Flowering time plasticity and global adaptation in rice

by Aalt DJ van Dijk 
Wageningen University

Both genetic and physical characteristics of plants are nowadays studied at large scale using automated approaches. In order to understand how genotype and phenotype are related, computational approaches are indispensable. An important caveat of such analyses is that to obtain a meaningful connection between genotype and phenotype, the environment has to be taken into account. This certainly holds for flowering time related traits, given the strong impact of environmental signals on flowering. Hence, it is of major importance to consider the interplay between genes, phenotypes, and the environment when modelling the connection between genotype and flowering related traits.

This is both of fundamental interest and of practical relevance. Plant breeding has been very successful in creating and selecting genotypes that are well-adapted. Plant adaptation depends on the genotype, the environment, and on how sensitive the genotype is to the environmental conditions. This effect is known under the name of ‘genotype by environment interaction’: genotypes showing different sensitivities to the environment. This genotype by environment interaction can cause that the best genotype for one condition might not be the best for another condition, complicating the selection of superior individuals.

A paper by Guo et al. (2020) presents a study of flowering time adaptation to different temperature zones. In doing so, they provide an example of how to integrate data from genomics, phenotype measurements, and quantification of the environment. Guo et al. studied a rice genetic mapping population in nine different natural environments across Asia. To characterize the different environment, a so-called environmental index was defined. To that end, a measure called ‘growing degree days (GDD) from 9 to 50 days after planting’ was chosen. This basically represents the amount of heat accumulation in a certain timeframe (here, day 9-50). This specific index was chosen based on how well it correlated with the mean flowering time in the different environments. (see Fig. 1, left panel).


Fig. 1. Integrating genotype, environment, and flowering time. (Left) Environmental index: Growing degree days (GDD) is used as an index to characterize the environment. (Right) Joint genomic regression analysis: Regression of flowering time on GDD for each genotype results in intercept and slope parameter for each genotype. This is visualized for three out of 174 genotypes. These parameters were subsequently linked to genomic markers.

The idea of this index is that in subsequent analyses, each environment can be represented by a single number. This allows to predict flowering time between different environments, in a similar way as we can predict between different genotypes using genotypic information. These two sources of information (genotype and environment) were then combined in a so-called ‘joint genomic regression analysis’. This was performed in two scenarios, of which I will only describe one (see Fig. 1, right panel); the alternative approach basically contains the same two steps but in a different order. (1) In the first step of this analysis, flowering-time observations for each genotype were regressed on values of the environmental index (GDD). For each genotype, this results in two numbers. The first is an intercept, which quantifies the expected flowering time when GDD equals zero, i.e. without temperature effect. The other is the slope of the regression line, quantifying the sensitivity of flowering time to temperature change; in other words, how much does flowering time change when GDD is increased by a given amount. (2) In the second step of the analysis, the environment-related parameters obtained for each genotype in step 1 (intercept and slope) were now linked to the genotype. A model was fitted to predict these parameters using genotypic markers. The result is a model that can predict for unseen genotypes and/or unseen environments what the flowering time would be. In other words, this is a model that includes rice flowering time plasticity at the genotype level.

Subsequently, to zoom in on specific genes potentially involved in flowering time plasticity, Guo et al. (2020) focused in detail on four known rice flowering time genes: Hd1, Hd2, Hd5, and Hd6. Part of the motivation to focus on these came from a QTL mapping experiment which I will not describe here in detail. In choosing those genes, Guo and colleagues made use of the fact that the molecular mechanisms underlying the timing of the transition from vegetative to reproductive growth have been well-studied in rice. This allowed to select these four genes, known for their role in photoperiod response, as putative causal genes underlying QTLs detected in this study. In the mapping population, for each of the different haplotype combinations for those four genes, slopes of a model regressing flowering time to GDD were obtained, in a similar way as described above.

Next, the results were placed in a genomic context by analyzing the roughly 3000 available genomes from cultivated rice. From these, haplotypes were obtained and designated as wild type and non-wildtype. This allowed to categorize the haplotype combinations in 16 groups (2*2*2*2). The question then was if and how geographic distribution of these haplotypes would correspond with temperature response behaviour. This question was addressed by looking at the slopes obtained for each haplotype combination in the biparental population. It turned out that regions with lower mean annual temperature were dominated by haplotypes sensitive to temperature (large slope) and regions with higher temperature mostly had haplotypes less sensitive to temperature (smaller slope).

The research by Guo and colleagues allows to conceptualize the relationship between environment, flowering time, and either genotype or haplotype. It will be exciting to see further developments in this area. In particular, one could imagine that a next step could be to expand the representation of the genotype by taking into account known molecular modes of action. This could, for example, involve knowledge on how the different genes included are involved in a network of interactions. This would potentially allow further improvement in prediction performance but in particular would serve to enable further interpretation of the relationship between genes, environment, and flowering.


Guo T, Mu Q, Wang J, Vanous A, Onogi A, Iwata H, Li X and Yu J. 2020. Dynamic effects of interacting genes underlying rice flowering-time phenotypic plasticity and global adaptation. Genome Research 30, 673-68310. doi: 1101/gr.255703.119


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Turing’s reaction-diffusion mechanism explains dispersed petal spots in monkeyflowers

by Paula Elomaa
Department of Agricultural Sciences, Viikki Plant Science Centre, University of Helsinki, Finland

Colorful pigmentation patterns in plants – spots, blotches, stripes, or colored veins – are particularly fascinating. Anthocyanin pigments are synthesized via the well-known flavonoid pathway, and they form patterns that, especially in flowers, are associated with spatial developmental signals but may also be activated by environmental cues. Complex pigmentation patterns, including those not visible for human eye, have also evolved in response to specialized pollination strategies and contributed to speciation.

Studies in diverse model systems, including maize, petunia, snapdragon and Arabidopsis show that the anthocyanin regulators are highly conserved, and include MYB, bHLH, and WD40-repeat proteins functioning in a so-called MBW complex that activates the biosynthetic genes (Feller et al., 2011; Xu et al., 2015). In this complex, tissue specifically expressed R2R3-MYB proteins primarily define spatial pigmentation patterns. Anthocyanin regulation also involves repressor proteins. As shown in petunia, the EAR-domain containing R2R3-MYBs may function as a part of the DNA-binding MBW complex, but converts it from an activating into a repressing complex (Albert et al., 2014). Moreover, small R3-MYB proteins, with only a single MYB repeat, have been identified as repressors. They do not bind DNA but interact with bHLH activators, consequently sequestering them from the MBW complexes. R3-MYBs are suggested to provide feedback regulation that limits anthocyanin accumulation and affects pigmentation patterning (Albert et al., 2011, 2014).

The paper by Ding et al. (2020) focuses on anthocyanin spots in monkeyflowers (Mimulus) that dot the nectar guide of the ventral petal (Fig. 1A). These spots form in randomly dispersed patterns raising a question of how they emerge. Ding et al. (2020) refer to the classical reaction-diffusion (RD) model of patterning, originally proposed by Turing (1952) and Gierer and Meinhardt (1972). The model states that patterns in various biological systems arise autonomously through a network of reacting factors that combines a short-range positive feedback with a long-range negative feedback (reviewed in Kondo and Miura, 2010). In its most simple form, the model may include only two components; an activator that controls its own synthesis as well as the synthesis of a repressor that is able to diffuse or move to neighboring cells and, in turn, to inhibit the activator function. To test whether this model holds for spot formation in Mimulus, Ding et al. (2020) functionally validated the interactions of two MYB domain factors; a known, self-activating R2R3-type anthocyanin activator NECTAR GUIDE ANTHOCYANIN (NEGAN) and a newly identified R3-type inhibitor RED TONGUE (RTO).

The rto mutant with uniform red pigmentation in the nectar guide, was identified through EMS mutagenesis screen in M. lewisii. By independent mapping experiments, the authors identified the causal gene as an R3-MYB, and verified that mutations in the same gene were responsible for the naturally occurring rto phenotypes of M. guttatus found in Oregon and southern California (Fig. 1B). The repressor function of RTO was confirmed in transgenic Mimulus lines. Following the principle of the RD model, NEGAN, in a protein complex with bHLH and WD40 components, was shown to activate RTO expression in the nectar guides in both Mimulus species. On the other hand, analysis of the rto mutant, rto-like natural variants and RTO transgenic lines indicated that RTO in turn inhibits NEGAN. Repression was shown to occur through interaction with bHLH proteins sequestering them into inactive complexes.

Fig. 1. Dispersed anthocyanin spots in monkeyflowers. (A) Spots in Mimulus lewisii dot the nectar guide of the ventral petal. Photo courtesy of Jouko Rikkinen, Univ. of Helsinki, Finland. (B) Segregating variation at the RTO locus in a wild Mimulus guttatus population at the UC Davis McLaughlin Reserve. Uniform red pigmentation in nectar guide region is observed in rto mutants. Photo courtesy of Benjamin Blackman, Univ. of California, USA.

The RD model also assumes long-range movement of a repressor. Using fluorescent marker lines, Ding et al. (2020) showed that the RTO gene was transcribed in the pigmented spots but the RTO protein was localized in a broader domain indicating that it is moving from the source cell to the neighboring cells. Mobility of R3-MYBs has also previously been shown for the petunia MYBx (Albert et al., 2014). The observed regulatory interactions, and the movement of the repressor support a simple two-component RD model. The authors applied computer simulations to test patterning dynamics. Using diverse parameter values for degradation rate of the activator NEGAN as well as production and degradation rates of the inhibitor RTO, they were able to recapitulate the phenotypes observed  both in wild-type and transgenic lines. Finally, the authors showed that the rto alleles are retained in natural populations of M. guttatus. Using controlled laboratory experiments, they showed that naïve bumblebees preferred both the homozygote and heterozygote rto flowers over wild-type suggesting that the patterning impacts plant-pollinator interactions.

The paper by Ding et al. (2020) provides a real-life example of Turing’s mechanism operating in pattern formation in plants. The presented experimental data combined with computer simulations elegantly demonstrates the extremely fine-tuned interaction dynamics between the activator and the repressor molecules leading to dispersed anthocyanin spot patterns. It will be interesting to see how widely the two-component RD model underlies pigmentation patterns observed in diverse plant lineages, and how it is linked with spatial regulation of pigmentation, upstream of MYBs.


Albert NW, Lewis DH, Zhang H, Schwinn KE, Jameson PE, Davies KM. 2011. Members of an R2R3-MYB transcription factor family in Petunia are developmentally and environmentally regulated to control complex floral and vegetative pigmentation patterning. The Plant Journal 65, 771-784. https://doi.org/10.1111/j.1365-313X.2010.04465.x

Albert NW, Davies KM, Lewis DH, Zhang H, Montefiori M, Brendolise C, Boase MR, Ngo H, Jameson PE, Schwinn KE. 2014. A conserved network of transcriptional activators and repressors regulates anthocyanin pigmentation in eudicots. The Plant Cell 26, 962-980. https://doi.org/10.1105/tpc.113.122069

Ding B, Patterson EL, Holalu SV, Li J, Johnson GA, Stanley LE, Greenlee AB, Peng F, Bradshaw Jr. HD, Blinov ML, Blackman BK, Yuan Y-W. 2020. Two MYB proteins in a self-organizing activator-inhibitor system produce spotted pigmentation patterns. Current Biology 30, 1-13. https://doi.org/10.1016/j.cub.2019.12.067

Feller A, Machemer K, Braun EL, Grotewold E. 2011. Evolutionary and comparative analysis of MYB and bHLH transcription factors. The Plant Journal 66, 94-116. https://doi.org/10.1111/j.1365-313X.2010.04459.x

Gierer A, Meinhardt H. 1972. A theory of biological pattern formation. Kybernetik 12, 30-39. https://doi.org/10.1007/BF00289234

Kondo S, Miura T. 2010. Reaction-diffusion model as framework for understanding biological pattern formation. Science 329, 616-620. DOI: 10.1126/science.1179047

Turing AM. 1952. The chemical basis of morphogenesis. Philosophical Transactions of the Royal  Society of London. Series B, Biological Sciences 237, 37-72. https://doi.org/10.1098/rstb.1952.0012

Xu W, Dubos C, Lepinic L. 2015. Transcriptional control of flavonoid biosynthesis by MYB-bHLH-WDR complexes. Trends in Plant Science 20, 176-185.https://doi.org/10.1016/j.tplants.2014.12.001

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Flowering plants return to the sea…

by Charles P Scutt

Laboratoire de Reproduction et Développement des Plantes, Université de Lyon, ENS de Lyon, UCB Lyon-1, CNRS, INRA, F-69342 Lyon, France

From 18 to 22 June this year, around 120 fans of floral biology met at Hyères-les-Palmiers on the French Côte d’Azur for the latest in a series of two-yearly workshops on ‘Molecular Mechanisms Controlling Flower Development’. These international meetings usually take place somewhere on the Mediterranean coast, and Hyères provided an idyllic setting in which to meet up with colleagues and discuss science on the beach as well as in the lecture hall or poster room. One flowering plant that really did return to the sea, Posidonia oceanica, was to be found growing in the shallow coastal waters surrounding the conference venue, and many delegates could be seen splashing about and (no doubt) thoroughly investigating this marine monocot between scientific sessions.

The workshop, of which details can be found at: http://www.ens-lyon.fr/RDP/FlowerWorkshop2019/, was divided into seven oral sessions and formatted so that a maximum number of speakers could present their data in 20 min highlighted talks, 15 min standard talks, or 4 min ‘flash posters’. In all, 74 oral presentations were made, involving nearly two-thirds of delegates, who came mostly from Europe, but also from as far afield as China, Japan, Vietnam, Mexico, and Washington State. It is impossible in a short review to cover all the subjects addressed at the meeting, so the following is a personal view of some of the highlights and novel themes to emerge.

The workshop got underway with its traditional starting topic of ‘Flowering’, namely the complex and interconnecting networks used by flowering plants to initiate their reproductive phase in response to numerous environmental and endogenous cues. Much of what is known about flowering comes from the model plant Arabidopsis thaliana, which is a typical annual species that flowers once in its life cycle, promptly sets seed, and dies. It has been known for several years that multiple cues for flowering in Arabidopsis converge on the FLOWERING TIME (FT) protein, which moves from its site of synthesis in the leaves to the shoot apex where flower production is initiated. However, two recent research trends were evident from this year’s workshop. First, several mobile signals other than FT are coming to the fore, including a potentially large list of metabolites that can promote flowering, as highlighted in a talk by Reyes Benlloch (IBMCP, Valencia, Spain). Secondly, several research groups are now focusing on alternative models with contrasting life cycles and flowering requirements. For example, the work of George Coupland’s laboratory (MPI-Cologne, Germany) demonstrates that endogenous cues that depend on the age of the plant are of particular importance to flowering in perennial Brassicaceae (Hyun et al., 2019). Indeed, although the workshop included a devoted ‘evo-devo’ session, the prominent presence of new and emerging models (Fig.1) in all seven scientific sessions meant that evolutionary considerations were discussed throughout the meeting.

fig1 CS JXB

Fig. 1. Some of the many plant models featured in ‘Molecular Mechanisms Controlling Flower Development, 2019’. Clockwise from bottom left: Chara braunii (image: G. Theißen), Oryza sativa (Asian cultivated rice; image: H. Adam), Erycina pusilla (image: B. Gravendeel), Petunia hybrida (image: M. Vandenbussche), Rosa chinensis ‘Old Blush’ (image: M. Bendahmane), Arabidopsis thaliana (image: C. Scutt), Thalictrum thalictroides (image: V. Di Stilio), Trithuria submersa (image: C. Scutt), and Amborella trichopoda (image: C. Scutt). The ANA grade comprises the three most basally diverging extant angiosperm lineages of Amborellales (containing only A. trichopoda), Nymphaeales (water lilies and their allies, including T. submersa), and Austrobaileyales.

Many species require vernalization: cold treatment as a necessary prelude to flowering. However, the need for vernalization can cause unwanted delays in plant breeding programs, which is why Richard Immink’s group (Wageningen University, The Netherlands) recently made a screen of some 9000 chemical structures to find a compound that could bypass the vernalization requirement (Fiers et al., 2017). Two compounds were found to have the desired effect in Arabidopsis, one of which might prove of practical use in crop species. Interestingly, this compound did not appear to act via any of the known dedicated components of the vernalization pathway, but instead induced FT by a novel mechanism, possibly involving the production of hydrogen peroxide.

After deciding to flower, a plant must then determine how its flowers should be arranged, and so the workshop turned its attention to the subject of inflorescence architecture. The enlargement of model species from Arabidopsis was clearly apparent in this theme too, with several impressive talks featuring cereals, legumes, Solanaceae, and Asteraceae. One particularly interesting model for inflorescence architecture is cultivated rice, which has undergone the domestication process twice from distinct wild species in Asia and Africa. These domestication events appear to have resulted in parallel changes to the rice panicle, and a collaboration involving Stefan Jouannic and Helene Adam (IRD-Montpellier, France) and Ngang Giang Khong (Agricultural Genetic Institute, Hanoi, Vietnam) is taking a multidisciplinary approach to discover whether similar genetic targets and developmental processes were affected in these two events. Working on another cereal crop, Scot Boden (John Innes Centre, Norwich, UK) has revealed a previously unsuspected role for an amino acid transporter in wheat spikelet formation, suggesting that small metabolites may influence inflorescence architecture as they do flowering. Inflorescences in Asteraceae are showy, flower-like platforms consisting of many small flowers that develop in both left- and right-turning spirals, often defined by pairs of quite large Fibonacci numbers (Elomaa et al., 2018). Using a combination of reporter lines, imagery, and modeling, Teng Zhang (Helsinki University, Finland) elegantly showed how growth and hormone dynamics contribute to generate these patterns in the inflorescence meristem of Gerbera hybrida.

Moving on from inflorescence architecture, a major part of the workshop was devoted to a detailed discussion of flower and fruit development. Several talks and associated posters addressed the central question of how quaternary complexes of MADS-box transcription factors, also known as floral quartets, act on specific subsets of direct target genes to control floral patterning and organ identity. Three major ideas came to the fore, all of which may be involved in this process. Kerstin Kaufmann (Humboldt University, Berlin, Germany) emphasized the importance of the width of the minor DNA groove in the CArG box motifs to which MADS proteins are known to bind, and which can vary between motifs. Cezary Smaczniak from the same research group talked about the role of complexes formed between MADS proteins and numerous other classes of transcription factors and chromatin remodeling enzymes, which can also affect the specificity and the positive or negative transcriptional effects of the complexes formed. Chloe Zubieta (CEA-Grenoble, France) has been addressing the specificity of MADS complexes mainly using a structural biology approach. Her work indicates that differential flexibility occurs within quaternary MADS complexes of different compositions. As these complexes are hypothesized to bind simultaneously and cooperatively to two CArG boxes, the spacing between adjacent motifs may also be a determining factor in the specificity of the interaction.

An interesting presentation in a further session of the workshop, given by Günter Theißen (Friedrich Schiller University, Jena, Germany), returned to the subject of quaternary MADS complexes, but took an evo-devo approach that might also provide insights into their mechanism of action. By analysing homologs of classic MIKC-class MADS proteins from early-diverging land plants and their charophyte green algal relatives (Nishiyama et al., 2018), he showed that quaternary binding behavior evolved following the duplication of an exonic segment of an ancestral MIKC-class gene in a common ancestor of land plants. It seems therefore that earlier MADS complexes bound only as dimers to single CArG boxes, and so any influence of motif spacing on the specificity and cooperativity of the DNA–protein interaction would presumably have been a later mechanistic addition.

Many other presentations in the meeting focused on the precise roles of MADS-box factors in different systems. These included both well-developed core eudicot models such as Petunia, tomato, and Cardamine, and basal eudicots in which some methods of functional genetic analysis are becoming available, as explained by Verónica Di Stilio (Washington University, Seattle, USA), who has used virus-induced gene silencing and horticultural mutants to uncover processes of neo- and subfunctionalization in the evolution of MADS-box genes in the ranunculid Thalictrum thalictroides (Galimba et al., 2018). Among the talks featuring core eudicot models, Michiel Vandenbussche (ENS-Lyon, France) gave a thorough analysis of the SEPALLATA (SEP) and APETALA1 (AP1) MADS-box subfamilies in Petunia, showing that the roles of these genes in organ identity and floral meristem determination have evolved along substantially different lines in the asterids compared with the rosids (Morel et al., 2017). A further impressive feat of functional genetics was presented by Vojtech Hudzieczek (Institute of Biophysics, Brno, Czech Republic), who has developed gene editing in the dioecious angiosperm Silene latifolia and used this to demonstrate a role in male fertility for a Y chromosome-linked AP3-class MADS-box gene. This technical breakthrough might pave the way to solving long-standing questions on the mechanism of sex determination in Silene. A further talk in this section by Silvia Moschin (Padua University, Italy) explored the presence and expression of MADS-box floral homeotic genes in the basal angiosperm Trithuria submersa, a novel experimental system for which functional genetic methods have yet to be developed.

Numerous speakers focused on developmental processes within specific floral organs that rely on classes of regulators other than MADS-box proteins. These included an analysis by Stefan de Folter (LANGEBIO-Irapuato, Mexico) of crosstalk between the basic helix–loop–helix transcription factor SPATULA and hormone signaling in the Arabidopsis gynoecium (Reyes-Olalde et al., 2017), and a transcriptomics-based analysis of the evolution of multiple families of transcriptional regulators in the gynoecium by Annette Becker (Justus-Liebig University, Giessen, Germany). Maura Cardarelli’s laboratory (Sapienza University, Rome, Italy) provided two interesting talks on Arabidopsis stamen development, one of which showed how this process is coordinated by alternative transcripts of the transcription factor gene AUXIN RESPONSE FACTOR8, while the other, given by Davide Marzi, demonstrated a developmental effect of light quality. Again moving away from classical models, Barabara Gravendeel (Leiden University, The Netherlands) gave a fascinating talk on specialized floral organs that facilitate pollen transfer in orchids. Protein–protein interaction and gene expression data suggest that several transcription factors whose orthologs function in the development of the dehiscence zone in Arabidopsis may contribute to the formation of these unique floral structures in Orchidaceae (Dirks-Mulder et al., 20172019). At a higher level of regulation, flower development is ultimately coordinated by the master transcriptional regulator LEAFY (LFY), and Leonie Verhage (CEA-Grenoble, France) delved way back in evolutionary time to show how LFY’s DNA-binding behavior became more uniform before the radiation of extant vascular plants. Bifunctional LFY proteins, possessing an alternative binding behavior, are still present in some bryophyte (sensu lato) and green algal lineages.

Aspects of flower development revealed through genomic-scale approaches were particularly featured in the workshop. For example, a pair of talks by Jeremy Just and Lea Francois (both of ENS-Lyon, France), respectively, detailed the production of a high-quality assembly of the rose genome (Raymond et al., 2018) and the use of this resource to identify the gene behind the double-flowered phenotype in modern rose cultivars (François et al., 2018): an miR172-resistant variant of an AP2-family transcription factor. François Parcy (CEA-Grenoble) presented novel methods to combine large in vivo and in vitro data sets with the objective of determining the rules that govern transcription factor binding to DNA, using the master regulator LFY as an example.

Another fascinating session of the workshop looked at functional aspects of the reproductive system in flowering plants. Lucia Colombo (Milan University, Italy) focused on a female-expressed histone deacetylase in Arabidopsis that is required for pollen tube guidance, while Thomas Dresselhaus (Regensburg University, Germany) gave a wide-ranging presentation on the multiple defense-related pathways that operate along the pollen tube’s journey from the stigma to the micropyle in both grasses and Arabidopsis (Zhou and Dresselhaus, 2019). From the same university, Stefanie Sprünck provided evidence for the widespread conservation of multiple mechanisms that regulate fertilization, even between Arabidopsis and the probable sister to all other living flowering plants, Amborella trichopoda. This work involved the impressive characterization of the egg apparatus transcriptome in Amborella (Flores-Tornero et al., 2019). Finally in this section, Daphné Autran (IRD-Montpellier, France) described sophisticated imaging, modeling, and experimental approaches to investigate spatial relationships in the Arabidopsis ovule during megaspore mother cell formation.

The effect of music on plants remains controversial, but its effect on plant scientists was clearly apparent at the workshop. This year’s event coincided with the annual Fête de la Musique in France, and a Balkan sextet of immense talent travelled from the town of Sète to help us close the meeting in style. Delegates made their way home the following day, safe in the knowledge that the next Flower Development Workshop will be organized by Cristina Ferrandiz, Paco Madueño, and colleagues in Spain in 2021. Hope to see you there!


Dirks-Mulder A, Ahmed I, Broek MUH, et al. 2019. Morphological and molecular characterization of orchid fruit development. Frontiers in Plant Science 10, 137. https://doi.org/10.3389/fpls.2019.00137

Dirks-Mulder A, Butot R, van Schaik P, et al. 2017. Exploring the evolutionary origin of floral organs of Erycina pusilla, an emerging orchid model system. BMC Evolutionary Biology 17, 89. https://doi.org/10.1186/s12862-017-0938-7

Elomaa P, Zhao Y, Zhang T. 2018. Flower heads in Asteraceae—recruitment of conserved developmental regulators to control the flower-like inflorescence architecture. Horticulture Research 5, 36. https://doi.org/10.1038/s41438-018-0056-8

Fiers M, Hoogenboom J, Brunazzi A, Wennekes T, Angenent GC, Immink RGH. 2017. A plant-based chemical genomics screen for the identification of flowering inducers. Plant Methods 13, 78. https://doi.org/10.1186/s13007-017-0230-2 

Flores-Tornero M, Proost S, Mutwil M, Scutt CP, Dresselhaus T, Sprunck S. 2019. Transcriptomics of manually isolated Amborella trichopoda egg apparatus cells. Plant Reproduction 32, 15–27. https://doi.org/10.1007/s00497-019-00361-0 

Francois L, Verdenaud M, Fu X, Ruleman D, Dubois A, Vandenbussche M, Bendahmane A, Raymond O, Just J, Bendahmane M. 2018. A miR172 target-deficient AP2-like gene correlates with the double flower phenotype in roses. Scientific Reports 8, 12912. https://doi.org/10.1038/s41598-018-30918-4

Galimba KD, Martinez-Gomez J, Di Stilio VS. 2018. Gene duplication and transference of function in the paleoAP3 lineage of floral organ identity genes. Frontiers in Plant Science 9, 334. https://doi.org/10.3389/fpls.2018.00334

Hyun Y, Vincent C, Tilmes V, Bergonzi S, Kiefer C, Richter R, Martinez-Gallegos R, Severing E, Coupland G. 2019. A regulatory circuit conferring varied flowering response to cold in annual and perennial plants. Science 363, 409–412. DOI: 10.1126/science.aau8197

Morel P, Heijmans K, Rozier F, Zethof J, Chamot S, Bento SR, Vialette-Guiraud A, Chambrier P, Trehin C, Vandenbussche M. 2017. Divergence of the floral A-function between an asterid and a rosid species. Plant Cell 29, 1605–1621. https://doi.org/10.1105/tpc.17.00098

Nishiyama T, Sakayama H, de Vries J, et al. 2018. The Chara genome: secondary complexity and implications for plant terrestrialization. Cell 174, 448–464.E24. https://doi.org/10.1016/j.cell.2018.06.033

Raymond O, Gouzy J, Just J, et al. 2018. The Rosa genome provides new insights into the domestication of modern roses. Nature Genetics 50, 772–777. https://doi.org/10.1038/s41588-018-0110-3

Reyes-Olalde JI, Zuniga-Mayo VM, Serwatowska J, et al. 2017. The bHLH transcription factor SPATULA enables cytokinin signaling, and both activate auxin biosynthesis and transport genes at the medial domain of the gynoecium. Plos Genetics 13, e1006726. https://doi.org/10.1371/journal.pgen.1006726

Zhou L, Dresselhaus T. 2019. Friend or foe: signaling mechanisms during double fertilization in flowering seed plants. Current Topics in Developmental Biology 131, 453–496. https://doi.org/10.1016/bs.ctdb.2018.11.013

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Flowering in response to cold in annual and perennial plants

by Jens Sundström
Department of Plant Biology, Swedish University of Agricultural Sciences, Uppsala, Sweden.

In relation to annual plants, perennial plants initiate flowering after a more extended juvenile period. Also, perennial plants can often initiate flowering during repeated seasonable cycles and maintain vegetative shoot apical meristems during flowering. A recent study by Hyun and co-workers demonstrate that flowering in response to cold temperatures is regulated by distinct parallel pathways in the annual Arabidopsis thaliana and in the closely related perennial species Arabis alpina (Hyun et al., 2019).

In current discussions about future agriculture, perennial crops are frequently proposed as a possible solution for some of the problems associated with modern agriculture (Crews et al., 2018). In contrast to annual plants, which flower within one growing season, perennial plants typically go through a more extended juvenile period that may last several growth-seasons before flowering and seed production first is initiated. During the juvenile period, the plants invest in vegetative growth, which allows the plants to grow deeper root systems and more extensive foliage. Evergreen fields and plants with deeper root systems will increase carbon sequestration to the soil and provide crops with enhanced drought tolerance. Once the juvenile period has ended, perennial plants may go through repeated rounds flowering, which, if implemented in crop plants would allow for repeated harvesting without tillage, thus reducing nutrient leakage and fossil fuel usage. Hence, even though most crops presently are annuals, perenniality may become an important trait in future agriculture. Increased understanding of how annuals and perennials regulate flowering is therefore timely.

Figure 1. Schematic drawing on the genetic pathways regulating flowering in the annual Arabidopsis thaliana and in the perennial plant Arabis alpina (Illustration by Cajsa Lithell, RedCap design, 2019)

Described in general terms, the time to flower or flowering is determined by a combination of environmental and developmental signals. In temperate regions, an extended period of cold temperatures is often needed for flowering to occur, even if day-length and temperatures are permissive. This process is called vernalization, and in the annual crucifer A. thaliana winter temperatures lead to a stable down-regulation of the floral repressor FLOWERING LOCUS C (FLC) (Michaels and Amasino, 1999), which allows for flowering to occur when day length conditions are favourable (Figure 1). Studies of the perennial crucifer A. alpina have demonstrated that the FLC ortholog is only transiently down-regulated upon cold treatments (Wang et al., 2009), allowing for repeated cycles of floral initiation and floral repression, which is characteristic of the cyclic life history of perennials. 

One additional important distinction between annual and perennial plants is that in annuals all shoot meristems initiate reproductive development at the same time point, whereas perennial plants maintain vegetative after flower initiation. In perennial plants vegetative growth after flowering is maintained either by keeping some meristems in a vegetative state during flowering or by reverting meristems back to vegetative development after flowering.

In a recent study published in Science, Hyun and co-workers demonstrate that the characteristic perennial flowering properties of A. alpina depends on the floral integrator SQUAMOSA PROMOTER BINDING PROTEIN-LIKE 15 (SPL15) (Hyun et al., 2019). In A. thaliana, SPL15 promotes flowering under short-day conditions. Using a combination of genetic and molecular methods, Hyun and co-workers showed that transcription of the SPL15 ortholog in A. alpina is negatively regulated by the FLC ortholog. Hence, temporal down-regulation of the FLC ortholog in shoot apical meristems in response to cold treatments leads to an up-regulation of AaSPL15 and subsequent flowering.

However, flowering is only initiated in shoot apical meristems of a certain age, suggesting an additional age-dependent regulation of AaSPL15. AaSPL15 belongs to a family of transcription factor-coding genes that are negatively regulated by microRNAs of the miR156 family. In diverse plant lineages, miR156 is expressed at high levels in young plants and the expression of miR156 gradually decreases as plants ages. miR156 binds to a specific motif in the AaSPL15 transcript, leading to a reduction in its levels (Bergonzi et al., 2013). By expressing a miR156-resistant form of AaSPL15 in transgenic A. alpina plants, Hyun and co-workers show that miR156 suppresses AsSPL15 accumulation in young shoot apical meristems. In line with this, cold-treated plants that express the miR156-resistant form of AaSPL15 display a shortened juvenile period and flower prematurely. In addition, shoot meristems that would be maintained as vegetative in the wild type initiate flowers in the transgenic plants, suggesting that AaSPL15 is negatively regulated by miR156 in an age-dependent manner.

Central to the seasonal reproductive cycles in A. alpina is the transient down-regulation of the FLC ortholog in response to cold (Wang et al., 2009). In the annual A. thaliana transcription of FLC is permanently down-regulated after cold treatment (Hepworth and Dean, 2015). FLC acts as a repressor of both SPL15 and genetic factors that promote flowering in response to day length. Hence, in the annual A. thaliana, long-day conditions after vernalization induce flowering, whereas restored expression of the FLC ortholog in A. alpina represses flowering in response to day length conditions.

Taken together, this suggests that AaSPL15 integrates cues derived from winter cold and age in shoot apical meristems of A. alpina. The transient down-regulation of the FLC ortholog in A. alpina and subsequent up-regulation of AaSPL15 transcription allows for seasonal induction of flowering after winter vernalization, whereas expression of miR156 suppresses AaSPL15 in juvenile plants and in meristems that maintain vegetative identity during flowering.

Distinctions between annual and perennial growth have arisen independently in many different plant lineages (Thomas et al., 2000), suggesting that distinct mechanisms may regulate flowering in different plant lineages. However, the central genetic components of the flowering pathways are in many cases well conserved even between distantly related plant lineages indicating that it is, in part, the regulation of the flowering pathways that determine an annual or perennial life cycle. It will be interesting to see if the knowledge gained in these studies can facilitate further development of perennial crops plants.


Bergonzi S, Albani MC, Ver Loren van Themaat E, Nordstrom KJ, Wang R, Schneeberger K, Moerland PD, Coupland G. 2013. Mechanisms of age-dependent response to winter temperature in perennial flowering of Arabis alpina. Science 340, 1094-1097. DOI: 10.1126/science.1234116

Crews TE, Carton W, Olsson L. 2018. Is the future of agriculture perennial? Imperatives and opportunities to reinvent agriculture by shifting from annual monocultures to perennial polycultures. Global Sustainability 1, e11. https://doi.org/10.1017/sus.2018.11

Hepworth J, Dean C. 2015. Flowering locus C’s lessons: conserved chromatin switches underpinning developmental timing and adaptation. Plant Physiology 168, 1237-1245. DOI: https://doi.org/10.1104/pp.15.00496

Hyun Y, Vincent C, Tilmes V, Bergonzi S, Kiefer C, Richter R, Martinez-Gallegos R, Severing E, Coupland G. 2019. A regulatory circuit conferring varied flowering response to cold in annual and perennial plants. Science 363, 409-412. DOI: 10.1126/science.aau8197

Michaels SD, Amasino RM. 1999. FLOWERING LOCUS C encodes a novel MADS domain protein that acts as a repressor of flowering. Plant Cell 11, 949-956. DOI: https://doi.org/10.1105/tpc.11.5.949

Thomas H, Thomas HM, Ougham H. 2000. Annuality, perenniality and cell death. Journal of Experimental Botany 51, 1781-1788.

Wang R, Farrona S, Vincent C, Joecker A, Schoof H, Turck F, Alonso-Blanco C, Coupland G, Albani MC. 2009. PEP1 regulates perennial flowering in Arabis alpina. Nature 459, 423-427. https://doi.org/10.1038/nature07988

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Modelling vernalization response – fast and slow

by  Aalt-Jan van Dijk

Wageningen University, The Netherlands

In one of my previous commentaries (see ‘A hitchhiker guide to modelling’), I discussed some issues related to the development of computational models for the molecular regulation of flowering. One aspect that I did not mention was how the different timescales at which biological processes take place have repercussions for the models built to represent these processes, and vice versa, how different models deal with time in different ways. One example is that in some modelling approaches (e.g. Boolean Networks) time is represented as a discrete variable. In other modelling approaches, time is continuous. Ordinary differential equations (ODEs) are the most prominent example of such models. ODEs can model processes that take place ‘fast’ as well as ‘slow’. This is encoded in the model either by the structure of the equations or by the value of specific parameters in the equations. One example from my own work is transport of FT which can be included in an ODE model using a so-called delay term in the equations (Leal Valentim et al., 2015).aaltjan image

Different time scale are clearly present during vernalization. As it is well-known, FLC is downregulated by prolonged cold and epigenetically silenced to allow the plant to be maximally responsive to floral-promoting long-day photoperiods in spring. The regulatory network controlling FLC must distinguish a seasonal signal over months, despite daily temperature fluctuations that can exceed average seasonal differences. A recent publication by Antoniou-Kourounioti et al. 2018 analyse the regulation of FLC by temperature using a mathematical model for vernalization that operates on multiple timescales: long term (month), short term (day), and ‘immediate’ response. Here, I describe how this model was built and how it describes fast and slow response of plants to temperature.
The model contains two main modules: one consisting of a model for VIN3 regulation, and the other for FLC regulation. In both models, temperature sensing itself is not directly modelled; rather, heuristic functions are defined which reflect how temperature changes would affect VIN3 and FLC. In addition, temperature-induced changes in VIN3 also affect FLC because of the regulation of FLC by VIN3.
For VIN3 regulation, two thermoregulatory processes were already known experimentally, and both of these were included in the model. (i) One temperature-sensitive pathway holds the memory of the duration of the cold. This process was described by a component that would be produced only in the cold and degrades very slowly in both the cold and the warm, thereby integrating over the period of cold that the plant has experienced. (ii) For a second temperature-sensitive pathway a component was used which measures current temperature and has fast-acting dynamics. This component is responsible for the rapid reduction in VIN3 levels observed at high temperatures. In addition to these two temperature-dependent pathways, a temperature-independent function was used to represent the circadian clock.
After performing experiments with temperature spikes which I do not describe here, a vernalization model was constructed, representing the dynamics of FLC, incorporating both VIN3-dependent (derived from the VIN3 model above) and VIN3-independent pathways. The FLC model consists of three states of the FLC gene, together with transitions between these states. One of the states is transcriptionally active. Gene copies in this state can switch to a transcriptionally inactive state through a VIN3-independent pathway. From the inactive state, there is an irreversible switch to an epigenetically stable off state. The rate at which this switch takes place depends on the cold-induced VIN3 level. An additional VIN3-dependent transition directly from the active stage to the off state allows epigenetic silencing of FLC in the absence of VIN3-independent FLC downregulation. In addition to the temperature dependence of VIN3 dynamics, the transitions to the off state are also directly temperature dependent in this model.
To build the model described above, experimental data were used for model construction and for parameter fitting. The fact that the model can reproduce these data does not tell us whether it is capable of predicting anything. To test this, the authors used additional field experiments: measured temperature profiles from these experiments were used as input to predict VIN3 and FLC, and these predictions were compared with measurements. Such comparison is less straightforward than one might think. For example, the time of the day at which sampling takes place could clearly have an influence. It is mentioned that the diurnal pattern of VIN3 was shifted by several hours between different experimental conditions. This change meant that the peak of VIN3 expression was much later than the sampling time in some conditions, and therefore, compared to the model, the experiments had much lower amplitude for these samples. To me this demonstrates one key reason why models can be useful: not only when they are ‘correct’ but in particular when there is a mismatch between model predictions and experimental data, which helps to make knowledge gaps very explicit.
Having established that the VIN3/FLC combined model can predict responses to field conditions, it was examined to which features of the field temperature profile the model was most sensitive. First, the full temperature profile was replaced by the mean temperature of each day. According to the model, because of lower activation of the VIN3-independent pathway, the absence of cold temperatures in the day-mean profile initially lead to slower FLC downregulation. However, later in winter, the absence of daily warm spikes caused simulated VIN3 levels to be higher, leading to lower simulated FLC levels. In subsequent simulations, a higher mean temperature as well as the same mean temperature with more fluctuations were tested.
One main conclusion from the paper is that temperature sensing is broadly distributed, with various thermosensory processes responding to specific features of the plants’ history of exposure to warm and cold. Less strongly stated, the paper presents a model describing distributed thermosensing, which is in accordance with the available experimental data. Apart from what these results mean for our understanding of thermosensing, I found it particularly interesting that the paper demonstrates how a computational model can be used to improve our understanding of fast and slow response during vernalization.


Antoniou-Kourounioti RL, Hepworth J, Heckmann A, Duncan S, Qüesta J, Rosa S, Säll T, Holm S, Dean C, Howard M. 2018. Temperature sensing is distributed throughout the regulatory network that controls FLC epigenetic silencing in vernalization. Cell 7, 643-655.e9. https://doi.org/10.1016/j.cels.2018.10.011

Espinosa-Soto C, Padilla-Longoria P, Alvarez-Buylla ER. 2004. A gene regulatory network model for cell-fate determination during Arabidopsis thaliana flower development that is robust and recovers experimental gene expression profiles. The Plant Cell, (16) 2923–2939. DOI: https://doi.org/10.1105/tpc.104.021725

Leal Valentim F, van Mourik S, Posé D, Kim MC, Schmid M, van Ham RCHJ, Busscher M, Sanchez-Perez GF, Molenaar J, Angenent GC, Immink RGH, van Dijk ADJ. 2015. A quantitative and dynamic model of the Arabidopsis flowering time gene regulatory network. PLoS ONE 2015 doi: 10.1371/journal.pone.0116973. https://doi.org/10.1371/journal.pone.0116973

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Polyploidy – from evolution to development?

by Rainer Melzer
School of Biology and Environmental Science and Earth Institute, University College Dublin

A large variety of molecular mechanisms regulate developmental processes. Transcription factors entered the stage already decades ago and are still widely considered master regulators of development (Wray et al., 2003). However, in the past years it has been demonstrated that many other biomolecules play pivotal roles in development. This includes histone modifying complexes, small RNAs and long          noncoding RNAs (Holoch and Moazed, 2015, Venkatesh and Workman, 2015). In almost all those cases, the developmental importance of a regulatory mechanism is demonstrated first. This is often then followed by evolutionary considerations and phylogenetic studies. For example, transcription factors now play a central role in evo-devo research, and changes in transcription factor function are proposed to have played important roles in the evolution of plant and animal body plans (Cheatle Jarvela and Hinman, 2015, Rodríguez-Mega et al., 2015). Likewise, microRNAs have been implicated in major evolutionary transitions (Peterson et al., 2009).

In contrast to those regulatory molecules, another mechanism that has far-reaching consequences for cellular processes has mainly been discussed in the ecological and evolutionary arena: polyploidy (Ramsey and Ramsey, 2014). The doubling of the entire genome has long been implicated in the origin of new species or even entire new clades. Polyploidy has recently gained even more attention with the discovery that many plant lineages underwent polyploidization events in the past. Current evidence indicates that polyploidization is associated with an increase in diversification rates and possibly also with the origin of evolutionary novelties (Schranz et al., 2012, Landis et al., 2018).

OrchidFlower Rainer Melzer Compared to the importance of polyploidy in evolutionary research, it has gained relatively little attention in plant developmental biology. This is to some extent surprising, as it is well established that different somatic cells of a plant can differ in their ploidy level (Traas et al., 1998). At the same time, it is also known that there is a direct relationship between e.g. the size of a cell and its ploidy level (Traas et al., 1998). Thus, heterogeneity at the cell level and phenotypic output might be directly linked.               Bateman et al., (2018) explored this link between polyploidy and developmental patterning further by analysing orchid flowers. Orchids are an especially interesting study subject as the floral structure of many orchids is quite elaborate, showing extensive micromorphological diversification. Thus, one question that arises is whether those intricate morphologies are in part brought about by different ploidy levels in different cells.
Bateman and colleagues used microscopy to measure the size of nuclei in different regions of the labellum – a highly specialized petal-like organ of orchids. They complemented those studies with flow cytometry, which provides an independent and more quantitative estimate of nuclear genome sizes in different organs, but lacks the cellular resolution of microscopic measurements (Bateman et al., 2018).

Intriguingly, the authors find significant variations of the ploidy level depending on the labellum region. Polyploidy was especially pronounced in trichomes of the labellum. Those trichomes were also among the largest and most complex cells analysed, confirming that there might be a direct link between cell size and ploidy level. In addition, the flow cytometry data provided some evidence that the ratio between different ploidy levels is tissue- and organ-specific, with leaf material showing a relatively high percentage of diploid and tetraploid nuclei whereas in the labellum diploid nuclei were very rare and higher ploidy levels dominated (Bateman et al., 2018). Also, different labellum regions showed different flow cytometry signatures, indicating that some kind of ‘ploidy code’ might be applicable to distinguish different organs and tissues.

The authors further speculate that the size increase often observed in polypoid cells might, if cells are embedded in a tissue, lead to local distortions that may in turn contribute to the elaborate three-dimensional structure of many orchid labella. Interestingly, data from giant polyploid cells in Arabidopsis sepals support this notion. In this case, the large cells are required for the curvature of the sepals (Roeder et al., 2012).

The study of Bateman et al., (2018) provides an exciting starting point for further research. After polyploidy research has taken evolutionary biology by storm the time seems right to explore its relevance for developmental mechanisms.


Bateman RM, Guy JJ, Rudall PJ, Leitch IJ, Pellicer J, Leitch AR. 2018. Evolutionary and functional potential of ploidy increase within individual plants: somatic ploidy mapping of the complex labellum of sexually deceptive bee orchids. Annals of Botany 122: 133-150. doi: 10.1093/aob/mcy048

Cheatle Jarvela AM, Hinman VF. 2015. Evolution of transcription factor function as a mechanism for changing metazoan developmental gene regulatory networks. EvoDevo 6: 3. https://doi.org/10.1186/2041-9139-6-3

Holoch D, Moazed D. 2015. RNA-mediated epigenetic regulation of gene expression. Nature Reviews Genetics 16: 71. doi: 10.1038/nrg3863

Landis JB, Soltis DE, Li Z, Marx HE, Barker MS, Tank DC, Soltis PS. 2018. Impact of whole-genome duplication events on diversification rates in angiosperms. American Journal of Botany 105: 348-363. https://doi.org/10.1002/ajb2.1060

Peterson KJ, Dietrich MR, McPeek MA. 2009. MicroRNAs and metazoan macroevolution: insights into canalization, complexity, and the Cambrian explosion. Bioessays 31: 736-47. https://doi.org/10.1002/bies.200900033

Ramsey J, Ramsey TS. 2014. Ecological studies of polyploidy in the 100 years following its discovery. Philosophical Transactions of the Royal Society B: Biological Sciences 369.

Rodríguez-Mega E, Piñeyro-Nelson A, Gutierrez C, García-Ponce B, Sánchez MDLP, Zluhan-Martínez E, Álvarez-Buylla ER, et al. 2015. Role of transcriptional regulation in the evolution of plant phenotype: A dynamic systems approach. Developmental Dynamics 244: 1074-1095. https://doi.org/10.1002/dvdy.24268

Roeder AHK, Cunha A, Ohno CK, Meyerowitz EM. 2012. Cell cycle regulates cell type in the Arabidopsis sepal. Development 139: 4416-4427.

Schranz EM, Mohammadin S, Edger PP. 2012. Ancient whole genome duplications, novelty and diversification: the WGD Radiation Lag-Time Model. Current Opinion in Plant Biology 15: 147-153. https://doi.org/10.1016/j.pbi.2012.03.011

Traas J, Hülskamp M, Gendreau E, Höfte H. 1998. Endoreduplication and development: rule without dividing? Current Opinion in Plant Biology, 1: 498-503. https://doi.org/10.1016/S1369-5266(98)80042-3

Venkatesh S, Workman JL. 2015. Histone exchange, chromatin structure and the regulation of transcription. Nature Reviews Molecular Cell Biology 16: 178. http://dx.doi.org/10.1038/nrm3941

Wray GA, Hahn MW, Abouheif E, Balhoff JP, Pizer M, Rockman MV, Romano LA. 2003. The evolution of transcriptional regulation in eukaryotes. Molecular Biology and Evolution 20: 1377-1419. https://doi.org/10.1093/molbev/msg140

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A tribute to Lars Hennig (1970–2018)

by Iva Mozgova, Cristina Alexandre, Yvonne Steinbach, Maria Derkacheva, Eberhard Schäfer and Wilhelm Gruissem

         Lars Hennig_photoLars Hennig, Professor of Genetics at the Swedish University of Agricultural Sciences in Uppsala, Sweden, passed away on 17 May 2018, at the early age of 47. Lars was a passionate plant scientist who had a profound knowledge of biology and the determination to address fundamental questions using state-of-the-art methods. His research focused on plant developmental epigenetics, in particular the role of Polycomb group proteins and other chromatin-modifying complexes in modulating plant development and environmental responses. His extensive work is documented in over 100 scientific publications.

Lars was born in Rostock, Germany. After graduating from the Martin Luther University of Halle-Wittenberg he moved to the Albert Ludwigs University in Freiburg in 1996. There he joined the laboratory of Eberhard Schäfer to study the dynamic behaviour and complex interactions of plant photoreceptors. Lars obtained his PhD degree in 1999. He then moved for his postdoctoral research to the ETH in Zurich where he first studied cell cycle-regulated gene expression in Wilhelm Gruissem’s laboratory. In 2003, Lars started his own research group at the ETH focusing on chromatin-based regulation of flowering time. His career as an independent researcher continued to flourish, and in 2010, he and his wife and scientific collaborator Claudia Köhler accepted full professorships at the Swedish University of Agricultural Sciences. Together with their two children they moved to Uppsala. Uprooting his research group was not without challenges, but Lars navigated the move with tact and diplomacy, from accommodating the personal circumstances of all his group members to managing the logistics of doing research during this transition period. Coming to Sweden, Lars set out to combine the best of ETH’s scientific traditions with his new cultural and scientific environment.

Although his research career was cut short by illness, Lars mentored 11 PhD students and 11 postdoctoral fellows who all successfully continued their own careers. His research led to several seminal contributions to the fields of chromatin biology and plant development.

Lars’ postdoctoral research on the different roles of MULTICOPY SUPRESSOR OF IRA 1 (MSI1) in plant development kindled his long lasting interest in chromatin dynamics and the role of chromatin-modifying complexes in regulating developmental transitions. His early work helped establish MSI1 as a subunit of two distinct chromatin-modifying complexes, CHROMATIN ASSEMBLY FACTOR 1 (CAF-1) and POLYCOMB REPRESSIVE COMPLEX 2 (PRC2). He showed that their functions were genetically separable (Hennig et al., 2003; Kohler et al., 2003). Later on, a significant body of work in Lars’ own group was centred on the multiple functions of MSI1, which by then he affectionately called the ‘Swiss-army-knife’. He discovered the function of MSI1-containing complexes in the control of flowering time (Bouveret et al., 2006; Steinbach and Hennig, 2014), cell differentiation and reprogramming (Exner et al., 2006; Mozgová et al., 2017; Nakamura and Hennig, 2017), and modulation of biotic and abiotic stress responses (Alexandre et al., 2009; Mehdi et al., 2016; Mozgová et al., 2015). Lars searched for binding partners of MSI1, and found that it linked the H3K27me3-binding LIKE HETEROCHROMATIN PROTEIN 1 (LHP1) (Turck et al., 2007; Exner et al., 2009) to the PRC1-PRC2 functional network. As a PRC2 component, LHP1 was proposed to be involved in the inheritance of H3K27me3 marks during cell division (Derkacheva et al., 2013). LHP1 immunoprecipitation further revealed its direct interaction with PRC2 subunits, including MSI1, and identified the histone H2A deubiquitinases UBP12 and UBP13 to be physically and functionally associated with PRC2 (Derkacheva et al., 2016).

Lars was enthusiastic about exploring global chromatin structure, mapping genome-wide patterns of DNA accessibility and non-canonical histone variant distribution (Shu et al., 2012, 2014), developing protocols for profiling of DNA accessibility (Shu et al., 2013), and identifying secondary DNA structures in intact chromatin (Gentry and Hennig, 2016). Using purified histones from cauliflower, his group identified two novel histone modifications in plants, the pericentromeric heterochromatin-associated H3K23me1 (Trejo-Arellano et al., 2017) and H3K36ac associated with actively transcribed genes (Mahrez et al., 2016).

While pursuing his research interests, Lars was always an active member of the plant science community. As a skilled biostatistician and bioinformatician, he and his colleagues at ETH Zurich developed pioneering functional genomic tools and established benchmarks for plant researchers. Examples include the powerful search engine Genevestigator for mining and comparative analysis of gene expression data (Zimmermann et al., 2004, 2005), the AGRONOMICS1 Affymetrix microarray that expanded options for Arabidopsis transcriptomics and ChIP-chip experiments (Rehrauer et al., 2010), the MIAME annotation standards for plant genome-wide profiling (Zimmermann et al., 2006), and PlantDB (Exner et al., 2008), a database for managing plant experiment documentation and stocks.

Together with Valérie Gaudin and Claudia Köhler, Lars initiated the successful biannual European Workshop Series on Plant Chromatin. He had an enduring fascination with the beauty and complexity of flowers. In his laboratory, flowering time reigned supreme as the developmental phenotype of choice. Outside his lab, Lars was an associate editor and the Flowering Newsletter editor of the Journal of Experimental Botany from 2012 to 2017, and established the Flowering Highlights blog.

Lars had an unwavering scientific curiosity, an astounding breadth of knowledge spanning different research fields, and the uncanny ability to remember seemingly all pertinent published data. As a mentor, Lars was dedicated and caring: he knew how to motivate students and postdocs at times of frustration but he also made them pause and reflect on exciting but preliminary results. His insistence on multiple experimental controls as well as the critical judgement of all data and the distinction between facts and interpretations became tenets for students and postdocs alike. Lars was committed to training the next generation of curious and rigorous scientists. He actively encouraged them to explore their career opportunities, not only by providing them with the freedom to pursue their own scientific questions but also by helping them to hone their manuscript and grant-writing skills. He wanted to see them grow as scientist and spent many hours discussing and proof-reading manuscripts.

The atmosphere around Lars was always lively and enjoyable: he liked to mingle with group members, get to know their personality and cultural background, promote discussions, and facilitate collaboration. There were laboratory lunches sweetened with Swiss chocolates, many outings, accepted manuscript celebrations, and regular after-lab beer meetings. All the BBQs, hikes in the mountains, kayaking on the Baltic Sea, and even the visit to a moose farm in the gushing rain will be fondly remembered.

We were fortunate to have worked with Lars as mentors, colleagues, collaborators, students, and postdocs. Despite his conviction that ‘life is not designed to be fair’ and his doubt about the ‘absolute truth’ in biology, Lars’ passionate quest for fairness and truth was inspiring. His sharp mind, his wisdom, his sense of humour and his friendship will be greatly missed.


We would like to thank the following colleagues for suggestions and insights: Claudia Köhler, Miyuki Nakamura, Jordi Moreno Romero, Minerva Trejo Arellano, Jennifer de Jonge, and Thomas Wildhaber.


Alexandre C, Moller-Steinbach Y, Schonrock N, Gruissem W, Hennig L. 2009. Arabidopsis MSI1 is required for negative regulation of the response to drought stress. Molecular Plant 2, 675–687. doi: 10.1093/mp/ssp012

Bouveret R, Schonrock N, Gruissem W, Hennig L. 2006. Regulation of flowering time by Arabidopsis MSI1. Development 133, 1693–1702.

Derkacheva M, Liu S, Figueiredo DD, Gentry M, Mozgova I, Nanni P, Tang M, Mannervik M, Kohler C, Hennig L. 2016. H2A deubiquitinases UBP12/13 are part of the Arabidopsis polycomb group protein system. Nature Plants 2, 16126. http://dx.doi.org/10.1038/nplants.2016.126

Derkacheva M, Steinbach Y, Wildhaber T, Mozgova I, Mahrez W, Nanni P, Bischof S, Gruissem, Wilhelm2 3, Hennig L. 2013. Arabidopsis MSI1 connects LHP1 to PRC2 complexes. EMBO Journal 32, 2073–2085. doi: 10.1038/emboj.2013

Exner V, Aichinger E, Shu H, Wildhaber T, Alfarano P, Caflisch A, Gruissem W, Kohler C, Hennig L. 2009. The chromodomain of LIKE HETEROCHROMATIN PROTEIN 1 is essential for H3K27me3 binding and function during Arabidopsis development. PLoS ONE 4, e5335. https://doi.org/10.1371/journal.pone.0005335

Exner V, Hirsch-Hoffmann M, Gruissem W, Hennig L. 2008. PlantDB – a versatile database for managing plant research. Plant Methods 4, 1. https://doi.org/10.1186/1746-4811-4-1

Exner V, Taranto P, Schonrock N, Gruissem W, Hennig L. 2006. Chromatin assembly factor CAF-1 is required for cellular differentiation during plant development. Development 133, 4163–4172.doi: 10.1371/journal.pone.0005335

Gentry M, Hennig L. 2016. A structural bisulfite assay to identify DNA cruciforms. Molecular Plant 9, 1328–1336. https://doi.org/10.1016/j.molp.2016.06.003

Hennig L, Taranto P, Walser M, Schonrock N, Gruissem W. 2003. Arabidopsis MSI1 is required for epigenetic maintenance of reproductive development. Development 130, 2555–2565.

Kohler C, Hennig L, Bouveret R, Gheyselinck J, Grossniklaus U, Gruissem W. 2003. Arabidopsis MSI1 is a component of the MEA/FIE polycomb group complex and required for seed development. EMBO Journal 22, 4804–4814.

Mahrez W, Trejo Arellano MS, Moreno-Romero J, Nakamura M, Shu H, Nanni P, Köhler C, Gruissem W, Hennig L. 2016. H3K36ac is an evolutionary conserved plant histone modification that marks active genes. Plant Physiology 170, 1566–1577. https://doi.org/10.1104/pp.15.01744

Mehdi S, Derkacheva M, Ramström M, Kralemann L, Bergquist J, Hennig L. 2016. MSI1 functions in a HDAC complex to fine-tune ABA signaling. The Plant Cell 28, 42–54. https://doi.org/10.1105/tpc.15.00763

Mozgová I, Muñoz-Viana R, Hennig L. 2017. PRC2 represses hormone-induced somatic embryogenesis in vegetative tissue of Arabidopsis thaliana. PLOS Genetics 13, e1006562. https://doi.org/10.1371/journal.pgen.1006562

Mozgová I, Wildhaber T, Liu Q, Abou-Mansour E, L’Haridon F, Metraux JP, Gruissem W, Hofius D, Hennig L. 2015. Chromatin assembly factor CAF-1 represses priming of plant defence response genes. Nature Plants 1, 15127. http://dx.doi.org/10.1038/nplants.2015.127

Nakamura M, Hennig L. 2017. Inheritance of vernalization memory at FLOWERING LOCUS C during plant regeneration. Journal of Experimental Botany 68, 2813–2819. https://doi.org/10.1093/jxb/erx154

Rehrauer H, Aquino C, Gruissem W, Henz SR, Hilson P, Laubinger S, Naouar N, Patrignani A, Rombauts S, Shu H, Van de Peer Y, Vuylsteke M, Weigel D, Zeller G, Hennig L. 2010. AGRONOMICS1: a new resource for Arabidopsis transcriptome profiling. Plant Physiology 152, 487–499. https://doi.org/10.1104/pp.109.150185

Shu H, Gruissem W, Hennig L. 2013. Measuring Arabidopsis chromatin accessibility using DNase I-polymerase chain reaction and DNase I-chip assays. Plant Physiology 162, 1794–1801. https://doi.org/10.1104/pp.113.220400

Shu H, Nakamura M, Siretskiy A, Borghi L, Moraes I, Wildhaber T, Gruissem W, Hennig L. 2014. Arabidopsis replacement histone variant H3.3 occupies promoters of regulated genes. Genome Biology 15, R62. https://doi.org/10.1186/gb-2014-15-4-r62

Shu H, Wildhaber T, Siretskiy A, Gruissem W, Hennig L. 2012. Distinct modes of DNA accessibility in plant chromatin. Nature Communications 3, 1281. http://dx.doi.org/10.1038/ncomms2259

Steinbach Y, Hennig L. 2014. Arabidopsis MSI1 functions in photoperiodic flowering time control. Frontiers in Plant Science 5, 77. https://doi.org/10.3389/fpls.2014.00077

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Flowering and dormancy in temperate perennials

by Maria C. Albani
Botanical Institute, University of Cologne, Germany.
Max Planck Institute for Plant Breeding Research, Cologne, Germany.

In most temperate environments one can see trees flowering very early in the spring. For most perennials flowering in the spring marks the event of floral emergence instead of the time of the induction of flowering and flower bud initiation as it is for many annual species.


A flowering cherry tree in Cologne, Germany, Spring 2018

Thus, trees can flower very early because most of them had initiated the flower buds already the previous year during the summer, autumn or winter. To survive the winter, many perennials also cease growth in the autumn and become dormant during the winter.  Environmental cues such as photoperiod and cold regulate growth cessation and bud dormancy release.  For example, short photoperiods in the autumn are required to induce growth cessation whereas prolonged cold is required to break bud dormancy.
A recent study in hybrid aspen, which is a cross between the European Populus tremula and the American aspen P. tremuliodes, highlights the role of photoperiod in setting the dormant state independently of growth cessation (Tylewicz et al., 2018). Short days block cellular communication through plasmodesmata closure and this process involves the phytohormone ABA. The authors created transgenic aspen with reduced ABA response, overexpression lines of the PDLP1 gene, which impairs trafficking via plasmodesmata, and DsRNAi lines of the chromatin remodelling factor PICKLE (PKL). These transgenics were used to demonstrate the ABA-dependent pathway for plasmodesmata closure and their role in bud dormancy. The authors also used grafting to show that closure of the plasmodesmata regulates the inability of the bud to grow. For this, they grafted scions of wild type and transgenics plants with reduced ABA response grown in short days (so that only scions of the transgenics will have open plasmodesmata) onto rootstocks of lines overexpressing the aspen FLOWERING LOCUS T 1 (FT1) gene. Under these conditions, buds of wild type scions did not reactivate growth whereas buds from scions of the transgenics that had compromised ABA response showed bud outgrowth. These results lead to the conclusion that plasmodesmata closure induced by short days blocks the FT1-derived growth promoting signals to access the meristem. The authors also suggested that re-opening of the plasmodesmata occurs slowly and only after exposure to low temperatures.

The study of Tylewicz et al., 2018 is not about flowering as it has been performed using juvenile/vegetative plants. It however raises interesting questions if one takes into account the flowering patterns in perennials. In P. deltoides trees grown in Starville (Mississippi, USA) it has been demonstrated that flower buds are initiated during the winter when plants are exposed to short day length and low temperatures. In this Populus species, flowering and the return to vegetative development is regulated by two paralogues of FT, FT1 and FT2 (Hsu et al., 2011).  FT1 expression was increased during the winter in many tissues including the reproductive buds, whereas FT2 trancripts were only  up-regulated in the leaves after the return to warm temperatures. These results suggested that FT1 regulates reproductive onset in response to winter temperatures whereas FT2 promotes vegetative growth after the winter in response to warm temperatures and long days.

In the study of Hsu et al., 2011, although trees were considered to undergo the dormant phase, obviously things still happen during prolonged exposure to cold as flower buds were initiated. Thus it would be interesting to study how the model on bud dormancy in vegetative buds, described by Tylewicz et al., 2018 can be translated to the regulation of flowering in perennials. Is plasmodesmata closure also important in the flowering buds? Does flower bud initiation need open or closed plasmodesmata? Do plasmodesmata also play a role in the outgrowth of these flower buds in the spring? Although these are interesting questions to answer, it is probably technically difficult to address due to the long juvenile phase of trees.


Tylewicz S, Petterle A, Marttila S, Miskolczi P, Azeez A, Singh RK, Immanen J, Mähler N, Hvidsten TR, Eklund DM, Bowman JL, Helariutta Y, Bhalerao RP. 2018. Photoperiodic control of seasonal growth is mediated by ABA acting on cell-cell communication. Science 360(6385): 212-215. doi: 10.1126/science.aan8576.

Hsu CY, Adams JP, Kim H, No K, Ma C, Strauss SH, Drnevich J, Vandervelde L, Ellis JD, Rice BM, Wickett N, Gunter LE, Tuskan GA, Brunner AM, Page GP, Barakat A, Carlson JE, DePamphilis CW, Luthe DS, Yuceer C. 2011. FLOWERING LOCUS T duplication coordinates reproductive and vegetative growth in perennial poplar. Proceedings of the National Academy of Sciences, USA. 108(26):10756-61. doi: 10.1073/pnas.1104713108.

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TCP functions branching out

by Paula Elomaa
Department of Agricultural Sciences, Viikki Plant Science Centre, University of Helsinki, Finland

Since their discovery about 20 years ago, the TCP domain transcription factors have been shown to control diverse aspects of plant growth and development (reviewed in Nicolas and Cubas, 2015). The functions of class II TCP proteins, including the CINCINNATA and CYCLOIDEA/TEOSINTE BRANCHED1–like proteins, have been attributed to leaf development, floral symmetry patterns (zygomorphy) as well as outgrowth of lateral shoots. Many of these genes have been targeted during domestication and by adaptation under natural conditions. Emerging data emphasizes the importance of TCP proteins, and particularly their fine-tuned regulation, in integrating hormonal and environmental signals affecting development (Li et al., 2015; Nicolas and Cubas, 2015; 2016).

One of the founding members of the TCP protein family was the TEOSINTE BRANCHED1 (TB1) in maize found to suppress lateral branching, a major trait contributing to its domestication from teosinte (Doebley et al., 1997). Shoot branching control by TB1 orthologs is highly conserved among angiosperms, and a key trait also from an agronomic perspective (Nicolas and Cubas, 2015). A recent paper by Dixon et al. (2018) demonstrates a functional role for a TB1 ortholog of bread wheat (Triticum aestivum L.) in regulation of inflorescence architecture, providing potential to affect grain production. The inflorescence development in grasses involves complex branching events. While the indeterminate raceme in Arabidopsis elongates and develops individual pedicellate flowers in its axils, the basic unit in a grass inflorescence is the branched spikelet, a terminal unit capable of producing florets (Fig. 1A). In case of wheat, single spikelets develop in alternate phyllotaxis along the central rachis and each of them produce multiple florets. In their paper, Dixon et al. demonstrate that TB1 regulates the paired spikelet trait in wheat where two spikelets are formed in individual rachis nodes instead of a single one (Fig. 1B).

The highly-branched (hb) wheat line, analyzed in this paper, showed altered growth of lateral organs by developing multiple paired spikelets in their inflorescences as well as fewer tillers. The hb lines were delayed in their transition to reproductive development, and showed delayed inflorescence growth especially during early developmental stages (leaf stages L5-L7). However, the final length of the mature inflorescences was not altered. Analysis of the QTL region contributing to the paired spikelet trait revealed the presence of the TB1. The hb line showed increased (tetrasomic) dosage of chromosome 4D, and specifically the TB1 expression originating from the wheat D genome (TB-D1) was significantly upregulated both in hb tillers (including tiller buds) as well as in inflorescences during the stages when their growth was delayed. The dosage dependent TB-D1 regulation was confirmed by modifying the number of functional copies through crossings to tb-d1 mutant line, by analyzing the revertant phenotypes of hb plants as well as by overexpression of TB-D1 in transgenic plants. Intriguingly, Dixon et al. linked the TB1 function with regulation of flowering by showing that TB-D1 directly interacts with the major flowering regulator FT1, and that increased dosage of TB-D1 reduces the transcript levels of several meristem identity genes. The allelic diversity of TB1 in both B and D genomes was further associated with paired spikelet development in modern wheat cultivars.

This work connects branching control with regulation of flowering, and demonstrates a fine-tuned regulatory link between these major developmental events. The established model by the authors propose that increased dosage of TB1 reduce the availability of FT to activate spikelet meristem identity genes, and facilitates inflorescence branching by modifying the temporal timing or rate of spikelet meristem maturation. As the number of spikelets determines the seed number and crop yield – keeping in mind the possible adverse effects due to altered sink-source relations – this work adds a valuable gene from the TCP family among the breeding targets, not only in wheat but potentially also in other cereals as discussed by Dixon and colleagues.


Dixon LE, Greenwood JR, Bencivenga S, Zhang P, Cockram J, Mellers G, Ramm K, Cavanagh C, Swain SM, Boden SA. 2018. TEOSINTE BRANHCED1 regulates inflorescence architecture and development in bread wheat (Triticum aestivum L.). The Plant Cell, doi: 10.1105/tpc.17.00961
Doebley J, Stec A, Hubbard L. 1997. The evolution of apical dominance in maize. Nature 386, 485-488. doi:10.1038/386485a0
Li S. 2015. The Arabidopsis thaliana TCP transcription factors: a broadening horizon beyond development. Plant Signaling & Behavior 10, e1044192-2. doi: 10.1080/15592324.2015.1044192
Nicolas M, Cubas P. 2015. The role of TCP transcription factors in shaping flower structure, leaf morphology, and plant architecture. In: Gonzalez DH, ed. Plant transcription factors. Evolutionary, structural, and functional aspects. Academic Press, Elsevier Inc. doi: 10.1016/B978-0-12-800854-6.00016-6
Nicolas M, Cubas P. 2016. TCP factors: new kids on the signaling block. Current Opinion in Plant Biology 33: 33-41. doi: 10.1016/j.pbi.2016.05.006

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A surprising role for ethylene in the regulation of petal cell shape

Beverley Glover
Department of Plant Sciences, University of Cambridge, Cambridge CB2 3EA, UK bjg26@cam.ac.uk

The different shapes of plant epidermal cells are always fascinating. One of the first experiences many students have of scanning electron microscopy is the excitement of seeing the amoeba-shaped cells of leaves, interspersed with stomata. Leaf epidermal cell shape is particularly intriguing because the amoeboid cells are so unexpected – something about the smooth flat surface of a leaf suggests that a much more regular arrangement of cells is likely. Recent work by Sapala et al. (2018) has suggested a function for these unusually shaped cells – in the dispersal of mechanical stresses as the cell grows. Accordingly, the differentiation of these cells can be thought of as a product of the need to minimise stress on the growing cell wall, and the pattern we observe may simply be the outcome of a series of mechanical compromises.

Petal epidermal cells, in contrast, have a very different shape. In most plant species they are regular at the base (loosely rounded, or hexagonal), but with significant expansion in the Z plane, perpendicular to the main surface of the tissue. This results in a conical growth form, and these cells are often called conical cells or conical-papillate cells (as they resemble short papillae). The function of this particular cell morphology has been very well studied, and they are known to play roles in light focusing (which enhances pigmentary colour), surface wettability and floral temperature (Whitney et al., 2011). Their most significant function is thought to be in providing grip to pollinating insects – a series of studies using mutant lines of Antirrhinum majus with flat petal epidermal cells revealed that bees preferred conical-celled flowers, but only when they were made difficult to handle (by being presented vertically, or made to move as if in the wind). The conical cells are thought to provide an opportunity for the pairs of cleft claws on the tarsae of bees to interlock with the petal surface, reducing energy expenditure and improving foraging efficiency (Whitney et al., 2009).

Figure 1. Beverley may2018 c

Fig. 1. Petal conical epidermal cells differ in size and shape in characteristic ways between closely related species. A. Scanning electron micrograph of petal of Veronica chamaedrys. B. Veronica officinalis. C. Veronica prostrate. D. Veronica spicata. All scale bars = 50 µm.

Since conical cells have an important function in pollination and therefore plant fitness, it is perhaps not surprising that they appear to be under tight developmental control. The size and shape of conical cells is subtly different in every plant species, and the detail of petal epidermal cell morphology can be diagnostic in species identification (Fig. 1). Although we have known for over 20 years that the outgrowth of petal epidermal cells into a conical form is regulated by MIXTA-like transcription factors from subgroup 9 of the MYB family (Noda et al., 1994), the detail of how specific parameters of conical cell shape and size are controlled is poorly understood.

In a recent paper van Es et al. (2018) reveal a surprising role for the plant growth regulator ethylene in the differentiation of petal epidermal cells. The authors set out to investigate the control of overall petal cell shape and size, observing that, unlike most vegetative tissues, ‘petals … have a morphology that requires differential regulation of cell proliferation and expansion in the basal and distal parts’. To better understand how this differential regulation of the primary drivers of development occur, they studied the three members of the TCP5-like transcription factor family in Arabidopsis. These proteins represent a sister group to the 5 members of the JAW subfamily of TCP proteins, and together these two groups form the CIN clade of the type II TCP family. Previous studies have shown that the TCP5-like proteins play a role in determining petal size and shape, and also in regulating petal epidermal cell shape (Huang and Irish, 2015).

van Es et al. (2018) began by describing the expression profiles of the three TCP5-like genes. TCP5 itself is expressed during cell elongation stages of petal development, TCP13 later in petal development, and TCP17 at a generally low level. Ectopic expression of TCP5 fused to GFP in the petal epidermis (using an L1-specific promoter) produced smaller petals, suggesting that the epidermis itself is regulating final shape and size of the whole petal. The conical petal epidermal cells of these transgenic lines were bigger and less regular than those of wild type plants. However, a very surprising result was that a tcp5 mutant line, and a tcp5 tcp13 tcp17 triple mutant line, showed similarly perturbed petal epidermal cells – although still loosely cone-shaped they were larger and less regular than wild type, and could not be easily distinguished from each other or from the transgenic line ectopically expressing TCP5.

The mystery of why the loss of function and ectopically expressing lines produced the same phenotype was solved by a transcriptomic analysis, using wild type, the three lines described above, and an inducible epidermis-specific ectopic expression line. The authors discovered that genes encoding enzymes of ethylene synthesis (ACS2 and ACO2) were always down-regulated when the TCP5-like genes were up-regulated, and always up-regulated when the TCP5-like genes were mutated. Similarly, the activity of ethylene response factor genes (ERFs) was down-regulated in the ectopic expression lines and up-regulated in the mutant lines. To confirm these findings the authors showed that ethylene itself was present at higher concentrations in the inflorescences of mutant lines and at lower concentrations in an ectopic expression line. The hypothesis that ethylene was directly regulating petal epidermal cell shape was tested by inhibiting ethylene response using silver thiosulphate application in the mutant lines – this returned the petal epidermal cells to a normal size and shape. Finally, the authors showed that TCP5 binds directly to the ACS2 locus, suggesting a direct regulatory role for this transcription factor family in the ethylene response pathway of Arabidopsis petals.

So, why were the ectopic expression and mutant lines phenotypically so similar? The authors hypothesise that wild type petal epidermal cell shape is a product of wild type levels of ethylene production and perception. When the ethylene pathway is perturbed, in either direction, the tight developmental control of cell differentiation is lost and the epidermal cells grow in a less controlled way, producing larger and less regular shapes. In this scenario ethylene is not a specific regulator of any particular cell shape – less ethylene does not mean smaller cells and more ethylene larger cells, for example – but is instead a signal of ‘normal’, which allows tight regulatory control of cell shape. When perturbation of ethylene signalling tells the plant that all is not well, that tight regulatory control is lost, perhaps in part because the plant’s energies may switch to other activities downstream of ethylene signalling, such as induction of defence responses. It is surprising to find a hormone implicated in such a specific developmental process, but the idea that its role is as a signal of general well-being, allowing development to proceed in a coordinated fashion, fits well with recent developments in understanding plant hormone signalling. It will be interesting to see whether this deregulation of petal epidermal cell differentiation has consequences for pollinator attraction and plant fitness in an animal-pollinated system.

Huang T. and Irish V. 2015. Temporal control of plant organ growth by TCP transcription factors. Current Biology 25, 1765-1770. https://doi.org/10.1016/j.cub.2015.05.024
Noda K, Glover BJ, Linstead P and Martin C. 1994. Flower colour intensity depends on specialized cell shape controlled by a Myb-related transcription factor. Nature 369, 661-664. doi:10.1038/369661a0
Sapala A, Runions A, Routier-Kierzkowska A, Gupta M, Hong L, Hofhuis H, Verger S, Mosca G, Li C, Hay A, Hamant O, Roeder A, Tsiantis M, Prusinkiewicz P and Smith R. 2018. Why plants make puzzle cells and how their shape emerges. eLife 2018;7:e32794 DOI: 10.7554/eLife.32794
Van Es S, Sylveira S, Rocha D, Bimbo A, Martinelli A, Dornelas M, Angenent G and Immink R. 2018. Novel functions of the Arabidopsis transcription factor TCP5 in petal development and ethylene biosynthesis. The Plant Journal doi: 10.1111/tpj.13904
Whitney H, Chittka L, Bruce T and Glover BJ. 2009. Conical Epidermal Cells Allow Bees to Grip Flowers and Increase Foraging Efficiency. Current Biology 19, 1-6. https://doi.org/10.1016/j.cub.2009.04.051
Whitney H, Bennett KMV, Dorling MW, Sandbach L, Prince D, Chittka L and Glover BJ. 2011. Why do so many petals have conical epidermal cells? Annals of Botany 108, 609-616. doi:  10.1093/aob/mcr065

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