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.

References

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

Advertisements
Posted in flowering | Tagged , , , , | Leave a comment

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.

Acknowledgements

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.

References

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

Trejo-Arellano MS, Mahrez W, Nakamura M, Moreno-Romero J, Nanni P, Köhler C, Hennig L. 2017. H3K23me1 is an evolutionary conserved histone modification associated with CG DNA methylation in Arabidopsis. The Plant Journal 90, 293–303. https://doi.org/10.1111/tpj.13489

Turck F, Roudier F, Farrona S, Martin-Magniette ML, Guillaume E, Buisine N, Gagnot S, Martienssen RA, Coupland G, Colot V. 2007. Arabidopsis TFL2/LHP1 specifically associates with genes marked by trimethylation of histone H3 lysine 27. PLoS Genetics 3, 0855–0866. https://doi.org/10.1371/journal.pgen.0030086

Zimmermann P, Hennig L, Gruissem W. 2005. Gene-expression analysis and network discovery using Genevestigator. Trends in Plant Science 10, 407–409. https://doi.org/10.1016/j.tplants.2005.07.003

Zimmermann P, Hirsch-Hoffmann M, Hennig L, Gruissem W. 2004. GENEVESTIGATOR. Arabidopsis microarray database and analysis toolbox. Plant Physiology 136, 2621–2632. https://doi.org/10.1104/pp.104.046367

Zimmermann P, Schildknecht B, Craigon D, Garcia-Hernandez M, Gruissem W, May S, Mukherjee G, Parkinson H, Rhee S, Wagner U, Hennig L. 2006. MIAME/Plant – Adding value to plant microarrray experiments. Plant Methods 2, 1–3. https://doi.org/10.1186/1746-4811-2-1

Posted in flowering | Tagged , , | Leave a comment

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.

Maria-Albani-Figure-may-2018

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.

References

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.

Posted in flowering | Leave a comment

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).

Figure2
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.

References

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

Posted in flowering | Tagged , , , , , , | Leave a comment

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.

References
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

Posted in flowering | Tagged , , , , | Leave a comment

ACME system allows in vivo quantification of cellular mechanical properties within developing plant organs

by Robert G. Franks

North Carolina State University, Raleigh, NC., USA

How organ size and shape are determined in developing organisms remains a key question of interest to developmental biologists. An understanding of the relationships between gene expression and the organ shape requires analyses of relationships between the subcellular, cellular, and organ levels and is often approached through mathematical modeling techniques (Boudon et al., 2015Heer and Martin 2017; Eder et al., 2017). One mechanism that links these multiple organizational scales is the biomechanics of the cells and tissues of the organ itself. In the developing plant it has become clear that the biomechanical properties of the cells, and particularly the cell walls, can be key determinants of both cell and organ shape. However, the relationships between gene expression and cell wall properties and organ shape currently are poorly understood.

The ACME system. Image kindly provided by Sarah Robinson (University of Bern)

In a new article Robinson et al. (2017) describe a new method to measure the mechanical properties of plant tissues. This method has several advantages over previously utilized methods; the measurements can be made in live tissues, have cellular scale resolution, and can be measured in the plane of growth. To measure the mechanical properties of an organ or tissue, the deformation of the tissue must be measured in response to a known, externally applied force. One method to achieve this is to use extensometers to apply a known force to a structure and then measure the deformation.  One disadvantage of this method is that the mechanical properties can only be measured at the tissue level. Robinson et al. have paired a micro-extensometer with a confocal microscope to measure the mechanical properties of the cells within an organ at single-cell resolution. They call this system Automated Confocal Micro-Extensometer or ACME. The authors have made available custom software and instructions for the 3D printing of custom parts required for outfitting a confocal microscope with the ACME system.

Robinson and colleagues use the ACME system to measure the changes in the mechanical properties of the individual cells of the developing hypocotyl in an Arabidopsis seedling upon the application of the plant hormone gibberellic acid that promotes growth in the hypocotyl (Robinson et al. 2017). Changes in the mechanical properties of the hypocotyl cells were found to occur in a gradient along the main axis of the hypocotyl in a pattern that correlated with the observed cellular growth patterns. The experimental determination of the mechanical properties of cells is critical for the successful construction of mathematical models of plant organ growth that are useful for understanding the complex interactions between the cellular and tissue scales that can significantly affect the shape of the mature organ.

The application of this technology to developing floral organs is promising. Although the relatively small size of the Arabidopsis flower may make it unsuitable for mounting on this system without modifications, species that generate larger flowers would provide suitable material for analysis of mechanical properties of cells of the developing floral organs. The quantitative biophysical data at cellular resolution that can be resolved with the ACME technology is well suited for developing mathematical models of floral organ size and shape determination.

References

Boudon F, Chopard J, Ali O, Gilles B, Hamant O, Boudaoud A, Traas J, Godin C. 2015. A computational framework for 3D mechanical modeling of plant morphogenesis with cellular resolution. PLoS Computational Biology 11 (1), e1003950. https://doi.org/10.1371/journal.pcbi.1003950

Eder D, Aegerter C, Basler K. 2017. Forces controlling organ growth and size. Mechanisms of Development 144, 53–61. https://doi.org/10.1016/j.mod.2016.11.005

Heer NC, Martin AC. 2017. “Tension, contraction and tissue morphogenesis. Development  144 (23), 4249–60. 

Robinson S, Huflejt M, Barbier de Reuille P, Braybrook SA, Schorderet M, Reinhardt D, Kuhlemeier C. 2017. An automated confocal micro-extensometer enables in vivo quantification of mechanical properties with cellular resolution. The Plant Cell. https://doi.org/10.1105/tpc.17.00753.

 

Posted in flowering | Leave a comment

DELLA proteins restrict cell divisions by distinct mechanisms

by Jens Sundström
Swedish University of Agricultural Sciences

Breeding cereal crops with reduced stem length, in so-called semi-dwarf varieties, greatly contributed to yield increases associated with the green revolution. However, the semi-dwarf genotype was also associated with reduced inflorescence size. Now, researchers at the John Innes Centre in Norwich, UK, have demonstrated that these traits are regulated by distinct pathways (Serrano-Mislata et al., 2017).  This finding opens up new venues for breeding of semi-dwarf crops without compromising yields by reducing inflorescence size.

Breeding efforts between 1960 and 1985, largely carried out at international public goods institutions such as the International Maize and Wheat Improvement Centre in Mexico (CIMMYT), contributed to a massive increase in crop yields (Pingali, 2012). For instance, yields for wheat in many developing countries have increased almost 200% since the mid -1960s. One of the key traits introgressed in many high yielding varieties is the semi-dwarf genotype. Reduced stem elongation aids reduction of lodging and allows more resources to be allocated to other parts of the plant. Typically, varieties harbouring this trait are mutated in genes affecting responses to the plant hormone gibberellin (GA) (Daviere and Achard, 2013).

Jens blog Dec 2017 3


Fig. 1. Rye usually grow shoulder high. Field of rye-wheat in which the semi-dwarf genotype found in modern wheat varieties has been introgressed. The work by Serrano-Mislata et al. indicates that the semi-dwarf trait can be uncoupled from the reduction of inflorescence size.

Genetic analyses of GA response mutants, primarily carried out in the model species Arabidopsis thaliana have contributed to a working model for GA activity, in which GA acts as an “inhibitor of an inhibitor” (Harberd et al., 2009). DELLA-proteins, which belong to a sub-family of the plant specific GRAS family, act as key repressors of GA responses (Daviere and Achard, 2013). In the absence of GA, DELLA-proteins bind other transcription factors and inhibit their activity. In the presence of GA, the DELLA proteins are degraded and transcription of GA-responsive genes can occur. Plants with mutated DELLA-proteins have pleiotropic phenotypes; for example, reduced seed germination and reduced stem length.

In a recent report, Serrano-Mislata and co-workers (2017) demonstrated that DELLA proteins inhibit shoot growth by negatively regulating cell division rather than cell expansion. The authors provided evidence for this by expressing a stabilized form of DELLA proteins in either the internodes of a stem or in the apical segment of an inflorescence (Fig. 1).

In both cases, expression of the stabilized DELLA-protein resulted in fewer dividing cells as compared to the wild type. These results suggested that DELLA-proteins act as inhibitors of genes involved in cell-cycle regulation or cell division. To test this hypothesis, the authors performed a chromatin immunoprecipitation (ChIP) experiment that allowed them to identify promoters to which the DELLA proteins bind. One of the candidate genes identified in the ChIP experiment encodes a protein belonging to a family of cell cycle inhibitors. Next, the authors made a cross between a knock-out mutant of the cell cycle inhibitor and the line expressing the stabilized form of the DELLA-proteins. Interestingly, the resulting line retained the semi-dwarf phenotype, but the number of cells in the shoot apical meristem was similar to that in the wild type. Hence, cell division were inhibited in the stem internodes but unaffected in the shoot apical meristem, suggesting that DELLA proteins, at least in part, control growth through the activity of cell cycle inhibitors and that this regulation occurs through distinct pathways in different parts of the plant.

While the mechanistic and genetic insights revealed by Serrano-Mislata et al., (2017) are based on work done in Arabidopsis, their study also provides evidence for the presence of  conserved mechanisms in cereals. Hence, their findings may provide future tools for breeding high yielding semi-dwarf cereal varieties, without compromising growth in the seed-bearing parts of the plants.

References
Serrano-Mislata A, Bencivenga S, Bush M, Schiessl K, Boden S, Sablowski R. (2017). DELLA genes restrict inflorescence meristem function independently of plant height. Nature Plants 3(9):749-754. doi:10.1038/s41477-017-0003-y

Pingali PL. (2012). Green revolution: impacts, limits, and the path ahead. Proccedings of the Natural Academy of Sciences, USA 109(31):12302-12308. doi:10.1073/pnas.0912953109

Daviere JM and Achard P. (2013). Gibberellin signaling in plants. Development 140(6):1147-1151. doi: 10.1242/dev.087650

Harberd NP, Belfield E, Yasumura Y. (2009). The angiosperm gibberellin-GID1-DELLA growth regulatory mechanism: how an “inhibitor of an inhibitor” enables flexible response to fluctuating environments. The Plant Cell 21(5):1328-1339. https://doi.org/10.1105/tpc.109.066969

Posted in flowering, Plant breeding | Tagged , , , , , , | Leave a comment

Does gibberellin regulate the trade-off between flowering and runnering in strawberry?

by Timo Hytönen
Department of Agricultural Sciences, University of Helsinki, Helsinki, Finland

Strawberry is one of the most economically-important berry crops in the world. It is a rosette plant that reproduces both generatively and vegetatively through stolons called runners. There is a strong trade-off between flowering and runnering, but runners are important because fruit production is based on clonally propagated plants. In a diploid woodland strawberry, two classical mutants affecting flowering and runnering are known. Recessive mutations in Seasonal flowering locus (SFL) and Runnering locus (R) cause perpetual flowering and runnerless phenotypes, respectively (Figure 1; Brown and Wareing 1965). SFL encodes a major floral repressor, the woodland strawberry homolog of TERMINAL FLOWER1 (Koskela et al. 2012; Iwata et al. 2012), which mediates photoperiodic and temperature signals to control seasonal flowering (Rantanen et al. 2015). The molecular nature of R, however, has remained elusive.

TFL1mutantFigure 1. Classical mutations in woodland strawberry. Recessive mutations in SFL and R genes cause perpetual flowering and inability to produce runners, respectively (left), whereas the plant with dominant alleles (right) is seasonal flowering and produces runners.

Guttridge and Thompson (1964) showed that exogenous gibberellin (GA) treatment induces runner formation and suppresses flowering in a runnerless perpetual flowering mutant of woodland strawberry, indicating that GA may play a role in the trade-off between flowering and runnering. Now, over 50 years later, two studies provided molecular evidence for the role of GA in the control of axillary bud differentiation to runners or branch crowns. Tenreira et al. (2017) identified a gene encoding a GA biosynthetic enzyme, GA20-oxidase, as a plausible candidate for R (see also commentary by Lockhart 2017), and Caruana et al. (2017) reported a single functional DELLA protein that suppresses runner formation in woodland strawberry.

Tenreira et al. (2017) identified FvGA20ox4 as a candidate gene for R by genetic mapping and whole-genome sequencing of a pooled mutant sample. They found a 9-bp deletion in the second exon of the gene and showed by enzyme assays that only non-mutated FvGA20ox4 was able to convert GA12 to GA20 which is the precursor of active GA1. Furthermore, in situ hybridization experiments showed FvGA20ox4 expression in axillary meristems. Together with previous growth regulator experiments (e.g. Guttridge and Thompson 1964; Hytönen et al. 2009), these new data provide strong evidence for FvGA20ox4 being the R gene. However, the role of four other GA20-oxidase encoding genes in axillary bud differentiation remains unresolved. Mutant complementation or targeted mutagenesis of FvGA20ox4 is still required to obtain a final proof.

In another recent study, Caruana et al. (2017) performed a mutagenesis screen in a runnerless woodland strawberry. They found a mutant that continuously produced runners and, using a mapping-by-sequencing strategy, they identified a gene encoding a DELLA growth repressor FvRGA1, as the prime candidate. Next, they generated an inducible dominant negative version of the corresponding DELLA protein and showed that it was able to suppress the formation and elongation of runners in woodland strawberry indicating that a single DELLA protein controls axillary bud fate.

Does GA regulate the trade-off between flowering and runnering in strawberry then? The studies discussed here provide solid evidence for a role of the GA pathway in the control of axillary bud fate, and based on the presented evidence the following working model can be proposed: FvGA20ox4 likely encodes a rate-limiting enzyme of the GA biosynthetic pathway in the axillary bud. In the presence of an active GA20ox enzyme, GA20 is produced and further converted to GA1 by GA3-oxidases; GA1 then causes the degradation of FvRGA1 leading to runner growth, whereas the reduction of GA1 level leads to the accumulation of this DELLA protein and the differentiation of axillary buds to branch crowns. GA also indirectly affects flowering by controlling the number of shoots capable of producing an inflorescence (Tenreira et al. 2017; Caruana et al. 2017), but additional signals are required for floral induction in apical meristems of the crowns.

References

Brown T, Wareing PF. 1965. The genetical control of the everbearing habit and three other characters in varieties of Fragaria vesca. Euphytica 14: 97-112. https://doi.org/10.1007/BF00032819 

Caruana JC, Sittmann JW, Wang W, Liu Z. 2017. Suppressor of Runnerless encodes a DELLA protein that controls runner formation for asexual reproduction in strawberry. Molecular Plant http://dx.doi.org/10.1016/j.molp.2017.11.001

Guttridge CG, Thompson PA. 1964. The effect of gibberellins on growth and flowering of Fragaria and Duchesnea. Journal of Experimental Botany 15: 631–646. https://doi.org/10.1093/jxb/15.3.631.

Hytönen T, Elomaa P, Moritz T, Junttila O. 2009. Gibberellin mediates daylength controlled differentiation of vegetative meristems in strawberry (Fragaria x ananassa Duch.). BMC Plant Biology 9:18. doi:  10.1186/1471-2229-9-18

Iwata H, Gaston A, Remay A, Thouroude T, Jeauffre J, Kawamura K, Oyant LHS, Araki T, Denoyes B, Foucher  F. 2012. The TFL1 homologue KSN is a regulator of continuous flowering in rose and strawberry. Plant Journal 69: 116–125. http://doi.org/10.1111/j.1365-313X.2011.04776.x

Koskela E, Mouhu K, Albani MC, Kurokura T, Rantanen M, Sargent D, Battey NH, Coupland G, Elomaa P, Hytönen T. 2012. Mutation in TERMINAL FLOWER1 reverses the photoperiodic requirement for flowering in the wild strawberry, Fragaria vesca. Plant Physiology 159: 1043-1054. http://doi.org/10.1111/tpj.12809

Lockhart J. 2017. Flowering versus runnering: uncovering the protein behind a trait that matters in strawberry. Plant Cell 29: 2080-2081. https://doi.org/10.1105/tpc.17.00709

Rantanen M, Kurokura T, Jiang P, Mouhu K, Hytönen T. 2015. Strawberry homolog of TERMINAL FLOWER1 integrates photoperiod and temperature signals to inhibit flowering. Plant Journal 82: 163-173. http://doi.org/10.1111/tpj.12809

Tenreira T, Lange MJP, Lange T, Bres C, Labadie M, Monfort A, Hernould M, Rothan C, Denoyes B. 2017. A specific gibberellin 20-oxidase dictates the flowering-runnering decision in diploid strawberry. Plant Cell 29: 2168-2182. https://doi.org/10.1105/tpc.16.00949

Posted in flowering | Tagged , , , , , , | Leave a comment

Enhancing the possibilities of promoter research

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

Developmental regulatory genes have played a pivotal role during the evolution and domestication of plants. From a molecular genetics perspective, our understanding of those regulators is largely driven by the analysis of full loss-of-function mutants. This has provided profound insights into gene regulatory circuits governing developmental processes. However, from the analysis of evolutionary and domestication processes it is also evident that in many cases not loss-of-function mutations but variations in gene expression patterns and expression strength caused phenotypic change (Meyer and Purugganan, 2013). This is a least partly due to the pleiotropic functions of many developmental regulators: complete loss-of-function mutations often yield so dramatic phenotypes that the fitness of the plant is severely impaired, hence the relevance of null mutants during evolution and domestication is limited.

From the analysis of a few exemplary cases we know that the modularity of promoter architectures is one important component of pleiotropic gene functions. For example, APETALA3, a transcription factor coding gene that controls petal as well as stamen development, has specific enhancer regions that are required for AP3 expression in stamens (Hill et al., 1998). Studying promoter functions at the molecular level is therefore extremely valuable for our understanding of evolutionary and domestication processes. Detailed promoter studies are usually quite laborious, however, and are largely limited to a few genetic model plants. Promoter studies often encompass the identification of putative enhancer elements using in silico approaches and the expression of a reporter gene under control of a mutated promoter version lacking those elements (or containing multiples of them). Altered expression pattern of the reporter gene informs about the function of putative enhancer elements (Hernandez-Garcia and Finer, 2014). Although this approach has been very successful, in silico predictions do not always identify critical promoter elements (Hong et al., 2003). In many cases, a more unbiased approach to study promoter functions might be at least equally promising.

Figure_Tomatoes

Traditional breeding generated a large variation in tomato fruit size. The method presented by Rodríguez-Leal et al. (2017) has the potential to modify and fine-tune quantitative traits in just a few generations.

A recent paper by Rodríguez-Leal et al. (2017) presents a method that will substantially facilitate such an unbiased molecular genetic analysis. Using tomato fruit size as a model system, the authors target the promoter of Solanum lycopersicum CLAVATA3 (SlCLV3), a gene know to be involved in fruit size regulation, using CRISPR-Cas9. However, unlike more conventional CRISPR-Cas9 approaches that aim to generate full loss-of-function mutants, the authors designed not just one but eight guide RNAs that spanned a 2 kb range of the putative promoter region of SlCLV3. Because of variations in guide RNA directed cleavage activities and subsequent repair processes, many types of mutations can be induced, ranging from large promoter deletions to inversions, small deletions and single nucleotide substitutions (Rodríguez-Leal et al., 2017).

As the analysis of the mutant plants proved difficult because different mutations are introduced in the two alleles of the target gene, the authors devised a genetic screen in which they crossed a wild-type plant with a transgenic plant that expressed the eight guide RNAs and Cas9, and also carried a full loss-of-function allele of SlCLV3. In the resulting progeny, mutations were induced by CRISPR-Cas9 in the wild-type allele. In this set-up, the phenotypic effects of even mutations causing only subtle phenotypic changes are relatively easy to detect as the second SlCLV3 allele is a null allele. Thus, the authors essentially developed an elegant yet simple procedure to randomly mutagenize one particular locus and screen for phenotypic consequences of the mutations. Indeed, using this system, it was possible to create an allelic series of 14 SlCLV3 mutant alleles that showed a continuous variation in carpel number (thus leading to a variation in fruit size) (Rodríguez-Leal et al., 2017).

This method is potentially of huge importance for dissection promoter functions. It may be possible to quickly generate a large number of random mutations and analyse their phenotypic effect in a streamlined way. This will prove to be very powerful to untangle the functional elements in promoters. In turn, our understanding of how master regulators of development may have contributed to morphological evolution may be substantially increased. For example, it has been proposed that alterations in the expression of floral homeotic transcription factors contributed to floral diversity (reviewed by Theißen and Melzer, 2007). The approach presented by Rodríguez-Leal et al. (2017) constitutes a promising avenue to test whether and how promoter mutations can induce such phenotypic changes. Theoretically, a large number of mutant alleles can be generated from one transgenic plant. Thus, the approach may also be suitable for phylogenetically informative non-model plants that are difficult to transform.
Last but not least, plant domestication and crop improvement also often proceeds via variations in quantitative traits. Rodríguez-Leal et al. (2017) demonstrated that a substantial variation in tomato fruit size can be obtained in just a few generations, bypassing years of breeding efforts. It will be interesting to see to which extent the same method can be applied to other quantitative traits in crops.

References

Hernandez-Garcia CM, Finer JJ. 2014. Identification and validation of promoters and cis-acting regulatory elements. Plant Science 217, 109-119. https://doi.org/10.1016/j.plantsci.2013.12.007

Hill TA, Day CD, Zondlo SC, Thackeray AG, Irish VF. 1998. Discrete spatial and temporal cis-acting elements regulate transcription of the Arabidopsis floral homeotic gene APETALA3. Development 125, 1711-1721.

Hong RL, Hamaguchi L, Busch MA, Weigel D. 2003. Regulatory elements of the floral homeotic gene AGAMOUS identified by phylogenetic footprinting and shadowing. The Plant Cell 15, 1296-1309. https://doi.org/10.1105/tpc.009548

Meyer RS, Purugganan MD. 2013. Evolution of crop species: genetics of domestication and diversification. Nature Reviews Genetics 14, 840-852. https://doi.org/10.1038/nrg3605

Rodríguez-Leal D, Lemmon ZH, Man J, Bartlett ME, Lippman ZB. 2017. Engineering quantitative trait variation for crop improvement by genome editing. Cell 171, 470-480. https://doi.org/10.1016/j.cell.2017.08.030

Theißen G, Melzer R. 2007. Molecular mechanisms underlying origin and diversification of the angiosperm flower. Annals of Botany 100, 603-619. https://doi-org./10.1093/aob/mcm143

Posted in flowering | Tagged , , , , , , , | Leave a comment

Untangling complexity: shedding a new light on LEAFY and APETALA1 interactions

by Leonie Verhage and Francois Parcy
Institut de Biosciences et Biotechnologies de Grenoble (France)

Ever since their discovery almost 30 years ago, the transcription factors LEAFY (LFY) and APETALA1 (AP1) (together with its paralog CAULIFLOWER (CAL)) have been extensively studied for their roles in floral transition. Early genetic and molecular experiments indicated that LFY and AP1/CAL were partly redundant and partly complementary in the process of floral initiation, and numerous subsequent studies fit this model (see Denay et al., 2017 and Wils and Kaufmann, 2017 for recent reviews). However, combining a set of new experiments with published datasets, Goslin and colleagues manage to stir up the prevalent view (Goslin et al., 2017).

FrancoisParcyImage2

Scanning electron micrograph of an ap1 cal mutant. Floral meristems are transformed into proliferative inflorescence meristems. This mutant background was used by Goslin and colleagues. (Image courtesy of Marie le Masson and Christine Lancelon Pin)

To unravel the redundancy of LFY and AP1/CAL, the authors utilized a mutant line harboring a 35S:LFY-GR construct in an ap1/cal background (Wagner et al., 1999). With this line, they performed induction experiments and microarray analysis, in the same way as was previously performed with 35S:AP1-GR in the ap1/cal background (Kaufmann et al., 2010), to make the datasets comparable. This allowed them to compare the downstream genes that are regulated by LFY in the absence of AP1/CAL to genes that are regulated by AP1 in the presence of LFY.

Among the many things uncovered by these analyses, a few were expected and many completely unanticipated.

As already reported by Winter et al., 2015, there is a large overlap between the genes that are differentially regulated upon induction of LFY-GR or AP1-GR. It is likely that this represents true redundancy, where LFY and AP1 can regulate genes in the same way, independent of each other. However, due to a lack of experiments where AP1 is induced in the absence of LFY, it cannot be excluded that this set of genes can be regulated by LFY alone, or by LFY and AP1 together.

More surprisingly, many direct targets of LFY were found to be down-regulated, whereas most of the well-known targets are induced (such as the floral organ identity genes or the LATE MERISTEM IDENTITY genes).

Interestingly, a subset of genes showed differential expression in ap1 cal upon AP1 induction but not upon induction of LFY. By comparing these genes with previously published ChIP-seq data of LFY, the authors could identify a set of genes to which LFY is able to bind, but that are not differentially regulated in absence of AP1. This was the case for APETALA3 (AP3) and AGAMOUS (AG), consistent with a previous report showing that AP1 can act on these genes (Ng and Yanofsky, 2001). Hence, for regulation of these B- and C- type floral organ identity genes, LFY and AP1 appear to act interdependently.

The most surprising result, however, was the presence of genes that are differentially expressed upon LFY or AP1 induction, but in different directions. Apparently, besides acting redundantly or interdependently, LFY and AP1 can also act antagonistically. Notably, this turned out to be the case for several genes involved in inflorescence meristem identity, including TERMINAL FLOWER1 (TFL1). Contrary to the longstanding belief that AP1 and LFY are both repressors of TFL1, only AP1 repressed TFL1, whereas LFY actually activates this gene. It is not completely clear why LFY would up-regulate a gene that inhibits floral meristem identity. The authors speculate that it might be a way to better define the floral transition, so that it occurs only when AP1 is expressed high enough to overcome TFL1.

Goslin et al.  paper is a nice example of how to combine new experiments and existing datasets in a time with ever growing amounts of genome-wide data, with a surprising outcome. Two transcription factors that were long thought to function similarly in initiation of flower formation suddenly turn out to have a much more intriguing relationship, posing many new questions. When LFY and AP1 act together, the biochemical basis of their interaction is elusive. They might be part of the same regulatory complex, especially since their binding sites have been reported to be adjacent (Winter et al., 2015), but a direct interaction between the two proteins has not been observed. Analysis by targeted proteomics has uncovered AP1 interactors in floral tissue (Smaczniak et al., 2012), but has never been analyzed in earlier tissue in which LFY is expressed. Another question is how LFY and AP1 sometimes work together, and sometimes do not, sometimes activate and other times repress. One possibility is that there might be spatio-temporal differences in expression of interaction partners of LFY and AP1 (see also the previous Flowering Highlight on Spatially resolved floral transcriptome profiling by Aalt-Jan van Dijk). Altogether, there is still a lot to be understood about these two ‘very well known’ regulators!

References

Denay G, Chahtane H, Tichtinsky G, Parcy F. 2017. A flower is born: an update on Arabidopsis floral meristem formation. Current Opinion in Plant Biology 35, 15–22. https://doi.org/10.1016/j.pbi.2016.09.003

Goslin K, Zheng B, Serrano-Mislata A, et al. 2017. Transcription Factor Interplay between LEAFY and APETALA1/CAULIFLOWER during Floral Initiation. Plant Physiology 174, 1097–1109. https://doi.org/10.1104/pp.17.00098

Kaufmann K, Wellmer F, Muiño JM, et al. 2010. Orchestration of floral initiation by APETALA1. Science 328, 85–89.  https://doi.org/10.1126/science.1185244

Ng M, Yanofsky MF. 2001. Activation of the Arabidopsis B class homeotic genes by APETALA1. The Plant Cell 13, 739–753. https://doi.org/10.1105/tpc.13.4.739

Smaczniak C, Immink RGH, Muiño JM, et al. 2012. Characterization of MADS-domain transcription factor complexes in Arabidopsis flower development. Proceedings of the National Academy of Sciences 109, 1560–1565. https://doi.org/10.1073/pnas.1112871109

Wagner D, Sablowski RW, Meyerowitz EM. 1999. Transcriptional activation of APETALA1 by LEAFY. Science 285, 582–584.  https://doi.org/10.1126/science.285.5427.582

Wils CR, Kaufmann K. 2017. Gene-regulatory networks controlling inflorescence and flower development in Arabidopsis thaliana. BBA – Gene Regulatory Mechanisms 1860, 95–105. https://doi.org/10.1016/j.bbagrm.2016.07.014

Winter CM, Yamaguchi N, Wu M-F, Wagner D. 2015. Transcriptional programs regulated by both LEAFY and APETALA1 at the time of flower formation. Physiologia Plantarum 155, 55–73. https://doi.org/10.1111/ppl.12357

Posted in flowering | Tagged , , , | Leave a comment