Plant Cell 25, 4469–4478
59. Iglesias, F.M. et al. (2015) The Arabidopsis DNA polymerase delta has a role in the deposition of transcriptionally active epigenetic marks, development and flowering. PLoS Genet. 11, e1004975
35. Heyman, J. et al. (2013) ERF115 controls root quiescent center cell division and stem cell replenishment. Science 342, 860–863 36. Cruz-Ramirez, A. et al. (2013) A SCARECROW-RETINOBLASTOMA protein network controls protective quiescence in the Arabidopsis root stem cell organizer. PLoS Biol. 11, e1001724 37. Vilarrasa-Blasi, J. et al. (2014) Regulation of plant stem cell quiescence by a brassinosteroid signaling module. Dev. Cell 30, 36–47 38. Forzani, C. et al. (2014) WOX5 suppresses CYCLIN D activity to establish quiescence at the center of the root stem cell niche. Curr. Biol. 24, 1939–1944 39. Matos, J.L. et al. (2014) Irreversible fate commitment in the Arabidopsis stomatal lineage requires a FAMA and RETINOBLASTOMA-RELATED module. Elife 3, e03271 40. Nowack, M.K. et al. (2012) Genetic framework of cyclin-dependent kinase function in Arabidopsis. Dev. Cell 22, 1030–1040
60. Liu, X. et al. (2014) DNA topoisomerase I affects polycomb group protein-mediated epigenetic regulation and plant development by altering nucleosome distribution in Arabidopsis. Plant Cell 26, 2803–2817 61. Alabert, C. et al. (2015) Two distinct modes for propagation of histone PTMs across the cell cycle. Genes Dev. 29, 585–590 62. Talbert, P.B. and Henikoff, S. (2013) Phylogeny as the basis for naming histones. Trends Genet. 29, 499–500 63. Otero, S. et al. (2016) Histone H3 dynamics uncovers domains with distinct proliferation potential in the Arabidopsis root. Plant Cell http://dx.doi.org/10.1105/tpc.15.01003 64. Debernardi, J.M. et al. (2014) Post-transcriptional control of GRF transcription factors by microRNA miR396 and GIF co-activator affects leaf size and longevity. Plant J. 79, 413–426
41. Desvoyes, B. et al. (2014) Looking at plant cell cycle from the chromatin window. Front. Plant Sci. 5, 369
65. Ikeuchi, M. et al. (2015) Control of plant cell differentiation by histone modification and DNA methylation. Curr. Opin. Plant Biol. 28, 60–67
42. Henriques, R. et al. (2010) Arabidopsis S6 kinase mutants display chromosome instability and altered RBR1-E2F pathway activity. EMBO J. 29, 2979–2993
66. Polyn, S. et al. (2015) Cell cycle entry, maintenance, and exit during plant development. Curr. Opin. Plant Biol. 23, 1–7
43. Magyar, Z. et al. (2012) Arabidopsis E2FA stimulates proliferation and endocycle separately through RBR-bound and RBR-free complexes. EMBO J. 31, 1480–1493
832
Trends in Plant Science, October 2016, Vol. 21, No. 10
67. Edgar, B.A. et al. (2014) Endocycles: a recurrent evolutionary innovation for post-mitotic cell growth. Nat. Rev. Mol. Cell Biol. 15, 197–210
68. Kato, N. and Lam, E. (2003) Chromatin of endoreduplicated pavement cells has greater range of movement than that of diploid guard cells in Arabidopsis thaliana. J. Cell Sci. 116, 2195–2201 69. Hu, Z. et al. (2016) Mechanisms used by plants to cope with DNA damage. Ann. Rev. Plant Biol. 67, 439–462 70. Yoshiyama, K. et al. (2009) Suppressor of gamma response 1 (SOG1) encodes a putative transcription factor governing multiple responses to DNA damage. Proc. Natl. Acad. Sci. U.S.A. 106, 12843–12848 71. Yoshiyama, K.O. et al. (2013) ATM-mediated phosphorylation of SOG1 is essential for the DNA damage response in Arabidopsis. EMBO Rep. 14, 817–822 72. Cools, T. et al. (2011) The Arabidopsis thaliana checkpoint kinase WEE1 protects against premature vascular differentiation during replication stress. Plant Cell 23, 1435–1448 73. Fulcher, N. and Sablowski, R. (2009) Hypersensitivity to DNA damage in plant stem cell niches. Proc. Natl. Acad. Sci. U.S.A. 106, 20984–20988 74. Xiong, Y. et al. (2013) Glucose–TOR signalling reprograms the transcriptome and activates meristems. Nature 496, 181–186 75. Tsukagoshi, H. et al. (2010) Transcriptional regulation of ROS controls transition from proliferation to differentiation in the root. Cell 143, 606–616 76. Manzano, C. et al. (2014) The emerging role of reactive oxygen species signaling during lateral root development. Plant Physiol. 165, 1105–1119 77. Miyagishima, S.Y. et al. (2014) Translation-independent circadian control of the cell cycle in a unicellular photosynthetic eukaryote. Nat. Commun. 5, 3807 78. Hamdoun, S. et al. (2016) Differential roles of two homologous cyclin-dependent kinase inhibitor genes in regulating cell cycle and innate immunity in Arabidopsis. Plant Physiol. 170, 515–527 79. Eichmann, R. and Schafer, P. (2015) Growth versus immunity – a redirection of the cell cycle? Curr. Opin. Plant Biol. 26, 106–112
80. Bassel, G.W. and Smith, R.S. (2016) Quantifying morphogenesis in plants in 4D. Curr. Opin. Plant Biol. 29, 87–94 81. Ortiz-Gutierrez, E. et al. (2015) A dynamic gene regulatory network model that recovers the cyclic behavior of Arabidopsis thaliana cell cycle. PLoS Comput. Biol. 11, e1004486 82. von Wangenheim, D. et al. (2016) Rules and self-organizing properties of post-embryonic plant organ cell division patterns. Curr. Biol. 26, 439–449 83. Gonzalez-Garcia, M.P. et al. (2015) Single-cell telomere-length quantification couples telomere length to meristem activity and stem cell development in Arabidopsis. Cell Rep. 11, 977–989 84. Efroni, I. and Birnbaum, K.D. (2016) The potential of single-cell profiling in plants. Genome Biol. 17, 65 85. Efroni, I. et al. (2015) Quantification of cell identity from single-cell gene expression profiles. Genome Biol. 16, 9 86. Buettner, F. et al. (2015) Computational analysis of cell-to-cell heterogeneity in single-cell RNA-sequencing data reveals hidden subpopulations of cells. Nat. Biotechnol. 33, 155–160 87. McDavid, A. et al. (2016) The contribution of cell cycle to heterogeneity in single-cell RNA-seq data. Nat. Biotechnol. 34, 591–593 88. Chaiwanon, J. et al. (2016) Information integration and communication in plant growth regulation. Cell 164, 1257–1268 89. Zhu, W. et al. (2015) Natural variation identifies ICARUS1, a universal gene required for cell proliferation and growth at high temperatures in Arabidopsis thaliana. PLoS Genet. 11, e1005085 90. Ding, Y. et al. (2016) Recent advances in genome editing using CRISPR/Cas9. Front. Plant Sci. 7, 703 91. Puchta, H. (2016) Genome engineering using CRISPR/Cas: getting more versatile and more precise at the same time. Genome Biol. 17, 51 92. Khatodia, S. et al. (2015) The CRISPR/Cas genome-editing tool: application in improvement of crops. Front. Plant Sci. 7, 506 93. Dobzhansky, T. (1973) Nothing in biology makes sense except in the light of evolution. Am. Biol. Teacher 35, 125–129
Trends in Plant Science, October 2016, Vol. 21, No. 10
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