Plant Physiology

Ort, Donald R.; Grennan, Aleel K.

doi: 10.1104/pp.111.900428pmid: 22213247

Plant Physiology is proud to present in our first issue of 2012 our annual Founders Review article, this year authored by Detlef Weigel. Our goal for the Founders Review is to highlight the work of an internationally eminent plant biologist and to present an in-depth current perspective of a highly active area of research, which was met last year with a review from Christine Foyer entitled “Ascorbate and Glutathione: The Heart of the Redox Hub” (Foyer and Noctor, 2011). This year's article by Detlef Weigel, entitled “Natural Variation in Arabidopsis: From Molecular Genetics to Ecological Genomics,” explores the potential that insights gained from the study of genetic variation hold to inform not only Arabidopsis (Arabidopsis thaliana) biology but plant biology in general. Highlighted is the power as well as the tools available for the study of interactions among divergent genomes, which can act to either enhance or reduce the partitioning of genetic diversity into different lineages and in turn into different species. With potentially important practical applications to hybrid crop development, Dr. Weigel shows how the explosion in information about the genome-wide and population-specific distribution of sequence polymorphisms enables a more informed and systematic basis for choices of genotype combinations that have great promise to accelerate the pace with which hybrid performance can be improved. Detlef Weigel is the director of the Department of Molecular Biology at the Max Planck Institute for Developmental Biology in Tuebingen, Germany. He also holds adjunct professorships at The Salk Institute for Biological Studies (La Jolla, CA) and Eberhard Karls University (Tuebingen, Germany), where he did his PhD work with Herbert Jäckle on Drosophila development, teasing apart the role of the homeotic gene fork head. His entry into plant research came in Elliot Meyerowitz's group at the California Institute of Technology (Pasadena, CA), where he started his work on plant development. He continued his study on plant development as a faculty member at The Salk Institute for Biological Studies (La Jolla, CA), where he made important inroads into understanding floral patterning, initiation, and induction. Now as director of the Department of Molecular Biology at the Max Planck Institute (Tuebingen, Germany), he is also focusing on evolutionary genomics, which has powerfully complemented his group's continuing work in development. The 1001 Genomes Project (http://1001genomes.org/) for Arabidopsis was born out of this project with the “goal to discover the whole-genome sequence variation in 1001 strains (accessions) of the reference plant Arabidopsis thaliana.” Detlef's research career has garnered many honors, including the National Science Foundation Young Investigator Award and American Society of Plant Biologists Charles Albert Shull Award. He has been elected as a member of the European Molecular Biology Organization, Academia Europaea, German National Academy of Sciences Leopoldina, U.S. National Academy of Sciences, and the Royal Society of London. He has also been elected a corresponding member of the Heidelberg Academy of Sciences and Humanities as well as a fellow of the American Association for the Advancement of Science. Detlef has received the Dieter Rampacher Award of the Max Planck Society, the Gottfried Wilhelm Leibniz Award of the German Research Council (Deutsch Forschungsgemeinschaft), the Otto Bayer Award of the Bayer Foundations, and the State Research Award of Baden-Württemberg. Plant Physiology is exceptionally pleased to recognize Dr. Weigel by selecting him to author the 2012 Founders Review. LITERATURE CITED Foyer CH Noctor G ( 2011 ) Ascorbate and glutathione: the heart of the redox hub . Plant Physiol 155 : 2 – 18 Google Scholar Crossref Search ADS PubMed WorldCat © 2012 American Society of Plant Biologists. All rights reserved. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Weigel, Detlef

doi: 10.1104/pp.111.189845pmid: 22147517

One of the most remarkable biological insights in the past 30 years has been that many genetic programs for complex traits, such as flower or limb development, are shared across broad groups of organisms. These conserved pathways in turn can be tuned to produce tremendous phenotypic differences, not only between, but also within species. Intraspecific variation is often quantitative, one example being the onset of flowering, although there is also qualitative variation, such as in the ability to resist pathogens. While many tools for quantitative genetics were developed by breeders, the model plant Arabidopsis (Arabidopsis thaliana) was adopted for studying the genetic architecture of quantitative traits soon after molecular markers for mapping became available (Chang et al., 1988; Nam et al., 1989). The species belongs to a small genus with nine members. Different from most of its congeners, Arabidopsis is self-compatible, and its life cycle can be as short as 6 weeks, both properties that greatly facilitate genetic studies. Its native range is considered to be continental Eurasia and North Africa (Al-Shehbaz and O'Kane, 2002), but it has been introduced throughout much of the rest of the world, especially around the northern hemisphere. The potential of genetic variation to inform many different areas of Arabidopsis biology was most strongly advocated by Maarten Koornneef and his students. From the mid-1990s, they published both an impressive number of original research articles on this subject and a series of influential review articles that advertised the impact that the study of natural genetic variation could have on questions of both development and physiology (Alonso-Blanco and Koornneef, 2000; Koornneef et al., 2004). Today, the study of natural variation in Arabidopsis continues to reveal new biology. In addition, the entire genus is increasingly being used to address fundamental questions of evolution (Mitchell-Olds and Schmitt, 2006; Bergelson and Roux, 2010). Some of the problems studied are: How, and how frequently, do new variants arise? Why do some variants rise to high frequency, while others are eliminated? And why are certain combinations of new variants incompatible with each other? Here, I will first give an overview of the tools and resources available for the study of natural variation in Arabidopsis. Next, I will present a few examples of how our knowledge of important biological processes has been improved through insights obtained from varieties other than the common laboratory accessions. Where similar or contrasting findings have been made in other species of the Brassicaceae, to which Arabidopsis belongs, I will mention these. The article concludes with a discussion of recent work that aims to integrate evolutionary and ecological studies with functional tests. A final introductory note: Natural accessions of Arabidopsis have in the past often been referred to as “ecotypes.” This term implies that a line has a unique ecology and is adapted to specific environments, as opposed to differing only in genotype from other varieties (Turesson, 1922b). Preferable is the neutral term accession, which merely means that a unique identifier in a collection has been assigned (Alonso-Blanco and Koornneef, 2000). GENETIC TOOL KIT FOR THE STUDY OF NATURAL VARIATION Experimental Populations for Genetic Mapping Accessions of Arabidopsis vary in a number of traits (Fig. 1; Table I). The most general way to identify genes is by crossing two accessions, which may or may not have a different phenotype, but produce nonuniform F2 progeny. In the F2 or later generations, specific phenotypes are then associated with segregating genetic markers that distinguish the contributions from the parental genomes. When phenotypic classes are not discrete, this is done using the methods of quantitative trait locus (QTL) mapping (Falconer and Mackay, 1996). Figure 1. Open in new tabDownload slide Gross morphological variation in Arabidopsis and relatives. A, Variation between Arabidopsis accessions. On top, vegetative rosettes of accessions grown for 4 weeks in long days are shown. They vary in rosette diameter and compactness, leaf shape, and tissue necrosis or onset of senescence. Similarly, variation in size and shape of individual leaves, in this case the sixth in the rosette, is apparent in the 10 examples shown on the bottom left. Finally, differences in overall architecture are illustrated with five plants. On the left is an early flowering accession with few rosette leaves. The next two flower later, but the second one from the left has reduced apical dominance. Finally, the two accessions on the right have similarly tall main inflorescences but differ in the number of secondary inflorescences. The appearance on the far right is common among wild-grown plants. B, Some characters, such as flower size and fruit shape, vary relatively little within Arabidopsis, but more dramatic variation is found in comparison with closely related taxa, such as Capsella rubella (left) and A. lyrata (right). Images courtesy of Eunyoung Chae, Sang-Tae Kim, and George Wang. Figure 1. Open in new tabDownload slide Gross morphological variation in Arabidopsis and relatives. A, Variation between Arabidopsis accessions. On top, vegetative rosettes of accessions grown for 4 weeks in long days are shown. They vary in rosette diameter and compactness, leaf shape, and tissue necrosis or onset of senescence. Similarly, variation in size and shape of individual leaves, in this case the sixth in the rosette, is apparent in the 10 examples shown on the bottom left. Finally, differences in overall architecture are illustrated with five plants. On the left is an early flowering accession with few rosette leaves. The next two flower later, but the second one from the left has reduced apical dominance. Finally, the two accessions on the right have similarly tall main inflorescences but differ in the number of secondary inflorescences. The appearance on the far right is common among wild-grown plants. B, Some characters, such as flower size and fruit shape, vary relatively little within Arabidopsis, but more dramatic variation is found in comparison with closely related taxa, such as Capsella rubella (left) and A. lyrata (right). Images courtesy of Eunyoung Chae, Sang-Tae Kim, and George Wang. Traits studied by natural variation in Arabidopsis Table I. Traits studied by natural variation in Arabidopsis For references, see Supplemental Table S1. Trait Gene(s) Cloned?a Aluminum content N Autonomous endosperm development N Auxin response N Carbohydrate availability and content N Cell wall composition N Chiasma frequency N Chromatin compaction N Circadian clock C Copper tolerance Y Crowding response C Disease resistance Y Drought response N Editing and processing of mitochondrial transcripts Y Elemental composition Y/N Flowering time Y Freezing tolerance Y Fruit number C Genetic robustness N Glucosinolate content Y Inflorescence replacement (mimicking grazing) N Jasmonate response N Leaf senescence N Leaf, inflorescence, and flower morphology Y Lethality in interploidy crosses N Life history traits other than flowering and growth N Light response Y Molybdenum content Y Nitrogen availability response N Oil content N Osmotic and salt stress tolerance N Phosphate content N Phytate content N Recruitment of bacterial rhizosphere communities N Root hydraulics N Root system size N Salicylic acid response N Salinity tolerance N Seed dormancy Y Seed germination, longevity N Seed lipids N Seed mucilage composition Y Sinapoylmalate biosynthesis Y Sodium accumulation Y Stomata density N Submergence tolerance N Sulfate content Y Terpene biosynthesis Y Thermal dissipation N Trichome density Y Zinc response Y Trait Gene(s) Cloned?a Aluminum content N Autonomous endosperm development N Auxin response N Carbohydrate availability and content N Cell wall composition N Chiasma frequency N Chromatin compaction N Circadian clock C Copper tolerance Y Crowding response C Disease resistance Y Drought response N Editing and processing of mitochondrial transcripts Y Elemental composition Y/N Flowering time Y Freezing tolerance Y Fruit number C Genetic robustness N Glucosinolate content Y Inflorescence replacement (mimicking grazing) N Jasmonate response N Leaf senescence N Leaf, inflorescence, and flower morphology Y Lethality in interploidy crosses N Life history traits other than flowering and growth N Light response Y Molybdenum content Y Nitrogen availability response N Oil content N Osmotic and salt stress tolerance N Phosphate content N Phytate content N Recruitment of bacterial rhizosphere communities N Root hydraulics N Root system size N Salicylic acid response N Salinity tolerance N Seed dormancy Y Seed germination, longevity N Seed lipids N Seed mucilage composition Y Sinapoylmalate biosynthesis Y Sodium accumulation Y Stomata density N Submergence tolerance N Sulfate content Y Terpene biosynthesis Y Thermal dissipation N Trichome density Y Zinc response Y a Y, Yes; N, no; C, likely candidates. Open in new tab Table I. Traits studied by natural variation in Arabidopsis For references, see Supplemental Table S1. Trait Gene(s) Cloned?a Aluminum content N Autonomous endosperm development N Auxin response N Carbohydrate availability and content N Cell wall composition N Chiasma frequency N Chromatin compaction N Circadian clock C Copper tolerance Y Crowding response C Disease resistance Y Drought response N Editing and processing of mitochondrial transcripts Y Elemental composition Y/N Flowering time Y Freezing tolerance Y Fruit number C Genetic robustness N Glucosinolate content Y Inflorescence replacement (mimicking grazing) N Jasmonate response N Leaf senescence N Leaf, inflorescence, and flower morphology Y Lethality in interploidy crosses N Life history traits other than flowering and growth N Light response Y Molybdenum content Y Nitrogen availability response N Oil content N Osmotic and salt stress tolerance N Phosphate content N Phytate content N Recruitment of bacterial rhizosphere communities N Root hydraulics N Root system size N Salicylic acid response N Salinity tolerance N Seed dormancy Y Seed germination, longevity N Seed lipids N Seed mucilage composition Y Sinapoylmalate biosynthesis Y Sodium accumulation Y Stomata density N Submergence tolerance N Sulfate content Y Terpene biosynthesis Y Thermal dissipation N Trichome density Y Zinc response Y Trait Gene(s) Cloned?a Aluminum content N Autonomous endosperm development N Auxin response N Carbohydrate availability and content N Cell wall composition N Chiasma frequency N Chromatin compaction N Circadian clock C Copper tolerance Y Crowding response C Disease resistance Y Drought response N Editing and processing of mitochondrial transcripts Y Elemental composition Y/N Flowering time Y Freezing tolerance Y Fruit number C Genetic robustness N Glucosinolate content Y Inflorescence replacement (mimicking grazing) N Jasmonate response N Leaf senescence N Leaf, inflorescence, and flower morphology Y Lethality in interploidy crosses N Life history traits other than flowering and growth N Light response Y Molybdenum content Y Nitrogen availability response N Oil content N Osmotic and salt stress tolerance N Phosphate content N Phytate content N Recruitment of bacterial rhizosphere communities N Root hydraulics N Root system size N Salicylic acid response N Salinity tolerance N Seed dormancy Y Seed germination, longevity N Seed lipids N Seed mucilage composition Y Sinapoylmalate biosynthesis Y Sodium accumulation Y Stomata density N Submergence tolerance N Sulfate content Y Terpene biosynthesis Y Thermal dissipation N Trichome density Y Zinc response Y a Y, Yes; N, no; C, likely candidates. Open in new tab Because marker analysis used to be very tedious and expensive, substantial efforts were invested early on into producing recombinant inbred lines (RILs), which constitute immortal populations in which recombinant chromosomes have been fixed through inbreeding (Reiter et al., 1992; Lister and Dean, 1993; Fig. 2). RILs, which were first developed in mice (Bailey, 1971), have the advantage that they need to be genotyped only once but can be phenotyped repeatedly for many different traits and under different environmental conditions. An advantage of Arabidopsis is its self-compatibility, so that inbred lines can be easily generated by selfing and single-seed descent. Around 60 RIL populations are available from the stock centers as of the time this article is written (end of 2011; http://www.inra.fr/internet/Produits/vast/RILs.htm, http://www.arabidopsis.org/ and http://www.arabidopsis.info/). Importantly, the lengthy inbreeding process can now be bypassed through a revolutionary technology introduced by the laboratory of Simon Chan. This method allows the facile production of doubled haploid plants from recombinant populations (Ravi and Chan, 2010). Figure 2. Open in new tabDownload slide Populations for mapping genes causing trait variation. Colors indicate contribution from different parental accessions. Only one chromosome pair is shown for each individual. HIF individuals are derived from RILs, in which a small portion of the genome is still heterozygous. Figure 2. Open in new tabDownload slide Populations for mapping genes causing trait variation. Colors indicate contribution from different parental accessions. Only one chromosome pair is shown for each individual. HIF individuals are derived from RILs, in which a small portion of the genome is still heterozygous. Even after five or six generations of inbreeding, which is customary for RILs, a small percent of the genome remains heterozygous. This turns out to have its own benefits. In such a heterogeneous inbred family (HIF), only a small portion of the genome segregates for the two parental alleles (Tuinstra et al., 1997). Additional recombinants that further reduce an interval of interest are easily derived from heterozygous HIF individuals, as are near isogenic lines (NILs) that are homozygous for either parental allele at this locus. A disadvantage of HIF-derived NILs is that each HIF has a unique genome composition and that one can therefore not easily place several QTL in a common genetic background. NILs that carry only a small genomic region from one parent in a background that is otherwise composed of the genome of the other parent can also be generated directly by repeated backcrosses (Fig. 2). Such NILs, pioneered in crops where they are also called introgression lines (Seevers et al., 1971; Rhodes et al., 1989; Eshed and Zamir, 1995), are powerful for systematic analyses of interactions between genes from different genomes, although epistatic interactions among alleles from the introgressed genome are mostly lost. The properties of NIL sets are in many ways complementary to those of RILs, and they are particularly useful when introgression is performed in two directions (Törjék et al., 2008). NILs can identify QTL of smaller effect but with lower resolution than RIL populations (Falconer and Mackay, 1996; Keurentjes et al., 2007). Although the genomes of RILs already contain more recombination events than F2 populations and therefore afford higher mapping resolution, this can be further increased with advanced intercross RILs, in which individuals from the F2 and later generations are intermated before inbred lines are derived (Darvasi and Soller, 1995; Balasubramanian et al., 2009). Other approaches involve the use of multiple parents, as in the MAGIC (for multiple advanced generation intercross) and AMPRIL (for Arabidopsis multiparent RIL) populations (Fig. 2; Kover et al., 2009; Huang et al., 2011). The MAGIC design is more elaborate and generates more recombination events per line than the AMPRIL strategy, but the founder genomes are less evenly represented in the final lines. Mapping in either population is more complex than with RILs, but with a sufficiently high density of intermediate frequency markers, one can infer the most likely local founder genotype. Even more so than simple F2 or RIL populations, AMPRILs and MAGIC lines are likely to contain genotypic combination not found in the wild. QTL mapping accuracy increases with the MAGIC and AMPRIL populations, but not all possible QTL that can be found in pairwise crosses between some of the parents are detected. An alternative would be to combine the most informative subsets of RIL populations and to perform a joint QTL analysis. Especially when genotyped with common markers, a joint analysis can confirm common QTL (Bentsink et al., 2010; Salomé et al., 2011b). Some of the advantages of using RIL-type populations will continue to apply in the future. Trait values, especially those with low heritability, can be estimated more precisely due to replication (Soller and Beckmann, 1990; Mackay, 2001). Perhaps most importantly, one can study correlations between different traits, which can reveal fitness trade-offs, and reaction norms, the response of a specific genotype to different environments. However, not every geographic region where Arabidopsis is found is fairly represented in the available RIL populations because geographic sampling of Arabidopsis has so far been rather uneven (Fig. 3). Thus, forward genetics in additional material, even if composed mostly of F2 populations, will likely be informative. Fortunately, with reduced representation approaches such as restriction-associated DNA sequencing (RAD-seq) or genotyping-by-sequencing (Baird et al., 2008; Elshire et al., 2011) and multiplexing of genomic DNA from many individuals (currently, at least 96), costs for interrogating thousands of markers have dropped to a few U.S. dollars. Figure 3. Open in new tabDownload slide Distribution of over 7,000 Arabidopsis accessions collected from the wild and available in the stock center or soon-to-be-released collections. Western and southern Europe, including Great Britain, is heavily overrepresented, although sampling is not even. Accessions from the presumed native range are in yellow and likely introductions in red. Whether the distribution across China to Japan is continuous with the native range is unclear. Arabidopsis has been reported in additional locales, such as South Korea, and several African countries (Alonso-Blanco and Koornneef, 2000). Maps courtesy of George Wang. Figure 3. Open in new tabDownload slide Distribution of over 7,000 Arabidopsis accessions collected from the wild and available in the stock center or soon-to-be-released collections. Western and southern Europe, including Great Britain, is heavily overrepresented, although sampling is not even. Accessions from the presumed native range are in yellow and likely introductions in red. Whether the distribution across China to Japan is continuous with the native range is unclear. Arabidopsis has been reported in additional locales, such as South Korea, and several African countries (Alonso-Blanco and Koornneef, 2000). Maps courtesy of George Wang. Finally, a general caveat when performing conventional genetic mapping is that chiasma frequencies differ between accessions (Sanchez-Moran et al., 2002). Data from F2 populations also support the conclusion that recombination rates vary depending on the cross (Salomé et al., 2011a). Thus, the ease with which loci are mapped will differ from cross to cross, even more so if structural variants interfere with recombination near the loci of interest. Identification and Validation of Causal Genes and Polymorphisms After a genomic interval underlying phenotypic differences has been identified, there are various options to track down the responsible gene, assuming that only a single gene is causal. Different from induced mutations, simply resequencing a region with dozens or more genes is on its own generally not informative because of the high number of polymorphisms that distinguish an arbitrary pair of accessions, about 1 in every 200 bp. Fortunately, compared to other multicellular organism in which natural variation is studied, Arabidopsis has the enormous advantage that almost all accessions are quite easily transformed by dipping flowering plants into a suspension of Agrobacterium tumefaciens containing a T-DNA vector with the transgene of interest (Clough and Bent, 1998). If the final mapping interval does not contain a gene previously implicated in the trait of interest, one of the first steps will often be to investigate whether null alleles affect this trait. For the vast majority of genes, T-DNA insertion lines in the reference Columbia-0 (Col-0) background are available from the stock centers (http://arabidopsis.org, http://arabidopsis.info; for review, see Alonso and Ecker, 2006). The most straightforward approach to investigate the activity of individual genes in other genetic backgrounds is gene silencing, and collections of vectors for knocking down a large fraction of genes present in the reference genome are available, both for conventional hairpin RNA interference and artificial microRNAs (amiRNAs; for review, see Ossowski et al., 2008b). Gene silencing is a convenient tool to test the relative activity of alleles, an approach that we have called quantitative knockdown (Schwartz et al., 2009). It is conceptually related to quantitative complementation, where different alleles are examined in the hemizygous state, by crossing a homozygous strain to a tester that carries a knockout allele of the gene of interest (Mackay, 2001; Fig. 4). Figure 4. Open in new tabDownload slide Quantitative complementation and knockdown to determine whether QTL are allelic to a candidate gene. Both tests rely on quantitative comparisons between genotypes; the dashed boxes indicate phenotypic differences to the genotype to the left. In a quantitative complementation test, one determines whether the two QTL alleles, Q1 and Q2, are differentially affected when heterozygous with the wild-type (wt) or mutant (mut) allele of a candidate gene (Mackay, 2001). If the QTL alleles respond differently, i.e. if in this example only Q1 complements the mutant phenotype, the candidate gene and the QTL are probably allelic. Similarly, in a quantitative knockdown experiment, a differential effect of an amiRNA (amiR) against the candidate gene indicates that the Q1 allele has lower activity than Q2 and that the candidate gene is likely responsible for the QTL. Figure 4. Open in new tabDownload slide Quantitative complementation and knockdown to determine whether QTL are allelic to a candidate gene. Both tests rely on quantitative comparisons between genotypes; the dashed boxes indicate phenotypic differences to the genotype to the left. In a quantitative complementation test, one determines whether the two QTL alleles, Q1 and Q2, are differentially affected when heterozygous with the wild-type (wt) or mutant (mut) allele of a candidate gene (Mackay, 2001). If the QTL alleles respond differently, i.e. if in this example only Q1 complements the mutant phenotype, the candidate gene and the QTL are probably allelic. Similarly, in a quantitative knockdown experiment, a differential effect of an amiRNA (amiR) against the candidate gene indicates that the Q1 allele has lower activity than Q2 and that the candidate gene is likely responsible for the QTL. As an alternative, one can introduce genomic fragments spanning the region of interest to identify the gene(s) affecting the trait under investigation. Transgenic complementation also allows the examination of chimeric genes in different backgrounds to pinpoint the causal region, or even nucleotide, within an allele. A possible complication arises from the fact that the addition of an extra wild-type copy of an independent gene in the same pathway can quantitatively affect the phenotype and thus confound the interpretation of the observed phenotypes. An attractive feature of amiRNAs is that one can engineer transgenes that do not change the encoded protein but do not respond to silencing by a specific amiRNA anymore (Palatnik et al., 2003). One can thus use an amiRNA to knock out the endogenous gene and at the same time introduce a variant copy of the gene that is not affected by the amiRNA. This allows in essence the functional replacement of one allele with another. A final word of caution: Spontaneous mutations are not as rare as one might think, with direct measurements indicating about one new single base pair mutation per haploid genome and generation (Ossowski et al., 2010). Thus, not every genetic variant that distinguishes accessions must be a natural variant in the sense that it was present in nature. Indeed, there are now several reports of mutations with large phenotypic effects that were segregating in an accession and may only have arisen after the accession was collected. Two of these cases affect parents of commonly used RIL populations, Landsberg erecta-0 and Bayreuth-0 (Doyle et al., 2005; Loudet et al., 2008; Laitinen et al., 2010). Thus, even if misidentification of an accession has been ruled out, which is not uncommon (Anastasio et al., 2011; Simon et al., 2011), there can be true genetic and phenotypic differences between accessions that share recent common ancestry. WHOLE-GENOME RESOURCES FOR THE STUDY OF NATURAL VARIATION Enabling Genome-Wide Association Studies Genetic mapping in crosses is greatly facilitated when genome-wide polymorphisms, or better yet the entire genome sequences, of the investigated accessions are known. If a sufficient number of genome sequences is available, one can even dispense with experimental crosses and exploit shared ancestry to directly identify common alleles that are responsible for phenotypic variation in the entire population. This approach was first proposed for human, already before the first finished human genome sequence was in sight (Lander, 1996; Risch and Merikangas, 1996). Because obtaining complete genome sequences for many individuals of the same species was out of question at the time, it was proposed to rely on linkage disequilibrium (LD). LD refers to the fact that in most species there has not been enough historic recombination to produce all possible combinations of physically adjacent polymorphisms, but rather that sequence variants are normally found in haplotype blocks of various lengths. Thus, a causal polymorphism can in principle be identified indirectly through its association with any of the other sequence variants in its haplotype block (Kruglyak, 1999; Jorde, 2000). The term that is normally used today for this experimental strategy is genome-wide association study (GWAS). A shortcut that reduces the required genotyping effort has been to make use of prior information and to first focus on genes already shown to affect a trait of interest (Long et al., 1998; Caicedo et al., 2004; Olsen et al., 2004; Balasubramanian et al., 2006; Ehrenreich et al., 2009), but this has become largely obsolete today. While the principles of GWAS are easy to understand, important limitations arise from population structure, that is, not all investigated individuals being equally distantly related to each other. Powerful methods have been developed to correct for population structure, but how to reliably detect alleles that are largely fixed between populations remains a challenge. Other issues are allelic heterogeneity, that is, alleles at a single locus with similar effects on gene function having arisen repeatedly; or complex genetic architecture, where many different genes affect the same trait. A recent article by Myles et al. (2009) provides an excellent primer of the challenges for GWAS. As with RIL analyses, the selfing nature of Arabidopsis is a boon for GWAS, since each accession needs to be genotyped or sequenced only once but can be phenotyped many times. Magnus Nordborg almost single-handedly convinced the Arabidopsis community of the feasibility and usefulness of GWAS approaches, even before high-density genotype information was available (Aranzana et al., 2005; Zhao et al., 2007). While initial estimates of LD in Arabidopsis were too high (Nordborg et al., 2002, 2005), it finally turned out that LD in the global population extends over not more than about 5 to 10 kb, or one to two genes, which is very convenient for GWAS (Kim et al., 2007). It is thought that the relatively low LD reflects a history of frequent outcrossing together with rapid dispersal enabled by the selfing mode of reproduction. The first enterprise with the goal of finding a large fraction of sequence variants used high-density custom arrays with almost one billion unique oligonucleotides to interrogate the genomes of 20 accessions, including the Col-0 reference accession (Clark et al., 2007). This set was chosen to be maximally diverse based on a previous analysis of 96 accessions, from which about 1,000 short fragments distributed throughout the genome had been dideoxy sequenced (Nordborg et al., 2005). The most important information to come from the array-based resequencing study was a collection of hundreds of thousands of nonsingleton single nucleotide polymorphisms (SNPs) that could be used for GWAS (Kim et al., 2007). About 216,000 SNPs, or one every 0.5 kb, have been subsequently typed in over 1,000 accessions (Horton et al., 2012), chosen from a larger panel of more than 5,000 accessions for which information from 149 intermediate frequency markers was available (Platt et al., 2010). The high density of SNPs meant that a typical haplotype block was tagged with several SNPs, which made GWAS in Arabidopsis right away more powerful than in humans. Despite similar LD characteristics, GWAS in human initially used only about 1 SNP per 6 kb (Wellcome Trust Case Control Consortium, 2007). Prospects of GWAS in Arabidopsis Several proof-of-concept examples have now been published, indicating that GWAS will often be successful in Arabidopsis. In the first comprehensive study, over 100 different morphological, physiological, and molecular traits were analyzed in 96 to 192 accessions (Atwell et al., 2010). In several cases, known genes were rediscovered, and in many others, plausible candidates were identified with high precision. The most impressive results, in agreement with previous pilot studies (Aranzana et al., 2005), were obtained for disease resistance, which is often controlled by single genes with very large effects. This is in contrast with humans, where effect sizes of QTL detected by GWAS are often small (McCarthy et al., 2008; Manolio et al., 2009). The utility of GWAS can be increased by making use of prior information, such as functional data from mutant studies, gene annotation, or membership of genes in specific regulatory networks to prioritize GWAS candidates (Aranzana et al., 2005; Schadt et al., 2005; Atwell et al., 2010; Chan et al., 2011). Similarly, QTL mapping in experimental populations can greatly reduce the portion of the genome that one has to consider for the location of GWAS QTL (Brachi et al., 2010; Nemri et al., 2010). This approach becomes particularly powerful when both strategies are directly integrated using experimental populations with several parents, so that alleles pinpointed by GWAS are represented in multiple founder backgrounds. The term nested association mapping has been coined for this approach, which was pioneered in maize (Zea mays; Yu et al., 2008; McMullen et al., 2009). Arabidopsis populations, such as the MAGIC lines and AMPRILs, serve a similar purpose (Kover et al., 2009; Huang et al., 2011). An alternative will be to examine several independent RIL populations. An advantage of using RIL sets over F2 individuals in this case is that for each set of founders, the lines can be chosen to be maximally informative in terms of contribution of the founder genomes, thus greatly reducing phenotyping efforts (Xu et al., 2005; Simon et al., 2008). Because of the plasticity of plant development and physiology, the influence of genes on the phenotype is very often dependent on the environment, often codified as gene-by-environment or GxE interaction. Similarly, the effects of individual genes are often modified by other genes in the genome because genes do not act on their own but form more or less complex functional networks. When genes have nonadditive effects, this is called GxG or more commonly an epistatic interaction. While the identification of epistatic QTL is standard fare for mapping in experimental populations (Mackay, 2001), this continues to be a major challenge for GWAS. This has been suggested to be computationally and statistically feasible several years ago (Marchini et al., 2005), and several computational strategies have been developed since (Mitchell-Olds, 1995; Cordell, 2009; Kam-Thong et al., 2011). However, I am not aware of an example where all variants were used in a GWAS to detect epistatic loci. Here again, mapping in experimental populations, perhaps in combination with network reconstruction (Rowe et al., 2008; Jiménez-Gómez et al., 2010; Kerwin et al., 2011), should help to reduce the search space for GWAS of epistatic loci. A Proliferation of Genome Sequences In addition to the anonymous SNPs for the first generation of GWAS in Arabidopsis, array-based resequencing revealed tens of thousands of amino acid replacements along with hundreds of more drastic mutations that are likely to eliminate the function of many genes in various accessions. In addition, a large percentage of the reference genome was found to be missing in each accession (Borevitz et al., 2007; Clark et al., 2007; Zeller et al., 2008; Plantegenet et al., 2009). This implied that, conversely, the reference accession Col-0 likely lacked a substantial portion of genes present in other accessions. The analysis of individual loci had already shown that some gene families could differ greatly between accessions. Foremost are the disease resistance genes of the nucleotide-binding site-Leu-rich repeat (NB-LRR) class, with both presence/absence polymorphisms and highly divergent alleles in different accessions (Grant et al., 1995; Caicedo et al., 1999; Noël et al., 1999; Stahl et al., 1999; Rose et al., 2004). A logical next step was therefore to scrutinize the genomes of accessions for sequences not represented in the reference genome. With the advent of new sequencing technologies, this goal became attainable at a reasonable cost. Even before these methods were exploited to the same end for human genomes, it was shown that they not only gave an accurate account of small-scale polymorphisms in Arabidopsis genomes but that they could also be used to detect copy number variants and to assemble sequences absent from the reference (Ossowski et al., 2008a). The 1001 Genomes Project for Arabidopsis was announced in 2007 (Nordborg and Weigel, 2008; Weigel and Mott, 2009). The initial proposal was to pursue a two-pronged hierarchical strategy for defining the pangenome of Arabidopsis. The first hierarchical aspect was a sampling of accessions throughout the range of Arabidopsis such that diversity could be analyzed at global, regional, and local scales. Thus, rather than equidistant distribution of samples, it was envisioned that the project would include regional populations separated by distances measured in kilometers as well as individuals from within local stands spaced only meters apart. The second hierarchical aspect was to produce genome sequences at different levels of accuracy and completeness such that a relatively small number of highly accurate and complete genomes would inform the analysis of a much larger number of genomes that had not been completely assembled. The rationale behind this proposal was that mere lists of sequence variants that result from simple resequencing approaches, in which sequence reads are only aligned to a target genome, can be misleading. Specifically, because of false-negative problems, trying to reconstruct contiguous sequences by superimposing known isolated polymorphisms on the reference genome information can be problematic. To overcome these limitations, two groups have introduced reference-guided assembly approaches (Gan et al., 2011; Schneeberger et al., 2011), in which the Col-0 reference genome (Arabidopsis Genome Initiative, 2000) is first used to identify portions of the genome that are conserved in other accessions. Gaps are then filled in by assembling sequence reads and anchoring them to the known bits. As expected, multiple out-of-phase insertions or deletions in coding sequences can combine to restore open reading frames (Schneeberger et al., 2011). Similarly, additional mutations can make up for defects in splice acceptor or donor sites, as can be inferred from transcriptome analysis by RNA sequencing (Gan et al., 2011). The error rates of these reference-guided assemblies in single-copy regions were close to what was deemed as the lower acceptable bound in the initial reference genome sequencing project, about 1 in 10,000 bp (although final error rates in the reference genome were probably only about one-fifth; Ossowski et al., 2008a). As expected from previous resequencing studies, up to 2% of reference positions were judged to be absent from the new assemblies. Conversely, up to 0.6% of the new assemblies represented sequences not found in the reference genome (Gan et al., 2011; Schneeberger et al., 2011). Because the new sequencing technologies generate more error-prone and shorter reads, and the insert sizes for paired-end sequencing libraries are generally smaller as well (Metzker, 2010), there are limits to closing gaps between regions that are well conserved relative to the reference genome. That bases present in the reference, but missing from a nonreference accession, outnumber the opposite class severalfold indicates the shortcomings of the reference-guided assemblies, since it should be equally likely that insertions and deletions occur on either lineage. We are thus currently faced with a paradox: >90% of the euchromatic portion of an accession’s genome can be sequenced for a few hundred dollars, but the remainder can only be recovered when investing many hundred or thousand times that amount. This is particularly relevant because some of the most interesting genes in the genome, such as many disease resistance genes, reside in highly variable gene clusters with often nearly identical tandem repeats that are even challenging for assembly from dideoxy sequenced bacterial artificial chromosomes or fosmid clones (Noël et al., 1999). While the most common approach for the identification and annotation of variants has been comparison against the reference, a multiple alignment consensus benefits the evaluation of complex alleles (Gan et al., 2011). However, with the rapid increase in the number of genome sequences, simple all-against-all comparisons will soon not be feasible anymore because of the time required to perform them. It has therefore been proposed to represent the pangenome, that is, the collection of all possible sequence variants along each chromosome, in a single data structure as a graph, which would both facilitate the identification of polymorphisms in newly sequenced genomes and their classification as shared or unique (Schneeberger et al., 2009). Insights from Comparing Genome Sequences Apart from supporting forward genetic studies in Arabidopsis, genome sequences have increased our understanding of the evolutionary history of the species. Array-based comparison of 20 accessions revealed only a single large region in the genome that was shared by the majority of accessions, indicative of this region having experienced recent and strong selection in many different populations (Clark et al., 2007). Remarkably, the much more fine-grained information from short-read sequencing of 80 lines did not substantially change this picture of strong selective sweeps being rare, even though population differentiation along the genome is not uniform (Cao et al., 2011). In addition to local polymorphism patterns that are shaped by selection and demography, there are consistent chromosomal-scale differences that are probably caused by molecular and genetic factors, such as mutation, recombination, and biased gene conversion. One of these is an excess of polymorphisms in regions adjacent to the centromeres (Borevitz et al., 2007; Clark et al., 2007), which has also been reported in Medicago truncatula and rice (Oryza sativa), but not in maize (Gore et al., 2009; Huang et al., 2010c; Branca et al., 2011). The interpretation of polymorphism patterns in Arabidopsis has also benefited from the high-quality reference sequence available now for the close relative Arabidopsis lyrata (Hu et al., 2011). In agreement with lack of conservation between the two species reflecting either that sequences are dispensable or subject to species-specific positive selection, regions found only in Arabidopsis are more polymorphic than shared regions (Cao et al., 2011). Finally, Arabidopsis accessions harbor extensive variation in mitochondrial genomes (Forner et al., 2005; Arrieta-Montiel et al., 2009), in subtelomeric regions (Kuo et al., 2006; Wang et al., 2010), and in heterochromatic repeats, including retrotransposons and rDNA (Fransz et al., 2000; Davison et al., 2007; Ito et al., 2007). Structural differences between mitochondrial genomes can be revealed relatively easily by new sequencing methods (Davila et al., 2011). Furthermore, although read lengths and insert sizes are insufficient for long-range reconstruction of highly repetitive regions of the genome, read coverage and sequence variation in individual reads can be exploited to determine differences in genome size and repeat content (James et al., 2009; Tenaillon et al., 2011). Utility of Genome Sequences As of the time that this article was written (end of 2011), over 100 genome sequences for Arabidopsis had been published. In addition, sequence data for over 300 additional accessions were already publicly available. In aggregate, commitments for over 700 accessions had been made, indicating that the initial goal of 1,001 genome sequences would be reached well before the end of 2012 (http://1001genomes.org). Several of the Arabidopsis genome sequences were immediately useful. For example, the Landsberg erecta accession is commonly used for mutant screens, and its genome sequence is facilitating the mapping and analysis of induced mutations. Similarly, several of the accessions are parents of RIL populations (Ossowski et al., 2008a; Schneeberger et al., 2009, 2011; Gan et al., 2011), and their genome sequences are aiding the identification of polymorphisms responsible for QTL. Genome sequences also provide an inventory of potential knockout mutations, which is informative given that a considerable fraction of natural genetic variation is due to loss-of-function alleles. Examples are new alleles of PHYTOCHROME D (PHYD) and FRIGIDA LIKE1 (FRL1), for which before only single alleles were known (Aukerman et al., 1997; Schläppi, 2006; Cao et al., 2011). In addition, the 1001 Genomes Project is advancing GWAS. As discussed above, the first phase of GWAS in Arabidopsis has been based on a set of 216k tag SNPs, which were estimated to predict >90% of all common variants (Kim et al., 2007; Horton et al., 2012). It is simple to call the same SNPs in any of the accessions of the 1001 Genomes Project and to include any line that has not been array genotyped into GWAS projects that makes use of the 216k tag SNP array data. Furthermore, it is possible to accurately impute common variants identified by whole-genome sequencing in array genotyped accessions and GWAS with imputed data detects additional polymorphisms linked to traits under consideration (Cao et al., 2011). Apart from increasing the chances that sequence differences directly responsible for trait variation are found by GWAS, a major advantage of complete genome sequences is that they support the prediction of activity differences between potentially causal alleles. For example, in coding regions, mutations that disrupt the open reading frame or affect splicing are more likely to affect gene function than codon or silent changes. And among amino acid substitutions, one can estimate how probable it is that a mutation has deleterious effects based on conservation of that amino acid in other species (Ng and Henikoff, 2006). Complete genome sequences will thus help to tackle one of the major challenges of GWAS, allelic heterogeneity, where several different alleles have similar effects on the trait of question. That independent alleles at the same locus can have the same phenotypic consequences has been known for a quarter of a century, since the first genes responsible for genetic disorders or cancer in humans were cloned (Royer-Pokora et al., 1986; Clark et al., 1989; Botstein and Risch, 2003). In Arabidopsis, the flowering regulators FRIGIDA (FRI) and FLOWERING LOCUS C (FLC) are often partially or completely inactivated, with many of these alleles being found only in single accessions (Johanson et al., 2000; Le Corre et al., 2002; Gazzani et al., 2003; Michaels et al., 2003; Lempe et al., 2005; Shindo et al., 2005; Méndez-Vigo et al., 2011). Drastic mutations that prematurely terminate or partially delete the same open reading frame are found more often than expected by chance in the genomes of different accessions (Cao et al., 2011; Fig. 5). This might be the outcome of positive selection, as is the case for FRI and FLC (Toomajian et al., 2006), or purifying selection being weak or absent. In either case, the presence of multiple alleles with similar effects on a particular phenotype makes the detection of such loci in GWAS analyses difficult since each polymorphism is considered separately (Myles et al., 2009). If, instead, all alleles with similar predicted activity differences were combined or, better yet, if alleles were considered according to their relative degree of activity, this hurdle could be overcome. Figure 5. Open in new tabDownload slide Comparison of expected and observed occurrences of 8,133 independent premature stops in 4,263 protein coding genes, considering all genes with >90% coverage in 75 out of 80 accessions. Data are from Cao et al. (2011). Figure 5. Open in new tabDownload slide Comparison of expected and observed occurrences of 8,133 independent premature stops in 4,263 protein coding genes, considering all genes with >90% coverage in 75 out of 80 accessions. Data are from Cao et al. (2011). The methods discussed in the preceding paragraph would be a considerable improvement over the strategy that is gaining popularity in humans: the search for an excess of rare variants in candidate genes. In rare-variant-burden methods, rare variants are combined for the purposes of contrasting phenotypically distinct classes of individuals, but functional effects of alleles are ignored, and these methods are not integrated into standard GWAS (Asimit and Zeggini, 2010). Epigenomic Variation GWAS in humans, where it is not unusual that tens of thousands of individuals are analyzed, has been successful in detecting many alleles, even with very small effects, but the fraction of the total variation explained by these variants is often only small. This also has been the case for traits such as height that are known to be highly heritable from family studies. Some possibilities are that genetic architecture may be more complex, with many interacting loci, or that rare alleles are more important than anticipated (see above). An alternative explanation, which is en vogue in many circles, is that epigenetic variation unlinked to sequence variants and, hence, not detectable by conventional GWAS is responsible for many phenotypic differences (McCarthy et al., 2008; Manolio et al., 2009). Epigenetic differences can have obvious consequences in plants. In several species, including Arabidopsis, spontaneously occurring epialleles with overt phenotypes have been described (Jacobsen and Meyerowitz, 1997; Cubas et al., 1999; Hollick et al., 2000; Soppe et al., 2000; Stam et al., 2002; Manning et al., 2006; Martin et al., 2009). The epialleles often show increased cytosine methylation of the promoter and strongly reduced RNA expression. In several cases, the epialleles are associated with structural changes, such as the g mutation in melon, which is apparently caused by the insertion of a transposon and spread of DNA methylation into adjacent sequences. Tiling array analyses comparing two different pairs of Arabidopsis accessions have shown that these differ in the extent of methylation at individual cytosines. That there are fewer differences in transposable element than genic methylation between natural accessions (Vaughn et al., 2007) agrees with transposable element methylation being more stable in inbred lines (Becker et al., 2011; Schmitz et al., 2011). Methylation differences seem to be largely stable in F1 hybrids (Woo and Richards, 2008; Zhang et al., 2008; Groszmann et al., 2011), but methylation patterns can change at relatively high rates, around 1% or more, in subsequent generations (Vaughn et al., 2007). The fluidity of the genomic methylation landscape after crosses is consistent with RNA-dependent DNA methylation mediated by short interfering RNAs being able to target other loci in trans, as long as these harbor sufficient levels of sequence similarity (Melquist and Bender, 2003). This is substantiated by nonadditive expression levels of short interfering RNAs and correlated effects on DNA methylation in F1 hybrids (Groszmann et al., 2011). Importantly, although epialleles with phenotypic effects are largely stable and can be inherited over many generations, most revert occasionally to the wild-type form (Jacobsen and Meyerowitz, 1997; Cubas et al., 1999; Hollick et al., 2000; Soppe et al., 2000; Stam et al., 2002; Manning et al., 2006; Martin et al., 2009). The stability of DNA methylation in inbred Arabidopsis lines has recently been examined directly (Becker et al., 2011; Schmitz et al., 2011). While loss and gain of methylation at individual sites occurred much more often than mutations in the nucleotide sequence (Ossowski et al., 2010), changes in larger methylated regions similar to the ones that distinguish epialleles identified by forward genetics were rare. However, both types of methylation changes were distinguished from DNA mutations in that the same positions were affected in independent lines much more often than expected by chance and that there was an appreciable rate of reversions. Crosses of wild-type lines to mutant strains with largely demethylated genomes have also revealed a wide range in the stability of epialleles after the causal mutations had been segregated away (Reinders et al., 2009; Teixeira et al., 2009). Consistent with the more labile nature of epialleles, heritability estimates in such lines are considerably lower than they are in natural accessions for the same traits (Roux et al., 2011). Thus, while the large majority of DNA methylation differences is sufficiently stable to account for inheritance within a limited number of generations, it remains unclear how often epialleles can become subject to Darwinian selection and thus make a contribution to long-term evolution. If reversion rates exceed the selective advantage conferred by an epiallele, its frequency in the population will be largely determined by the equilibrium of forward and reverse epimutation rates (Slatkin, 2009; Johannes and Colomé-Tatché, 2011). In summary, although natural epialleles are often due to nearby structural variation, crosses between divergent accessions can induce new epialleles in trans. While the first class does not pose a problem for conventional GWAS, as such alleles should be tagged by linked sequence polymorphisms, the second class would only be revealed if GWAS would be extended to directly include information on DNA methylation profiles. A different question is whether epialleles are equally, more, or less likely than DNA alleles to reflect adaptation to the local environment. LEARNING NEW BIOLOGY FROM THE STUDY OF NATURAL VARIATION While knowledge about the origin and phenotypic effects of sequence polymorphisms is central to understanding how species adapt to their natural environment, most studies of genetic variation in Arabidopsis have probably been motivated by the desire to identify regulatory and other genes that are not present in the common laboratory accessions. An especially original use of natural variation has been the search for second site modifiers of ABA insensitive3 and leafy cotyledon1 mutant phenotypes. Both mutants suffer from impaired seed maturation, and seed viability declines much more rapidly than in wild-type plants. Introgression of the mutant alleles into other accessions identified natural modifiers that can partially suppress the mutant phenotypes, possibly pointing to new regulators of seed maturation (Sugliani et al., 2009). In a similar manner, the CAULIFLOWER (CAL) gene was discovered serendipitously as an enhancer of the apetala1 (ap1) mutant phenotype. CAL and AP1 turned out to be paralogs with an asymmetrical relationship: While AP1 can compensate for loss of CAL activity, the reverse is not true. Thus, in contrast with induced ap1 mutations, natural loss-of-function alleles of CAL have no overt phenotype on their own and are only noticed if AP1 is inactive as well (Bowman et al., 1993; Kempin et al., 1995). Arabidopsis was used early on to identify genes that control seed dormancy (van Der Schaar et al., 1997). For ease of cultivation, common laboratory accessions had been selected to be early flowering (more below) and to have little dormancy, meaning that seeds would germinate relatively quickly after harvest. The DELAY OF GERMINATION1 (DOG1) locus, the first dormancy QTL cloned, encodes the prototype of a small gene family of unknown molecular function. There is extensive variation in DOG1 expression levels between accessions, suggesting the presence of many functionally distinct alleles of DOG1 (Bentsink et al., 2006). Arabidopsis accessions also remain an important resource for functional and evolutionary analyses of large-effect resistance genes (Staskawicz et al., 1995). This is a large area for which there are several recent in-depth reviews (Nishimura and Dangl, 2010). Below, I will discuss three naturally variable traits in some more detail: trichome density, which provides a paradigm for how information from multiple genome sequences can be used to pinpoint causal polymorphisms; glucosinolate content, which has an underlying biochemical pathway with variation at almost every step; and the onset of flowering, a developmental trait with a well-understood molecular basis. Trichome Density Early studies by Rodney Mauricio and Mark Rausher came to the conclusion that both physical defenses in the form of leaf hairs (trichomes) and chemicals (glucosinolates) reduce herbivore damage to Arabidopsis in the field but that these are not without costs (Mauricio and Rausher, 1997; Mauricio, 1998). Several genes have been identified as affecting trichome density of natural Arabidopsis accessions. The most dramatic effects are seen in accessions that are glabrous, that is, lack trichomes completely, and at least two different loss-of-function mutations at GLABRA1 (GL1) have been found. Whether a fitness trade-off, as suggested for other defense traits, underpins the GL1 polymorphisms is unknown. Balancing selection, however, which is often taken as a sign of trade-offs, does not appear to be responsible for maintaining different GL1 alleles (Hauser et al., 2001). Glabrousness caused by inactivating mutations in GL1 also segregates in A. lyrata and Arabidopsis halleri populations (Hauser et al., 2001; Kärkkäinen and Ågren, 2002; Kivimäki et al., 2007; Kawagoe et al., 2011). A less extreme phenotype of reduced trichome density is caused in some Arabidopsis accessions by a nonsynonymous substitution in MYC1 (Symonds et al., 2011). As another warning to population geneticists, one of the exons was found to exhibit a strong signal of divergent selection, with many amino acid substitutions. However, this signal was not correlated with trichome density. Other accessions have increased trichome number relative to the Col-0 reference accession, and ENHANCER OF TRY AND CPC2 (ETC2) has been identified as the causal gene (Hilscher et al., 2009). ETC2, MYC1, and GL1 all encode transcription factors, with GL1 promoting and ETC2 repressing trichome formation by competing for interaction with common partners, a group of basic helix-loop-helix proteins that includes GL3 and MYC1 (Ishida et al., 2008). In contrast with MYC1, the high- and low-activity variants of ETC2 segregate at intermediate frequencies, indicating that ETC2 is a major determinant of natural variation in trichome number. ETC2 very likely corresponds to one of the first QTL that was mapped in Arabidopsis, REDUCED TRICHOME NUMBER (Larkin et al., 1996), and consistent with alleles of different activity being common, ETC2 can also be detected by GWAS (Atwell et al., 2010). Notably, it had initially been suggested that ETC2 has only a minor role in trichome formation, a conclusion that came from studies done with common accessions that have an ETC2 allele without obvious disruptions but with nevertheless low activity. The work on ETC2 is noteworthy because of how the causal polymorphism was first pinpointed using a strategy that should be broadly applicable. To triangulate the causal region in the final mapping interval, accessions with either very high or very low trichome densities were selected, and the extent of haplotype sharing in each group was compared, which identified a small region with only two candidate polymorphisms (Hilscher et al., 2009). Transformation with chimeric transgenes provided conclusive support that one of the variants, a nonsynonymous mutation, was reducing the activity of ETC2. With the resources of the 1001 Genomes Project, these types of local association studies should become a common strategy for the endgame in identifying QTL after conventional mapping in F2 or similar populations. Glucosinolate Content In addition to the gene-for-gene resistance loci that are effective against individual pathogen strains (for review, see Nishimura and Dangl, 2010), Arabidopsis accessions also show quantitative variation in resistance, in particular against herbivorous insects. As with trichomes, chemical defenses in the form of a Brassicaceae-specific class of secondary metabolites, the glucosinolates, can reduce herbivore damage (Blau et al., 1978). There are considerable inter- and intraspecific differences in the repertoire of glucosinolates, which are hydrolyzed by the enzyme myrosinase into the active defense compounds (Kliebenstein et al., 2005). In Arabidopsis, METHYLTHIOALKYLMALATE SYNTHASE (MAM) and AOP are the two major loci responsible for variation in glucosinolate biosynthesis, with additional contributions from the GSL-OH locus (Kliebenstein et al., 2001; Kroymann et al., 2001, 2003). Hydrolysis of the glucosinolates is further affected by the polymorphic EPITHIOSPECIFIER PROTEIN and EPITHIOSPECIFIER MODIFIER1 loci (Lambrix et al., 2001; Zhang et al., 2006). In other Brassicaceae, several of the same genes are responsible for intraspecific variation in glucosinolate content, including A. lyrata (Li and Quiros, 2003; Heidel et al., 2006). Notably, both the MAM and AOP loci are complex, with several tandem arrayed genes that vary in presence, enzyme activity, or expression level between accessions, giving rise to more than two alternative allelic states, processes that are apparently driven by positive selection (Kliebenstein et al., 2001; Kroymann et al., 2001, 2003). At least MAM shows a similar pattern of diversity created by gene duplication and neofunctionalization between other members of the Arabidopsis genus as well as closely related genera (Benderoth et al., 2006). The detailed understanding of the control of glucosinolate accumulation in turn supports research into broader questions of genetic variation, such as the importance of stochastic variation, which was found to be genetically encoded as well (Jimenez-Gomez et al., 2011). Flowering Time Seed production is one of the most important components of fitness, and to optimize seed set, plants need to flower at the right time of year. In agreement with Arabidopsis is found in places with very different growing seasons, natural accessions differ greatly in their flowering behavior. Beginning with Laibach (1943, 1951), several investigators reported flowering variation not only in inbred accessions, but also in individuals collected from the wild (Napp-Zinn, 1957; Cetl et al., 1968; Jones, 1971; Westerman, 1971). That this trait is under selection has also been inferred from population genomics analyses (Flowers et al., 2009) and from the finding of latitudinal and altitudinal clines, likely due to covariation of flowering time with climatic factors (Caicedo et al., 2004; Stinchcombe et al., 2004; Lempe et al., 2005). The first natural allele to be mapped with molecular markers in Arabidopsis was at the FRI locus, which segregates in a Mendelian manner in crosses between late- and early-flowering accessions (Lee et al., 1993; Clarke and Dean, 1994). The first QTL mapped in Arabidopsis were also ones controlling flowering (Kowalski et al., 1994; Clarke et al., 1995), followed by many additional QTL studies (for review, see Koornneef et al., 2004; Shindo et al., 2007). Mapping in crosses and GWAS have shown that flowering time variation can be explained by relatively few large-effect QTL (Atwell et al., 2010; Brachi et al., 2010; Li et al., 2010; Salomé et al., 2011b; Strange et al., 2011), which is very different from maize (Buckler et al., 2009). FRI and the epistatically acting FLC gene are responsible for a large fraction of flowering time variation in Arabidopsis accessions when these are not exposed to a winter-like vernalization treatment. FRI promotes expression of the FLC transcription factor, which directly represses genes with positive roles in flowering (Li et al., 2008; Deng et al., 2011). Allelic variation at FLC likely accounts for flowering time differences in other Brassicaceae as well, including Capsella bursa-pastoris and some, but not all, Brassica species (Long et al., 2007; Razi et al., 2008; Slotte et al., 2009; Zhao et al., 2010). A role for FRI in flowering time variation in A. lyrata and Brassica napus has been inferred from association studies (Kuittinen et al., 2008; Wang et al., 2011). Strikingly, there are many alleles at both FRI and FLC (Michaels and Amasino, 1999; Johanson et al., 2000; Le Corre et al., 2002; Gazzani et al., 2003; Lempe et al., 2005; Shindo et al., 2005; Méndez-Vigo et al., 2011). Because of the convenience of early flowering, commonly used laboratory accessions have a loss-of-function allele at one or both loci. However, while low-activity FRI alleles typically have disrupted open reading frames, FLC alleles are predominantly characterized by noncoding structural variation. During vernalization, FLC becomes epigenetically silenced, and natural alleles differ in the duration of vernalization needed for stably switching off FLC expression (Shindo et al., 2006). In addition to its repressive effects on flowering, high-activity alleles of FLC promote germination in the cold, which in turn allows plants to experience the longer cold period required for flowering when FLC is active (Chiang et al., 2009). The FRI homologs FRL1 and FRL2 along with the FLC homologs FLM/MAF1 and MAF2 provide additional routes to flowering time variation (Werner et al., 2005; Schläppi, 2006; Caicedo et al., 2009; Rosloski et al., 2010). Flowering time control is one of the most intensively investigated developmental processes in Arabidopsis, and well over 100 genes are known to affect flowering, with many having substantial pleiotropic effects on plant growth (Srikanth and Schmid, 2011). Remarkably, only one gene with very few nonflowering phenotypes, the central flowering activator FT, has been shown to contribute extensively to flowering time variation between Arabidopsis accessions (Schwartz et al., 2009; Li et al., 2010; Huang et al., 2011; Salomé et al., 2011b; Strange et al., 2011). QTL studies have implicated FT as being the cause of flowering time differences also in B. napus (Long et al., 2007). Several other genes responsible for flowering time variation in Arabidopsis have multiple functions during plant development, including the photoreceptor encoding genes CRYPTOCHROME2, PHYC, and PHYD (Aukerman et al., 1997; El-Din El-Assal et al., 2001; Balasubramanian et al., 2006; Méndez-Vigo et al., 2011). In addition, there is functional allelic variation at PHYA and PHYB. Both regulate flowering (Srikanth and Schmid, 2011), although the effects of the natural alleles on flowering have not been studied (Maloof et al., 2001; Filiault et al., 2008). Two other pleiotropically acting, naturally variable flowering regulators are FY (Adams et al., 2009) and HUA2. In addition to affecting flowering time, a natural HUA2 change-of-function allele has a dramatic effect on plant architecture that had not been anticipated from mutant studies (Alonso-Blanco et al., 1998a; Wang et al., 2007; Huang et al., 2011; Strange et al., 2011). Finally, additional loci responsible for flowering time regulation have been identified by growing plants under variable conditions (Weinig et al., 2002; Brachi et al., 2010; Li et al., 2010). TOWARD AN UNDERSTANDING OF THE FORCES SHAPING GENETIC VARIATION Apart from extending our knowledge of biological mechanisms and pathways in Arabidopsis, a major motivation for studying genetic variation is to understand how a species adapts to different local environments, which traces adaptation leaves in the genome, and how this leads to the formation of new species. In this section, I describe how genome analyses have provided insights into the history of the species, what is being learned about epistatic interactions between alleles from different genomes, and how evidence for local adaptation is emerging. Geographic Distribution of Population Diversity Until a decade ago, the vast majority of the few hundred Arabidopsis accessions available from the stock centers came from western Europe. In the past years, collections have been substantially expanded, with more than 2,000 genotypically distinct accessions having been described (Schmuths et al., 2006; Beck et al., 2008; Picó et al., 2008; Montesinos et al., 2009; Bomblies et al., 2010; Lewandowska-Sabat et al., 2010; Platt et al., 2010; Cao et al., 2011; Méndez-Vigo et al., 2011). With whole-genome data, the pattern of isolation-by-distance that had been deduced from more sparse data before came into even sharper focus. In addition, it was found that geographic regions differ greatly both with respect to the total number of polymorphisms distinguishing accessions within a region from each other and from other regions and the relative frequency of variants that are shared with other regions. There is an overall gradient from west to east: The greatest diversity is found at the western end of the native range, in the Iberian Peninsula, including North Africa, while the most uniform regions are in Central Asia. This is consistent with the view that Arabidopsis populations in the west are the oldest, with later expansion into the eastern end of its native distribution, along with recently colonized regions, such as the Alps, in the center of the range (Sharbel et al., 2000; Nordborg et al., 2005; Schmid et al., 2005; Ostrowski et al., 2006; Beck et al., 2008; Picó et al., 2008; Platt et al., 2010; Cao et al., 2011). In addition, there is also altitudinal stratification within regions, with populations from high altitude being overall less diverse than those from lower altitude (Montesinos et al., 2009; Lewandowska-Sabat et al., 2010; Gomaa et al., 2011). It has also been suggested that there is evidence for migration from east to west, accompanying the spread of agriculture (François et al., 2008); however, knowing that the Iberian Peninsula is the most diverse region, it is unclear what to make from this. The regional differences have certainly important implications for the design of GWAS, since LD extends further in less diverse regions (Cao et al., 2011). In continental Eurasia, identical multilocus genotypes are almost exclusively found only in the same local patches of Arabidopsis individuals (Picó et al., 2008; Bomblies et al., 2010; Lewandowska-Sabat et al., 2010; Platt et al., 2010). Exceptions are the British Isles and North America. In both regions, one specific genotype is found in many different places. For North America, recent and widespread, but uneven, introduction by European settlers has been suggested as the most likely cause; this scenario is compatible with the absence of genetic isolation by distance in North America (Platt et al., 2010). Epistatic Interactions between Genomes Despite its selfing nature, and contrary to what early analyses had suggested, stands of Arabidopsis plants can include several different multilocus genotypes. Moreover, outcrossing rates of Arabidopsis in nature can be several percent, and heterozygous individuals are thus not that rare (Stenøien et al., 2005; Bakker et al., 2006; Jorgensen and Emerson, 2008; Bomblies et al., 2010; Platt et al., 2010). Superior performance in heterozygous F1 hybrids is known as heterosis or hybrid vigor. Heterosis in Arabidopsis is generally not as dramatic as in other species, but heterotic QTL for biomass and metabolites have been identified by backcrossing RILs derived from two inbred accessions to the founders (Syed and Chen, 2005; Kusterer et al., 2007; Lisec et al., 2009; Meyer et al., 2010). There is also extensive evidence for nonadditive, or epistatic, effects on gene expression in intra- and interspecific hybrids (Wang et al., 2006; Zhang and Borevitz, 2009; Zhang et al., 2011). In both stable allotetraploids and F1 hybrids of Arabidopsis × arenosa, circadian gene expression programs are altered, and a similar trend is apparent in F1 hybrids between two Arabidopsis accessions that exhibit hybrid vigor. The heterotic effects are mediated by central regulators of the circadian clock (Ni et al., 2009), although the proximate causes that alter activity of these regulators, and their relationship to the heterosis QTL identified in the same cross before, remain unknown. Inferior performance of F1 hybrids is known as hybrid weakness or incompatibility, with extreme cases presenting as hybrid sterility or lethality. In addition, a decline in fitness of later generations is called hybrid breakdown or inbreeding depression (Hochholdinger and Hoecker, 2007; Charlesworth and Willis, 2009; Bomblies, 2010) A commonly observed incompatibility phenomenon is cytoplasmic male sterility (CMS), due to a mismatch between nuclear genes that encode proteins active in mitochondria and the mitochondrial genome (Fujii and Toriyama, 2008). Despite well over 1,000 different interaccession crosses having been examined (Bomblies et al., 2007), CMS has not yet been reported in Arabidopsis, even though weak CMS has been observed in A. lyrata (Leppälä and Savolainen, 2011). The most common obvious defect in F1 hybrids of Arabidopsis appears to be an autoimmune syndrome, hybrid necrosis, that is also known from many other plants. Hybrid necrosis can often be explained by one or two epistatically interacting loci (Bomblies et al., 2007; Bomblies and Weigel, 2007). At least one of the genes causal for hybrid necrosis in Arabidopsis encodes an immune receptor of the NB-LRR class (Bomblies et al., 2007), consistent with the identification of immune genes underlying hybrid necrosis in other species (Krüger et al., 2002; Jeuken et al., 2009; Yamamoto et al., 2010). The NB-LRR family is the most variable gene family in plants, with genes often being found in clusters that have a complex history of gene duplication, deletion, and gene conversion. NB-LRR genes are engaged in recognition of diverse proteins (Nishimura and Dangl, 2010), providing an intuitive explanation for why hybrid necrosis is so common. In a broader context, hybrid necrosis is a manifestation of the costs of disease resistance (Tian et al., 2003). In some instances, hybrid necrosis becomes only expressed in the F2 generation (Alcázar et al., 2009). In one such case, one of the causal genes encodes a receptor kinase homolog, with evidence of positive selection for disease resistance having increased the frequency of this allele in Central Asia (Alcázar et al., 2010). A receptor-kinase-like gene of a different class is responsible for an incompatibility that primarily causes growth defects. This specific case involves an interaction between alleles at a single locus with similar properties as many NB-LRR loci, namely being composed of a highly variable tandem array of genes (Smith et al., 2011). Notably, not every highly variable gene family appears to cause problems in hybrids. Cytochrome P450s, which are important for plant insect defense and are produced by one of the most highly variable gene families (Clark et al., 2007; Cao et al., 2011), have so far not been tied to hybrid weakness, perhaps because they are not designed to interact with a diverse set of other proteins. Most F2 incompatibilities were not discovered because of overt phenotypic effects but were deduced from segregation distortion, that is, the absence of certain genotypic combinations, in F2 or RIL populations (Lister and Dean, 1993; Mitchell-Olds, 1995; Alonso-Blanco et al., 1998b; Loudet et al., 2002; Werner et al., 2005; Törjék et al., 2006; Simon et al., 2008; Balasubramanian et al., 2009; Salomé et al., 2011a). For RILs, this can be due to inadvertent selection, e.g. because late-germinating lines are eliminated, but several cases are associated with lethality of specific segregants. One example involves a pair of paralogs that arose from a very recent ectopic duplication event and that independently sustained inactivating mutations in different lineages (Bikard et al., 2009). About three-quarters of accessions carry inactive copies of one or the other paralog, suggesting that increased dosage is disfavored. A similar situation of reciprocally mutated paralogs explains an epistatic interaction affecting shoot growth (Vlad et al., 2010). Both cases differ from other examples of complex duplication and mutation events, where the paralogs have become neofunctionalized and have now distinct activities (Kliebenstein et al., 2001; Kroymann et al., 2003; Huang et al., 2010a). Experimental Ecology and Ecological Genomics The worldwide distribution of Arabidopsis can be well described by climatic range boundaries; these indicate that laboratory conditions commonly used for growth of Arabidopsis are at the extreme end of its normal habitats, which are normally much cooler and drier (Hoffmann, 2002). This has important implications for interpreting phenotypic differences observed in the greenhouse. For example, strains with differential activity of the key flowering regulators FRI and FLC, known to vary in many accessions, only differ strongly in their flowering behavior outdoors when germinated at specific times of the year, with a critical period in early fall having a disproportionately large effect on flowering time, namely, whether plants overwinter (Wilczek et al., 2009). Such knowledge is essential if one wants to predict responses to a changing climate (Wilczek et al., 2010). Furthermore, by culturing plants in seminatural settings, in which either variable light and temperature conditions are reproduced in climate chambers or plants are germinated in the greenhouse, then transplanted outdoors, one can detect QTL that are not found when plants are grown in a uniform environment. Whether either type of QTL is more relevant is unclear and can only be addressed by phenotyping truly naturally growing individuals. Nevertheless, analysis in seminatural conditions provides insights into the genetic basis of traits considered to be indicative of fitness, such as germination, survival, fruit and seed number, or competitiveness (Weinig et al., 2002, 2003a, 2003b; Stinchcombe et al., 2004; Donohue et al., 2005; Li et al., 2006; Brachi et al., 2010; Huang et al., 2010b; Li et al., 2010; Fournier-Level et al., 2011). Different experimental approaches are beginning to reveal local adaptation in Arabidopsis. When 74 accessions were monitored in the greenhouse under different temperatures, it was found that accessions from cold regions respond in their growth more strongly to elevated temperatures than accessions from warm regions, which are only moderately inhibited by colder temperatures (Hoffmann et al., 2005). Systematic correlation of phenotypes with environmental gradients can indicate adaptation (Endler, 1977), and there are also latitudinal clines in light sensitivity and altitudinal clines in flowering-related traits (Maloof et al., 2001; Méndez-Vigo et al., 2011). It has been similarly proposed that populations of Arabidopsis near oceans or saline soils are more likely to carry an allele at the HKT1 locus that increases sodium accumulation in leaves (Baxter et al., 2010). However, the accessions investigated were unevenly sampled, information about soil salinity at the places of origin was not available, and the relationship between compromised activity of HKT1 and salt tolerance is complex (Mäser et al., 2002; Berthomieu et al., 2003). Thus, the conclusions about adaptation to salinity should be taken with the proverbial grain of salt. Reciprocal transplantation experiments have produced evidence for local adaptation in A. lyrata (Leinonen et al., 2009, 2011). Somewhat surprisingly, this approach, a gold standard in ecology (Turesson, 1922a), has so far only been sparingly applied in Arabidopsis. This has recently been remedied, with an impressive study in which hundreds of accessions were grown at several different places in the native range of the species (Fournier-Level et al., 2011). Alleles associated with superior fitness at each site were most likely to be found in accessions originating near that site. GWAS identified several candidates for survival and fruit number, although only one, the photoreceptor gene PHYB, which affects light response, can be easily connected to local adaptation based on prior knowledge. Additional evidence for local adaptation comes from GWAS for climate variables at the place of origin combined with fitness tests at a single site (Hancock et al., 2011). Both of these studies were carried out predominantly with accessions from the western European and Scandinavian part of the native range, and it will be interesting to repeat these experiments with a broader spectrum of accessions and test locales. OUTLOOK Our knowledge of natural variation in Arabidopsis has advanced tremendously in the past decade, with an impressive set of genetic and genomic approaches and resources that are now available (Fig. 6). In the near future, the simultaneous application of different strategies will lead to genetic variation increasingly informing basic plant biology. Combined analyses of global transcript and metabolite levels and biomass across accessions and RIL populations is supporting the reconstruction of functional networks (Wentzell et al., 2007; Lisec et al., 2008; Rowe et al., 2008; Sulpice et al., 2009, 2010). Integration of QTL data with such information has shown that in addition to biosynthetic and metabolic enzymes, upstream transcription factors of the MYB class contribute to diversity in glucosinolate content (Sønderby et al., 2007) and that the clock gene ELF3 has a role in shade avoidance (Jiménez-Gómez et al., 2010). Another instructive example of how natural variation can help to discover a new regulatory pathway comes from the study of xylem expansion (Sibout et al., 2008). The authors noted that the xylem expansion loci colocalized with flowering time QTL, which led them to hypothesize that the onset of flowering causes xylem expansion in both the shoot and the root. They subsequently confirmed such a model by transiently inducing the activity of a central floral regulator. There is similarly great promise in GWAS with the same material to identify cases of pleiotropic action of natural sequence variants. Figure 6. Open in new tabDownload slide Relationship between approaches to the study of genetic variation. Figure 6. Open in new tabDownload slide Relationship between approaches to the study of genetic variation. I have also highlighted the many opportunities Arabidopsis offers for the study of interactions between divergent genomes, which may both promote or reduce outcrossing, and thereby affect the partitioning of genetic diversity into different lineages (and ultimately into different species). So far, the parents for the investigated crosses have largely been chosen randomly. With increasing information about the genome-wide and population-specific distribution of sequence polymorphisms, more judicious and systematic choices of genotype combinations should accelerate the pace with which we can obtain insights into the fascinating questions of hybrid performance. Another important direction will be to phenotype naturally growing plants in situ over several years (Montesinos et al., 2009). Genotyping of very large numbers of wild plants has become very affordable with next-generation sequencing methods, which will facilitate linking genotype and phenotype even on an individual basis (Baird et al., 2008; Elshire et al., 2011). An example for such strategies is a study that monitored over 4 years the load of five different viruses that had been known before to infect wild Brassicaceae (Pagán et al., 2010). Such experiments are required to test claims about fitness trade-offs between disease resistance and growth (Tian et al., 2003; Todesco et al., 2010). Finally, selection experiments are a tool that should not be underestimated for their potential to provide insights into favorable allele combinations (Ungerer et al., 2003; Ungerer and Rieseberg, 2003; Scarcelli and Kover, 2009; Fakheran et al., 2010). Supplemental Data The following materials are available in the online version of this article. Supplemental Table S1. Full references for Table 1. ACKNOWLEDGMENTS I thank Eunyoung Chae, Sang-Tae Kim, and George Wang for plant images; Joy Bergelson, Carlos Alonso-Blanco, Jun Cao, Karl Schmid, and George Wang for help in producing the map of Arabidopsis accessions; and Annie Schmitt and Joy Bergelson for preprints. I am especially grateful to three anonymous reviewers, who provided insightful comments and helped to correct several oversights in the original manuscript. LITERATURE CITED Adams S Allen T Whitelam GC ( 2009 ) Interaction between the light quality and flowering time pathways in Arabidopsis . Plant J 60 : 257 – 267 Google Scholar Crossref Search ADS PubMed WorldCat Alcázar R García AV Kronholm I de Meaux J Koornneef M Parker JE Reymond M ( 2010 ) Natural variation at Strubbelig Receptor Kinase 3 drives immune-triggered incompatibilities between Arabidopsis thaliana accessions . 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[OA] Open Access articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.111.189845 © 2012 American Society of Plant Biologists. All rights reserved. © The Author(s) 2012. Published by Oxford University Press on behalf of American Society of Plant Biologists. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.

Cheung, Alice Y.; Reddy, Anireddy S.N.

doi: 10.1104/pp.111.900426pmid: 22213248

The nucleus is master of the cellular universe in eukaryotes and home to much of the genome and the machinery for its replication, maintenance, and expression. Although subnuclear compartments such as the nucleolus and Cajal bodies have been known for over a century, studies in the last 2 decades have clearly established that the nucleoplasm is organized into nonmembrane-bound subdomains and bodies. However, detailed understanding of nuclear organization and its dynamics in plants and how they impact DNA replication and gene expression remain to be elucidated. The goal of this Focus Issue is to examine, in the form of Updates, how recent advances contribute to the current understanding of nuclear architecture and dynamics in plant cells and their relevance to nuclear functions and developmental and physiological processes. The Updates also highlight areas that are ripe for mechanistic analysis and technical advances that will enable such studies. We hope that these articles will further stimulate research in this emerging area that has important implications for plant cell and developmental biology and plant responses to the environment. CHROMOSOME AND CHROMATIN ORGANIZATION AND DYNAMICS Pawlowski and his colleagues (Tiang et al., 2012) discuss recent advances in chromosome organization and dynamics made using improved fluorescent in situ hybridization protocols as well as live-cell imaging with fluorescent protein (FP)-tagged probes. In particular, they review the concept of chromosome territory in the interphase nucleus and chromosome organization and dynamics in other parts of the cell cycle. They also discuss how the various configurations of chromosome territory observed in different plant species may all achieve the result of bringing telomeric and subtelomeric regions into the vicinity of each other, impacting interchromosomal interactions. They also discuss recent studies that have allowed correlation of cytological phenomena, e.g. chromosome condensation, with molecular events, such as chromatin remodeling, that occur during meiotic prophase. These discussions underscore the progress of chromosome organization studies from descriptive to mechanistic dissection of its functional significance and regulation. Research on chromatin dynamics dramatically illustrates how nuclear dynamics on the molecular level contributes to gene expression, thereby controlling development. Holec and Berger (2012) compare and contrast the polycomb group complexes from plants and animals and focus on how chromatin modifications by these complexes confer epigenetic control of coordinated expression of gene networks to mediate major progressions in the plant life cycle. They use deposition, spreading, maintenance, and removal of the histone modification mark (methylation of Lys-27 of histone 3) that accompany the cycle of repression and derepression of the Flowering Locus C to illustrate how various protein and RNA factors mediate chromatin dynamics. They also highlight some major challenges in the field such as understanding the resetting mechanisms for chromatin modification during developmental transitions and how plants make the decision between prolonged maintenance of certain chromatin marks while remodeling others. Rapid advances have been made in the enzymology involved in regulating chromatin dynamics. Liu et al. (2012b) show that transposable element transcriptional reactivation in Arabidopsis (Arabidopsis thaliana) is accompanied by an increase in histone acetylation and methylation. They discovered that histone deacetylase6 and DNA methyltransferase physically interact; together, they mediate histone acetylation and modulate DNA methylation status, silencing the transposable element. NUCLEAR BODIES The proteins that control many nuclear processes are highly organized into discrete bodies within the nucleus (Misteli and Spector, 2011), which are also referred to as nuclear organelles, enriched in specific proteins and/or RNAs (Mao et al., 2011b). Several different types of nuclear bodies were identified and are thought to function in diverse nuclear activities including gene regulation, processing of different types of RNAs, proteolysis, and DNA replication (Mao et al., 2011b). Four Update articles in this Focus Issue discuss different nuclear bodies and their functions. Shaw and Brown (2012) provide a succinct overview of the classical and emerging knowledge about the nucleoli, the most prominent of nuclear compartments and the sites for rRNA synthesis, ribosome biogenesis, and a processing and assembly center for many RNA-based processes. They discuss the complexity of nucleolus RNAs and proteins in the context of the emerging understanding of the plurifunctional roles nucleoli play in the cell, including regulating cell cycle, cell death, and stress responses. Shaw and Brown also underscore the view of a dynamic nucleolus with a constant flux of proteins shuttling in and out and reflect on how the constant nucleation and deconstruction of the nucleolus may relate to some of the cellular processes it regulates. Much remains to be discovered regarding all the nucleolus activities and understanding why these activities are compartmentalized within a defined domain in the nucleus. Many aspects of plant growth and development from seed germination to flowering are regulated by light. Five classes of photoreceptors sense the light spectrum and elicit appropriate responses, mostly by altering gene expression (Franklin and Quail, 2010; Chen and Chory, 2011). Many of these photoreceptors move to the nucleus in a light-dependent manner and form discrete nuclear bodies called photobodies, which are unique to plants and are regulated by external light cues. The Update by Chen and his colleagues (Van Buskirk et al., 2012) presents recent developments pertinent to composition, dynamics, and potential roles of these bodies in light-regulated gene regulation. It also highlights the utility of forward genetic screens to isolate mutants that are defective in proper localization of photoreceptors to gain further insights into functions of photobodies. They also discuss different potential mechanisms by which photobodies regulate gene expression. A research article by Sokolova et al. (2012) describes how a missense mutation (Ala to Val at position 30) in the N-terminal domain of phytochrome A induces hyposensitivity to continuous low-intensity far-red light and reduces the very low fluency and far-red high irradiance responses. They have shown that this mutation results in reduced affinity to nuclear import facilitators, impairing its nuclear localization and, thereby, altering light responses. Noncoding small RNAs (micro-RNAs, small interfering RNA, and others) regulate gene expression both at the transcriptional levels by chromatin modifications and/or posttranscriptional level by regulating RNA stability or translation (Czech and Hannon, 2011; Liu et al., 2012a). The update article by Fang and his colleagues (Liu et al., 2012a) provides a nice overview of microRNA (miRNA) biogenesis pathways in plants. They summarize the current status of the composition and function of plant dicing bodies, which are distinct from other nuclear bodies and have been implicated in processing primary miRNA precursors into miRNA. They emphasize that further studies are needed to elucidate all potential roles of dicing bodies in various aspects of small RNA biogenesis and function. The dependence on constitutive splicing by a cell to generate functional transcripts and the prevalent use of alternative splicing to amplify the capacity of gene functions underscore the need to understand the mechanism of pre-mRNA processing. Reddy et al. (2012) provide an overview of conserved and unique aspects of mRNA processing in plants. Numerous splicing regulatory factors are known to shuttle between the nucleoplasm and the nuclear subdomains called nuclear speckles. Reddy and colleagues provide a systematic discussion of how a variety of recently developed approaches have revealed the temporal and spatial dynamics of these splicing factories. Developmental and environment signals may all affect the dynamics of the nuclear speckles, such as their sizes and the fluxes of some of the speckle-resident molecules in and out of these compartments. Fundamental aspects regarding these nuclear speckles, such as composition, biogenesis, and functional specialization of subpopulations of these dynamic nuclear structures remain to be elucidated. NUCLEAR ENVELOPE AND NUCLEOCYTOPLASMIC TRAFFICKING The nuclear envelope, which separates the nuclear compartment from cytoplasm, is implicated in the organization of chromatin and gene regulation. Nuclear pores regulate the movement of RNA, protein, and RNA-protein complexes into and out of the nucleus. The Update article from Meier and colleagues (Boruc et al., 2012) focuses on recent advances in elucidating the composition and dynamic organization of the nuclear envelopes and nuclear pore complexes. They further define their roles in selective nuclear import and export of macromolecules in plant cells. They also discuss the dynamics of nuclear envelope proteins during cell division and their additional roles in mitosis. Tiang et al. (2012) also discuss the involvement of the nuclear envelope in the formation of telomere bouquet during meiotic prophase. Most signaling pathways culminate in the nucleus, leading to regulation of expression of specific genes whose products are necessary for eliciting a signal-specific response. Regulated trafficking of proteins, RNAs, RNA-protein complexes, and other molecules into and out of the nucleus is important in diverse processes. The Update by Rivas (2012) covers emerging roles of nuclear trafficking in plant immune responses. She discusses how effectors from diverse pathogens are targeted to the nucleus by coopting the host nuclear import machinery to alter host transcription. She presents an overview of how regulated nuclear localization of pathogen effectors, R proteins, and other host defense signaling proteins modulate plant immunity. Mutations in genes encoding proteins that are part of the nuclear pore complex and nucleocytoplasmic trafficking machinery have been shown to impair plant defense responses, suggesting the importance of dynamic translocation of proteins into the nucleus in regulating the expression of plant defense genes. New kinds of FPs, called optical highlighters, such as photoactivatable FPs that can be activated to produce fluorescence from a quiescent state or photoconvertible FPs that can be converted from one fluorescent state to the other (e.g. orange to red or green to red), are serving as powerful probes to investigate dynamics of proteins and organelles (Shaner et al., 2007). Monomeric EosFP, a photoconvertible FP, changes its fluorescence irreversibly from green to red when it is exposed to violet-blue light. A research article contributed by Wozny et al. (2012) reports use of monomeric EosFP fused to a histone to monitor increase in DNA content during endoreduplication based on a shift of red fluorescent nuclei to a green fluorescence state. The use of such optical highlighters (Shaner et al., 2007) offers new avenues to investigate many aspects of plant nuclear architecture and dynamics in live cells. PERSPECTIVES Knowledge about the organization of the chromosomes, chromatin, and nuclear bodies in the nucleus and the role such organization plays in nuclear processes is vital to understand how plants grow, develop, and respond to a variety of internal and external cues. As summarized in the articles of this Focus Issue, we are beginning to gain insights into nuclear organization, nuclear compartments and their functions, and the mechanisms that regulate their dynamics in real time. Although targeting signals of some of the resident proteins of plant nuclear bodies are identified (Liu et al., 2012a; Reddy et al., 2012; Van Buskirk et al., 2012), the biogenesis of these subnuclear structures remains largely unknown. Although some clues and experimental approaches may be learned from their animal counterparts (Sharma et al., 2010; Mao et al., 2011a), unique mechanisms supporting the growth and developmental strategies specific to plants are likely to be discovered. The full composition of most plant nuclear bodies is not known. This will require isolation of each type of nuclear body using biochemical and/or microscopic methods followed by proteomic analysis. Also, little is known about the interrelationship(s) between different types of plant nuclear bodies. Simultaneous labeling of multiple nuclear bodies with different FPs fused to proteins that define individual types of nuclear bodies in a single nucleus may provide some clues about this. Precise functions of most of the plant nuclear bodies are still not known. Future studies using genetic, cytological, cell biological, and proteomic approaches and emerging tools are necessary to address these questions. Further elucidation of spatial and temporal organization of nuclear processes and associated factors is necessary to advance understanding of how nuclear architecture and dynamics impact nuclear functions. ACKNOWLEDGMENTS We thank all the authors who contributed to this Focus Issue for their efforts and the editorial staff, in particular Jon Munn, L. Ash Csiko, and A.K. Grennan, for their expert help on the editorial process and design of the cover. 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Tiang, Choon-Lin; He, Yan; Pawlowski, Wojciech P.

doi: 10.1104/pp.111.187161pmid: 22095045

Chromosomes are key building blocks of eukaryotic genomes. Studies on chromosome organization and dynamics not only address questions of how chromosomes behave and what mechanisms control this behavior but also examine how chromosome organization and dynamics affect gene expression and genome maintenance. A number of important studies on chromosome organization and dynamics have been conducted in plants in the past few years. Many of them have been made possible by recent advances in cytogenetics tools, including improvements in fluorescent in situ hybridization (FISH) protocols and development of live-imaging techniques. To the most significant discoveries belong elucidating the chromosome arrangement in interphase nuclei in Arabidopsis (Arabidopsis thaliana) and finding that interphase chromosome organization is controlled by both genetic and environmental factors. Other notable studies included elucidation of the role of the Pairing homoeologous1 (Ph1) locus in chromosome interactions in somatic and meiotic cells in wheat (Triticum aestivum), identification of the link between homologous chromosome pairing in meiosis and recombination, and discovery of rapid chromosome movements in meiotic prophase. Investigations of chromosome organization and arrangement in the nucleus have been conducted since the invention of the light microscope. With the development of molecular cytogenetics tools, these studies matured from mostly descriptive to more mechanism driven that aim to elucidate factors controlling chromosome organization and dynamics. Until very recently, chromosomes, particularly in plants and other multicellular eukaryotes, were mostly examined in fixed cells. These observations, although static themselves, provided indications that chromosome behavior is quite dynamic. Introduction of new microscopy methods that allow observations of chromosomes in live cells has confirmed the dynamic nature of chromosomes and enabled better understanding of the complexities of chromosome behavior. In this review, we mainly focus on two aspects of chromosome organization and dynamics that have received the most attention in the past few years in plant studies: chromosome organization in interphase nuclei and organization and dynamics of chromosomes during the prophase of meiosis. CHROMOSOME ORGANIZATION AND DYNAMICS IN THE INTERPHASE NUCLEUS Although organization of chromatin in interphase nuclei has been a subject of speculations for several decades, the past few years have brought much better understanding of this issue. In plants, studies conducted during the last 10 years resulted in elucidating interphase chromatin organization in Arabidopsis. A driving force behind this research is the desire to understand how organization of interphase chromosomes affects gene expression, although such studies are only now beginning in plants and other multicellular eukaryotes. Chromosome Territories During interphase, chromosomes assume a largely decondensed state. However, chromatin is still nonrandomly arranged within the nuclear space. Each chromosome occupies a limited, exclusive nuclear subdomain, known as a chromosome territory. The concept of chromosome territories was proposed by Carl Rabl in 1885, based on his observation of salamander cell division. Existence of chromosome territories was confirmed in the 1980s in human cells using FISH with chromosome-specific DNA probes (Manuelidis and Borden, 1988). Chromosome territories in plants were first visualized in Arabidopsis using chromosome-specific bacterial artificial chromosome FISH probes (Lysak et al., 2001). Rabl Configuration In many plant species with relatively large genomes, chromosomes during interphase adopt Rabl configuration (Cowan et al., 2001). This term describes an interphase chromosome arrangement in which centromeres and telomeres are located at opposite sides of the nucleus (Fig. 1A). This configuration is thought to be a remnant of a preceding anaphase. In some of these species, such as wheat or barley (Hordeum vulgare), Rabl configuration persists in all cells throughout the plant (Anamthawat-Jonsson and Heslop-Harrison, 1990). In rice (Oryza sativa), on the other hand, Rabl is only observed in certain tissues, such as xylem cells in the root and undifferentiated cells in the anther (Prieto et al., 2004a; Santos and Shaw, 2004). Other plant species, such as maize (Zea mays) and sorghum (Sorghum bicolor), despite having fairly large genomes, are not known to exhibit Rabl configuration at all (Dong and Jiang, 1998). In these species, chromosomes lose their polarized anaphase distribution of centromeres and telomeres after entering interphase. Figure 1. Open in new tabDownload slide Patterns of chromosome arrangement in the nucleus. A, Rabl configuration found in interphase nuclei of many large-genome plant species. B, Rosette-like organization of chromosomes in interphase nuclei in Arabidopsis. C, Telomere bouquet. Figure 1. Open in new tabDownload slide Patterns of chromosome arrangement in the nucleus. A, Rabl configuration found in interphase nuclei of many large-genome plant species. B, Rosette-like organization of chromosomes in interphase nuclei in Arabidopsis. C, Telomere bouquet. Interphase Chromosome Organization in Arabidopsis Interphase chromosomes in Arabidopsis do not display Rabl configuration but exhibit a strikingly different type of chromatin arrangement. In this species, telomeres cluster around the nucleolus while centromeres are located at the nuclear periphery (Armstrong et al., 2001; Fransz et al., 2002). Arabidopsis centromeric heterochromatin forms distinct, dense bodies called chromocenters. Chromocenters contain the majority of genomic repeats and exhibit epigenetic marks of inactive chromatin (Fransz et al., 2002). From the chromocenters, euchromatic loops of 0.2 to 2 Mb in length emanate, resulting in a rosette-like structure of Arabidopsis chromosome territories (Fig. 1B). Chromocenters of most Arabidopsis chromosomes do not seem to show preferential positioning relative to each other (Pecinka et al., 2004; Berr and Schubert, 2007; de Nooijer et al., 2009). Exceptions to this rule are chromosomes carrying nucleolar organizing regions (NORs), which contain tandemly arranged copies of rRNA genes (Pecinka et al., 2004). Physical association of NORs with the nucleolus is likely responsible for the nonrandom association of the NOR-bearing chromosomes. Although centromeres in Arabidopsis interphase nuclei do not cluster, telomeres show persistent clustering at the nucleolus (Armstrong et al., 2001). This phenomenon is not related to Rabl configuration but, similarly to Rabl, results in bringing certain chromosome regions into close vicinity of each other, which may have direct effects on interchromosome interactions and dynamics. Factors Affecting Interphase Chromosome Organization The arrangement of chromosome territories within the nucleus exhibits dynamic changes in response to various internal and external conditions. Histone modification and DNA methylation patterns are expected to affect chromosome organization, although data on this subject are still scarce. Nevertheless, it has been shown that in rice DNA demethylation causes chromatin decondensation and induces Rabl configuration in those tissues in which Rabl is not normally present (Santos et al., 2011). Changes in the ploidy level generated by endoreduplication have been shown to affect chromosome arrangement in Arabidopsis (Berr and Schubert, 2007). The shape and size of the nucleus is also related to chromosome arrangement, although it is not clear whether chromosome organization is the cause or a result of altered nuclear size and/or shape (Berr and Schubert, 2007; Dittmer et al., 2007). Chromosome organization has been shown to change during plant development and in response to the environment. Chromocenters become smaller in leaves prior to the transition to reproductive development and recover to their former size after the elongation of the floral stem (Tessadori et al., 2007). Both processes are affected by light conditions. Furthermore, Arabidopsis genotypes acclimated to different latitudes exhibit genetically programmed levels of chromatin compaction, depending on the light intensity of their original habitats (Tessadori et al., 2009). In rice, heat stress has been shown to induce chromatin decondensation (Santos et al., 2011). Functional Implications of Interphase Chromosome Organization In the past few years, there has been a growing interest in understanding how chromosome and chromatin arrangement in interphase nuclei affect gene activity. Arrangement of chromosome territories that brings certain chromosome regions together has the potential to contribute to regulation of gene expression. This notion has lead to development of the concept of transcriptional factories, discrete sites in the nucleus where gene transcription is particularly active (Sutherland and Bickmore, 2009). Hundreds of such factories are proposed to be present in each nucleus and they are thought to be anchored to a nuclear substructure. On the other hand, there might be also heterochromatic neighborhoods in which gene expression is silenced. It has been proposed that physical interactions between gene copies located on different chromosomes may contribute to gene silencing (Lanctôt et al., 2007). Effects of chromatin organization on gene expression are poorly understood in plants. In Arabidopsis, the majority of genes are located on euchromatic loops stretching out of the chromocenters (Fransz et al., 2002). However, it is unclear if there are particular nucleus regions that are occupied by highly expressed genes. We anticipate that the near future will bring a more complete picture of interphase chromosome arrangement and dynamics during growth and development as well as under various environmental conditions in plants. These data will be a starting point for understanding how interphase chromosome arrangement affects gene expression. Interphase Chromosome Dynamics In the past few years new tools have been developed to facilitate investigations of interphase chromosome dynamics in live cells in Arabidopsis (Fang and Spector, 2005; Matzke et al., 2005, 2008, 2010). However, so far, it appears that interphase chromosomes only display mostly limited, diffusive movements (Kato and Lam, 2003; Fang and Spector, 2005). Interstitial chromosome regions generally exhibit more movements than centromeres. Interestingly, endoreduplication-driven polyploidy has been found to reduce chromosome movement speed but increase the freedom of movement, i.e. the area within the nucleus to which movement of a chromosome segment is constrained (Kato and Lam, 2003). Overall, chromosome dynamics in interphase nuclei is still quite poorly understood. Further development of live-imaging tools should lead to substantial progress in this area, particularly in understanding the implications of interphase chromosome motility for gene activity as well as for genome maintenance processes, such as DNA replication or repair. CHROMOSOME DYNAMICS IN MITOSIS Even though chromosome segregation in mitosis is one of the most obvious and easily observable types of nuclear dynamics, patterns and mechanisms of mitotic chromosome segregation have, so far, been relatively poorly researched in plants. Nevertheless, live imaging of chromosomes during mitosis in root meristematic cells in Arabidopsis have yielded several interesting data (Fang and Spector, 2005). Chromosomes in mitosis show the most dynamic behavior during their congression to metaphase plate at the transition from prophase and metaphase and during their segregation in anaphase. During the prophase to metaphase transition, after breakdown of the nuclear envelope, condensed chromosomes relocate to the center of the cell and their centromeric regions gradually rotate to become oriented perpendicular to the metaphase plate (Fang and Spector, 2005). In anaphase, chromosomes move, centromere first, toward the opposite poles. This movement is not synchronous among all centromeres in the cell. Furthermore, a centromere may first start moving to one of the poles and later change direction and move to the other pole (Fang and Spector, 2005). Following anaphase, chromosomes assume the interphase configuration. However, chromosome positions and chromocenter arrangement in the nucleus in the daughter cells are not the same as in the mother cell (Fang and Spector, 2005; Berr and Schubert, 2007). On the other hand, chromosome positions in the two daughter cells often show mirror symmetry immediately after mitosis (Berr and Schubert, 2007). CHROMOSOME DYNAMICS IN MEIOTIC PROPHASE Prophase of the first division of meiosis is a period of some of the most dynamic chromosome behavior. During this time, chromatin undergoes major reorganization that includes: (1) chromosome condensation and establishment of meiotic chromosome structure, (2) pairing of homologous chromosomes, and (3) dynamic chromosome movements. The result of these activities is formation of stable chromosome pairs, the bivalents, which is essential for ensuring correct chromosome segregation at the end of meiosis. Chromosome Condensation Condensation is the most noticeable change in chromosome appearance in early meiosis and serves as the main criterion for dividing meiosis prophase into substages (Fig. 2). Chromatin condensation in leptotene, in addition to making chromosomes more compact, leads to establishment of a meiosis-specific chromosome structure. Adoption of meiosis chromosome structure is required for key processes of meiotic prophase I (Dawe et al., 1994). Studies in maize lacking ABSENCE OF FIRST DIVISION1, an α-kleisin participating in formation of the chromosome axis at the onset of meiosis, showed that proper chromosome structure is essential for meiotic recombination as well as chromosome pairing (Golubovskaya et al., 2006). Meiosis-specific patterns of chromatin remodeling have been also implicated in preconditioning specific chromosome regions to become sites of meiotic recombination events in mouse (Mus musculus) and budding yeast (Saccharomyces cerevisiae) studies (Borde et al., 2009; Baudat et al., 2010). Transcriptome analyses of Arabidopsis meiocytes showed that a staggering number of genes are expressed during meiotic prophase I (Chen et al., 2010). These gene expression patterns are also presumably results of chromatin remodeling in early meiosis. It remains to be seen whether all these genes are indeed needed for meiosis progression and their large number reflects the complexity of meiotic prophase, or whether the massive gene expression is a by-product of genome-wide chromatin opening to facilitate chromosome pairing and recombination. Figure 2. Open in new tabDownload slide A diagram showing major events and processes of meiotic prophase I. Only two chromosomes, each in different color, are shown in the diagram on the left. Figure 2. Open in new tabDownload slide A diagram showing major events and processes of meiotic prophase I. Only two chromosomes, each in different color, are shown in the diagram on the left. Pairing of Homologous Chromosomes Chromosome pairing is a process in which two homologous chromosome copies find each other among all chromosomes in the nucleus and juxtapose. Pairing includes bringing chromosomes together into a close proximity as well as an intimate homology search to recognize the correct pairing partner. Pairing interactions at select locations are followed by alignment along the entire length of the chromosomes. Homologous chromosome pairing in plants generally proceeds de novo at the onset of meiotic prophase and there is little evidence for persistent pairing of homologous chromosomes prior to meiosis. Some elements of interphase chromosome arrangement may, however, facilitate meiotic pairing. The Rabl-induced interphase centromere clustering in polyploid wheat affects progression of homologous pairing (Martinez-Perez et al., 2001). Similarly, the interphase telomere association with the nucleolus in Arabidopsis has been hypothesized to act in prealigning chromosomes and aiding pairing interactions in telomeric and subtelomeric regions (Armstrong et al., 2001). The basis of chromosome homology recognition in most species, including plants, is the DNA sequence along the entire chromosome. However, this rule does not exclude a potential for a role of chromatin states and modification patterns in chromosome pairing. Although considerable progress has been made during the past decade in understanding the biological nature of chromosome pairing, it is still one of the least-explored aspects of meiosis. Several meiotic processes are known to contribute to homologous chromosome pairing, including meiotic recombination, chromosome motility in early substages of meiotic prophase, and formation of the telomere bouquet (see below). Homologous Chromosome Pairing and Recombination Studies in a variety of eukaryotes, including plant model species Arabidopsis and maize, suggest that successful completion of homologous chromosome pairing is tightly linked to the progression of meiotic recombination (Franklin et al., 1999; Li et al., 2004; Pawlowski et al., 2004; Ronceret et al., 2009). This intimate dependence of pairing on recombination exists also in fungi and mammals (Baudat et al., 2000; Peoples-Holst and Burgess, 2005), but, interestingly, not in Drosophila melanogaster or Caenorhabditis elegans (Dernburg et al., 1998; McKim et al., 1998). Meiotic recombination is universally initiated by formation of double-strand breaks (DSBs) in chromosomal DNA (Fig. 2) by a conserved topoisomerase-like protein SPO11 (Lichten, 2001). Subsequently, the DSBs are resected, leading to formation of single-stranded DNA overhangs. Single-stranded DNA ends, which are several-hundred base pair long, invade dsDNA regions on the homologous chromosomes. This process, known as single-end invasion (SEI), is thought to be the basis of homology recognition during chromosome pairing in plants, fungi, and mammals (Bozza and Pawlowski, 2008). Defects in chromosome pairing have been observed in plant mutants in genes controlling DSB formation and resection, as well as SEI (Pawlowski and Cande, 2005). In most eukaryotes with relatively large genomes, including plants, the number of SEI sites is far greater than the number of crossovers. In maize, there are, on average, about 20 crossover sites per meiocyte, but as many as 500 SEI sites. These sites can be identified by immunolocalizing proteins that facilitate the SEI process, such as RAD51 (Fig. 2). RAD51 forms distinct foci on chromosomes in zygotene and pachytene. Studies in maize suggest that most, if not all, of the SEI sites are required to facilitate correct chromosome pairing. Mutants that exhibit reduced number of RAD51 foci show chromosome pairing defects as well (Pawlowski et al., 2003; Ronceret et al., 2009). Chromosome Pairing and Genome Complexity Although the dependence of pairing on recombination has been recognized, the exact nature of this link is not yet fully understood. Particularly, it is not clear how ectopic pairing is prevented between repetitive genome regions. For example, about 85% of the maize genome consists of repetitive DNA elements, many of which are several kilobase pair long (Schnable et al., 2009). These data suggest that there must be mechanisms that coordinate pairing along the entire length of chromosomes so that bivalents are only formed between homologous chromosome partners. However, the nature of these mechanisms remains unknown. Polyploidy, which is frequent in many plant families, adds another level of complication to the process of pairing. While in autopolyploids chromosome pairing is generally disturbed and may lead to formation of multivalents, allopolyploid species have evolved mechanisms that can distinguish between homologous and homeologous chromosomes (i.e. chromosomes derived from different progenitors that are similar but not identical). Studies in polyploid wheat have demonstrated that homeologous associations between chromosomes from different genomes are suppressed by the Ph1 locus (Moore and Shaw, 2009). Absence of Ph1 leads to incorrect chromosome pairing (Al-Kaff et al., 2008). Ph1 is proposed to act by controlling remodeling of chromatin structure at the onset of meiosis (Prieto et al., 2004b; Colas et al., 2008). The chromatin conformational change affects the homology search and, in particular, the specificity of interactions between wheat centromeres (Martinez-Perez et al., 2001). In the presence of Ph1, associations between centromeres of homeologous chromosomes become disrupted and only homologous centromere interactions remain. The Ph1 locus has been defined to a single wheat chromosome region that contains a cluster of genes related to the cell cycle regulator Cyclin-dependent kinase2 (Cdk2) gene (Griffiths et al., 2006). Cdk2 is known to control meiosis progression, expression of meiotic genes, meiotic DSB formation, as well as chromatin structure (Yousafzai et al., 2010). The function of Ph1 can be mimicked by application of okadaic acid, a drug known to induce chromosome condensation and affect meiosis progression by altering phosphorylation of the H1 histone (Knight et al., 2010). These data imply that histone phosphorylation and chromosome condensation may affect the chromosome pairing dynamics. The presence of the Cdk2 gene cluster appears to be specific to tetraploid and hexaploid wheat (Griffiths et al., 2006). This observation implies that, even though they could exploit the same aspect of chromosome dynamics, mechanisms for preventing homeologous pairing have likely evolved many times independently in different polyploid taxa. Although the mechanism of Ph1 function still remains to be fully elucidated, it suggests existence, at least in some plant species, of chromatin-level homology recognition mechanisms that operate in addition to the DSB-dependent mechanism of homology search (Moore and Shaw, 2009). Chromosome Motility in Meiotic Prophase I Live-imaging observations in a number of species of plants, animals, and fungi, most of them conducted during the past few years, have demonstrated that early stages of meiotic prophase are a period of extremely dynamic chromosome movements (Koszul and Kleckner, 2009; Sheehan and Pawlowski, 2009; Baudrimont et al., 2010). In plants, studies using intact live anthers of maize showed that meiotic chromosomes exhibit complex and stage-specific motility patterns in zygotene and pachytene (Sheehan and Pawlowski, 2009). During zygotene, short chromosome segments adjacent to chromosome ends exhibit robust short-range movements, while movements of interstitial chromosome segments are more restrained. At the same time, the entire chromatin in the nucleus rotates back and forth in a coordinated manner at angles ranging from 7° to 10°, but sometimes as much as 90°. In pachytene, the rapid short-range chromosome end movements are replaced by slower but long-distance movements of much larger chromosome segments. The rotational movements, in contrast, persist through pachytene. Prophase chromosome movements in maize appear more complex in comparison to other species, as they include both coordinated chromatin rotations as well as movements of individual chromosome segments. In contrast to maize, only uncoordinated movements of individual chromosomes or chromosome segments are seen in budding yeast, while only coordinated movements of the entire chromatin have been reported in fission yeast (Schizosaccharomyces pombe) and rat (Rattus norvegicus) spermatocytes (Sheehan and Pawlowski, 2009). The significance of the prophase chromosome movements is not yet entirely understood. It has been suggested that zygotene chromosome movements may aid homologous chromosome pairing by facilitating interchromosome interactions and disrupting associations of nonhomologous chromosomes (Koszul and Kleckner, 2009; Sheehan and Pawlowski, 2009). The pachytene movements, on the other hand, may help resolve chromosome entanglements (known as interlocks) that form during chromosome pairing in zygotene. Interlocks have to be disentangled prior to further chromosome condensation and segregation or chromosome breakage may occur. The Role of Telomeres during Early Meiotic Prophase Chromosome ends (telomeres) play a critical role in chromosome dynamics during meiotic prophase. In many species of plants, animals, and fungi, telomeres attach to the nuclear envelope and cluster within a small region, leading to formation of the telomere bouquet (Figs. 1C and 2; Harper et al., 2004). In yeast and mammals, telomeres cluster at the microtubule-organizing center. Plants do not have microtubule-organizing centers but immunocytological studies in rye (Secale cereale) showed that telomeres cluster to form the bouquet opposite a band of microtubules in early zygotene nucleus (Cowan et al., 2002). While telomeres cluster, centromeres do not, although they are generally located on the opposite side of the nucleus from the bouquet. This organization results in an overall polarization of the nucleus that is somewhat similar to Rabl configuration. However, the mechanism of the bouquet formation and the function of the bouquet are very different than those of Rabl configuration. Based on analyses of mutants defective in the bouquet formation, it has been speculated that telomere clustering facilitates homologous pairing by bringing chromosome ends together (Harper et al., 2004). The plural abnormalities of meiosis1 (pam1) mutant of maize is one of the best-studied bouquet mutants in plants (Golubovskaya et al., 2002). Cytological analysis of pam1 showed that in early meiotic prophase telomeres in mutant meiocytes associate with the nuclear envelope but fail to cluster. The mutant also exhibits a reduction in pairing of homologous chromosomes and shows unresolved chromosome interlocks, all presumably resulting from the telomere bouquet formation defect. Arabidopsis belongs to a small group of species (also including C. elegans and Drosophila) that do not form telomere bouquets (Harper et al., 2004). However, the clustering of Arabidopsis telomeres at the nucleolus present in interphase tends to persist into early meiosis, although the telomeres dissociate from the nucleolus during the course of leptotene and become widely dispersed within the nucleus (Armstrong et al., 2001). Subtelomeric chromosome regions begin to homologously pair prior to telomere detachment from the nucleolus. Based on this sequence of events, Armstrong et al. (2001) suggested that the premeiotic and early meiotic association of telomeres with the nucleolus in Arabidopsis may play a role similar to that of the bouquet in other species. Although Arabidopsis telomeres are not attached to the nuclear envelope during their nucleolus association, they do become transiently associated with the nuclear envelope during zygotene and occasionally exhibit loose clustering, although not classical bouquet formation. Telomere Attachment to the Nuclear Envelope in Meiotic Prophase Attachment of telomeres to the nuclear envelope during formation of the telomere bouquet is the basis of meiotic prophase chromosome dynamics. The telomere-nuclear envelope attachment is mediated by a multiprotein complex (Fig. 3). Proteins involved in this complex bridge the double-membrane nuclear envelope, tethering telomeres on the inner side of the nuclear envelope and linking them to the cytoskeleton on the outside. Several proteins involved in this complex have been identified in a variety of species. The best studied of them are proteins containing the conserved SUN (for Sad1p, UNC-84) domain. Homologs of these proteins have been identified in budding yeast and fission yeast mammals, C. elegans, as well as plants, maize, and Arabidopsis. SUN domain proteins bridge the inner membrane of the nuclear envelope. On their N terminus, they interact with telomere-binding proteins, while the C terminus is located in the lumen between the inner and outer nuclear membrane (Schmitt et al., 2007). The Arabidopsis genome encodes two SUN domain proteins, AtSUN1 and AtSUN2 (Graumann et al., 2010). Similarly to the SUN domain proteins from other in species, AtSUN1 and AtSUN2 localize to the inner nuclear membrane. However, this localization pattern has only been demonstrated so far in somatic cells and it remains to be shown whether the two proteins also function in meiosis. Figure 3. Open in new tabDownload slide A diagram showing the telomere-nuclear envelope attachment involved in formation of the telomere bouquet. Figure 3. Open in new tabDownload slide A diagram showing the telomere-nuclear envelope attachment involved in formation of the telomere bouquet. Several telomere-binding proteins that interact with SUN domain proteins have been identified in fission yeast (Chikashige et al., 2006). However, homologs of these proteins have not been found yet in plants or other species as their sequences are fairly poorly conserved. It is also unclear what specific role the actual telomeres play in chromosome end attachment to the nuclear envelope and the bouquet formation. In the mouse, lack of the telomerase enzyme, which maintains telomeres and preserves their length, leads to defects in telomere bouquet formation and chromosome behavior (Liu et al., 2004), suggesting that presence of telomeric DNA repeats is important for bouquet function. In yeast and mammals, it has been shown that the C-terminal part of SUN domain proteins interacts with another type of transmembrane proteins known as KASH (for Klarsicht, ANC-1, Syne Homology; Chikashige et al., 2007). On their cytoplasmic sides, KASH proteins interact with proteins that bind the cytoskeleton. Presumably, KASH proteins are also present in plants, although so far, this fact has not been directly demonstrated. The amino acid sequence of KASH proteins is much less conserved that the sequence of SUN domain proteins so identifying KASH protein homologs by sequence alone is difficult. The Role of Cytoskeleton in Chromosome Dynamics The SUN-KASH protein complex provides a physical link between chromosomes and the cytoskeleton. Analyses of meiotic prophase chromosome dynamics in a number of species, including maize, indicate that the physical forces responsible for chromosome movements are generated in the cellular cytoskeleton (Koszul and Kleckner, 2009; Sheehan and Pawlowski, 2009). From there, they are conveyed onto the nuclear envelope and then, by the virtue of telomere attachment to the nuclear envelope, further onto chromosome ends. Sheehan and Pawlowski showed that treating maize anthers with cytoskeleton-disrupting drugs, colchicine, which prevents tubulin polymerization, and latrunculin B, an inhibitor of actin polymerization, leads to complete cessation of prophase chromosome movements, as well as movements of the nuclear envelope, which accompany chromosome motility (Sheehan and Pawlowski, 2009). These data suggest that both actin and tubulin cytoskeletons play critical roles in meiotic prophase chromosome dynamics. Interestingly, the link between cytoskeleton and prophase chromosome dynamics in plants had been identified indirectly even before the development of methods for live imaging of meiotic chromosomes. In the 1970s, Driscoll and Darvey showed that colchicine disrupts homologous chromosome pairing (Driscoll and Darvey, 1970). Furthermore, Cowan and Cande demonstrated that colchicine forestalls the bouquet formation (Cowan and Cande, 2002). Studies using cytoskeleton-disrupting drugs have also been conducted in species outside of plants. Interestingly, these studies uncovered that different cytoskeletal components are involved in chromosome motility in different species. In fission yeast and mammals, chromosome movements require the microtubule cytoskeleton (Salonen et al., 1982; Ding et al., 1998) whereas in budding yeast, the actin cytoskeleton is used for this purpose (Scherthan et al., 2007; Koszul et al., 2008). How exactly the cytoskeleton generates the various types of nuclear and chromosomal movements during meiotic prophase is not yet clear. Further studies to elucidate the organization of the meiocyte cytoskeleton are needed to elucidate these dynamics. However, observations of the effects of cytoskeleton-disrupting drugs on chromosome movements, bouquet formation, and meiotic prophase progression already shed new light on the function of the telomere bouquet. These studies suggest that, rather than brining chromosome ends together, the main role of the bouquet is facilitating chromosome motility by conveying movement-generating forces from the cytoskeleton to chromosomes. Future studies combining genetic dissection of bouquet components with live-imaging observations will help elucidate this issue. CONCLUSION AND PERSPECTIVES Several important advances have been made in the past few years in studies on chromosome organization and dynamics in plants. These studies have addressed various aspects of chromosome organization and behavior in interphase cells, as well as mitosis and meiosis. They also examined effects that chromosome organization and dynamics have on the key nuclear functions of maintenance, transcription, and transmission to progeny of genetic material. Results of some of the studies have shown that plants exhibit patterns of chromosome organization and behavior that are similar to those found in animals and fungi, such as existence of chromosome territories or the telomere bouquet formation. Other studies, however, have revealed plant-specific modifications of universal mechanisms, or existence of mechanisms that are entirely plant specific, specific to a certain group of plants, or even to individual species. An excellent example of the latter is the proposed mechanism of the Ph1 locus function, which employs a unique way of regulating activity of the widely conserved CDK2 protein to accomplish a function specifically needed in a polyploid species with highly similar ancestral genomes. More important than the individual discoveries, however, has been the fact that chromosome research in plants has moved from mostly descriptive studies to hypothesis-driven research addressing the mechanisms of chromosome behavior. 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Plant J 50 : 848 – 857 Google Scholar Crossref Search ADS PubMed WorldCat Tessadori F van Zanten M Pavlova P Clifton R Pontvianne F Snoek LB Millenaar FF Schulkes RK van Driel R Voesenek LA et al. ( 2009 ) Phytochrome B and histone deacetylase 6 control light-induced chromatin compaction in Arabidopsis thaliana . PLoS Genet 5 : e1000638 Google Scholar Crossref Search ADS PubMed WorldCat Yousafzai FK Al-Kaff N Moore G ( 2010 ) Structural and functional relationship between the Ph1 locus protein 5B2 in wheat and CDK2 in mammals . Funct Integr Genomics 10 : 157 – 166 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by the National Science Foundation (grant no. IOS–1025881) and the U.S. Department of Agriculture-National Institute of Food and Agriculture (grant no. NYC–149562). 2 These authors contributed equally to the article. * Corresponding author; e-mail [email protected]. www.plantphysiol.org/cgi/doi/10.1104/pp.111.187161 © 2012 American Society of Plant Biologists. All rights reserved. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Holec, Sarah; Berger, Frédéric

doi: 10.1104/pp.111.186445pmid: 22086420

The modulation of transcription by chromatin modifications participates in the coordination of gene networks regulating development. Chromatin marks deposited by Polycomb group (PcG) complexes induce a repressive state of the transcription, which is propagated through cell division. Here, we focus on the implications of this epigenetic regulation in the development of flowering plants like Arabidopsis. We present the mechanism of chromatin modification by PcG and its modulation by other factors. We discuss in detail the mechanisms leading to flowering and illustrate how PcG controls major progressions through the life cycle. The life cycle of eukaryotes is marked by developmental phases characterized by specific spatial and temporal regulation of genome expression. Dynamic regulation of chromatin state is crucial to ensure proper regulation of gene expression. The mechanisms involved in this regulation comprise nuclear localization, DNA methylation, histone variant replacement, and histone posttranslational modifications that recruit or release various chromatin remodeling factors and govern nucleosome occupancy on the DNA (Margueron and Reinberg, 2010). Among the regulators of chromatin state and transcription, Polycomb group (PcG) genes were first discovered in Drosophila melanogaster as repressors of the homeotic Hox genes involved in embryo segmentation (Lewis, 1978; Jürgens, 1985). Four PcG proteins form the core Polycomb Repressive Complex2 (PRC2): Enhancer of Zeste [E(z)], Suppressor of Zeste12 [Su(z)12], Extra sex combs (Esc), and p55. When properly assembled in a protein complex by p55 and Esc, E(z) trimethylates histone H3 Lys-27 (H3K27me3) at target loci (Nekrasov et al., 2005). This mark induces a durable transcriptional silencing (Fig. 1). In the absence of PcG activity, the initial pattern of Hox genes breaks down, causing embryo developmental arrest. Figure 1. Open in new tabDownload slide Action of Polycomb and Trithorax complexes. PRC2 complexes trimethylate the Lys-27 residue of H3K27me3 and induce a silenced state of the chromatin at the target locus. This mark is recognized by the PRC1 complex. In animals, this complex then monoubiquitinates H2A and triggers a compaction of the chromatin into a heterochromatin state that stably represses expression. The components of PRC1 complex are not fully described in plants so far. PRC2 activity of repression is antagonized by the function of Trithorax (TRX). Trithorax trimethylates the histone H3 Lys-4 residue (H3K4me3) and induces the expression of target loci. Although Trithorax function has been found in plants, the complex has not been identified yet. Figure 1. Open in new tabDownload slide Action of Polycomb and Trithorax complexes. PRC2 complexes trimethylate the Lys-27 residue of H3K27me3 and induce a silenced state of the chromatin at the target locus. This mark is recognized by the PRC1 complex. In animals, this complex then monoubiquitinates H2A and triggers a compaction of the chromatin into a heterochromatin state that stably represses expression. The components of PRC1 complex are not fully described in plants so far. PRC2 activity of repression is antagonized by the function of Trithorax (TRX). Trithorax trimethylates the histone H3 Lys-4 residue (H3K4me3) and induces the expression of target loci. Although Trithorax function has been found in plants, the complex has not been identified yet. PcG proteins were likely present in the last common ancestor of eukaryotes and are conserved from unicellular organisms to metazoans, and plants but were lost in yeast (Shaver et al., 2010; Margueron and Reinberg, 2011). Here, we summarize the nature of plant PcG complexes and their mode of action. We describe how PcG complexes interact with other chromatin-modifying complexes and influence gene expression. Furthermore, we detail the specific role of plant PcG complexes in developmental transitions that require coordinated regulation of gene networks. THE MOLECULAR ACTORS OF PCG-MEDIATED REPRESSION In Arabidopsis (Arabidopsis thaliana), PcG proteins form a family of eight homologs of PRC2 components. The homologs of E(z) are MEDEA, CURLY LEAF (CLF), and SWINGER. Su(z)12 also has three homologs, EMBRYONIC FLOWER2 (EMF2), FERTILIZATION INDEPENDENT SEED2 (FIS2), and VERNALIZATION2 (VRN2). Esc and p55 have the single homologs FERTILIZATION INDEPENDENT ENDOSPERM (FIE) and MULTICOPY SUPPRESSOR OF IRA1 (MSI1), respectively (Fig. 2). Figure 2. Open in new tabDownload slide Components of complexes associated with PRC2, PRC1, or Trithorax functions in Arabidopsis. While PRC2 complexes are well established, the other complexes are currently hypothetical. Figure 2. Open in new tabDownload slide Components of complexes associated with PRC2, PRC1, or Trithorax functions in Arabidopsis. While PRC2 complexes are well established, the other complexes are currently hypothetical. FIE and MSI1 are expressed in all cell types, but the expression of other PRC2 homologs is more restricted, and molecular and genetic evidences support the idea of at least three forms of PRC2 in plants: FIS, VRN, and EMF, as per the different Su(z)12 homologs (Spillane et al., 2000; Yoshida et al., 2001; Köhler et al., 2003; Chanvivattana et al., 2004; De Lucia et al., 2008). The FIS complex, for instance, is active in the female gametophyte and developing seed endosperm (Köhler et al., 2003; Guitton et al., 2004), whereas the EMF complex acts in the embryo and during subsequent sporophytic development (Goodrich et al., 1997; Yoshida et al., 2001). The VRN complex activity is triggered upon association with PHD proteins VERNALIZATION INSENSITIVE3 (VIN3), VERNALIZATION LIKE1 (VEL1; also known as VIN3-LIKE2 [VIL2]), and VERNALIZATION5 (VRN5; also known as VIL1) and represses targets of vernalization, the prolonged cold period that allows annual plants to flower in spring (Fig. 2). A comparable association of PRC2 with PHD proteins was observed in animals (Nekrasov et al., 2007; Sarma et al., 2008). In Drosophila, stable transcriptional repression by the PRC2 complex requires the activity of other proteins complexes: PRC1, Pleiohomeotic Repressive Complex, and a newly identified module named PR-DUB (Zheng and Chen, 2011). PRC1, in animals, binds to H3K27me3 marks and ubiquitinates H2A Lys-119. As a consequence, chromatin is compacted in a stable heterochromatin state, and transcription is durably repressed (Sawarkar and Paro, 2010). In plants, homologs have been identified for the PRC1 components BMI1 and RING1. Monoubiquitination of H2A Lys-121 in Arabidopsis by AtBMI1A and AtBMI1B is implicated in repression of embryonic and stem cell regulators (Bratzel et al., 2010). AtBMI1C physically interacts with AtRING1A/B and may be involved in flowering regulation (Li et al., 2011). AtRING1A and AtRING1B also associate with LIKE HETEROCHROMATIN PROTEIN1 (LHP1) to form a complex similar to the animal PRC1 (Xu and Shen, 2008; Fig. 1). There are no clear homologs of other members of PRC1 defined in animals. However, proteins with an activity comparable to PRC1 have been identified in Arabidopsis. LHP1 binds H3K27me3 (Turck et al., 2007) and shares a subset of PRC2 targets in a manner similar to Pc, a PRC1 component in animals (Bracken et al., 2006). With the exception of an impact on flowering time (Mylne et al., 2006; Sung et al., 2006), the lhp1 phenotype is quite distinct from the phenotypes of mutants affecting PRC2 (Gaudin et al., 2001; Takada and Goto, 2003). Two other proteins, VRN1 and EMF1, which bind and act together with LHP1 and AtBMI1A/B (Bratzel et al., 2010), have been proposed to be involved in PRC1-like functions (Mylne et al., 2006; Calonje et al., 2008). Hence, several proteins with an activity similar to PRC1 components are specific to plants. Because PRC1 homologs are not conserved between animal species, it may not be surprising that PRC1 function in plants is mediated by mechanisms distinct from that described in mammals. POLYCOMB AND TRITHORAX COMPLEXES ARE ANTAGONIZING EACH OTHER As opposed to PRC2, Drosophila Trithorax group proteins cause trimethylation of H3 Lys-4 and positively regulate transcription. The Arabidopsis genome encodes five ARABIDOPSIS TRITHORAX proteins (ATX1 to ATX5), with a SET domain and a PHD domain, and seven ARABIDOPSIS TRITHORAX-RELATED proteins (ATXR1 to ATXR7; Tamada et al., 2009). ATXR5 and ATXR6 do not appear to be directly related to ATXR function defined as antagonists to PRC2 and rather cause H3K27 monomethylation associated with constitutive heterochromatin (Jacob et al., 2009; Roudier et al., 2011). ATX1 and ATXR7 are required for the expression of homeotic genes involved in flower organogenesis and the floral repressor FLOWERING LOCUS C (FLC; Pien et al., 2008; Tamada et al., 2009). Besides its H3K4 methylation activity, ATX1 influences transcription by recruiting the TATA binding protein and RNA Polymerase II at promoters (Ding et al., 2011). Although ULTRAPETALA1 (ULT1) has no homology with animal Trithorax group components, it functions as a Trithorax factor regulating in an opposed manner to PRC2 a common set of loci silenced by CLF. However, ULT1 does not have a methlytransferase activity but rather helps recruiting ATX members on the DNA (Carles and Fletcher, 2009). Hence, it is possible that ATX1, ATXR7, and ULT1 associate to form a Trithorax complex that may contain other proteins (Fig. 1). Similarly, the chromatin remodeling factors PICKLE and PICKLE RELATED2 antagonize CLF for the regulation of root meristematic growth (Aichinger et al., 2011). In conclusion, PRC2 is conserved in plants, but the regulatory PRC1 and TRX activities also are mediated by nonconserved protein complexes that evolved independently from their functional counterparts in metazoans. THE INS AND OUTS OF H3K27ME3 MARK ON CHROMATIN As many other annual species, Arabidopsis flowers in spring after an obligate period of cold in winter (vernalization). However, some natural accessions of Arabidopsis flower late in summer and do not require vernalization. Such natural variations rely primarily on the regulation of two factors, FRIGIDA and FLC (Koornneef et al., 1994; Lee et al., 1994). FLC encodes a MADS box transcription factor that represses flowering. At the FLC locus, H3K27me3 marks accumulate during vernalization, and FLC expression remains repressed throughout the life cycle until seed development initiates. Then, FLC becomes expressed again during early embryogenesis (Sheldon et al., 2008; Choi et al., 2009; Kim et al., 2009). The mode of deposition and removal of H3K27 methylation that accompanies the cycle of FLC expression have been studied in much detail and provide a good study case. The cycle can be divided in four steps: deposition, spreading, maintenance, and removal. Deposition How PRC2 is recruited on specific targets to deposit the H3K27me3 mark is still largely unknown. Some DNA sequences of the Hox loci in Drosophila are recognized by PRC2 complexes and are called Polycomb response elements (PREs). However, the degree of conservation of these elements is low across animal species, and no PRE has been found in plants to date. Nonetheless, at the FLC locus, some cis-sequences in the promoter and intron 1 are required for the establishment of repression (Sheldon et al., 2002). Some of these sequences play a role in assisting the recruitment of the noncoding RNA COLDAIR (for COLD ASSISTED INTRONIC NONCODING RNA) that is required for establishing stable repressive chromatin by recruiting PRC2 (Heo and Sung, 2011). In addition, COOLAIR (for COLD INDUCED LONG ANTISENSE INTRAGENIC RNA) is an antisense transcript initiated after the polyadenylation site of FLC that is induced by cold early in the vernalization process and silences FLC (Swiezewski et al., 2009). The role of long noncoding RNAs is reminiscent of the silencing by the XIST noncoding RNA and PRC2 in mammals (Margueron and Reinberg, 2011), and further analyses may show a generalization of this mechanism (Spitale et al., 2011). Hence, it is likely that PRE represents only the sequence complementary to a specific element of the long noncoding RNA that forms a complex with PRC2, and if true, this hypothesis would explain the lack of conservation of PREs. It is also possible that other mechanisms participate to PRC2 recruitment. In animals, the JumonjiC (JmjC) domain protein Jarid2 anchors PRC2 but inhibits its repressive action. It remains to be discovered whether plant homologs of Jarid2 fulfills the same role (Zheng and Chen, 2011). In addition, a combination of histone modifications could also be read by PRC2 complexes or associated proteins to specify a target. Spreading Analyses of transgenes carrying FLC and a fusion protein have shown that PRC2 can spread from an initial entry site to methylate histones at adjacent sequences (Schubert et al., 2006). However, genome-wide studies showed that H3K27me3 marks do not extend over large regions in plants as it does in flies and mammals but are restricted to discrete domains (Turck et al., 2007; Zhang et al., 2007). The PHD proteins VIN3, VRN5 (VIL1), and VEL1 (VIL2; Sung et al., 2006) that enhance the VNR complex activity at vernalization were shown to play a role in the spreading of H3K27me3 marks on FLC locus upon return of warm temperature (De Lucia et al., 2008). At the tissue level, spreading of H3K27me3 marks takes place only in dividing cells (Finnegan and Dennis, 2007). In root meristems, a pFLC-GUS reporter is expressed in a salt and pepper pattern after a limited period of vernalization. This is interpreted as two populations of cells, one with FLC still expressed and another with FLC expression suppressed. The gradual reduction of FLC expression would result from an increased proportion of cells with FLC expression turned off, and a mathematical model has been proposed to back up this hypothesis. The model proposes a switching mechanism involving the local nucleation of opposing histone modifications (Angel et al., 2011). Maintenance The mark H3K27me3 is generally heritable through cell division for a stable repression. The PRC2 complex may directly bind to H3K27me3, most likely through the WD40 domain of FIE (Xu et al., 2010), thus ensuring a semiconservative mode of maintaining the PRC2 mark (Hansen et al., 2008). It was hypothesized that the spreading of H3K27me3 also plays a role in this mitotic inheritability, as a critical number of modified nucleosomes at one locus may be necessary to ensure the fidelity of the epigenetic information being passed along mitosis (Schubert et al., 2006). Possible additional mechanisms for this controlled inheritance have been proposed in mammals where it was shown that PcG proteins have the ability to stay bound on replicating DNA. As noncoding RNAs have been observed to play a role in PRC2 recruitment, RNA could be the molecule passing along the information (Sawarkar and Paro, 2010). In addition, H3K27me3 recognition by PRC1 induces chromatin compaction, thus enforcing a stable repression and cellular memory of PRC2 marks. But in the case of FLC, none of these mechanisms has been identified and the maintenance of H3K27me3 at FLC locus remains to be understood. Removal FLC expression resumes after flowering, during the early stages of seed development in the embryo but not in the endosperm (Sheldon et al., 2008; Choi et al., 2009). This period coincides with the time when histones H3 transmitted by the gametes are actively removed after fertilization and replaced by newly synthesized histone H3 variants in the zygote but not in the endosperm (Ingouff et al., 2010). Thus, resetting the H3 proteins provides a potential global mechanism to discard H3K27me3 as well as other marks carried by the nucleosomes at the FLC locus (Fig. 3). This model is supported by the association between histones and FLC reactivation (Yun et al., 2011). Figure 3. Open in new tabDownload slide Possible mechanisms causing H3K27me3 removal. A, Dilution by successive cell divisions: At each DNA replication cycle, the H3K27me3 mark is lost if not added actively on the newly formed DNA strand. B, Removal in a transcription factor-dependent manner. For flowering after vernalization, the transcription factor AG binds on the KNU locus, and H3K27me3 marks are gradually removed in an AG-dependent manner. The precise mechanism of this removal is still unknown. C, Demethylation by an H3K27me3 demethylase, such as REF6. D, Replacement of the H3 histone that bears the H3K27me3 mark by a variant H3 (H3v) that doesn’t carry the H3K27me3 mark. Figure 3. Open in new tabDownload slide Possible mechanisms causing H3K27me3 removal. A, Dilution by successive cell divisions: At each DNA replication cycle, the H3K27me3 mark is lost if not added actively on the newly formed DNA strand. B, Removal in a transcription factor-dependent manner. For flowering after vernalization, the transcription factor AG binds on the KNU locus, and H3K27me3 marks are gradually removed in an AG-dependent manner. The precise mechanism of this removal is still unknown. C, Demethylation by an H3K27me3 demethylase, such as REF6. D, Replacement of the H3 histone that bears the H3K27me3 mark by a variant H3 (H3v) that doesn’t carry the H3K27me3 mark. A second possible mechanism of removal of H3K27m3 is biochemical demethylation by a bone fide demethylase (Fig. 3). In animals, H3K27me3 is removed by KDM6a and KDM6b, two Jumonji domain-containing Lys demethylases. In Arabidopsis, RELATIVE OF EARLY FLOWERING6 (REF6), also known as JUMONJI12 (JMJ12), demethylates H3K27me3 and H3K27me2, whereas its metazoan counterparts, the KDM4 proteins (homologous in the JmjN and JmjC domains), are H3K9 and H3K36 demethylases (Lu et al., 2011). However, other proteins might be required for the demethylation of all H3K27me3 targets as there are at least two close homologs of REF6 in Arabidopsis, EARLY FLOWERING6 and JMJ13, that could act redundantly with REF6. The reactivation of FLC remains to be understood at the mechanistic level. PCG COMPLEXES PLAY A CENTRAL ROLE IN DEVELOPMENTAL TRANSITIONS The transition toward flowering represents one of the many sharp transitions between different phases of development that mark the plant life cycle. All plant species alternate a haploid life form (gametophyte) and a diploid form (sporophyte). The gametophyte produces gametes, which undergo fertilization resulting in a zygote. Zygotic division initiates a sporophyte that produces meiotic haploid spores. In flowering plants, the sporophytic life is divided into further transitions, including, for example, the differentiation of meristem cells into vegetative organs and the switch between an indeterminate vegetative meristem to a determinate inflorescence meristem. Here, we detail how PcG is involved in several phase developmental transitions (Fig. 4). Figure 4. Open in new tabDownload slide PcG complexes mediate transitions from one stage of development to another. PRC2 complexes have been found to play an essential role in major transitions from one phase to another of plant development. The inner cycle marks the transition between the gametophytic to the sporophytic phase. Orange arrows mark PRC2-dependent transitions that take place during the sporophytic development. Figure 4. Open in new tabDownload slide PcG complexes mediate transitions from one stage of development to another. PRC2 complexes have been found to play an essential role in major transitions from one phase to another of plant development. The inner cycle marks the transition between the gametophytic to the sporophytic phase. Orange arrows mark PRC2-dependent transitions that take place during the sporophytic development. DEVELOPMENTAL TRANSITION DURING SEXUAL REPRODUCTION Gametophyte-to-Sporophyte Transition In mosses, the gametophytic life is the predominant vegetative phase, and the sporophyte is a short-lived structure specialized in meiosis and spore production. In the moss Physcomitrella patens, mutants were obtained for the homologs of FIE and CLF (Mosquna et al., 2009; Okano et al., 2009). In the absence of PpFIE and PpCLF, meristems overproliferate and are unable to develop leafy gametophytes or reach the reproductive phase. Instead, gametophytes produce proliferating clumps of cells that express sporophytic markers, showing that PRC2 prevents a precocious transition from the gametophytic to the sporophytic stage. Similarly in Arabidopsis, mutants for the FIS complex produce an overproliferating central cell in the gametophyte (Ohad et al., 1996; Chaudhury et al., 1997). This phenotype was interpreted as a prevention of fertilization or endosperm development by the FIS complex. However, in the light of the impact of PRC2 on gametophytic-to-sporophytic transition in mosses, it is more likely that in Arabidopsis, FIS prevents the arrest of gametophytic life. Hence, the tissue proliferating from the fis central cell would represent abnormal gametophytic tissue. This suggests an ancestral role of PRC2 in the control of the transition between the gametophytic and sporophytic life. Endosperm Maturation Embryo development requires the function of the second product of fertilization, the endosperm. Endosperm development is initiated by a series of syncytial divisions leading to a large multinucleate cell. After a specific number of synchronous nuclear divisions, cytokinesis takes place and cellular endosperm is formed (Berger and Chaudhury, 2009). Seed reserves accumulate in the cellular endosperm and, depending on the species, are either retained in the endosperm or transferred to the embryo. The transition to cellular endosperm is controlled by PRC2 activity. Endosperms deprived of FIS function hyperproliferate and never cellularize. The retention of a syncytial endosperm in fis mutants is confirmed by an abnormal expression of molecular markers of syncytial endosperm and the absence of expression of genes normally expressed in wild-type cellular endosperm (Ingouff et al., 2005). This conclusion is further strengthened by the abundance of H3K27me3 on genes involved in cell wall synthesis and other markers of late endosperm development (Weinhofer et al., 2010). Hence, PRC2 controls the transition from syncytial to differentiated cellular endosperm. Embryo Maturation When the strict requirement of the FIS complex for gametophytic development and endosperm is overcome, it is possible to study the function of PRC2 during plant embryogenesis (Kinoshita et al., 2001; Bouyer et al., 2011). fie embryos never mature but keep proliferating and eventually produce tissues that resemble calluses. PRC2 not only prevents the transition between early seed to dry seed but also represses entire networks of genes involved in the reproductive transition. Interestingly, fie mutant seedlings are initially indistinguishable from wild-type plants, suggesting that the initial patterning does not rely on PRC2 but rather does the maintenance of this pattern (Bouyer et al., 2011). TRANSITIONS DURING POSTEMBRYONIC LIFE Transition from Juvenile to Adult Stage The shoot apical meristem continuously initiates primordia that exit the stem cell self-renewal program and engage toward leaf development (Fig. 4). The genome-wide distribution of H3K27me3 marks and the transcriptome were compared between meristematic cells and leaves to gain insight into the function of PRC2 in tissue-specific differentiation (Lafos et al., 2011). During wild-type leaf development, H3K27me3 marks are removed at several hundreds of loci. A large fraction of these loci encode transcription factors, some of which are already identified as key regulators of leaf development. Interestingly, many of the transcriptional networks identified are also targeted by microRNAs, which also become derepressed during differentiation. For instance, genes involved in the entire pathway of the phytohormone auxin regulation are targeted by H3K27me3, including microRNAs repressing auxin response factors. The mechanism leading to coordinated removal of the H3K27me3 mark remains unclear. Vegetative-to-Reproductive Transition The shoot apical meristem keeps producing leaf primordia until the transition to flowering. Then, the meristem becomes determinate and becomes an inflorescence meristem that, in turn, produces floral meristems (Fig. 4). We discussed above the predominant role of PRC2 in the induction of flowering in response to vernalization. This role also extends to the control of meristem determinacy via other major targets of PRC2: LEAFY (LFY) and the transcription factors AGAMOUS (AG) and KNUCKLES (KNU). LFY is essential for the transition to an inflorescence meristem. Its expression is repressed by PRC2 during embryo development (Kinoshita et al., 2001). The MADS box transcription factor AG and the zinc finger domain transcriptional repressor KNU ensure a determinate number of organs is produced by the floral meristem. Their expression is repressed by H3K27me3 in a highly regulated temporal manner (Sun et al., 2009). AG expression is initiated first, and AG binds to the upstream promoter of KNU, which is still repressed by H3K27me3. Gradually, the repressive mark is removed in a cell cycle in an AG-dependent manner, allowing the basic transcriptional machinery to access the KNU locus and induce transcription. This mechanism provides a delay that is key to ensure that a proper cell number is produced by the floral meristem before the flower organ differentiation. CONCLUSION PcG complexes PRC2 are conserved. However, there is limited conservation of the activities that read the H3K27me3 marks they deposit. As shown in animals, long noncoding RNAs certainly play a role in PRC2 targeting (Zhao et al., 2010), but it remains unclear whether this is also the case in plants where very few long noncoding RNAs have been isolated so far. Several other mechanisms might recruit PRC2 and explain PRC2 targeting. Unlike in animals, PRC2 in plants does not appear to play a major role in patterning but is essential in the control of developmental phase transitions (Fig. 4). This implies cycles of maintenance and reprogramming, and the nature of these mechanisms is still not understood. Certain phases are long lasting (for example, the sporophytic phase in flowering plants) and contain cascades of more specific transitions (for example, vegetative to inflorescence to floral meristem). This implies that developmental transitions controlled by PRC2 become gradually more refined, while they do not necessarily rely on a cascade of repressive marks. Understanding resetting mechanisms and how plants maintain certain marks specific for long-lasting developmental status while other PRC2 marks are remodeled during a transition contained within this period are major challenges of this research field. ACKNOWLEDGMENTS We thank Laurent Pieuchot for help with the figures. LITERATURE CITED Aichinger E Villar CBR Di Mambro R Sabatini S Köhler C ( 2011 ) The CHD3 chromatin remodeler PICKLE and polycomb group proteins antagonistically regulate meristem activity in the Arabidopsis root . 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Shaw, Peter; Brown, John

doi: 10.1104/pp.111.188052pmid: 22082506

RDNA AND NUCLEOLAR CHROMATIN rDNA Structure The nucleus is a highly organized structure. However, unlike cytoplasmic organelles, nuclear substructures are not bounded by membranes, but rather are held together by interactions between their component proteins and nucleic acids, and are thus probably best regarded as giant, extended multimolecular complexes. The nucleolus is the most prominent of these structures. It is the site of transcription by RNA polymerase I (Pol I) of the rDNA, tandemly repeated copies of the genes for three of the four ribosomal RNAs (rRNAs: s-rRNA, 5.8S-rRNA, and l-rRNA), which are called the nucleolar organizer regions (NORs) of the chromosomes (Raska et al., 2006a, 2006b). The rDNA repeats are transcribed as a single precursor RNA, which is edited into the three rRNAs by excising leading, internal, and trailing transcribed spacer sequences. The rDNA repeats are separated by untranscribed intergenic sequences, which are much shorter in plants (2–3 kb) than in mammals and many other vertebrates (10–30 kb; Hadjiolov, 1985). In many cases the size of the intergenic spacers is heterogeneous, as, for example, in Xenopus laevis (3–9 kb). The fourth rRNA, 5S, is transcribed by RNA Pol III from tandem repeats elsewhere in the nucleus (Highett et al., 1993). The nucleolus is also the location at which most of the steps of ribosome assembly and maturation occur. This means about 90 ribosomal proteins, as well as many processing and accessory factors, need to be imported from the cytoplasm to the nucleolus, and the nearly complete small and large ribosomal subunits separately exported to the cytoplasm. Since a rapidly growing cell may require millions of ribosomes to be synthesized, the nucleolus is by far the major destination and origin of nucleocytoplasmic transport in the most active cells. Nucleolar Organization When nuclei are stained with fluorescent DNA dyes such as 4′,6-diamidino-2-phenylindole, the nucleolus is seen as a dark region within the more brightly stained nuclear chromatin. This is an indication that the active rDNA is highly dispersed within the nucleolus. On the basis of their appearance in electron microscope (EM) images, the nucleoli in many animal cells have been described in terms of a tripartite structure, with small, lightly staining regions called fibrillar centers (FCs), surrounded by densely stained material called the dense fibrillar component (DFC); the remainder of the nucleolus contains what appear to be densely packed granules (the granular component [GC]; Shaw et al., 1995; Fig. 1). Originally these granules were assumed to be entirely preribosomal particles in various stages of processing, but recent studies have indicated other types of complexes may occupy specific parts of the GC, which may be segregated into distinct regions of different compositions (Politz et al., 2005). What appear to be FCs are often seen in plants, but the regions assumed to be related to the DFC are much more extensive in plant nucleoli and often no more densely stained than the surrounding granules (Shaw and Jordan, 1995; Fig. 1). In reality the organization of the nucleolus is likely to be more complex than previously thought. A recent article has shown that within the chordates there are species that display a bipartite organization, in essence lacking FCs (Thiry et al., 2011). These authors suggest that the FCs originated with the emergence of the amniotes. A problem with this hypothesis is that some anamniote nucleoli, such as in Xenopus, do contain FCs, as indeed do plants (Raska et al., 2006b). The controversies about nucleolar organization will probably not be fully resolved until we have a better understanding of what these electron microscopic structures represent in molecular and functional terms. Figure 1. Open in new tabDownload slide Comparison of the structure of animal (A) and plant (B) nucleolar organization. TS, Transcription sites. In plants (B) a central region is often present and is called the nucleolar cavity. Figure 1. Open in new tabDownload slide Comparison of the structure of animal (A) and plant (B) nucleolar organization. TS, Transcription sites. In plants (B) a central region is often present and is called the nucleolar cavity. The molecular interpretation of these structures has been the subject of a long controversy. Immunogold EM studies showed concentrations of RNA Pol I in the FCs, which were consequently suggested to be the sites of transcription (Scheer and Rose, 1984). In situ labeling, whether at the optical or EM level, showed most rDNA in condensed heterochromatin at the nucleolar periphery, and smaller foci of rDNA labeling corresponding to FCs (Shaw et al., 1995). With more sensitive in situ techniques, fainter, diffuse rDNA labeling was subsequently seen throughout the DFC. The problem with a simple interpretation of these labeling experiments is that most Pol I, and most rDNA copies are not transcriptionally active. It was not until fluorescence detection of bromo-uridine labeling was introduced that the sites of transcription could be clearly labeled. In plant cells, this showed the transcription sites to consist of many foci within the DFC, with the smallest foci representing individual rDNA gene repeats (Thompson et al., 1997; González-Melendi et al., 2001; Koberna et al., 2002). Additional labeling with probes to transcribed spacer regions in the prerRNA and with various proteins and other small RNAs known to be involved in ribosome maturation led to a radial model, where the newly formed transcripts moved away from the dispersed genes within the DFC and then subsequently moving into the GC for later processing stages (Brown and Shaw, 1998). Nucleolar Chromatin and Epigenetics The number of rDNA repeats varies greatly among eukaryotes; many plants have thousands of copies, of which only a small proportion seems to be transcriptionally active at any one time. There is evidence that some repeats in human rDNA are inverted and may not be functional (Caburet et al., 2005). In rice (Oryza sativa), fiber fluorescence in situ hybridization has suggested that the repeats are regular and do not contain inversions or rearrangements (Mizuno et al., 2008). Furthermore, physical mapping studies in Arabidopsis (Arabidopsis thaliana) using pulsed field gel electrophoresis showed that restriction endonucleases that cut once per rDNA repeat gave only a single length of fragment (Copenhaver and Pikaard, 1996). Nevertheless the rDNA has not been fully sequenced in any higher eukaryote, as with current technology it is very difficult to analyze such highly repetitive sequences. We therefore do not know absolutely whether all the rDNA repeats are intact functional genes or whether other functional sequences are interspersed with them, and it is merely an assumption that all rDNA repeats are potentially transcribable. As with other genomic regions, current models suggest that rDNA chromatin can adopt three different states: inactive and condensed (heterochromatin), which corresponds to the condensed chromatin at the nucleolar periphery as well as to some intranucleolar condensed regions; active genes in an extended conformation within the DFC; and a poised or potentiated state, available for transcription but not currently being transcribed (McKeown and Shaw, 2009). This latter state may correspond to rDNA within the FCs. The balance between these states, and thus Pol I loading and transcription, is determined by DNA methylation, differences in the histone variants associated with the DNA, remodeling of the DNA, particularly the promoter regions, and the presence of histone modifications (Grummt and Pikaard, 2003). Loading of Pol I and some other factors is maintained through mitosis, and thus the chromatin state can be epigenetically inherited. A particularly clear example of epigenetic control of rDNA is provided by nucleolar dominance in hybrids where the NORs of one parental genome are active, whereas those of the other are inactive (Tucker et al., 2010). An RNAi approach has been used with Arabidopsis suecica, a hybrid of Arabidopsis and Arabidopsis arenosa, to determine factors involved in nucleolar dominance. This has shown the involvement of the histone deacetylases Histone Deacetylase1 and Histone Deacetylase6, the de novo DNA methyltransferase Domains Rearranged Methyltransferase2 (DRM2), and the methylcytosine binding domain proteins Methyl CpG Binding Domain6 (MBD6) and MBD10 (Preuss et al., 2008). Further confirmation that nucleolar dominance in plants involves RNA-directed DNA methylation was obtained by knockdown of RNA Dependent RNA Polymerase2, Dicer-like3, and DRM2, which disrupted the silencing of the Arabidopsis-derived rRNA genes in A. suecica (Preuss et al., 2008). In mammals, rRNA genes are silenced by the nucleolar remodeling complex, NoRC, which is recruited to rRNA genes by 200- to 300-nt RNA species, termed pRNA, derived from intergenic regions of the rDNA (Guetg et al., 2010; Santoro et al., 2010). PLURIFUNCTIONAL NUCLEOLUS Over the last 10 to 15 years, it has become clear that the nucleolus is involved in numerous other functions than ribosome biogenesis (Pederson, 1998; Rubbi and Milner, 2003; Olson, 2004; Raska et al., 2006a; Boisvert et al., 2007). Many are RNA-related functions such as RNA processing and assembly of ribonucleoproteins (RNPs). For example, the nucleolus (and associated bodies, particularly Cajal bodies [CBs]) is involved in the maturation, assembly, and export of RNP particles such as the signal recognition particle, telomerase RNP, and processing of precursor transfer RNAs and U6 small nuclear RNA. In addition, the nucleolus has roles in cellular functions such as regulation of the cell cycle, stress responses, telomerase activity, and ageing (Pederson, 1998; Tsai and McKay, 2002; Rubbi and Milner, 2003; Olson, 2004; Raska et al., 2006a; Boisvert et al., 2007; Boulon et al., 2010). Sequestration of specific proteins in the nucleolus or their release is one mechanism by which processes such as the cell cycle or cell death are regulated. The multifunctional nature of the nucleolus is therefore reflected in the complexity of the protein and RNA composition of the nucleolus and in the dynamic composition changes in response to cellular conditions. Protein Composition of the Nucleolus Initial analyses of the proteome of human cells identified around 450 proteins including ribosomal proteins and proteins known to be involved in ribosome biogenesis (fibrillarin, nucleolin, B23, etc.; Andersen et al., 2002; Scherl et al., 2002). Even at this stage, unexpected proteins like splicing factors, spliceosomal proteins, and translation factors were identified. The increasing resolution of mass spectrometry techniques had led to the current characterization of the nucleolar proteome from human cells of circa 4,500 proteins (Ahmad et al., 2009). Quantitative proteomic analyses have demonstrated the dynamic behavior of nucleolar proteins such as ribosomal proteins and of protein complexes such as Pol I (Andersen et al., 2005; Lam et al., 2007). For example, quantitative changes in the relative levels of nucleolar components (reflecting accumulation or loss) were apparent upon transcriptional inhibition. The degree of change varied greatly for different proteins and showed that ribosomal proteins were highly expressed and either incorporated into ribosomal subunits or rapidly degraded (Andersen et al., 2005; Lam et al., 2007). It is also intriguing that ribosomal protein complexes associate with chromosomes and in particular transcription start sites of tRNAs, although the nature of these complexes and their function are as yet unknown (De et al., 2011). In contrast, characterization of the proteome of plant nucleoli lags significantly behind. In an initial proteomic analysis of purified Arabidopsis nucleoli 217 proteins were identified. In addition to the expected ribosomal and nucleolar proteins, a range of nonribosomal and nonnucleolar proteins including plant-specific proteins, proteins with unknown function, and splicing and translation factors was observed (Pendle et al., 2005). In particular, exon junction complex proteins (known in animal systems to associate with mRNAs after splicing) were identified and their nucleolar localization was confirmed by confocal microscopy of GFP fusions (Pendle et al., 2005; Fig. 2). One of the core exon junction complex components, eIF4A-III, was shown to redistribute from the nucleoplasm to the nucleolus and finally to splicing speckles under hypoxia stress conditions (Koroleva et al., 2009). Although the inference is that the relocalization of eIF4A-III might cause redistribution of mRNAs to the nucleolus, the exon junction complex has not yet been formally identified in plants and its association with mRNAs when in the nucleolus has not been demonstrated. A similar redistribution of an SR protein splicing factor to the nucleolus under ATP depletion has also been demonstrated (Tillemans et al., 2006). To date, this SR protein, RSZ22, is the only SR protein to show such a relocalization and again whether it relocalizes mRNAs is not known. A number of small nuclear RNP proteins were identified in the nucleolar proteome (Pendle et al., 2005) and CBs/nucleolus may be involved in production of the spliceosomal U1snRNP in plants (Lorković and Barta, 2008). Our knowledge of the nucleolar proteome in plants is still relatively limited and given the multifunctionality of this nuclear compartment, a full analysis is overdue and would be likely to provide supporting evidence for different functions or even identify new functions. Figure 2. Open in new tabDownload slide Examples of transient expression of GFP fusions in suspension culture cells to proteins identified as nucleolar components. A to C, RNA binding proteins with known animal homologs. D and E, Uncharacterized, plant-specific proteins. The nucleolus is indicated in each section by an arrow. Bar = 5 μm. Figure 2. Open in new tabDownload slide Examples of transient expression of GFP fusions in suspension culture cells to proteins identified as nucleolar components. A to C, RNA binding proteins with known animal homologs. D and E, Uncharacterized, plant-specific proteins. The nucleolus is indicated in each section by an arrow. Bar = 5 μm. RNA Complexity in the Nucleolus The multiple functions of the nucleolus in processing of various RNAs and assembly of different RNPs are reflected in the different species of RNA. Besides rRNAs, animal cell nucleoli contain the extensive families of small nucleolar RNAs (snoRNAs) as well as snRNAs, tRNAs, 7SL RNA (signal recognition pathways), and telomerase RNA. In plants, analyses of nucleolar RNAs, excluding rRNAs, suggest that the plant nucleolus is also involved in many RNA/RNP events. Moreover, the presence of mRNAs and small-regulatory RNAs in the plant nucleolus and nucleolar-associated bodies is intriguing in terms of potentially novel functions in plants, broadening the roles of the nucleolus still further. Cloning and sequencing of RNAs from purified Arabidopsis nucleoli identified tRNAs, snRNAs, and small cajal body-specific RNAs along with many known and novel snoRNAs (Kim et al., 2010). In particular, in addition to the expected conserved snoRNAs (U3 and MRP), orthologs of human U13 snoRNAs were discovered. The function of U13 has not been determined but it contains complementarity to the 3′ end of s-rRNA. All eukaryotes contain orphan snoRNAs, which do not have complementarity to rRNAs or snRNAs. In animals, such snoRNAs have been found to be involved in RNA editing, alternative splicing, and regulation of gene expression (Vitali et al., 2005; Kishore and Stamm, 2006; Ender et al., 2008; Saraiya and Wang, 2008; Ono et al., 2010). Recently, three mammalian snoRNAs (all encoded in the introns of a ribosomal protein gene) were shown to be involved in regulation of metabolic stress response (Michel et al., 2011). Although plants have a number of orphan snoRNAs, to date no noncanonical function in, for example, modulating gene expression has been described. The identification of mRNA-associated proteins in the plant nucleolus suggests a function in mRNA biogenesis. In animals, only a very few mRNAs have ever been identified in the nucleolus (Pederson, 1998; Olson, 2004). In contrast, mRNAs from a wide range of genes were identified in a cDNA library from purified Arabidopsis nucleoli. Of particular interest was that aberrantly spliced mRNAs were enriched in the nucleolus and the vast majority contained premature termination codons, and therefore were likely to be turned over by the nonsense-mediated decay (NMD) pathway (Kim et al., 2009). Further, the localization of proteins involved in NMD—UPF2 and UPF3—to the nucleolus suggested a novel function for the plant nucleolus in mRNA surveillance/NMD and thereby in mRNA biogenesis (Kim et al., 2009). However, we have recently shown that mRNA transcripts with retained introns (or containing unspliced introns) are not turned over by NMD and appear to avoid the NMD pathway (Kalyna et al., 2011). While other aberrant transcripts are targeted to the nucleolus as part of the NMD pathway, intron-containing transcripts may accumulate in the nucleolus for a different function (e.g. degradation by a different pathway) or may be targeted there instead of being exported. In this regard, an association between the nucleolus and export of particular human virus RNAs is well established (Hiscox, 2007; Hiscox et al., 2010) and has been suggested to function for cellular mRNAs. In yeast (Saccharomyces cerevisiae) and animal cells the question of whether specific mRNAs/mRNPs have a nucleolar phase is still open (discussed in Jellbauer and Jansen, 2008). Besides the novel finding of mRNAs and aberrant mRNAs in the plant nucleolus, heterochromatic small interfering RNAs, which are involved in transcriptional silencing, are produced in a region of the nucleolus or in bodies often found associated with the nucleolus called d-bodies (Pontes and Pikaard, 2008). The localization of precursor microRNAs (miRNAs) and Dicer-like1 to d-bodies also suggests a role for the nucleolus in the maturation of miRNAs (Pontes and Pikaard, 2008). Some precursor and mature miRNAs are enriched in the nucleolus of mammalian cells possibly for modification, assembly, or to regulate snoRNA activity (Politz et al., 2009; Scott et al., 2009). There is also an evolutionary relationship between miRNA precursors and snoRNAs, with some miRNAs being processed from snoRNA precursors and some miRNA precursors retaining snoRNA features (Saraiya and Wang, 2008; Politz et al., 2009; Scott et al., 2009; Ono et al., 2011). In addition, snoRNA-derived small RNAs were found to be associated with Argonaute proteins of RNA silencing pathways in both animals and Arabidopsis (Taft et al., 2009) and small RNAs from a human snoRNA reduced expression of gene targets (Ender et al., 2008). The complexity of RNA/RNP processes involving the nucleolus suggests that it is a center of RNA activity. For plants, the presence of mRNAs and small regulatory RNAs in the nucleolus allows us to speculate that the nucleolus is involved in regulation of expression, possibly in response to cellular conditions. The Nucleolus and Virus Infection From the integral nature of the nucleolus to many RNA processing and RNP assembly pathways, it is not surprising that many animal and plant viruses exploit the nucleolus in production and transport of viral RNPs. The involvement of the nucleolus in infection cycles of animal and human viruses is well established (Hiscox, 2007; Hiscox et al., 2010; Taliansky et al., 2011). In plants, a growing number of viruses show some interaction with the nucleolus (and other nuclear bodies such as CBs) and roles for the nucleolus and nucleolar proteins are now emerging (Taliansky et al., 2011). For example, plant viruses can recruit nucleolar proteins for assembly of viral RNP particles, virus replication and movement, and to counteract host-viral defense systems. In the best-studied system to date, the ORF3 long-distance movement protein of Groundnut rosette virus trafficks to the nucleolus via CBs, causing reorganization of CBs into multiple CB-like structures that fuse with the nucleolus. ORF3 then recruits fibrillarin (an abundant nucleolar RNA binding protein, known to be an RNA methylase) for assembly of cytoplasmic infectious viral particles (Kim et al., 2007a, 2007b; Canetta et al., 2008). The nucleolar localization of ORF3 is essential for systemic infection. Viral proteins from other plant viruses also target the nucleolus and the function of this localization and their interactions with nucleolar proteins are being established. For example, the coat protein and coat protein read-through proteins of Potato leaf roll virus (PLRV) are targeted to the nucleolus (Haupt et al., 2005) and systemic infection of PLRV is inhibited in fibrillarin-silenced plants, suggesting that fibrillarin is also involved in long-distance movement of PLRV. Another aspect of nuclear/nucleolar targeting of viral proteins is to interfere with host defenses. Most plant viruses encode silencing suppressor proteins to counteract the silencing triggered by infection. The NIa/VPg protein of Potato virus A has suppressor activity dependent on the localization of VPg to the nucleolus and CBs, raising the question of whether this protein targets components or pathways involved in RNA silencing that are found in the nucleus or nucleolus and associated bodies (Pontes and Pikaard, 2008). The VPg domain of NIa interacts with fibrillarin in the nucleolus and CBs, and depletion of fibrillarin reduces accumulation of the virus, suggesting that fibrillarin is involved in the infection process (Rajamäki and Valkonen, 2009). The silencing suppressor protein of Cucumber mosaic virus, CMV 2b, also localizes to the nucleus and nucleolus where it interacts with Argonaute1 and Argonaute4 (González et al., 2010). However, these interactions are not sufficient for suppression of RNA silencing and hence their biological relevance remains so far unclear (González et al., 2010). Other viral proteins are also found in the nucleus/nucleolus (e.g. the P3 protein of Tobacco etch virus and the P6 protein of Cauliflower mosaic virus) but the function of the nucleolar localization is unknown (Taliansky et al., 2011). How different viral proteins target CBs and the nucleolus, their interactions with host proteins like fibrillarin, and the impact of usurping normal functions on nucleolar structure and function are important questions for the future that are likely to provide insights into nucleolar biology. DYNAMICS OF THE NUCLEOLUS The nucleolus, like the other parts of the nucleus, is dynamic at a number of levels. First, it breaks down and reforms during the cell cycle; second, the nucleolus structure itself is dynamic, changing shape, size, and position within the nucleus; third, the nucleolar constituents undergo exchange with pools both within the nucleolus and outside in the nucleoplasm and cytoplasm. Cell Cycle Dynamics The nucleolus disassembles at the end of G2 as most transcription ceases and the nuclear envelope breaks down, and then reassembles with the onset of rDNA transcription at the beginning of the following G1 (Hernandez-Verdun, 2011). During disassembly, the GC components are lost first, followed by the DFC components. Certain proteins, such as Pol I subunits and upstream binding factor, which modulates DNA conformation, remain with the rDNA arrays; the presence of upstream binding factor alone is sufficient to produce a secondary constriction in the mitotic chromosome (Prieto and McStay, 2008). Some nucleolar components diffuse throughout the mitotic cytoplasm, whereas others, such as the protein B23, associate with the periphery of the mitotic chromosomes as chromosomal passengers. When rDNA transcription is halted during mitosis, unprocessed pre-rRNA transcripts persist through the mitotic cell, demonstrating that prerRNA transcript processing is also halted. The nucleolus reforms at the end of mitosis. First, small round bodies, called prenucleolar bodies are formed (Hernandez-Verdun, 2011). When transcription of the rDNA is reinitiated, the prenucleolar bodies disappear as new nucleoli are formed. Where more than one active NOR is present in the nucleus, separate nucleoli generally initially form at each active NOR. In plants, these small nucleoli then often fuse together to a single nucleolus as interphase progresses (Shaw and Jordan, 1995). Protein Mobility For many years cell biologists visualized fixed cells and thus had a tendency to regard the structures seen as stationary and long lived. Live-cell imaging studies, however, have revealed a much more dynamic picture. Studies using fluorescence recovery after photobleaching of nucleolar and nuclear proteins fused to GFP have shown that virtually all nucleolar and nuclear proteins are in constant flux, exchanging between the nucleolus and cytoplasm. The mean nucleolar residence time of even well-characterized nucleolar proteins is only a few tens of seconds (Phair and Misteli, 2000; Chen and Huang, 2001; Olson and Dundr, 2005). The distinction between nuclear and nucleolar proteins is that nucleolar proteins spend a greater proportion of their time in the nucleolus. The structure and even the existence of the nucleolus as a discrete structure must depend on the rDNA nucleating a small subpopulation of proteins that then form a structure on which all the other proteins assemble and disassemble dynamically. The nucleolus (and other nuclear bodies such as CBs) thus represents a steady-state flux of proteins in rapid equilibrium with the surrounding nucleoplasm (Raska et al., 2006a). CBs and Intranucleolar Bodies A number of dynamic nuclear bodies are either associated with the nucleolus or contained within it or both (Mao et al., 2011). The most familiar are the CBs. These bodies were first identified more than 100 years ago by Ramon y Cajal in neuronal cells, were originally called accessory bodies, and were proposed to have a connection with the nucleolus (Gall, 2000). They were later rechristened coiled bodies because of their appearance in the EM, but have now been renamed in honor of their original discoverer. Live-cell imaging of CBs by GFP showed that CBs move, fuse, and split within the nucleus of both plant (Boudonck et al., 1999) and animal (Platani et al., 2002) cells, often migrating dramatically to the nucleolar periphery or even being contained within the nucleolus. The detailed function of CBs is still not well understood, but their role is in maturation of RNA complexes such as spliceososomal subcomplexes, small RNAs, and RNA complexes involved in silencing (Stanek and Neugebauer, 2006). CBs share many components with the nucleolus, particularly a number of small nucleolar RNAs, and RNA processing proteins such as fibrillarin, which methylates various RNA species, and dyskerin/Cbf5p, which isomerizes uridine to pseudouridine in RNAs. A new intranucleolar body (INB) has recently been described, based on colocalization of about 20 well-characterized components (Hutten et al., 2011). The composition of INBs seems distinctly different from CBs; in particular they contain small ubiquitin-like modifier, a peptide modifier, probably conjugated to other substrate proteins. The INBs seem to have an involvement in DNA damage response, since treatments that caused DNA damage induced INBs. The INBs are completely enveloped in the nucleolus, and may be contained in or overlap with a central region of the nucleolus often called the nucleolar cavity, a structure that is particularly prominent in many plant cells and that has been shown to swell and contract dynamically. THE NUCLEOLUS, STRESS, AND DNA DAMAGE SENSING There are several lines of evidence suggesting that the nucleolus has a role in sensing and responding to stresses (Boulon et al., 2010). In mammals there is considerable evidence linking the P53 DNA damage-sensing pathway to the nucleolus. P53 is normally held at a low level by MDM2, an E3 ubiquitin ligase that ubiquitinates P53 and targets it for degradation. MDM2 can be inhibited by ARF, which is normally sequestered in the nucleolus. In one model, release of ARF from the nucleolus would then allow it to inhibit MDM2, with p53 levels consequently rising (Olson, 2004; Raska et al., 2006a). In an elegant series of experiments it was shown that the P53 pathway is induced by many different treatments that targeted the nucleolus (Rubbi and Milner, 2003), suggesting that the nucleolar structure itself is a direct sensor of DNA damage. In this respect the induction of INBs by DNA damage as described above is very interesting and may contribute to the sensing mechanism. rDNA has been associated with stress response to DNA damage and with aging, the pioneering studies of this going back several decades (Johnson and Strehler, 1972). rDNA copies are multiplied by repeated recombination, and their homogeneity is maintained by gene conversion events between the tandem repeats (Kobayashi, 2008). Recombination is induced by Fob1, which causes double-strand break formation. In yeast, at least, the histone deacetylase Sir2p also has a role in rDNA copy number regulation; in a sir2 mutant copy number fluctuated wildly, whereas in a fob1 mutant rDNA repeat fluctuation was reduced or eliminated (Kobayashi, 2008). In yeast, SIR1 and FOB1 affect cellular aging, sir1 mutants having a shorter lifespan and fob1 mutants a longer lifespan (Kobayashi, 2008). It was earlier shown that budding yeast cells accumulate extrachromosomal circles from the rDNA repeats preferentially in the mother cells as they age and that this accumulation in fact is a cause of aging (Sinclair and Guarente, 1997). It is possible that there are cell types or developmental stages in some organisms that require many more rDNA copies than are normally transcribed, but even in the yeast only about half the (approximately 150) copies are transcribed, and in a number of organisms, including yeast, viable mutants have been made with only a fraction of the normal number of rDNA copies (Takeuchi et al., 2003). There is a broad correlation between genome size and number of rDNA copies (Prokopowich et al., 2003), and this has led to a hypothesis that the rDNA may be acting as a sensor for DNA damage, protecting the rest of the genome by inducing DNA repair mechanisms or apoptosis. The extra copies present in the rDNA repeats would initially presumably buffer such damage, ensuring that sufficient undamaged copies were available for ribosome biosynthesis (Kobayashi, 2008). rDNA chromatin also seems to be able to affect the stability of heterochromatic repeats in trans, as loss of the NoRC component TIP5 leads to instability of microsatellite repeats (Guetg et al., 2010) and reduction in the number of Drosophila rDNA repeats themselves leads to a general release of heterochromatin silencing throughout the nucleus (Paredes and Maggert, 2009). Similarly, loss of chromatin assembly factor 1 activity in Arabidopsis led to loss of telomeric and rDNA repeats in successive generations, as well as enhanced sensitivity to DNA damage (Mozgová et al., 2010). As rDNA repeats are the most common gene in the genome, these effects may occur due to disrupted balance between euchromatin and heterochromatin. However, at least in yeast, there is evidence that rDNA organization can also affect the wider genome by regulating the global distribution of condensin (Wang and Strunnikov, 2008). CONCLUSION After many years in which the nucleolus was believed to have a well-understood and limited function in ribosome biogenesis, many novel results in recent years have pointed to a wide range of biological activities being localized to this region of the nucleus. As a great variety of species from different kingdoms has been used for these studies, we cannot as yet tell whether all these various functions are carried out in the nucleoli of all species. For the most part, what unifies these activities is the involvement of RNA at some level, usually in the biogenesis or assembly of RNA/protein machinery. Thus the nucleolus may be better regarded as an RNA processing center, rather than as purely a ribosome factory. There is clearly much to be done to clearly define all the activities of the nucleolus, and further to explain why these activities need to be partitioned together within a defined nuclear domain. The answer may lie in efficiency—increasing the local concentrations of limiting factors by sequestering them to the nucleolus. Alternatively, the answer may lie in evolutionary history; the RNA biosynthetic activities may have colocalized for reasons such as the use of common factors in different pathways. This may have left the various processes inextricably linked. LITERATURE CITED Ahmad Y Boisvert F-M Gregor P Cobley A Lamond AI ( 2009 ) NOPdb: Nucleolar Proteome Database—2008 update . Nucleic Acids Res ( Database issue ) 37 : D181 – D184 Google Scholar PubMed OpenURL Placeholder Text WorldCat Andersen JS Lam YW Leung AKL Ong SE Lyon CE Lamond AI Mann M ( 2005 ) Nucleolar proteome dynamics . 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This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Van Buskirk, Elise K.; Decker, Peter V.; Chen, Meng

doi: 10.1104/pp.111.186411pmid: 21951469

Plant development and growth are extremely plastic in response to changes in ambient light conditions. Light is not only the ultimate energy source for photosynthesis; its physical parameters, such as quality, intensity, direction, and duration, also serve as key environmental and time cues (Chen et al., 2004). Therefore, it is vital for plants to closely monitor and precisely respond to changes in light properties in order to optimize growth under a wide range of ecological environments and to synchronize developmental transitions with diurnal and seasonal time. Plants have evolved to “see” the light spectrum between 280 and 750 nm, which spans UV-B, UV-A, and visible light, through five classes of photoreceptors. In the reference plant Arabidopsis (Arabidopsis thaliana), they include the newly determined UV-B receptor UV RESISTANCE LOCUS8 (UVR8; Kaiserli and Jenkins, 2007; Rizzini et al., 2011); three types of UV-A/blue light receptors, including cryptochromes (cry1 to 3; Kleine et al., 2003; Yu et al., 2010; Chaves et al., 2011), phototropins (phot1 and 2; Briggs and Christie, 2002), and the family of ZEITLUPE/FLAVIN-BINDING, KELCH, F-BOX1 (FKF1)/LOV KELCH PROTEIN2 (LKP2; Nelson et al., 2000; Somers et al., 2000; Schultz et al., 2001); and the red and far-red sensing phytochromes (phyA to E; Quail, 2010). Collectively, these photoreceptors regulate almost every facet of plant development and growth from seed germination to floral initiation (Franklin and Quail, 2010; Kami et al., 2010). Many light responses are mediated by alterations in gene expression. Among the 15 photoreceptors discovered so far, 10 of them, UVR8 (Kaiserli and Jenkins, 2007; Favory et al., 2009), cry1 (Wu and Spalding, 2007), cry2 (Kleiner et al., 1999; Yu et al., 2009), FKF1 (Sawa et al., 2007; Fornara et al., 2009), LKP2 (Yasuhara et al., 2004), and phyA to E (Sakamoto and Nagatani, 1996; Yamaguchi et al., 1999; Kircher et al., 2002), have been shown to localize to the nucleus, where they regulate gene expression in a light-dependent manner (Jiao et al., 2007). An emerging common mechanism for such light-dependent gene expression is through regulating the stability of key transcriptional regulators (discussed below; Yi and Deng, 2005; Brown et al., 2009; Chen and Chory, 2011; Leivar and Quail, 2011). At the cellular level, photoactivation of photoreceptors triggers the rapid localization of a number of them, including phyA to E, cry2, and possibly cry1 and UVR8, to discrete subnuclear foci or nuclear bodies (Kleiner et al., 1999; Yamaguchi et al., 1999; Wang et al., 2001; Kircher et al., 2002; Favory et al., 2009; Yu et al., 2009; Gu et al., 2011; Lian et al., 2011; Liu et al., 2011). Nuclear bodies are morphologically distinct subnuclear domains that provide microenvironments for the regulation of protein dynamics, gene expression, and DNA replication and repair in both plant and animal cells (Shaw and Brown, 2004; Spector, 2006). The general principles of nuclear body function and assembly are still largely unknown. The photoreceptor-containing nuclear bodies, or “photobodies,” found in plants are a unique type of subnuclear domain whose size, number, and potentially function are directly regulated by external light cues (Chen and Chory, 2011). Observations of these speckle-shaped photobodies have raised many questions. How is the formation of photobodies regulated? What are the functions of the photobodies? What are the factors required for photobody formation? These are some of the key questions that are being actively investigated. Further understanding of photobody function and regulation will not only be important for understanding the subcellular organization of light signaling events in plants but also could potentially uncover general principles governing subnuclear domains in higher eukaryotes. In this review, we will summarize recent developments related to phy-containing photobodies, touch briefly on photobodies containing crys, and discuss the potential functions of photobodies in relationship to protein degradation and gene expression. For more comprehensive coverage of light signaling in plants, the reader is referred to the following recent reviews (Henriques et al., 2009; Jenkins, 2009; Chory, 2010; Franklin and Quail, 2010; Kami et al., 2010; Lau and Deng, 2010; Möglich et al., 2010; Nagatani, 2010; Rockwell and Lagarias, 2010; Yu et al., 2010; Chaves et al., 2011; Chen and Chory, 2011; Leivar and Quail, 2011; Ulijasz and Vierstra, 2011). LIGHT-DEPENDENT DYNAMICS OF PHYTOCHROME PHOTOBODY FORMATION Phys in higher plants are red (R) and far-red (FR) light receptors that use a linear tetrapyrrole, phytochromobilin, as their chromophore. Phys can interconvert between two relatively stable conformers: an R light-absorbing inactive Pr form (λmax = 660) and a FR light-absorbing active Pfr form (λmax = 730; Rockwell et al., 2006). Although it was initially thought that phys localized and functioned primarily in the cytoplasm, a series of studies performed over a decade ago using GUS-tagged and fluorescent protein-tagged phys convincingly demonstrated that photoactivation from the Pr to the Pfr form results in the rapid translocation of phys from the cytoplasm to the nucleus (Sakamoto and Nagatani, 1996; Kircher et al., 1999, 2002; Yamaguchi et al., 1999; Kim et al., 2000). This change in localization is one of the earliest phy responses to light; for both phyA and phyB, the most prominent phys in Arabidopsis, nuclear accumulation is required for most of their downstream responses (Huq et al., 2003; Genoud et al., 2008). It was first reported by Akira Nagatani and colleagues that photoactivated phyB-GFP was not only localized to the nucleus but also further compartmentalized to subnuclear speckle-like photobodies (Yamaguchi et al., 1999). Parallel studies by Eberhard Schäfer, Ferenc Nagy, and colleagues demonstrated that all five Arabidopsis phys, phyA to E, localize to photobodies in the light and that phy photobody localization is conserved in both dicotyledonous and monocotyledonous plants (Kircher et al., 1999, 2002; Kim et al., 2000). Although most studies on phy photobodies were conducted using transgenic lines that overexpressed fluorescent protein-tagged phys, both native pea (Pisum sativum) phyA and Arabidopsis phyB photobodies have been observed using immunocytochemistry, suggesting that the formation of photobodies is not an artifact of phy overexpression (Hisada et al., 2000; Kircher et al., 2002). The translocation of phys to photobodies happens very quickly during the dark-to-light transition; photobodies containing both phyA and phyB can be observed after 1 to 2 min of R light exposure (Bauer et al., 2004). PhyB photobody localization is triggered by R light (Yamaguchi et al., 1999; Kircher et al., 2002). In contrast, phyA photobody localization is triggered by R, FR, and blue light (Kim et al., 2000). These “early” photobodies are transient and disappear after 1 h of light exposure (Bauer et al., 2004). Phy photobodies reappear after 2 h in R light and remain present in the light (Yamaguchi et al., 1999; Bauer et al., 2004). These “late” photobodies contain mainly phyB, because phyA is rapidly degraded in R light (Kim et al., 2000; Kircher et al., 2002). Joanne Chory and colleagues showed that the steady-state pattern (size and number) of phyB photobodies under continuous R light is determined by the percentage of phyB in the Pfr form at a given moment (Chen et al., 2003). Light conditions that shift the Pr/Pfr equilibrium to the Pfr side or stabilize the Pfr form will promote large phy photobody formation. Consistent with this notion, under high-intensity R light, which drives the equilibrium to the Pfr form, phyB appears to be localized exclusively to a few large photobodies with diameters between 1 and 2 μm (Chen et al., 2003, 2010b). By contrast, under dim R light or light with a low R-to-FR ratio, where more phyB stays in the Pr form, phyB tends to localize to many smaller photobodies or localizes diffusely in the nucleoplasm (Fig. 1; Chen et al., 2003). The formation of large phyB photobodies correlates tightly with the light-dependent hypocotyl inhibition response. The fact that the steady-state pattern of phyB-GFP is predictable and can be precisely manipulated by external light quantity and quality makes it an excellent visible readout for genetic screens (discussed below; Chen, 2008). Although phyB photobodies appear to be morphologically stable, they are actually quite dynamic subnuclear domains; fluorescence recovery after photobleaching experiments on phyB-yellow fluorescent protein (YFP) photobodies showed that photobody-associated phyB-YFP is rapidly exchanged with nucleoplasmic phyB-YFP (Rausenberger et al., 2010). Figure 1. Open in new tabDownload slide The morphology of phy photobodies is directly regulated by light. Confocal images of phyB-GFP localization patterns and corresponding PBG seedlings under increasing intensities of red light are shown. PhyB-GFP is evenly distributed under dim (0.5 μmol m−2 s−1) R light. With increasing R light intensity (1 μmol m−2 s−1 and 2 μmol m−2 s−1), phyB-GFP starts to form exclusively small, or both small and large photobodies, respectively. Under strong R light (above 8 μmol m−2 s−1), phyB-GFP localizes exclusively to large photobodies. The localization of phyB-GFP correlates with the degree of hypocotyl inhibition in the light. Figure 1. Open in new tabDownload slide The morphology of phy photobodies is directly regulated by light. Confocal images of phyB-GFP localization patterns and corresponding PBG seedlings under increasing intensities of red light are shown. PhyB-GFP is evenly distributed under dim (0.5 μmol m−2 s−1) R light. With increasing R light intensity (1 μmol m−2 s−1 and 2 μmol m−2 s−1), phyB-GFP starts to form exclusively small, or both small and large photobodies, respectively. Under strong R light (above 8 μmol m−2 s−1), phyB-GFP localizes exclusively to large photobodies. The localization of phyB-GFP correlates with the degree of hypocotyl inhibition in the light. Some phy photobodies might also contain the blue light receptor cry2. Crys are photolyase-like photoreceptors that use FAD as the chromophore. When phyB-GFP and cry2-red fluorescent protein were coexpressed in BY-2 protoplasts, not only did they colocalize on photobodies, they could also be coimmunoprecipitated (Más et al., 2000). Because cry2 is photolabile and rapidly degraded in blue light, it could be colocalized with phyB in early photobodies during the dark-to-light transition. Consistent with this hypothesis, cry2 photobody localization is also a rapid light response, as Arabidopsis cry2 is translocated to photobodies within 15 min after blue light exposure (Yu et al., 2009). Activated cry1 has also been suggested to localize to nuclear bodies (Wang et al., 2001; Gu et al., 2011; Lian et al., 2011; Liu et al., 2011). Because cry1 and cry2 were colocalized with COP1 and SPA1 on nuclear bodies (Gu et al., 2011; Lian et al., 2011; Liu et al., 2011; Zuo et al., 2011), it is likely that cry1, similar to cry2, could also localize to phy-containing photobodies in the light. STRUCTURAL BASIS OF PHYTOCHROME PHOTOBODY LOCALIZATION The intramolecular requirements for photobody localization have been most extensively studied using Arabidopsis phyB. Two general approaches have been taken to dissect the phy subdomains required for photobody localization: one approach is to examine the localization pattern of phy truncation fragments; the other approach is to characterize the localization pattern of loss-of-function or gain-of-function alleles of phy. The domain structure of phys has been well defined. Phys can form either homodimers or heterodimers (Clack et al., 2009); each monomer is an approximately 125-kD polypeptide. The phy protein can be divided into two domains: an N-terminal photosensory and signaling domain and a C-terminal dimerization domain, with a hinge region connecting the two (Fig. 2; Nagatani, 2010). The N-terminal domain comprises four subdomains: an N-terminal extension (NTE); a PAS (for PER, ARNT, and SIM) domain; a GAF (for cGMP phosphodiesterase, adenylate cyclase, and FhlA) domain, which contains a conserved Cys residue forming a thioether linkage with the A ring of the chromophore phytochromobilin; and a PHY (for phytochrome-specific GAF-related) domain (Rockwell et al., 2006). The crystal structure of the PAS/GAF domain of the bacteriophytochrome DrBph1 from Deinococcus radiodurans revealed the presence of a light-sensing knot, which plays an important role in signaling. For example, amino acid residues located in the knot are involved in the interaction with phytochrome-interacting factors (PIFs; Oka et al., 2008; Kikis et al., 2009; Nagatani, 2010; Ulijasz and Vierstra, 2011). The C-terminal half of phys contains two subdomains, a PAS-related domain (PRD) containing two PAS domains (PAS-A and PAS-B) and a His kinase-related domain (HKRD; Rockwell et al., 2006). Figure 2. Open in new tabDownload slide Structural basis for phyB localization and signaling. Phys can be divided into the N-terminal light-sensing and signaling domain and the C-terminal dimerization and localization domain (Nagatani, 2010). The GAF and PHY domains of phyB physically interact with the PRD to mask nuclear localization signals located in the PRD (Chen et al., 2005). Additionally, although the PRD alone is sufficient for nuclear import, the PRD and HKRD together are required for normal phyB photobody localization (Matsushita et al., 2003; Chen et al., 2005). Mutations in red are those that have apparently normal photobody localization but fail to complement a null phyB mutant, possibly due to reduced interaction with PIF proteins (Oka et al., 2008; Kikis et al., 2009). The Y276H mutation, in green, causes constitutive phyB localization to large photobodies and constitutive phytochrome signaling (Su and Lagarias, 2007). The G767R mutation, in brown, results in the inability of phyB to localize to the nucleus (Matsushita et al., 2003). All other mutations, represented in blue, show abnormal photobody localization as well as impaired light signaling (Kircher et al., 2002; Chen et al., 2003; Oka et al., 2008). Specifically, mutant G118R does not incorporate the chromophore; mutants C327Y and A587T show slightly faster dark reversion; and mutant A372T shows a slightly red-shifted spectrum in addition to highly accelerated dark reversion. The chromophore is represented by four consecutive squares. Figure 2. Open in new tabDownload slide Structural basis for phyB localization and signaling. Phys can be divided into the N-terminal light-sensing and signaling domain and the C-terminal dimerization and localization domain (Nagatani, 2010). The GAF and PHY domains of phyB physically interact with the PRD to mask nuclear localization signals located in the PRD (Chen et al., 2005). Additionally, although the PRD alone is sufficient for nuclear import, the PRD and HKRD together are required for normal phyB photobody localization (Matsushita et al., 2003; Chen et al., 2005). Mutations in red are those that have apparently normal photobody localization but fail to complement a null phyB mutant, possibly due to reduced interaction with PIF proteins (Oka et al., 2008; Kikis et al., 2009). The Y276H mutation, in green, causes constitutive phyB localization to large photobodies and constitutive phytochrome signaling (Su and Lagarias, 2007). The G767R mutation, in brown, results in the inability of phyB to localize to the nucleus (Matsushita et al., 2003). All other mutations, represented in blue, show abnormal photobody localization as well as impaired light signaling (Kircher et al., 2002; Chen et al., 2003; Oka et al., 2008). Specifically, mutant G118R does not incorporate the chromophore; mutants C327Y and A587T show slightly faster dark reversion; and mutant A372T shows a slightly red-shifted spectrum in addition to highly accelerated dark reversion. The chromophore is represented by four consecutive squares. Truncation studies have revealed that the C-terminal half of phyB localizes to photobodies independently of light (Chen et al., 2003; Matsushita et al., 2003). Similarly, the photobody localization of phyA also requires its C-terminal half (Wolf et al., 2011). Within the C-terminal domain of phyB, the PRD is both required and sufficient for nuclear localization, suggesting that it either possesses a nuclear localization signal (NLS) or is able to bind to an unidentified shuttle protein containing a NLS (Matsushita et al., 2003; Chen et al., 2005). Both the PRD and HKRD are required for normal photobody localization (Chen et al., 2005). Consistent with this notion, several missense mutations that result in defective nuclear or photobody localization have been identified within the PRD (Fig. 2; Kircher et al., 2002; Chen et al., 2003; Matsushita et al., 2003). The role of the HKRD in photobody localization is still a mystery; although truncations lacking the entire HKRD do not localize to photobodies, a phyB truncation lacking a portion of the HKRD could still form smaller photobodies (Chen et al., 2005). Because nuclear and photobody localization are both light dependent, it raises the question of how the N-terminal photosensory domain regulates the C-terminal NLS/photobody localization signals. The current model is that, in the Pr form, C-terminal localization signals are masked by the N-terminal domain through an interaction between the GAF-PHY subdomains and the PRD, whereas both the putative NLS and photobody localization signals are exposed in the Pfr form as a result of light-dependent conformational changes (Chen, 2008; Fankhauser and Chen, 2008). The “open” conformation of the Pfr form could also expose domains required for interacting with other signaling components. Consistent with the notion that the conformation of Pfr is important for photobody localization, the NTE, which plays a role in stabilizing the Pfr form, is also required for phyB photobody formation (Chen et al., 2005). Localization studies of missense phyB alleles have further demonstrated that the Pfr form of phyB is required for photobody localization. N-terminal missense loss-of-function phyB alleles that are defective in photobody localization also have less stable Pfr, and some have abnormal light absorption spectra (Fig. 2; Chen et al., 2003; Oka et al., 2008). By contrast, YHB, a constitutively active phyB mutant, localizes to photobodies regardless of light conditions (Su and Lagarias, 2007). Moreover, loss-of-function mutations that only affect signaling but not the absorption properties of phyB have normal photobody localization patterns (Oka et al., 2008; Kikis et al., 2009), which further suggests that the photobody localization depends on phyB being in the Pfr form and is not a consequence of phy signaling. POSSIBLE FUNCTIONS OF PHOTOBODIES IN LIGHT SIGNALING Ever since the initial observation of phy photobodies, there has been much speculation about their function. One hypothesis is that the photobodies are storage depots for active photoreceptors but are not required for light signaling (Fig. 3A). In this model, photobodies serve as a valve to regulate the amount of active phy in the nucleoplasm. Consistent with this model, the N terminus of phyB fused to a dimerization domain and a NLS is active in mediating light responses but does not localize to photobodies (Matsushita et al., 2003; Palágyi et al., 2010). However, accumulating evidence from localization and colocalization studies on light signaling components supports the idea that photobodies might be the sites of light signaling. Figure 3. Open in new tabDownload slide Alternative models of photobody function. A, The storage depot model. In this model, the amount of photoactivated nucleoplasmic phys is regulated by sequestering them within photobodies. These photobodies serve as storage depots that later release the phys into the nucleoplasm to carry out their signaling functions, resulting in the regulation of light-responsive genes. B, The degradation model. In this model, the photobodies are sites for the ubiquitylation and degradation of key transcriptional regulators. C, The transcription model. Transcriptional regulators localize to photobodies, bringing their target DNA with them. The expression of the target genes is regulated within or in the vicinity of the photobodies. E3, E3 ubiquitin ligase; TR, transcriptional regulator. Figure 3. Open in new tabDownload slide Alternative models of photobody function. A, The storage depot model. In this model, the amount of photoactivated nucleoplasmic phys is regulated by sequestering them within photobodies. These photobodies serve as storage depots that later release the phys into the nucleoplasm to carry out their signaling functions, resulting in the regulation of light-responsive genes. B, The degradation model. In this model, the photobodies are sites for the ubiquitylation and degradation of key transcriptional regulators. C, The transcription model. Transcriptional regulators localize to photobodies, bringing their target DNA with them. The expression of the target genes is regulated within or in the vicinity of the photobodies. E3, E3 ubiquitin ligase; TR, transcriptional regulator. Most phy-mediated responses require global reprogramming of the transcriptome (Tepperman et al., 2006; Jiao et al., 2007; Hu et al., 2009; Leivar et al., 2009; Shin et al., 2009). Two emerging signaling mechanisms suggest that the key signaling events regulating gene expression work by modulating the stability of either positively or negatively acting transcription factors. The positively acting transcription factors include the basic leucine zipper (bZIP) transcription factor HY5 (for elongated hypocotyl 5; Koornneef et al., 1980; Oyama et al., 1997), the MYB factor LAF1 (for long after far-red light 1; Ballesteros et al., 2001), the helix-loop-helix (HLH) factor HFR1 (for long hypocotyl in far-red 1; Fairchild et al., 2000), and some members of the B-box zinc finger family (BBX; Kumagai et al., 2008; Khanna et al., 2009), including CONSTANS (CO)/BBX1 (Laubinger et al., 2006; Liu et al., 2008), COL3 (for CONSTANS-like 3)/BBX4 (Datta et al., 2006), LZF1 (for light-regulated zinc finger protein 1)/STH3 (for salt tolerance homolog 3)/BBX22 (Chang et al., 2008, 2011; Datta et al., 2008), and BBX21/STH2 (Datta et al., 2007). These proteins are degraded in the dark by the E3 ubiquitin ligase COP1 (for constitutively photomorphogenic 1) and/or the cullin4-DDB1 (for damaged DNA-binding protein 1)-COP1-SPA (for suppressor of phytochrome A-105) E3 ubiquitin ligase complex, where COP1 and members of the SPA family of proteins form the substrate recognition complex (Osterlund et al., 2000; Seo et al., 2003; Duek et al., 2004; Jang et al., 2005; Yang et al., 2005; Datta et al., 2006, 2007; Laubinger et al., 2006; Liu et al., 2008; Chen et al., 2010a; Chang et al., 2011). COP1 is also involved in the turnover of both phyA and phyB, partly as a mechanism to attenuate phy signaling in the light (Seo et al., 2004; Jang et al., 2010). The current model is that phys promote the stability of this group of positively acting transcription regulators by repressing E3 ubiquitin ligases (Chen and Chory, 2011). Although the molecular mechanism of how phys repress COP1 and/or the COP1 E3 complex is still unclear, recently it has been shown that crys directly regulate either the formation of the substrate receptor COP1/SPA1 complex or the interaction between the substrate receptor COP1/SPA complex and its target proteins (Lian et al., 2011; Liu et al., 2011; Zuo et al., 2011). It is quite possible that phys could utilize a similar mechanism to regulate the activity of COP1. Besides positively acting transcriptional regulators, there are also transcriptional regulators that are antagonistic to phy signaling. Some of the well-studied members of this group are the bHLH transcription factor called PIFs (Ni et al., 1998; Oh et al., 2007; Shen et al., 2008; Lorrain et al., 2009; Shin et al., 2009; Leivar and Quail, 2011). Phys bind directly to PIFs and trigger their phosphorylation and subsequent degradation in the light (Al-Sady et al., 2006; Leivar and Quail, 2011). The rapid turnover of PIFs in the light is a key mechanism to turn on phy-mediated responses (Leivar et al., 2009; Shin et al., 2009). One widely proposed hypothesis is that photobodies are sites for protein degradation (Fig. 3B). This model is supported by the fact that many signaling components are localized to photobodies prior to their degradation (Table I). For example, during the dark-to-light transition in seedling development, both phyA and PIF3 colocalize to early phy photobodies before their degradation (Al-Sady et al., 2006). In addition, the positively acting transcriptional regulators, including HY5, LAF1, HFR1, and some BBX proteins, also colocalize with COP1 on nuclear bodies (Table I). Moreover, members of the SPA protein family have also been colocalized with COP1 on nuclear bodies (Seo et al., 2003; Zhu et al., 2008). These results suggest that these transcriptional regulators could at least be ubiquitylated on photobodies. In addition, cry2-containing photobodies may be associated with cry2 degradation (Yu et al., 2009). Photobody constituents Table I. Photobody constituents A list of light signaling components that have been shown to localize to photobodies or photobody-like subnuclear domains. These include not only photoreceptors but a number of transcriptional regulators and their E3 ubiquitin ligases, suggesting that photobodies are involved in light-regulated protein degradation and/or transcription. Photobody Constituent Function Reference Photoreceptors  phyA to E R and FR receptors Yamaguchi et al. (1999); Kim et al. (2000); Kircher et al. (2002); Chen et al. (2003); Matsushita et al. (2003); Bauer et al. (2004)  cry1 and 2 UV-A/blue receptors Wang et al. (2001); Yu et al. (2009); Gu et al. (2011); Lian et al. (2011); Liu et al. (2011); Zuo et al. (2011)  UVR8 UV-B receptor Favory et al. (2009) Related to protein degradation  COP1 E3 ubiquitin ligase Stacey and von Arnim (1999); Wang et al. (2001); Seo et al. (2004); Subramanian et al. (2004)  SPA1 to 4 E3 ubiquitin ligase Seo et al. (2003); Laubinger et al. (2006); Zhu et al. (2008); Lian et al. (2011); Liu et al. (2011); Zuo et al. (2011)  HMR Structurally similar to RAD23 Chen et al. (2010b) Transcriptional regulators  PIFs (PIF3 and PIF7) bHLH transcription factor Bauer et al. (2004); Al-Sady et al. (2006); Leivar et al. (2008); Kidokoro et al. (2009)  HFR1 HLH transcriptional regulator Yang et al. (2005); Jang et al. (2007)  HY5/HYH bZIP transcription factor Ang et al. (1998); Holm et al. (2002)  LAF1 MYB transcription factor Ballesteros et al. (2001); Seo et al. (2003); Jang et al. (2007)  BBXs (CO, COL3, LZF1/STH3/BBX22, STH2/BBX21, STO/BBX24, DBB1a/BBX18, and DBB1b/BBX19) Transcriptional regulators Datta et al. (2006, 2007, 2008); Laubinger et al. (2006); Indorf et al. (2007); Jang et al. (2008); Liu et al. (2008); Yan et al. (2011)  ELF3 phyB signaling/clock associated Yu et al. (2008)  GI phyB signaling/clock associated Yu et al. (2008) Other proteins  FHY1/FHL phyA signaling component Hiltbrunner et al. (2005, 2006)  PAPP5 Phosphatase Ryu et al. (2005) Photobody Constituent Function Reference Photoreceptors  phyA to E R and FR receptors Yamaguchi et al. (1999); Kim et al. (2000); Kircher et al. (2002); Chen et al. (2003); Matsushita et al. (2003); Bauer et al. (2004)  cry1 and 2 UV-A/blue receptors Wang et al. (2001); Yu et al. (2009); Gu et al. (2011); Lian et al. (2011); Liu et al. (2011); Zuo et al. (2011)  UVR8 UV-B receptor Favory et al. (2009) Related to protein degradation  COP1 E3 ubiquitin ligase Stacey and von Arnim (1999); Wang et al. (2001); Seo et al. (2004); Subramanian et al. (2004)  SPA1 to 4 E3 ubiquitin ligase Seo et al. (2003); Laubinger et al. (2006); Zhu et al. (2008); Lian et al. (2011); Liu et al. (2011); Zuo et al. (2011)  HMR Structurally similar to RAD23 Chen et al. (2010b) Transcriptional regulators  PIFs (PIF3 and PIF7) bHLH transcription factor Bauer et al. (2004); Al-Sady et al. (2006); Leivar et al. (2008); Kidokoro et al. (2009)  HFR1 HLH transcriptional regulator Yang et al. (2005); Jang et al. (2007)  HY5/HYH bZIP transcription factor Ang et al. (1998); Holm et al. (2002)  LAF1 MYB transcription factor Ballesteros et al. (2001); Seo et al. (2003); Jang et al. (2007)  BBXs (CO, COL3, LZF1/STH3/BBX22, STH2/BBX21, STO/BBX24, DBB1a/BBX18, and DBB1b/BBX19) Transcriptional regulators Datta et al. (2006, 2007, 2008); Laubinger et al. (2006); Indorf et al. (2007); Jang et al. (2008); Liu et al. (2008); Yan et al. (2011)  ELF3 phyB signaling/clock associated Yu et al. (2008)  GI phyB signaling/clock associated Yu et al. (2008) Other proteins  FHY1/FHL phyA signaling component Hiltbrunner et al. (2005, 2006)  PAPP5 Phosphatase Ryu et al. (2005) Open in new tab Table I. Photobody constituents A list of light signaling components that have been shown to localize to photobodies or photobody-like subnuclear domains. These include not only photoreceptors but a number of transcriptional regulators and their E3 ubiquitin ligases, suggesting that photobodies are involved in light-regulated protein degradation and/or transcription. Photobody Constituent Function Reference Photoreceptors  phyA to E R and FR receptors Yamaguchi et al. (1999); Kim et al. (2000); Kircher et al. (2002); Chen et al. (2003); Matsushita et al. (2003); Bauer et al. (2004)  cry1 and 2 UV-A/blue receptors Wang et al. (2001); Yu et al. (2009); Gu et al. (2011); Lian et al. (2011); Liu et al. (2011); Zuo et al. (2011)  UVR8 UV-B receptor Favory et al. (2009) Related to protein degradation  COP1 E3 ubiquitin ligase Stacey and von Arnim (1999); Wang et al. (2001); Seo et al. (2004); Subramanian et al. (2004)  SPA1 to 4 E3 ubiquitin ligase Seo et al. (2003); Laubinger et al. (2006); Zhu et al. (2008); Lian et al. (2011); Liu et al. (2011); Zuo et al. (2011)  HMR Structurally similar to RAD23 Chen et al. (2010b) Transcriptional regulators  PIFs (PIF3 and PIF7) bHLH transcription factor Bauer et al. (2004); Al-Sady et al. (2006); Leivar et al. (2008); Kidokoro et al. (2009)  HFR1 HLH transcriptional regulator Yang et al. (2005); Jang et al. (2007)  HY5/HYH bZIP transcription factor Ang et al. (1998); Holm et al. (2002)  LAF1 MYB transcription factor Ballesteros et al. (2001); Seo et al. (2003); Jang et al. (2007)  BBXs (CO, COL3, LZF1/STH3/BBX22, STH2/BBX21, STO/BBX24, DBB1a/BBX18, and DBB1b/BBX19) Transcriptional regulators Datta et al. (2006, 2007, 2008); Laubinger et al. (2006); Indorf et al. (2007); Jang et al. (2008); Liu et al. (2008); Yan et al. (2011)  ELF3 phyB signaling/clock associated Yu et al. (2008)  GI phyB signaling/clock associated Yu et al. (2008) Other proteins  FHY1/FHL phyA signaling component Hiltbrunner et al. (2005, 2006)  PAPP5 Phosphatase Ryu et al. (2005) Photobody Constituent Function Reference Photoreceptors  phyA to E R and FR receptors Yamaguchi et al. (1999); Kim et al. (2000); Kircher et al. (2002); Chen et al. (2003); Matsushita et al. (2003); Bauer et al. (2004)  cry1 and 2 UV-A/blue receptors Wang et al. (2001); Yu et al. (2009); Gu et al. (2011); Lian et al. (2011); Liu et al. (2011); Zuo et al. (2011)  UVR8 UV-B receptor Favory et al. (2009) Related to protein degradation  COP1 E3 ubiquitin ligase Stacey and von Arnim (1999); Wang et al. (2001); Seo et al. (2004); Subramanian et al. (2004)  SPA1 to 4 E3 ubiquitin ligase Seo et al. (2003); Laubinger et al. (2006); Zhu et al. (2008); Lian et al. (2011); Liu et al. (2011); Zuo et al. (2011)  HMR Structurally similar to RAD23 Chen et al. (2010b) Transcriptional regulators  PIFs (PIF3 and PIF7) bHLH transcription factor Bauer et al. (2004); Al-Sady et al. (2006); Leivar et al. (2008); Kidokoro et al. (2009)  HFR1 HLH transcriptional regulator Yang et al. (2005); Jang et al. (2007)  HY5/HYH bZIP transcription factor Ang et al. (1998); Holm et al. (2002)  LAF1 MYB transcription factor Ballesteros et al. (2001); Seo et al. (2003); Jang et al. (2007)  BBXs (CO, COL3, LZF1/STH3/BBX22, STH2/BBX21, STO/BBX24, DBB1a/BBX18, and DBB1b/BBX19) Transcriptional regulators Datta et al. (2006, 2007, 2008); Laubinger et al. (2006); Indorf et al. (2007); Jang et al. (2008); Liu et al. (2008); Yan et al. (2011)  ELF3 phyB signaling/clock associated Yu et al. (2008)  GI phyB signaling/clock associated Yu et al. (2008) Other proteins  FHY1/FHL phyA signaling component Hiltbrunner et al. (2005, 2006)  PAPP5 Phosphatase Ryu et al. (2005) Open in new tab In mammalian cells, components of the ubiquitin-proteasome pathway have been shown to localize to subnuclear foci called clastosomes (Lafarga et al., 2002). However, it has not been demonstrated whether the proteasome colocalizes with photobodies in plants. In fact, the Arabidopsis proteasome components AtS6A and AtS9 were instead shown to localize to the nucleoplasm (Kwok et al., 1999). CUL4 and DDB1, the other two key components of the CUL4-DDB1-COP1-SPA complex, also localize to the nucleoplasm (Zhang et al., 2008). In addition, COP9, which is the key component of the COP9 signalosome that regulates the activity of cullin-based E3 ubiquitin ligases, localizes to the nucleoplasm in both light- and dark-grown Arabidopsis cotyledon and hypocotyl protoplasts (Chamovitz et al., 1996; Staub et al., 1996). Taken together, these results suggest the possibility that the protein substrates are modified on photobodies and are subsequently degraded in the nucleoplasm. Future investigation into the constituents of photobodies will further clarify this model. Although photobodies are likely involved in protein degradation, it is quite clear that not all of their constituents are subject to protein degradation. For example, PIF7 is localized to phy photobodies but is stable in the light (Leivar et al., 2008). What other functions could photobodies have besides protein degradation? One possibility is that photobodies are involved in transcriptional regulation (Fig. 3C). This is supported by the fact that many of the photobody constituents are transcriptional regulators, which could bring their targeted genes to the vicinity of photobodies. The link between nuclear bodies and transcriptional regulation is well documented (Zhao et al., 2009). For example, Promyelocytic Leukemia (PML) protein bodies have been shown to organize the higher order chromatin structures of the Major Histocompatibility Complex (MHC) class I locus, and they regulate the expression of MHC class I genes in mammalian cell lines by direct interactions between PML and Special AT-Rich Sequence Binding Protein1 (Kumar et al., 2007). Likewise, phy photobodies could serve as organization centers involved in the regulation of light-responsive genes (Fig. 3C). This model and the degradation model in Figure 3B are not mutually exclusive, as the degradation of some transcriptional activators has been shown to be coupled with their transcriptional activity (Lipford et al., 2005; Collins and Tansey, 2006). Therefore, it is also possible that photobodies are sites for both the degradation of transcriptional regulators and the regulation of transcription. GENETIC DISSECTION OF PHOTOBODY FUNCTION Although nuclear bodies have been extensively studied, particularly in mammalian systems, and key constituents of a few nuclear bodies have been successfully identified by cell biology and proteomic approaches (Gall, 2000; Bernardi and Pandolfi, 2007), the precise function and regulation of nuclear bodies are still poorly understood. Arabidopsis represents an ideal organism to dissect the function of nuclear bodies by the combination of molecular genetic and cell biological approaches (Shaw and Brown, 2004; Collier et al., 2006; Fang and Spector, 2010). Because their steady-state pattern can be precisely manipulated by external light quality and quantity, phyB-containing photobodies provide an excellent model system to investigate the function and regulatory mechanisms of nuclear bodies in relation to signaling events in the nucleus (Chen, 2008). We recently reported a forward genetic screen aimed at isolating mutants defective in phyB-GFP photobody localization in the light (Chen et al., 2010b). This screen identified a novel phy signaling component, HEMERA (HMR), which itself is localized to the periphery of phyB photobodies. The hmr mutant represents a new class of light signaling mutants that are albino and tall in R and FR light. In addition, phyB-GFP localizes to smaller photobodies in hmr (Fig. 4). More interestingly, in hmr mutants, phyA, PIF1, and PIF3 accumulate in the light (Chen et al., 2010b). HMR is predicted to be structurally similar to RAD23, which is a multiubiquitin receptor that delivers multiubiquitylated proteins to the proteasome for degradation, suggesting that HMR could play a similar role in phyA, PIF1, and PIF3 degradation in the light (Chen et al., 2010b). Taken together, these results provide genetic evidence supporting the model in which photobodies are sites for protein degradation (Fig. 3B; Chen et al., 2010b). Further investigation of the biochemical functions of HMR as well as the identification of other genes from the same genetic screen will likely provide greater insight into the link between photobodies and protein degradation. Figure 4. Open in new tabDownload slide The hmr mutant. Images of 4-d-old PBG (the parental type of hmr-1) and hmr-1 mutant seedlings grown under 8 μmol m−2 s−1 light. The hmr seedling has both long-hypocotyl and albino phenotypes. Confocal images show that phyB-GFP is localized to large photobodies in PBG seedlings. By contrast, phyB-GFP fails to form large photobodies and is instead localized to smaller photobodies in hmr-1. Figure 4. Open in new tabDownload slide The hmr mutant. Images of 4-d-old PBG (the parental type of hmr-1) and hmr-1 mutant seedlings grown under 8 μmol m−2 s−1 light. The hmr seedling has both long-hypocotyl and albino phenotypes. Confocal images show that phyB-GFP is localized to large photobodies in PBG seedlings. By contrast, phyB-GFP fails to form large photobodies and is instead localized to smaller photobodies in hmr-1. PERSPECTIVE Photoreceptor-containing photobodies in plants are unique and fascinating subnuclear domains whose assembly and function are directly regulated by light. The localization of photoreceptors, including phys and crys, to photobodies is triggered by a light-induced conformational switch to their active states. It is likely that the exposure of certain domains in the active state, such as the C-terminal domain of phyB, facilitates new protein-protein interactions and the “formation” or recruitment of photoreceptors and other signaling molecules to photobodies. The detailed mechanism of photobody assembly is still elusive. Localization studies of light signaling components and recent genetic evidence support the model that photobodies are sites for light signaling events, such as light-dependent turnover of key transcriptional regulators. However, we have only begun to understand the regulatory mechanisms and functions of photobodies. Many key questions remain to be answered. It is still not clear whether photobody-localized transcriptional regulators are degraded or only modified on photobodies. We still do not know whether photobodies are directly involved in transcriptional regulation and whether they are associated with chromatin. Accumulating evidence shows a convergence between the light signaling pathway and other signaling pathways, including those of temperature, hormones, and the circadian clock, on shared downstream signaling molecules (Kami et al., 2010; Lau and Deng, 2010; Leivar and Quail, 2011). Could photobodies serve as a hub for the interaction between these signaling pathways? 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Liu, Qi; Shi, Leilei; Fang, Yuda

doi: 10.1104/pp.111.186734pmid: 22025607

In eukaryotes, small noncoding RNAs of approximately 21 to 24 nucleotides function as guide molecules in many biological processes, including genome organization and stability, developmental timing and patterning, and antibacterial and antiviral defense (Carrington and Ambros, 2003; Poethig, 2009; Simon and Meyers, 2011). The small RNAs regulate the functions of target DNA or RNA in a sequence-specific manner at either the transcriptional or posttranscriptional level through an RNA-silencing mechanism (Hammond, 2005; Czech and Hannon, 2011). Based on whether RNase III family proteins participate in the biogenesis, the small RNAs are divided into at least two classes: RNase III family protein-dependent small RNAs, including microRNAs (miRNAs) and many small interfering RNAs (siRNAs); and RNase III family protein-independent small RNAs, including Piwi-RNAs and secondary siRNAs that are processed from single-stranded precursors in worms (Czech and Hannon, 2011). The mature miRNAs and siRNAs are sorted and loaded specifically with Argonaute (AGO) subfamily proteins, forming the RNA-induced silencing complexes (RISCs) that undergo a specific RNA-silencing mechanism (Ender and Meister, 2010; Fabian et al., 2010). Here, we briefly summarize the molecular basis of miRNA biogenesis pathways and provide an update on nuclear dicing bodies (D-bodies), structures involved in miRNA processing in plant cells. For an overview on siRNAs and other small RNAs, readers are referred to recent excellent articles (Li et al., 2006; Pontes et al., 2006; Ahmad et al., 2010; Chen, 2010a; Law and Jacobsen, 2010; Czech and Hannon, 2011; Simon and Meyers, 2011; Zhang and Zhu, 2011). THE MIRNA BIOGENESIS PATHWAYS IN ANIMALS AND PLANTS MiRNAs (approximately 21–22 nucleotides) are a class of small, regulatory RNAs that are found in almost all of the eukaryotes (Reinhart et al., 2000; Lau et al., 2001; Llave et al., 2002; Molnár et al., 2007; Zhao et al., 2007; Chen, 2010b). Like protein-coding genes, miRNA genes in both plants and animals are transcribed by RNA polymerase II into primary transcripts, known as pri-miRNAs. Animal miRNAs are often clustered on the same precursor. These pri-miRNAs are subject to 5′ capping, 3′ polyadenylation, and splicing, as some of the pri-miRNAs may contain introns. A pri-miRNA contains a stem-loop structure: an imperfect double-stranded (ds) RNA hairpin that harbors the mature miRNA (Bartel, 2004; Cai et al., 2004; Lee et al., 2004; Xie et al., 2005; Kim and Nam, 2006; Laubinger et al., 2008; Chen, 2010b). The pri-miRNAs are then processed by two substantial site-specific endonucleolytic events and eventually turned into miRNA duplexes. In animals, the initial step is converting the pri-miRNAs into precursor miRNAs (pre-miRNAs) in the nucleus through the microprocessor, a complex that contains the RNase III family protein Drosha and its partner Pasha/DiGeorge syndrome critical region gene 8, a dsRNA-binding (dsRBD) protein (Denli et al., 2004; Han et al., 2004a; Landthaler et al., 2004; Zeng et al., 2005). The specific sites of the pri-miRNA are recognized by the microprocessor and cleaved by Drosha to generate an approximately 60- to 70-nucleotide, folded pre-miRNA with a two-nucleotide overhang at the 3′ end (Han et al., 2006; Chen, 2010a). Mitrons, some intronic pri-miRNAs, are converted into pre-miRNAs by the RNA-splicing machinery rather than the microprocessor complex (Berezikov et al., 2007; Okamura et al., 2007; Ruby et al., 2007; Flynt et al., 2010). Recently, additional factors, including the nuclear export receptor Exportin1, the cap-binding complex, and ARSENITE-RESISITANCE PROTEIN2, were found to participate in this process (Laubinger et al., 2008; Gruber et al., 2009; Sabin et al., 2009; Buessing et al., 2010). The pre-miRNAs are then recognized by the nuclear export protein Exportin5 and transported to the cytoplasm in a Ran-GTP-dependent manner (Yi et al., 2003; Bohnsack et al., 2004; Lund et al., 2004). The secondary step is to generate the approximately 22- to 23-nucleotide miRNA/miRNA* duplex from the pre-miRNA through a cytoplasmic RNase III-like enzyme and its specific dsRBD partner (Bernstein et al., 2001; Hutvágner et al., 2001). In mammals, the RNase III enzyme Dicer functions in association with the cytoplasmic dsRBD protein TAR RNA-binding protein 2, whereas in Drosophila melanogaster, Dicer1 with its cofactor Loquacious plays a role in this transition (Chendrimada et al., 2005; Jiang et al., 2005; Saito et al., 2005; Park et al., 2007). The mature RNAs are sorted and loaded with specific AGO proteins to assemble RISCs. Sorting may depend on the interaction between components of biogenesis and effectors, 5′ end nucleotide and thermodynamics of the small RNA duplexes, and/or the structure biases of the AGO family (Khvorova et al., 2003; Schwarz et al., 2003; Gregory et al., 2005; Miyoshi et al., 2005; Rand et al., 2005; Mi et al., 2008; Okamura et al., 2009; Frank et al., 2010). In contrast to the nuclear cropping and cytoplasmic dicing of pri-miRNAs in animals, both of the steps in pri-miRNA processing occur in the nucleus of plant cells. In addition, a single plant RNase III family protein, DICER-LIKE1 (DCL1), plays similar roles to both nuclear Drosha and cytoplasmic Dicer in animals (Park et al., 2002; Reinhart et al., 2002; Schauer et al., 2002; Kurihara and Watanabe, 2004). Plant miRNA genes are also transcribed by RNA polymerase II, and the primary transcripts are 5′ capped and 3′ polyadenylated (Xie et al., 2005; Kim and Nam, 2006; Chen, 2010a). The pri-miRNAs are cropped into pre-miRNAs with a shorter stem-loop structure, which are further cut into the miRNA/miRNA* duplex. This process requires the interaction of DCL1 with its dsRBD protein partner HYPONSTIC LEAVES1 (HYL1), similar to Drosha and Dicer in animals, which are assisted by a specific dsRBD protein partner (Han et al., 2004a, 2004b; Vazquez et al., 2004; Kurihara et al., 2006; Yang et al., 2010). The C2H2 zinc finger protein SERRATE (SE) also interacts with DCL1 and HYL1 and participates in the transition process (Yang et al., 2006a; Laubinger et al., 2008; Montgomery and Carrington, 2008). HYL1 together with SE promotes the accuracy of miRNA processing (Dong et al., 2008). Many miRNAs are reduced in abundance, while their corresponding pri-miRNAs accumulate in the dcl1, hyl1, or se mutant (Han et al., 2004b; Kurihara and Watanabe, 2004; Vazquez et al., 2004; Yang et al., 2006a). Other proteins such as DAWDLE (an RNA-binding protein) and the nuclear cap-binding complex also act in this process, probably by facilitating DCL1 to access or recognize pri-miRNAs or the loading of miRNA processing factors onto pri-miRNAs (Laubinger et al., 2008; Yu et al., 2008). Recently, plant Mediator was found to participate in miRNA biogenesis by recruiting RNA polymerase II to MIR gene promoters and then promoting their transcription (Kim et al., 2011). After the miRNA/miRNA* duplex is released from the precursor, each strand of the duplex is methylated by the small RNA methyltransferase HUA ENHANCER1 (HEN1) in the 2′-OH of the 3′ terminal nucleotide (Li et al., 2005; Yu et al., 2005; Yang et al., 2006b). The methylation is a protection from further polyuridylation and from the subsequent degradation by the exonucleases of the Small RNA-Degrading Nuclease family (Li et al., 2005; Yu et al., 2005; Ramachandran and Chen, 2008). The 3′ methylated miRNA/miRNA* duplex may be transported by a plant Exportin5 homolog HASTY (HST) or through HST-independent mechanisms to the cytoplasm (Park et al., 2005; Eamens et al., 2009), where RISC can be assembled as in animals. However, the exact form of the exported miRNA and the subcellular localization of plant RISC loading and maturation are still not clear (Voinnet, 2009). Plant RISC may also be assembled in the nucleus. In this scenario, only mature AGO1 with a single-stranded miRNA can be exported to the cytoplasm (Eamens et al., 2009). Plant AGO1 is the major part of RISC and has an endonucleolytic cleavage activity that cleaves complementary mRNAs in the center of the miRNA-mRNA paired region (Vaucheret et al., 2004; Baumberger and Baulcombe, 2005; Qi et al., 2005). PLANT MIRNA PROCESSING PROTEINS CONCENTRATE IN DISCRETE D-BODIES Recent progress in live-cell imaging proposed that nuclear chromatin is packaged into a higher order three-dimensional structure that may correlate to the regulation of the genes (Hübner and Spector, 2010; Misteli, 2010). In addition, the interchromatin region in the cell nucleus is highly heterogeneous and contains various nuclear domains or bodies, for example, nuclear speckle, paraspeckle, nucleolus, perinucleolar compartment, Cajal body (CB), cleavage body, gemini of coiled bodies, OPT (for Oct1/PTF/transcription) domain, SAM68 (for Src associated in mitosis of 68 kD) nuclear body, polymorphic interphase karyosomal association, polycomb body, promyelocytic leukemia body (Mao et al., 2011), and plant-specific nuclear bodies, such as cyclophilin, phytochrome, or abscisic acid-activated protein kinase-containing nuclear bodies (Shaw and Brown, 2004; Chen et al., 2010). These bodies are present in the nucleus at steady state and dynamically respond to basic cellular processes as well as to diverse metabolic conditions, alterations in cellular signaling, and various forms of stress (Dundr and Misteli, 2010; Mao et al., 2011). Live-cell imaging of plant miRNA processing proteins DCL1 and HYL1, which were fused to fluorescent proteins and expressed in transgenic Arabidopsis (Arabidopsis thaliana) plants under the control of their endogenous promoters, revealed that DCL1 was enriched in round nuclear bodies measuring 0.2 to 0.8 μm in diameter as well as being diffusely distributed throughout the nucleoplasm, predominantly excluded from nucleoli (Fig. 1; Fang and Spector, 2007). The number of nuclear bodies present in each nucleus ranged from zero to four, with the majority of the nuclei having one nuclear body. A population of DCL1 bodies (approximately 60%) localize in close proximity to nucleoli in projection images, but three-dimensional deconvolution analysis revealed that they are not within nucleoli. The DCL1 partner protein HYL1 displays a similar localization pattern of nuclear bodies to DCL1 bodies. Colocalization analysis revealed that DCL1 bodies and HYL1 bodies are the same structures as they colocalize (Fang and Spector, 2007; Song et al., 2007). Unlike DCL1 and HYL1, SE was distributed in nuclear speckles or interchromatin granule clusters containing the Ser/Arg (SR) splicing factor SR33 (Fang et al., 2004; Fang and Spector, 2007). In a small population of cells, the SE signal was also present in HYL1 bodies in addition to its nucleoplasmic distribution. The dual localization patterns of SE both in nuclear speckles and DCL1/HYL1 nuclear bodies may correlate with its dual roles in both splicing and miRNA processing (Fang and Spector, 2007; Laubinger et al., 2008). Figure 1. Open in new tabDownload slide Arabidopsis nuclear D-bodies. A, HYL1-YFP signals in the nucleus of a leaf epidermal cell. B, 4′,6-Diamidino-2-phenylindole staining of the nucleus in A. C, Overlay of A and B. Two D-bodies are observed in the image, one of them close to the nucleolus as shown in C. Bar = 10 μm. Figure 1. Open in new tabDownload slide Arabidopsis nuclear D-bodies. A, HYL1-YFP signals in the nucleus of a leaf epidermal cell. B, 4′,6-Diamidino-2-phenylindole staining of the nucleus in A. C, Overlay of A and B. Two D-bodies are observed in the image, one of them close to the nucleolus as shown in C. Bar = 10 μm. The DCL1/HYL1-containing bodies are different from most known nuclear bodies, due to their round shape, size, and average number per nucleus (Shaw and Brown, 2004), but are similar to CBs, as they are round in shape and their distribution is frequently perinucleolar (Nizami et al., 2010). CBs contain components involved in the processing/assembly of small nuclear RNAs, small nucleolar RNAs, and possibly siRNAs (Li et al., 2006; Pontes et al., 2006; Nizami et al., 2010). However, colocalization analysis indicated that DCL1/HYL1-containing bodies are different from CBs (Fang and Spector, 2007; Song et al., 2007), since they show no overlay with the AtCoilin signal, a signature marker of CBs. DCL1/HYL1-containing bodies are called dicing bodies or D-bodies (Fang and Spector, 2007; Fig. 1). In living cells of Arabidopsis plants, D-bodies move in the nuclei in a constrained manner. HOW ARE D-BODIES FORMED? Nuclear bodies are membraneless subnuclear organelles. A specific nuclear body is formed in a stochastic or ordered assembly manner (Dundr and Misteli, 2010). In addition, a seeding mechanism has been proposed to assemble, maintain, and regulate particular nuclear bodies (Mao et al., 2011). DCL1 contains two C-terminal dsRBDs. The mutant DCL1-6, with truncation of its two dsRBDs, is embryo lethal, while DCL1-9, a mutant with truncation of only the second dsRBD, results in infertility and severe defects in the biogenesis of most miRNAs, suggesting an important role of these dsRBDs in miRNA processing (Schauer et al., 2002). Live-cell imaging revealed that DCL1-9 failed to localize to D-bodies but instead distributed diffusely in the nucleoplasm, demonstrating that the dsRBD of DCL1 is critical for its localization to D-bodies. HYL1 contains two N-terminal dsRBDs, and these two dsRBDs are sufficient for pre-miRNA processing and localization to D-bodies (Wu et al., 2007). These results suggested that the dsRBDs in these miRNA processing proteins are essential for their targeting to D-bodies, possibly forming the seed for the assembly of D-bodies. WHAT ARE THE FUNCTIONS OF D-BODIES? In vivo tracking of a pri-miRNA using 24 tandem MS2 translational operators (MS2 repeats) and the MS2 coat protein-yellow fluorescent protein (MS2-YFP) system demonstrated that an introduced pri-RNA concentrates in DCL1-containing D-bodies in addition to being present in a diffuse distribution in the nucleoplasm, indicating that the pri-miRNAs can be recruited to D-bodies, where the machinery for their processing is enriched (Fang and Spector, 2007). The precise and efficient pri-miRNA processing requires protein-protein interactions between the miRNA processing proteins (Kurihara et al., 2006). Using bimolecular fluorescence complementation, Fang and Spector (2007) found that DCL1, HYL1, and SE interact in the nuclear D-bodies in vivo, while the bimolecular fluorescence complementation signal in the surrounding nucleoplasm is very weak. In addition, DCL1 and HYL1 self-interact in the D-bodies. By contrast, DCL1-9 showed no interaction with SE, HYL1, or DCL1. Together, these results suggested a role of D-bodies in the dicing reaction of pri-miRNAs mediated by DCL1 and its interacting partner HYL1 (Fig. 2). Figure 2. Open in new tabDownload slide Plant nuclear D-bodies, which contain proteins for miRNA processing and are involved in miRNA biogenesis. In eukaryotic cells, the nucleus is encapsulated in two layers of membranes in which nuclear pore complexes are embedded for transport between the nucleus and the cytoplasm. Chromosomes in the nucleus are organized into chromosome territories. The interchromatin region of the cell nucleus is highly heterogeneous, containing various nuclear domains or bodies. In a plant cell, these nuclear bodies include nucleolus, CB, nuclear speckles, phytochrome nuclear body, AAPK-Interacting Protein1 (AKIP1)-containing nuclear body, and D-body. These bodies have different sizes, shapes, components, dynamics, and functions. D-bodies play a role in the biogenesis of miRNAs. Plant miRNA genes are transcribed by RNA polymerase II to generate pri-miRNAs. The RNA-binding protein DAWDLE presumably stabilizes pri-miRNAs and facilitates DCL1 to access or recognize pri-miRNAs. The nuclear cap-binding complex (CBC) likely facilitates the loading of miRNA processing factors onto pri-miRNAs. The pri-miRNAs are then recruited to D-bodies, which contain DCL1, the dsRBD protein HYL1, and the C2H2 zinc finger protein se. These pri-miRNAs are then processed into a shorter stem-loop structure called pre-miRNAs and then further into the miRNA/miRNA* duplex. The miRNA/miRNA* duplex is methylated by the small RNA methyltransferase HEN1 in the 2′-OH of the 3′ terminal nucleotide. The mature miRNA/miRNA* may be transported in an HST-dependent or -independent manner through the nuclear pore complex, or the guide strand of mature miRNA/miRNA* is probably selectively loaded into AGO1-RISC in the nucleus and the miRISC is transported into the cytoplasm. The miRISC carries out the silencing reactions through translation repression or mRNA cleavage in the cytoplasm. Figure 2. Open in new tabDownload slide Plant nuclear D-bodies, which contain proteins for miRNA processing and are involved in miRNA biogenesis. In eukaryotic cells, the nucleus is encapsulated in two layers of membranes in which nuclear pore complexes are embedded for transport between the nucleus and the cytoplasm. Chromosomes in the nucleus are organized into chromosome territories. The interchromatin region of the cell nucleus is highly heterogeneous, containing various nuclear domains or bodies. In a plant cell, these nuclear bodies include nucleolus, CB, nuclear speckles, phytochrome nuclear body, AAPK-Interacting Protein1 (AKIP1)-containing nuclear body, and D-body. These bodies have different sizes, shapes, components, dynamics, and functions. D-bodies play a role in the biogenesis of miRNAs. Plant miRNA genes are transcribed by RNA polymerase II to generate pri-miRNAs. The RNA-binding protein DAWDLE presumably stabilizes pri-miRNAs and facilitates DCL1 to access or recognize pri-miRNAs. The nuclear cap-binding complex (CBC) likely facilitates the loading of miRNA processing factors onto pri-miRNAs. The pri-miRNAs are then recruited to D-bodies, which contain DCL1, the dsRBD protein HYL1, and the C2H2 zinc finger protein se. These pri-miRNAs are then processed into a shorter stem-loop structure called pre-miRNAs and then further into the miRNA/miRNA* duplex. The miRNA/miRNA* duplex is methylated by the small RNA methyltransferase HEN1 in the 2′-OH of the 3′ terminal nucleotide. The mature miRNA/miRNA* may be transported in an HST-dependent or -independent manner through the nuclear pore complex, or the guide strand of mature miRNA/miRNA* is probably selectively loaded into AGO1-RISC in the nucleus and the miRISC is transported into the cytoplasm. The miRISC carries out the silencing reactions through translation repression or mRNA cleavage in the cytoplasm. PERSPECTIVES Apart from DCL1, HYL1, and SE, which localize predominantly or transiently to D-bodies, the miRNA/miRNA* duplex, the methyltransferase HEN1, and the slicer AGO1 also exhibited some localization to D-bodies in addition to their nucleoplasmic and cytoplasmic distribution patterns when examined by colocalization analysis with HYL1 (Fang and Spector, 2007). Therefore, it is of interest to investigate if HEN1 and AGO1 are recruited to D-bodies to methylate the miRNA/miRNA* duplex and load mature miRNAs to AGO1 to assemble the RISC complex in D-bodies. In this case, only mature AGO1 and miRNA containing RISC can be exported to the cytoplasm through nuclear pore complexes (Eamens et al., 2009; Fig. 2). In addition, HYL1 was observed to colocalize with its homolog DRB4 in D-bodies (Y. Fang and D.L. Spector, unpublished data). It was known that DRB4 interacts with DCL4 in vivo and is involved in the biogenesis of siRNAs (Fukudome et al., 2011). Therefore, more extensive studies are needed to learn about the potential roles of D-bodies in orchestrating the processing, sorting, RISC assembly, and functioning of small RNAs. ACKNOWLEDGMENTS We thank Dr. David L. Spector for critical reading of the manuscript and members of Y.F.’s laboratory for insightful discussions. We apologize to all colleagues whose relevant work could not be cited due to space limitations. LITERATURE CITED Ahmad A Zhang Y Cao XF ( 2010 ) Decoding the epigenetic language of plant development . Mol Plant 3 : 719 – 728 Google Scholar Crossref Search ADS PubMed WorldCat Bartel DP ( 2004 ) MicroRNAs: genomics, biogenesis, mechanism, and function . 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Genes Dev 21 : 1190 – 1203 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by the National Natural Science Foundation of China (grant nos. 31171168 and 91019006), the National Basic Research Program of China (grant no. 2012CB910500), and the Chinese Academy of Sciences (grant no. KSCX2–YW–N–099 to Y.F.). * Corresponding author; e-mail [email protected]. www.plantphysiol.org/cgi/doi/10.1104/pp.111.186734 © 2012 American Society of Plant Biologists. All rights reserved. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Reddy, Anireddy S.N.; Day, Irene S.; Göhring, Janett; Barta, Andrea

doi: 10.1104/pp.111.186700pmid: 22045923

The generation of mature mRNAs from most genes (about 80%–90%) in autotrophic eukaryotes requires the removal of noncoding sequences (introns) and splicing of the coding regions (exons; Labadorf et al., 2010). During splicing in some precursor messenger RNAs (pre-mRNAs), the same splice sites are always used, referred to as constitutive splicing (CS), resulting in a single transcript from a gene. However, from many pre-mRNAs, multiple mature mRNAs are generated from a single gene by alternative splicing (AS), where different combinations of splice sites are used. Both CS and AS are critical to the proper expression of intron-containing genes. Recent transcriptome-wide analysis of AS using high-throughput sequencing indicates that pre-mRNAs from up to 42% of intron-containing genes in Arabidopsis (Arabidopsis thaliana; Filichkin et al., 2010) and about 48% in rice (Oryza sativa; Lu et al., 2010) are alternatively spliced, whereas about 95% of human genes are alternatively spliced (Pan et al., 2008). In addition to pre-mRNAs, some primary microRNAs (pri-miRNAs) are also subject to CS and AS (Hirsch et al., 2006; Szarzynska et al., 2009; Mica et al., 2010). AS increases the protein-coding capacity of a genome and generates functionally different proteins from the same gene (Reddy, 2007). AS results in protein isoforms with loss or gain of function and altered subcellular localization, protein stability, and/or posttranslational modifications. Furthermore, AS plays an important role in gene regulation through regulated unproductive splicing and translation, leading to RNA degradation by mRNA surveillance mechanisms, differential recruitment of mRNAs to ribosomes, or translatability of splice variants (Kurihara et al., 2009; Licatalosi and Darnell, 2010; Palusa and Reddy, 2010). Numerous spliceosomal proteins either promote or suppress splicing by interacting with splicing regulatory elements on the pre-mRNAs. In recent years, the localization and dynamics of some splicing regulators in plants have been analyzed using a variety of approaches. This review summarizes the current status of research in this area, with emphasis on RNA-binding proteins (RBPs) that are involved in splicing regulation, discusses open questions, and presents some approaches to address these questions. SPLICING REGULATORS pre-mRNAs splicing takes place cotranscriptionally in the spliceosome, a large multicomponent complex composed of small nuclear RNAs (snRNAs) and about 170 proteins, many of which are involved in the regulation of splicing (Wahl et al., 2009; Valadkhan and Jaladat, 2010). The composition of the human and yeast spliceosome has been analyzed in great detail (Wahl et al., 2009); however, information on plant splicing has been scarce, as no in vitro assembly of a functional plant spliceosome has been possible to date. A detailed search for Arabidopsis orthologs of the human spliceosome proteome revealed that most splicing factors are present in the Arabidopsis genome, indicating a similar complexity (Barta et al., 2011). Interestingly, many of the plant orthologs have close or related homologs evolving from several genome duplication events with possible plant-specific functions. This is best exemplified by the Ser/Arg-rich (SR) protein subfamilies (see below). In Arabidopsis, there are over 200 RBPs, including many unique to plants (Lorković, 2009), suggesting plant-specific functions. Some of them are components of small nuclear ribonucleoprotein particles (snRNPs), while many others are not but are present in the spliceosome and hence are called non-snRNP proteins. A survey of RBPs including RNA recognition motif (RRM)-containing proteins, such as SR proteins, SR-like proteins, heterogeneous ribonucleoprotein particles (hnRNPs), and RNA-binding KH domain-containing proteins, revealed that many of them play key roles in regulating CS and AS. Some of these RBPs are relatively well studied in plants (Lorković, 2009; Reddy and Ali, 2011). SnRNP PROTEINS There are two types of spliceosomes in higher eukaryotes, the major (U2-type) and minor (U12-type) spliceosomes. Plants, like other eukaryotes, contain U1, U2, U4, U5, and U6 snRNAs in the major spliceosome, while the minor spliceosome contains U11, U12, U4atac, U5, and U6atac (Lorkovic et al., 2005; Ru et al., 2008). These snRNAs form complexes with proteins, including some proteins specific to each U-RNA, to form snRNPs. In animals, where the snRNP complexes have been extensively studied, all the major snRNPs except U6 snRNP have a core of seven Sm proteins and additional UsnRNP-specific proteins (Valadkhan and Jaladat, 2010). The U6snRNP has a different set of seven proteins termed LSm plus only one U6snRNP-specific protein. The minor spliceosome snRNPs also have Sm proteins and snRNP-specific proteins but are not as well studied as the major spliceosome proteins (Ru et al., 2008). Only one Sm protein (SAD1/LSm5) that is necessary for abscisic acid sensitivity and drought tolerance has been characterized in Arabidopsis (Xiong et al., 2001), but analysis of the Arabidopsis predicted proteome revealed that all the major snRNP proteins are conserved in Arabidopsis (Wang and Brendel, 2004). The counterparts of three U1snRNP-specific proteins (U1-A, U1-C, and U1-70K) identified in animals and yeast have been characterized in plants. U1-A in Arabidopsis has been shown to bind U1snRNA (Simpson et al., 1995), while Arabidopsis U1-70K has an RNA-binding domain and an Arg/Ser-rich (RS) domain responsible for interaction with SR proteins (see below; Golovkin and Reddy, 1996, 1998). An Arabidopsis U11 snRNP-specific 35K protein identified as a binding partner of an SR protein has been shown to incorporate into monomeric U11 snRNPs and U11/U12 di-snRNP complexes. These interactions, together with its sequence similarity to U1 70K in the U1snRNP, suggest that the U11-35K protein functions in the splicing of minor AT:AC introns (Lorkovic et al., 2005). In addition, other genes coding for minor snRNP proteins have been identified in both monocot and dicot genomes, indicating a high evolutionary conservation of minor spliceosome components and splicing function (Lorkovic et al., 2005). NON-SnRNP PROTEINS The non-snRNP proteins include splicing factors involved in splice site selection, U-snRNP-associated proteins, SR proteins, hnRNPs, and other splicing regulators. U2AF, composed of two subunits, U2AF65 and U2AF35, is a non-snRNP protein involved in 3′ splice site selection. Both subunits, each with two isoforms, have been identified in plants (Wang and Brendel, 2006). U2AF35 has been shown to interact with SR-like proteins SR45 and SR45a, which also interact with U1-70K, suggesting a role for these proteins in bridging of the 5′ splice site and 3′ splice site selection complexes (Tanabe et al., 2009; Reddy et al., 2011). Two subunits of the Cap Binding Complex (CBC20 and CBC80) and SERRATE, a zinc finger protein, regulate pre-mRNA splicing and pri-miRNA processing (Laubinger et al., 2008; Raczynska et al., 2010). MOS4, a protein involved in plant immunity, is thought to be a component of the spliceosome (Palma et al., 2007). An interesting group of splicing-related proteins is the RS domain-containing cyclophilins, termed atCypRS64 and atCypRS92, which were found in a yeast two-hybrid screen with Arabidopsis SR proteins. These proteins contain a prolyl-peptidyl isomerase domain important for protein structure and modification and have been shown to interact with SR proteins in vitro and in vivo, suggesting that they might regulate spliceosome assembly (Lorković et al., 2004). Another multidomain cyclophilin, atCyp59 (prolyl-peptidyl isomerase, RRM zinc knuckle, and RS/RD domain), has been found to interact with SR proteins as well as with the C-terminal domain of RNA polymerase II and was suggested to connect splicing and transcription (Gullerova et al., 2006). SR PROTEINS SR proteins have been redefined recently in plants (Barta et al., 2010) based on sequence features, including the number and location of RRMs and the RS domain. In plants, proteins with one or two N-terminal RRMs followed by a downstream RS domain of at least 50 amino acids and a minimum of 20% RS or SR dipeptides are considered as SR proteins (Barta et al., 2010). The RRM domain recognizes and binds to regulatory elements in pre-mRNAs, and the RS domain is involved in interactions with other proteins. SR proteins are important splicing regulators in both animals and plants. In animals, SRs are known to perform many other diverse functions in the nucleus as well as in the cytoplasm, including mRNA transport, localization, translation, and decay, as well as in genome stability and microRNA biogenesis (Twyffels et al., 2011). Interestingly, some SRs in animals are present at the cell surface and exhibit carbohydrate-binding activity (Hatakeyama et al., 2009). The SR proteins are major regulators of CS and AS (Barta et al., 2008; Long and Caceres, 2009; Reddy and Ali, 2011). Plants have more SR proteins (18 in Arabidopsis and 22 in rice) as compared with mammals (12 in humans; Barta et al., 2010). Plant SR proteins are grouped into six subfamilies: three subfamilies are orthologous to animal SRs, whereas 10 AtSRs in three subfamilies are plant specific, suggesting some differences in splicing regulation between plants and animals (Barta et al., 2010). The SR subfamily, which has two RRMs (the second RRM having a conserved SWQDLKD motif) followed by the RS domain, is orthologous to the mammalian SRSF1/SF2/ASF family. The RSZ subfamily members, with one RRM and one RS domain separated by a zinc (Zn) knuckle, are orthologous to the mammalian SRSF7/9G8 subfamily. The SC subfamily has one RRM and one RS domain and is orthologous to the SRSF2/SC35 subfamily. The SCL subfamily, while similar to the SRSF2 subfamily, has an RRM with an N-terminal charged extension. The RS2Z subfamily members have two Zn knuckles and an additional SP-rich region following the RS domain. Finally, the RS subfamily has two RRMs, but without the conserved SWQDLKD motif, followed by the RS domain. Based on the new definition, two plant proteins (SR45 and SR45a) previously classified as SR proteins are no longer included in the SR family. SR45 has two RS domains, one preceding and one following the RRM domain, with sequence similarity to an exon junction complex protein, RNPS1. SR45a is not related to SR45 at the sequence level, as the name implies, and is a homolog of metazoan tra-2. It is interesting that 14 of the 18 Arabidopsis SR proteins are alternatively spliced, as are SR45 and SR45a (Palusa et al., 2007; Tanabe et al., 2009). OTHER PLANT RBPS INVOLVED IN SPLICING An hnRNP-like protein that binds to U-rich or AU-rich sequences, called UBP1, with three RRMs enhances the splicing of suboptimal introns and enhances the steady-state levels of mRNA by preventing mRNA degradation (Lambermon et al., 2000). Two plant-specific proteins (UBA1 and UBA2) that interact with UBP1 also contain an RRM and increase mRNA levels but do not enhance splicing (Lambermon et al., 2002). RRM-containing RBPs with a C-terminal Gly-rich region (GRPs) are also involved in different aspects of RNA metabolism, including mRNA export and AS. In Arabidopsis, there are eight GRPs, and some of them (GRP7 and GRP8) are involved in regulating circadian rhythm by autoregulation and cross-regulation of AS of their pre-mRNAs coupled to nonsense-mediated decay (Schöning et al., 2008). Similarly, three polypyrimidine tract-binding proteins that belong to the hnRNP family are involved in autoregulation and cross-regulation of AS of their pre-mRNAs coupled to nonsense-mediated decay (Stauffer et al., 2010). EXPERIMENTAL APPROACHES USED TO STUDY THE LOCALIZATION AND DYNAMICS OF SPLICING REGULATORS Early approaches to localize splicing regulators used fixed cells or tissues with antibodies against mammalian splicing-related proteins using indirect immunofluorescence microscopy and electron microscopy (Ali and Reddy, 2008b), which provided static images of a protein at a particular point in time. The discovery of fluorescent proteins (FPs) has opened many new innovative methods to detect the localization and movement of individual proteins and RNA molecules and to monitor protein-protein, protein-RNA, and RNA-RNA interactions in living cells (Chudakov et al., 2010; Urbinati and Long, 2011). These methods are providing an unprecedented view of protein/RNA localization and their movement spatially and temporally (Larson et al., 2011; Urbinati and Long, 2011). Translational fusions of the protein of interest with GFP, its derivatives red (RFP), yellow (YFP), and cyan (CFP) FPs, or other FPs with different excitation and emission spectra are well suited for investigating the temporal and spatial distribution of proteins and have permitted the visualization of multiple proteins simultaneously (for review, see Chudakov et al., 2010). Fluorescence recovery after photobleaching (FRAP) is an imaging method where fluorophores from a FP-tagged protein are inactivated through bleaching of a defined area with intense laser pulses. Since bleaching permanently destroys fluorescence, the recovery of fluorescence to the bleached area depends on the mobility of the labeled protein from the unbleached area and is monitored over time for estimation of the dynamics of proteins (i.e. diffusion parameters or trafficking). FRAP provides information about how much of the total available protein is free to diffuse in live cells versus how much is restricted due to its association with complexes or partition into compartments or microdomains. Fluorescence loss in photobleaching (FLIP), a technique complementary to FRAP, also permits the analysis of the intracellular mobility of proteins. In FLIP analysis, an area of fluorescence is repeatedly bleached, and the progressive loss of fluorescence intensity in the unbleached region indicates the mobility of the labeled protein, providing the opportunity to monitor the direction of movement (for review, see Wang et al., 2008). Wild-type GFP exists in two forms, which lead to a minor and a major absorbance peak (395 and 475 nm, respectively). By activating the protein with intense illumination of 390 to 415 nm, the absorbance of the minor peak will increase dramatically and can be monitored. Photoactivatable GFP has only a negligible minor excitation peak at 475 nm; consequently, upon activation, this peak has a considerably greater proportional increase when compared with wild-type GFP, leading to a higher contrast (Patterson and Lippincott-Schwartz, 2002). This feature can be used for following activated proteins over time using time-lapse imaging and provides a simple and more direct alternative to standard molecular tracking procedures like FRAP and FLIP. A reversible photoswitchable fluorescent reporter protein, which allows repeated switching between a fluorescent and a nonfluorescent state, has been used to monitor intracellular protein trafficking. For instance, a variant of the photoswitchable fluorescent protein DRONPA, designed for use in transgenic Arabidopsis plants, was fused to the Arabidopsis RBP AtGRP7 under the control of the endogenous AtGRP7 promoter (Lummer et al., 2011). Fluorescence microscopy showed that AtGRP7 is a nucleocytoplasmic shuttling protein. To study the in vivo interaction/association of proteins, two methods involving a two-component system are feasible: bimolecular fluorescence complementation (BiFC)- and fluorescence resonance energy transfer (FRET)-based reporters. In BiFC, two halves of an FP are separately fused to two putative interacting proteins (Kerppola, 2009). When the two proteins interact, the two halves of the FP come close enough for fluorescence to be established. Because of the irreversible formation of the complex, weak or transient interactions can also be monitored. Reconstitution of fluorescence is also possible if the two proteins are in close proximity in a complex and do not interact directly. Interestingly, this method can also be adapted to monitor protein-RNA interactions (Rackham and Brown, 2004). FRET can be used to measure the proximity of two fluorophores, usually coupled to the proteins of interest. The fluorophore of one protein has an emission corresponding to the excitation of the other. Within 10 nm, a transfer of energy from the excited fluorophore (the donor) to the other fluorophore (the acceptor) can occur. The acceptor molecule enters an excited state from which it emits a longer wavelength light. By monitoring the donor and acceptor emissions, the efficiency of the resonance energy transfer and consequently the relative distance between the molecules can be calculated, with even higher accuracy than the optical resolution of a standard light microscope (for review, see Wang et al., 2008). RNA-protein interactions were also visualized in live cells using BiFC and FRET (Urbinati and Long, 2011). Fluorescence correlation spectroscopy monitors fluctuations of the fluorescent signal in a noninvasive fashion. It relies on changes in the specific signal provided by fluorescent particles to analyze their motions and interactions. In fluorescence correlation spectroscopy, a laser beam is focused on an area of fluorescence. Fluctuations in fluorescent signals are then recorded over time to determine the diffusion coefficients and binding constants of the protein (Krichevsky and Bonnet, 2002; Malchus and Weiss, 2010). During the last decade, FPs tagged to splicing regulatory proteins have been widely used in plants to monitor their location and dynamics to address issues about the spatiotemporal dynamics of splicing, which is discussed below. LOCALIZATION OF SPLICING REGULATORS In plant cells, only a limited number of splicing regulators have been investigated regarding their cellular localization, and most of them belong to the SR proteins, snRNP proteins, or RS-containing cyclophilins (Lorković and Barta, 2004; Ali and Reddy, 2008b). The nuclear space is compartmentalized by nuclear bodies, which are dynamic but relatively stable structures composed of proteins and RNAs without delineating membranes. Their morphology is dependent on cell differentiation and metabolic status. In animals, these include structures like the nucleoli, nuclear speckles, Cajal bodies, histone locus bodies, paraspeckles, and others (Misteli and Spector, 2011). Splicing regulators are detected in nucleoplasm, nuclear speckles, Cajal bodies, and sometimes the nucleolus. In plants, the best-investigated nuclear bodies are the nucleolus, photobodies, which contain phytochrome and proteins involved in photomorphogenesis, and nuclear speckles. As the nucleolus and photobodies are described in other Update articles in this Focus issue, we will focus on the occurrence and the dynamic aspects of nuclear speckles. Nuclear speckles are located in the interchromatin space and were detected as a storage place for splicing factors (Fang et al., 2004; Spector and Lamond, 2011). In animal tissues, nuclear speckles have also been termed SC35 bodies, as many SR proteins accumulate in these nuclear bodies. However, nuclear speckles also contain snRNPs, non-snRNP splicing proteins, transcription factors, and 3′ end processing factors. Speckles are often observed near active transcription sites, and pre-mRNAs were detected outside the speckles in fibrillar structures (for review, see Spector and Lamond, 2011). Hence, nuclear speckles are viewed as storage and assembly areas that procure splicing factors to active transcription sites. In line with this hypothesis, actively transcribing cells have smaller and more speckles, whereas inhibition of transcription or splicing leads to an accumulation of splicing factors and larger speckles. Like speckles, paraspeckles are subnuclear bodies found in interchromatin spaces and contain long non-protein-coding RNAs and novel proteins (Spector and Lamond, 2011); however, they have not been described yet in plant cells. The complex compartmentalization of plant nuclei is now an active field of research. Genomic and proteomic approaches are being used to investigate the nucleolus and the upcoming new nuclear dicing bodies. In plants, the RNA processing machinery was first described in experiments using antibodies to U2snRNP-specific proteins, which were detected diffusely in the nucleoplasm and in Cajal bodies (Beven et al., 1995). SRs representing each group of SR proteins in Arabidopsis have been shown to display a characteristic distribution pattern of high concentration in speckles and diffuse distribution in the nucleoplasm (Fang et al., 2004; Lorković and Barta, 2004; Lorković et al., 2004, 2008; Tillemans et al., 2005, 2006; Ali and Reddy, 2006). The speckle localization of proteins involved in pre-mRNA splicing is so characteristic that it is used for a diagnostic tool for splicing proteins (Spector, 2001; Lorković and Barta, 2004). Recently, a protein similar to transportin-SR (also called MOS14 in Arabidopsis) that is required for plant immunity was shown to be necessary for transporting SR proteins into the nucleus (Xu et al., 2011). Studies using GFP-labeled RRM and RS domains of plant proteins suggest that RS domains of the plant SR proteins seem to contain the nuclear localization and speckle targeting/retention signals, as the GFP-RRM fusions of SR45, AtRS31, and AtRSZ22 were localized to the nucleoplasm or cytoplasm (Tillemans et al., 2005; Ali and Reddy, 2006). Using fluorescently labeled Arabidopsis SR proteins, several groups have demonstrated that plant nuclei have nuclear speckles whose size and shape are dependent on cell type, metabolic state, and transcriptional activity (Fang et al., 2004; Lorković and Barta, 2004; Lorković et al., 2004, 2008; Tillemans et al., 2005, 2006; Ali and Reddy, 2006). An interesting phenomenon was observed when colocalization studies of Arabidopsis SR proteins were performed in plant protoplasts (Lorković et al., 2008). While representatives of all six subfamilies of plant SRs localize to speckles, it could be demonstrated that several of the SR proteins do not or only partly colocalize to the same nuclear area. Interestingly, the two families containing a Zn knuckle motif did not colocalize with AtSR34 (an SF2/ASF ortholog). Figure 1 shows an example of RS2Z33-GFP cotransfected with SR34-RFP, where the projection image almost does not have overlapping speckles (Lorković et al., 2008). Furthermore, AtSR34 only partly colocalized with AtSC35, although both proteins are used as genuine markers for nuclear speckles. In some cases, colocalization results were found to differ compared with other studies using different plants and conditions (Tillemans et al., 2005), suggesting that colocalization of SR proteins may depend on cell type and particular conditions. In addition, colocalization did not always correlate with in vitro protein-protein interaction studies (Lorković et al., 2008); therefore, these data are not straightforward to interpret. Other plant spliceosomal proteins (U1-70K, U2AF35a, U2AF35b) are also localized to speckles in the nucleus (Wang and Brendel, 2006; Ali et al., 2008). Speckle components have been shown to shuttle continuously between speckles and nucleoplasm (Ali and Reddy, 2008b; Rausin et al., 2010). Their biogenesis is proposed to occur through self-assembly processes whereby a transient binding interaction to macromolecules will determine the size and shape of the speckles (Dundr and Misteli, 2010). Recently, in animals, large scaffolding proteins have been implicated in the organization of nuclear speckles (Sharma et al., 2010). The colocalization data might indicate that binding interactions between some of the SR proteins are less strong than their self-assembly ability, resulting in different speckled areas in the nucleus. These data also indicate that nuclear speckles might exhibit different splicing factor compositions dependent on the different states of the cell and the needs for transcription/splicing of particular genes. Figure 1. Open in new tabDownload slide Plant SR proteins do not always colocalize in the same speckles. Maximum intensity projection images show RS2Z33-GFP and SR34-RFP transiently coexpressed in tobacco protoplasts. The merged image shows no or very little overlap in the speckled areas (Lorković et al., 2008). Figure 1. Open in new tabDownload slide Plant SR proteins do not always colocalize in the same speckles. Maximum intensity projection images show RS2Z33-GFP and SR34-RFP transiently coexpressed in tobacco protoplasts. The merged image shows no or very little overlap in the speckled areas (Lorković et al., 2008). DYNAMICS OF SPLICING REGULATORS Speckles and their components are dynamic structures. They change in size, in shape, and in number. These properties of a given nuclear body can be different in different cell types or at different times in the same cell type. At the same time, they are structurally stable, individual nuclear bodies persist during the entire interphase (entry to G1 to exit from G2), but there is an exchange of a large number of the major nuclear body components with the surrounding nucleoplasm (Dundr and Misteli, 2001). The morphological appearance of a nuclear body is determined by the components in the body and their interactions. Individual speckle components are able to shuttle in and out of the speckles. It was proposed that the size, and possibly the shape, of nuclear bodies is likely determined by the balance of the on rate relative to the off rate of its components, with an increase in on rate or a drop in off rate leading to an increase in size and a decrease in on rate or an increase in off rate leading to shrinkage (Dundr and Misteli, 2001). The intranuclear mobility of RSZ22-GFP was investigated using FRAP and FLIP (Tillemans et al., 2006). These experiments showed that this SR splicing factor shuttles rapidly between subnuclear compartments, including the nucleolus. A sensitive shuttling assay using FLIP and nuclear export inhibitors with cells transiently expressing FP fusions strongly suggested that RSZ22 is a nucleocytoplasmic shuttling SR splicing factor (Tillemans et al., 2006). The dynamics of RSZ22 was recently studied in stable transformants enabling tissue-specific expression of the transgene (Rausin et al., 2010). The results demonstrated that RSZ22 is a bona fide nucleocytoplasmic shuttling SR protein. RSZ22 mutants with mutated RNP1 or Zn knuckle domains were able to shuttle and localize to speckles, suggesting that neither RNP1 nor Zn knuckle motifs are key to nuclear localization and speckle-like organization. However, FRET studies established that molecular interactions between splicing factors were strongly destabilized, although not entirely inhibited, with Zn knuckle and RNP1 mutants, suggesting that both RNA-binding domains might be needed either for direct protein-protein interactions or for mRNA-mediated interactions independent of splicing factor preassembly. Studies using GFP-SR45 showed its localization to speckles and nucleoplasm, and the speckles exhibited intranuclear movements and changes in morphology (Ali et al., 2003; Zhang and Mount, 2009). Later, using FRAP, the effective diffusion coefficient (Deff), a measure of the apparent mobility of a protein, for GFP-SR45 (Fig. 2) was determined to be 1.01 μm2 s−1 in the nucleoplasm and 0.38 μm2 s−1 in the speckles (Ali and Reddy, 2006). Using GFP-labeled U1-70K, FRAP analyses were performed on speckles and the nucleoplasmic regions of the nucleus (Ali and Reddy, 2006). It was shown that U1-70K moves in the nucleus very rapidly, on a millisecond scale. The expected Deff value of uncomplexed GFP/U1-70K based on its size is approximately 12 (in nucleoplasm) to 31 (in speckles) times greater than the experimental values, suggesting that U1-70K interacts with other nuclear components or is in a larger snRNP complex (Ali and Reddy, 2006). Figure 2. Open in new tabDownload slide Mobility analysis of SR45 and SRp34 in control and ATP-depleted cells using FRAP. A speckle or a defined nucleoplasmic area in control or ATP-depleted cells expressing either GFP-SR45 (A) or SRp34-YFP (B) was bleached, and the recovery of fluorescence was quantified for 80 s. The mobility of these splicing factors was different in nucleoplasm as compared with speckles, and the mobility required ATP, as ATP depletion reduced the movement in both nucleoplasm and speckles (Ali and Reddy, 2006). Deff values (in μm2 s−1) are as follows: for GFP-SR45, in nucleoplasm, 1.01, −ATP 0.06; in speckle, 0.38, −ATP 0.05; for SR1/SRp34-YFP, in nucleoplasm, 1.34; −ATP 0.07; in speckle, 0.88, −ATP 0.04. Figure 2. Open in new tabDownload slide Mobility analysis of SR45 and SRp34 in control and ATP-depleted cells using FRAP. A speckle or a defined nucleoplasmic area in control or ATP-depleted cells expressing either GFP-SR45 (A) or SRp34-YFP (B) was bleached, and the recovery of fluorescence was quantified for 80 s. The mobility of these splicing factors was different in nucleoplasm as compared with speckles, and the mobility required ATP, as ATP depletion reduced the movement in both nucleoplasm and speckles (Ali and Reddy, 2006). Deff values (in μm2 s−1) are as follows: for GFP-SR45, in nucleoplasm, 1.01, −ATP 0.06; in speckle, 0.38, −ATP 0.05; for SR1/SRp34-YFP, in nucleoplasm, 1.34; −ATP 0.07; in speckle, 0.88, −ATP 0.04. Time-lapse microscopy studies of the mobility of speckles in different plant cells and under different physiological conditions showed that SR speckles in plant nuclei display limited movements in a constrained area (Ali and Reddy, 2008b, and refs. therein). The speckles are possibly loosely anchored to less mobile nuclear components or restrained from mobility by physical barriers such as chromatin. Besides these restricted movements, plant SR speckles also fuse with each other and bud off from speckles (Ali et al., 2003; Ali and Reddy, 2008b). Studies with animal cells have shown that the distribution pattern of SR proteins and other splicing factors changes with the cell cycle (for review, see Dundr and Misteli, 2010; Spector and Lamond, 2011). Changes in the distribution pattern of SR proteins during the cell cycle were also observed in plant cells (Fang et al., 2004). SR proteins became more diffuse, so that during metaphase they were distributed throughout the cytoplasm, with speckles reforming in late telophase. REGULATION OF THE LOCALIZATION AND DYNAMICS OF SPLICING REGULATORS The localization and dynamics of these splicing regulators are regulated by phosphorylation status, transcriptional activity of the cell, stages of the cell life cycle, such as M phase versus interphase, cell type, developmental stage, stresses imposed on plants, and hormone signals. A study of SR-FP fusion proteins found that meristematic cells or otherwise rapidly growing cells had the highest number of small speckles, with most of the FP fusion proteins found in a diffuse nucleoplasmic pool, while highly differentiated cells (e.g. leaf mesophyll or epidermal cells) showed larger speckles with fewer FP fusion proteins in the nucleoplasm (Lorković et al., 2008). Phosphorylation and Dephosphorylation SR proteins are phosphoproteins, with the RS domains being extensively phosphorylated in vivo (Reddy, 2007; Barta et al., 2008). In plants Clk/LAMMER type, SRPKs, and mitogen-activated protein kinases phosphorylate one or more SR proteins. Phosphorylation is thought to influence subcellular localization, protein-protein interactions, RNA-protein interactions, and splicing activities. Inside speckles, many SR proteins exist in a hypophosphorylated state; when phosphorylated by speckle-resident SR-protein kinases, the hyperphosphorylated SR proteins dissociate from speckles and are competent to participate in the splicing reaction (Dundr and Misteli, 2010). The SR proteins are dephosphorylated in the splicing process, and their affinity for speckles is reestablished. Early studies have shown that inhibition of phosphorylation in plant cells sequestered SR and SR-like proteins to large speckles and other altered localization patterns (Fig. 3; Ali et al., 2003; Tillemans et al., 2005). Not only is the localization pattern different but also the mobility is altered. FRAP and FLIP analyses with AtSR45 and AtSR34 demonstrated that their mobility is reduced by the inhibition of protein phosphorylation (Ali and Reddy, 2006). The use of staurosporine to inhibit protein kinases resulted in a significant decrease in the mobility of AtSR45 and AtSR34. The Deff values of AtSR45 were reduced four times, whereas the Deff values of AtSR34 were reduced 14 times. Their mobile fractions were also significantly reduced. The enlargement of speckles in the presence of staurosporine suggests that the release of SR proteins from speckles requires their phosphorylation (Ali et al., 2003; Tillemans et al., 2005). Studies by Tillemans et al. (2006) suggested that the motility of AtRSZ22 is mainly phosphorylation dependent and that the phosphorylation/dephosphorylation cycle of the RS domain may influence the subnuclear distribution and dynamics of RSZp22 into or out of the nucleolus. AtRSZ22-GFP was shown to concentrate in the nucleolus upon phosphorylation inhibition, while other tested AtSRs did not (Tillemans et al., 2006). Later studies in root and pollen cells of stably transformed plants confirmed the accumulation of AtRSZ22-GFP within nucleoli upon phosphorylation inhibition (Rausin et al., 2010). In transiently expressing root cells, AtRS31-GFP, unlike other tested SRs, accumulated into concave cap-like structures surrounding regions devoid of fluorescence corresponding to the nucleoli following treatment with staurosporine. This occurred in all cell types of the primary root, from meristematic cells to trichoblasts, and there was a similar redistribution in transiently expressing Arabidopsis leaf cells. The perinucleolar localization of AtRS31 proteins was shown to be reversible, because the AtRS31-GFP relocalized into nucleoplasmic speckles upon the return to physiological conditions. A cyclin-dependent kinase, CDKC2, colocalizes with AtSR34 and other spliceosomal components, and this association is dependent upon the CDKC2 kinase activity and transcriptional status of the cells (Kitsios et al., 2008), suggesting a role for this kinase in the dynamics of spliceosomal proteins. Overexpression of a LAMMER kinase in Arabidopsis, which colocalizes with AtSRp34, altered the splicing pattern of several genes and the plants showed developmental abnormalities, suggesting a role of phosphorylation in modulating AS (Savaldi-Goldstein et al., 2003). Figure 3. Open in new tabDownload slide Redistribution of an SR-related splicing regulator (SR45) in the nucleus in response to temperature stress and inhibition of transcription or phosphorylation. Arabidopsis plants expressing GFP-SR45 show its localization to characteristic speckles and nucleoplasm. Heat treatment (42°C for 24 h) resulted in the accumulation of SR45 into large irregularly shaped speckles, whereas in the cold (4°C for 12 h), speckles disappeared. Inhibition of transcription (actinomycin D for 1 h) resulted in the accumulation of SR45 into a few very large round speckles. Inhibition of protein phosphorylation (staurosporine for 1 h) also resulted in the appearance of large irregularly shaped speckles similar to heat (Ali et al., 2003). Figure 3. Open in new tabDownload slide Redistribution of an SR-related splicing regulator (SR45) in the nucleus in response to temperature stress and inhibition of transcription or phosphorylation. Arabidopsis plants expressing GFP-SR45 show its localization to characteristic speckles and nucleoplasm. Heat treatment (42°C for 24 h) resulted in the accumulation of SR45 into large irregularly shaped speckles, whereas in the cold (4°C for 12 h), speckles disappeared. Inhibition of transcription (actinomycin D for 1 h) resulted in the accumulation of SR45 into a few very large round speckles. Inhibition of protein phosphorylation (staurosporine for 1 h) also resulted in the appearance of large irregularly shaped speckles similar to heat (Ali et al., 2003). ATP FRAP and FLIP analyses with AtSR45 and AtSR34 in ATP-depleted cells showed a dramatic reduction in the mobile fraction of both proteins in the nucleoplasm as well as in speckles (Fig. 2), and this reduction in mobility was restored by ATP (Ali and Reddy, 2006), suggesting that the association/dissociation of these SRs with other SRs/spliceosomal proteins is dependent on ATP. The retarded mobility in ATP-depleted cells is not likely due to global nuclear rearrangement, as ATP depletion had no substantial effect on the mobility of GFP alone and of a nucleus-localized NLS-GFP-GUS fusion that is twice the size of SR45. In further support that ATP depletion does not result from nonspecific general perturbation of the cellular environment, FRAP analysis with the deletion mutants of SR45 exhibited different localization and kinetic properties in response to ATP depletion (Ali and Reddy, 2006). In other studies, ATP depletion resulted in an accumulation of AtRSZ22-GFP within nucleoli, as had been demonstrated in phosphorylation inhibition, suggesting that the absence of RSZp22 mobility upon ATP depletion is linked to its phosphorylation state (Tillemans et al., 2006; Rausin et al., 2010). In animals, ATP depletion did not change the mobility of SR proteins (Phair and Misteli, 2000). Transcription Several studies have shown that transcription sites are often found at the periphery of speckles and that the disassembly of speckles affects the coordination between transcription and pre-mRNA splicing (for review, see Lorković et al., 2008; Spector and Lamond, 2011). Generally, inhibition of transcription causes a decrease in the number of nuclear speckles and the redistribution of splicing factors into larger storage bodies (Ali et al., 2003; Tillemans et al., 2005; Kitsios et al., 2008). After release of the inhibition of transcription, nuclear speckles can form de novo, speckles expand from a condensed state and take on their typical irregular shape, and new speckles form (for review, see Dundr and Misteli, 2010). Inhibition of transcription sequestered AtSR45 to large speckles and other altered localization patterns (Fig. 3; Ali et al., 2003; Tillemans et al., 2005). Studies of AtSR34, AtSR30, and AtSCL33 also showed increased accumulation of splicing factors in speckles, eventually leading to increased speckle area and the cessation of all kinds of speckle movements, including budding off and peripheral movements (Ali and Reddy, 2008b, and refs. therein). The mobility of the SR and SR-like proteins is also affected by transcription. FRAP analysis revealed that the inhibition of transcription by actinomycin D resulted in a significant decrease in the mobility of AtSR45 and AtSR34. The mobile fraction of AtSR45 was also significantly reduced by actinomycin D treatment but not that of AtSR34. Treatment of plants expressing AtRS31 with okadaic acid and α-amanitin did not result in AtRS31 accumulation around the nucleolus, as with staurosporine treatment, but instead resulted in accumulation into static large speckles (Tillemans et al., 2006). Stresses Stresses have been shown to affect the splicing of pre-mRNAs from genes involved in stress responses, including the splicing of many SR genes (Iida et al., 2004; Reddy, 2007; Ali and Reddy, 2008a). The AS patterns of SR proteins have been shown to be affected by heat, cold, salt, drought, and osmotic stress (Palusa et al., 2007; Ali and Reddy, 2008a). Heat has also been shown to affect the distribution of AtSR45. Heat treatment redistributed AtSR45 into enlarged, irregularly shaped compartments, and cold relocalized AtSR45 mostly to the nucleoplasmic pool (Fig. 3; Ali et al., 2003). Studies have shown that heat shock leads to an accumulation of various RBPs and several SR proteins to form de novo a morphologically distinct nuclear stress body (for review, see Dundr and Misteli, 2010). Interestingly, the enlargement of speckles due to heat shock was inhibited by a phosphatase inhibitor, implying that heat shock-induced relocalization involves the dephosphorylation of SR proteins (Ali et al., 2003). As heat and cold also changed the AS pattern of pre-mRNAs of several SR genes, it is possible that regulation of the AS pattern of these genes is related to a change in the subnuclear reorganization of SR splicing factors (Ali and Reddy, 2008b). As with SR45 speckles, the size of nuclear bodies containing one of the protein kinases (CDKC2) implicated in regulating the dynamics of spliceosomal proteins is increased upon heat treatment, whereas cold treatment results in the disappearance of CDKC2-containing speckles (Kitsios et al., 2008). mRNA LOCALIZATION Splicing regulators often bind to their target RNA directly or are part of splicing complexes. Following the life cycle of an individual RNA molecule from transcription to processing to localization on both the RNA and protein levels would give insights into the actual parameters of RNA splicing and its regulation. Furthermore, it is of particular importance to determine the fate of alternatively spliced transcripts, as AS is a major regulator of gene regulation in plants (Reddy, 2007, Barta et al., 2011). The distribution and spatiotemporal dynamics of mRNA molecules have been an active field of research in the past few years, including experiments performed using neuronal cells, fibroblasts, Drosophila melanogaster oocytes, and Saccharomyces cerevisiae cells (Park et al., 2010). However, data on compartmentalized RNA localization in planta are scarce. Using in situ hybridization, the distribution and, hence, the intercellular transport of transcripts through plasmodesmata has been investigated in plant cells (Lucas et al., 1995; Sambade et al., 2008; Hyun et al., 2011). However, this technique gives a stagnant picture of the subcellular mRNA distribution. Recent technical advances allow the exact localization and also the tracking of RNA molecules in real time in their natural environment, the living cell (Christensen et al., 2010). An indirect approach for monitoring RNA targets in living cells is the tracing of a fluorescently labeled RBP. The disadvantage of this procedure is that it is not certain whether the correct RNA species is monitored, as the fluorescent protein might bind other RNAs as well. Other approaches alter the RNA target itself by introducing binding motifs for fluorescently labeled RBPs (e.g. MS2-FP system, or λN22) or by inserting dye-binding aptamers into the sequence (for discussion, see Christensen et al., 2010). For localization and tracing purposes, these approaches are not recommended, since the dynamics of the RNA molecules could be changed due to the fluorescent foreign proteins hindering the trafficking or due to alterations in the naturally occurring secondary and tertiary structures of the RNA molecule. In addition, in this case, the injection of directly labeled RNA transcripts is problematic, because these molecules would not proceed via their natural processing steps (e.g. cotranscriptional splicing, polyadenylation, capping, export, etc.) and hence localization and complex formation might be different. Nonetheless, the aforementioned in vivo techniques for monitoring RNA targets are powerful tools that have been used successfully for specialized purposes (for review, see Christensen et al., 2010). Therefore, to investigate RNA in its natural context, it is important to image endogenous, and therefore not altered, RNA transcripts. To accomplish this, hybridization probes in various forms have been used. The most promising ones are termed molecular beacons, which carry a fluorophore and a quencher group at the 3′ and 5′ end, respectively (Tyagi and Kramer, 1996). These probes have a special design allowing them to assume a secondary structure, which blocks fluorescence emission in the closed conformation. Upon hybridization to the target, the probes open and the fluorescence intensity rises up to 100 times above the background level. Since plant cells are known for high autofluorescence originating from plastids, the low background levels of molecular beacons and also the free choice of fluorophoric groups allow the usage of them in living plant cells. One main concern in using hybridization probes in living cells is their delivery method. Microinjection has been used before, but this technique is time consuming and very laborious in plants. One laboratory reports the transfection of molecular beacons into plant suspension cells via electroporation. Treated cells show a sufficient survival rate and transfection efficiencies that are adequate for confocal fluorescence microscopy and single particle tracing (J. Göhring and A. Barta, unpublished data). CONCLUSION AND PERSPECTIVE The organization of plant nuclei, once thought to be static, has been shown to be highly dynamic, with the components of splicing in a constant flux between various subnuclear domains. The mechanisms that control the mobility of splicing regulators and the morphology of speckles in plants are emerging (Ali and Reddy, 2008b). Plant SR proteins and other splicing regulators constantly exchange between speckles and the nucleoplasm under a steady-state equilibrium state. A balance of the influx and efflux of SR proteins into and out of the speckles determines the morphology and size of speckles. An enhanced influx and/or decreased efflux increases the size of the SR speckles, and a decrease in the influx and/or increased efflux decreases the size or dismantles the speckles. Inhibition of transcription leads to a decline of pre-mRNAs and, hence, splicing. The SR protein molecules are recycled for the next round of splicing, and in the absence of any pre-mRNA substrate, SR proteins would tend to stay in the speckles, eventually leading to an increased speckle size. Phosphorylated SR proteins leave the speckle and participate in splicing, and when dephosphorylated, they enter the speckles. In the absence of phosphorylation, they are not competent for recruitment to the splicing sites and remain in the speckles. The dynamics of only a few plant splicing regulators have been studied to date. However, with the increased use of in vivo imaging methods combined with high-throughput analyses, a rapid expansion in our understanding of how plant pre-mRNA splicing is spatially and temporally organized and how the interactions of various splicing components are regulated should be forthcoming. Many fundamental questions pertinent to speckles in plants have yet to be answered. What is the composition of speckles? What is the meaning of different types of speckles, and when do they occur? Are there subpopulations of speckles that differ in their global composition? Does the composition of speckles differ depending on the physiological status of the cells or cell type? To address these questions, technological advances are needed to isolate speckles either biochemically or microscopically, which can then be analyzed using the available highly sensitive proteomic methods. Recently, single lipid droplets from cytoplasm were isolated by combining visualization and microphase separation to analyze their composition (Horn et al., 2011). A similar approach might be useful in analyzing the composition of speckles. Also, we do not have a clear understanding of the relationship between speckle-like patterns observed with various other proteins, including transcription factors (Tao et al., 2005; Chen, 2008), light receptors (Chen, 2008), and microRNA processing proteins (Fang and Spector, 2007), in plant cells. The biogenesis of speckles in plants is also not known. What initiates the formation of a speckle? Is there a small set of proteins that function as scaffolding proteins and initiate speckle formation? Although some plant SRs are found to be shuttling proteins, their cytosolic functions are not known. AS is a prominent feature of plant gene regulation and provokes questions about the fate of alternative transcripts, their localization, and their dynamics. One promising approach to follow RNA-protein particles is to label specific RNAs and determine their fate during the maturation steps. Future hopes lie specifically on single particle tracing with molecular beacons combined with high-resolution techniques, which should provide insight into the diffusion constants of different subpopulations of mRNA molecules within the nucleus and the cytoplasm (Christensen et al., 2010; J. Göhring and A. Barta, unpublished data). Further studies that address these questions should lead to a better understanding of splicing and gene regulation. ACKNOWLEDGMENTS We thank current and former members in the Barta and Reddy laboratories for their contributions to splicing research. LITERATURE CITED Ali GS Golovkin M Reddy AS ( 2003 ) Nuclear localization and in vivo dynamics of a plant-specific serine/arginine-rich protein . Plant J 36 : 883 – 893 Google Scholar Crossref Search ADS PubMed WorldCat Ali GS Prasad KV Hanumappa M Reddy AS ( 2008 ) Analyses of in vivo interaction and mobility of two spliceosomal proteins using FRAP and BiFC . 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Plant Physiol 150 : 1450 – 1458 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by grants from the U.S. National Science Foundation, Department of Energy, and Department of Agriculture, by the European Union FP6 Programme Network of Excellence on Alternative Splicing (grant nos. LSHG–CT–2005–518238), the Austrian Science Fund (grant nos. SFB 1710, SFB 1711, DK W1207, and ERA-NET Plant Genomics I254), and the Austria Genomic Program. * Corresponding author; e-mail [email protected]. www.plantphysiol.org/cgi/doi/10.1104/pp.111.186700 © 2012 American Society of Plant Biologists. All rights reserved. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Boruc, Joanna; Zhou, Xiao; Meier, Iris

doi: 10.1104/pp.111.185256pmid: 21949214

The nucleus is the most prominent compartment of any eukaryotic cell and home to its genetic information. The nucleoplasm is surrounded by a double membrane system, the nuclear envelope (NE). The outer nuclear membrane (ONM) and the inner nuclear membrane (INM) are separated by the perinuclear space (or periplasmic space; Hetzer et al., 2005). The lipid bilayer of the ONM is continuous with the endoplasmic reticulum (ER), thus allowing for direct insertion of NE membrane proteins and translocation of proteins into the perinuclear space (Hetzer and Wente, 2009); however, the ONM protein composition differs from the ER (Hetzer et al., 2005). The INM has a distinct protein composition and specialized functions. The INM and ONM are fused at specific sites to form aqueous pores. Inserted at these sites are the nuclear pore complexes (NPCs), large protein conglomerates responsible for the selective nuclear import and export of macromolecules (D’Angelo and Hetzer, 2008; Brohawn et al., 2009). Chromatin association with the nuclear pores and the NE is involved in gene activation and repression, respectively (Akhtar and Gasser, 2007; Kalverda et al., 2008; Capelson and Hetzer, 2009). In higher organisms, the NE plays a role in the dissociation and reformation of the nucleus during cell division (Kutay and Hetzer, 2008). Proteins that interact in the perinuclear space connect the nucleoplasm and cytoplasm through the NE, thereby transmitting information from the cytoskeleton and giving rise to nuclear mobility (Burke and Roux, 2009). Like the ER, the NE lumen acts as a repository of calcium, and ion transporters in both the ONM and INM are involved in signal transduction (Erickson et al., 2006; Bootman et al., 2009). Together, the NE and NPCs are at the crossroad of communication between the nucleus and cytoplasm. Recent reviews have discussed the mechanism and relevance of nuclear import and export in plants (Merkle, 2009), the regulation of plant nuclear import in the context of signal transduction (Meier and Somers, 2011), and the plant NE during the cell cycle (Evans et al., 2011). Here, we focus on the dynamic organization of the NE and nuclear pore in quiescent and dividing plant cells. COMPONENTS OF THE NUCLEAR PERIPHERY The Nuclear Lamina A mesh of intermediate filament proteins, the nuclear lamina, lines the mammalian INM. Lamins mediate the attachment of chromatin to the NE during interphase and chromatin detachment during mitosis (Gant and Wilson, 1997; Dechat et al., 2010). Lamin mutations cause a variety of human diseases that are collectively termed laminopathies (Andrés and González, 2009). Lamins have not been found outside the metazoan lineage; however, early electron microscopy and immunohistochemistry suggested a nuclear lamina and lamin-like proteins in plants (Galcheva-Gargova and Stateva, 1988; Li and Roux, 1992; McNulty and Saunders, 1992; Mínguez and Moreno Díaz de la Espina, 1993). In contrast, no lamin-coding genes were found in the complete plant genome sequences (Meier, 2007). New ultrastructural studies now suggest that a lamina-like structure does indeed exist in plants. A meshwork of filaments underlying the inner NE in tobacco (Nicotiana tabacum) BY-2 cells was recently revealed, closely resembling the animal nuclear lamina both in terms of organization and filament thickness (Fiserova et al., 2009). The best candidates for plant lamin-like proteins are currently a family of coiled-coil proteins about twice the size of lamins but with similar overall structure. First identified as Nuclear Matrix Constituent Protein1 (NMCP1) in carrot (Daucus carota; Masuda et al., 1997), NMCP1-like proteins have been found in many plant species, and some localize exclusively to the nuclear periphery (Moriguchi et al., 2005; Fig. 1A). Mutants in two NMCP1-related proteins in Arabidopsis (Arabidopsis thaliana), LITTLE NUCLEI1 (LINC1) and LINC2, have reduced nuclear size and changes in nuclear morphology, suggesting an involvement in plant nuclear organization (Dittmer et al., 2007). Figure 1. Open in new tabDownload slide Identified NE and NPC components in higher plants and vertebrates. A, Comparison of the NE and NPC components between higher plants and vertebrates. Subcomplexes are grouped in single units. Units in contact indicate confirmed interactions. The NPC organization is modified after Tamura et al. (2010). In the higher plant NPC, boldface protein names indicate confirmed NE localization. Mutant phenotypes have been reported for the plant Nups indicated in red. Mammalian Nups, Nup358, Nup188, Nup37, Nup97, Nup45, and Pom121, appear to have no counterparts in plants. The positioning of plant Nups is based on their vertebrate counterparts. B, NE localization of putative Arabidopsis NDC1 in Arabidopsis root tip cells. Cell walls were counterstained with propidium iodide (PI). Bars = 5 μm. Figure 1. Open in new tabDownload slide Identified NE and NPC components in higher plants and vertebrates. A, Comparison of the NE and NPC components between higher plants and vertebrates. Subcomplexes are grouped in single units. Units in contact indicate confirmed interactions. The NPC organization is modified after Tamura et al. (2010). In the higher plant NPC, boldface protein names indicate confirmed NE localization. Mutant phenotypes have been reported for the plant Nups indicated in red. Mammalian Nups, Nup358, Nup188, Nup37, Nup97, Nup45, and Pom121, appear to have no counterparts in plants. The positioning of plant Nups is based on their vertebrate counterparts. B, NE localization of putative Arabidopsis NDC1 in Arabidopsis root tip cells. Cell walls were counterstained with propidium iodide (PI). Bars = 5 μm. It is conceivable that NMCP1-like proteins or other, unknown proteins form a lamina-like protein meshwork underneath the plant NE. It will be well worth unraveling the function of plant lamin-like proteins, given the exciting emerging connection between the animal nuclear lamina and gene regulation (see below). Nuclear Envelope Proteins Proteins of the animal INM have been related to several human genetic diseases (Ellis, 2006; Worman and Bonne, 2007; Wheeler and Ellis, 2008). They include Lamin B Receptor (LBR), Lamina-Associated Polypeptide1 (LAP1), the LEM (for LAP2, Emerin, MAN1) domain protein family, as well as the Spindle Architecture Defective1/UNC84 (SUN) domain proteins (Wilson, 2010). Proteome analyses have added more proteins that have not yet been functionally investigated (Schirmer and Gerace, 2005). Surprisingly, very few INM proteins have homologs in plants. There is no plant LBR, but a GFP-LBR fusion protein is located at the plant INM, suggesting that the INM targeting signal is conserved (Irons et al., 2003). The first bona fide plant INM proteins have recently been reported in Arabidopsis and maize (Zea mays; Graumann et al., 2010; Murphy et al., 2010; Graumann and Evans, 2011; Oda and Fukuda, 2011). While the maize genome encodes at least five different SUN domain proteins, there are only two of them in the Arabidopsis genome. AtSUN1 and AtSUN2 are the Arabidopsis homologs of animal and yeast INM proteins containing a conserved SUN domain. In animals, SUN proteins interact in the perinuclear space with KASH domain proteins (located at the outer NE) to form protein bridges that connect the nucleus to the cytoplasmic cytoskeleton. SUN-KASH protein bridges are involved in attaching centrosomes to the nuclear periphery, the alignment of homologous chromosomes, and their pairing and recombination in meiosis. They have been implicated in the regulation of apoptosis, the maturation and survival of the germline, nuclear location, and human diseases such as laminopathies and Emery-Dreifuss muscular dystrophy (Burke and Roux, 2009; Fridkin et al., 2009; Hiraoka and Dernburg, 2009). AtSUN1 and AtSUN2 form dimers and are located at the INM in tobacco BY-2 cells (Graumann et al., 2010) and at the NE in different cell types of Arabidopsis plants (Oda and Fukuda, 2011). Their only currently known in planta role is an involvement in root hair nuclear shape. Nuclei in mature root hairs, which are normally elongated, appear round in the mutant, suggesting an involvement of plant SUN proteins in nuclear morphology. No KASH proteins are known in plants; thus, it is of great interest to identify plant interaction partners of SUN proteins. There are now a significant number of proteins available to serve as markers for NE dynamics in plants: NMCP1/2 (LINC1/2), SUN1/2, WPP DOMAIN-INTERACTING PROTEIN1 (WIP1)/2/3, WPP DOMAIN-INTERACTING TAIL-ANCHORED PROTEIN1 (WIT1)/2, and Nuclear Pore Anchor (NUA; Dittmer et al., 2007; Jacob et al., 2007; Xu et al., 2007a, 2007b; Zhao et al., 2008; Graumann et al., 2010; Fig. 1A). Together with the nucleoporins (see below), this should allow for the first thorough investigation of the order of disassembly/reassembly of plant NE/NPC components, similar to the impressive studies performed in other model organisms (Onischenko et al., 2009). In addition to dual and multicolor labeling for real-time imaging, the requirement of individual proteins, protein families, and protein domains for the dynamic behavior of other NE/NPC components can now be tested. NPCs NPCs are 40- to 60-MD multiprotein complexes embedded in the NE and involved in the nucleocytoplasmic trafficking of macromolecules. They consist of multiple copies of about 30 different nucleoporins (Nups) organized in a structure of 8-fold symmetry (Brohawn et al., 2009; Brohawn and Schwartz, 2009; Elad et al., 2009). The actual transport barrier in the core is composed of unfolded, hydrophobic repeat regions (FG repeats) of FG-Nups, which bind to shuttling transport receptors moving through the NPC (Frey et al., 2006; Frey and Görlich, 2007; Jovanovic-Talisman et al., 2009). For recent reviews on the different models of passage through the nuclear pore, see Wälde and Kehlenbach (2010) and Kahms et al. (2011). For many years, plant biologists have relied on high-resolution images of yeast and vertebrate NPCs and on one early study of the plant NPC (Roberts and Northcote, 1970). An in-depth view of the tobacco BY-2 cell and onion (Allium cepa) NPC structure and organization has recently been provided, demonstrating that the plant NPC closely resembles the known yeast and vertebrate NPCs (Fiserova et al., 2009). Plant NPCs appear to be surprisingly densely spaced (approximately 50 NPCs μm−2 compared with 60 NPCs μm−2 for Xenopus laevis oocytes, considered very rich in NPCs). Interestingly, the NPCs are not randomly distributed but rather aligned in rows, similar to other higher eukaryotes but different from yeast (Belgareh and Doye, 1997; Maeshima et al., 2006). Several proteins with significant similarity to animal and yeast Nups have been identified in forward genetic screens for diverse pathways. In addition, reverse genetic approaches with Nup homologs have been performed (Zhang and Li, 2005; Dong et al., 2006; Kanamori et al., 2006; Jacob et al., 2007; Saito et al., 2007; Wiermer et al., 2007; Xu et al., 2007b; Zhao and Meier, 2011). In general, however, it has proven difficult to assign plant Nup identity solely based on sequence similarity. A comprehensive proteomic study of the Arabidopsis nuclear pore has now added several additional plant Nups (Tamura et al., 2010). Using nuclear pore-associated GFP-Rae1 as their starting point, the authors performed a series of immunoprecipitations coupled with mass spectrometry, added more thorough sequence similarity searches, and identified together eight known and 22 novel Nups (Fig. 1A). Only the homologs for human Nup358, Nup188, Nup153, Nup45, Nup37, NUCLEAR DIVISION CYCLE1 (NDC1), and Pore membrane protein121 (Pom121) were absent in both the immunoprecipitations and the genome data. A candidate for Arabidopsis NDC1, however, had been proposed by Stavru et al. (2006). AtNDC1 (At1g73240) has sequence similarity to yeast Ndc1p and is predicted to contain six transmembrane domains shared by all NDC1 proteins (Stavru et al., 2006). When fused N terminally to GFP, AtNDC1 is localized at the NE in Arabidopsis root tip cells (Fig. 1B), thus adding AtNDC1 to the list of likely Arabidopsis Nups (Fig. 1A). An FG-Nup identified both as Nup136 (Tamura et al., 2010) and as Nup1 (Lu et al., 2010) appears to be unique to plants. Its cell cycle dynamics include dispersal at metaphase, accumulation around the chromosomes in late anaphase/early telophase, and reestablishment at the NE in late telophase. Nup136 mutants have complex developmental phenotypes reminiscent of other Nup mutants (Zhang and Li, 2005; Parry et al., 2006; Xu et al., 2007b; Zhao and Meier, 2011). Together, Tamura et al. (2010) provide a copious amount of new and confirmatory data about the plant NPC that have the potential to spark a much-needed systematic, multiprong functional investigation of the plant nuclear pore. DYNAMIC INTERACTION OF CHROMATIN WITH THE NE AND NPC Electron micrographs have long shown that heterochromatin accumulates under the NE, with gaps at the NPCs, while euchromatin is more centrally localized. This is true for most higher eukaryotes, including plants (Solovei et al., 2009). Large areas of gene-poor chromatin in humans are associated with the nuclear lamina (lamina-associated domains [LADs]). Thousands of genes are present in LADs in a low-density arrangement, and most genes within LADs have very low expression levels (Guelen et al., 2008). The mammalian histone deacetylase HDAC3 accumulates at the nuclear periphery, binds to lamina-associated proteins, and induces histone deacetylation (Somech et al., 2005). Histone methylation marks involved in silencing are enriched at the NE (Yokochi et al., 2009). Depletion of lamins causes the large-scale misregulation of gene expression (Malhas et al., 2007). Several transcription factors directly interact with proteins of the nuclear lamina. The transcription factor Oct1, for example, binds Lamin B1 and is enriched at the NE, dependent on Lamin B1. In a Lamin B1 mutant, the expression of Oct1-dependent genes is deregulated, suggesting that the physical association of Oct1 with lamins is involved in gene regulation (Malhas et al., 2009; Malhas and Vaux, 2009). Interestingly, artificial tethering of genes to the NE has resulted in the repression of some, but not all, tested genes, suggesting that while the NE environment can be sufficient to repress genes, active transcription also can occur at the NE (Finlan et al., 2008; Kumaran and Spector, 2008; Reddy et al., 2008). In contrast to the NE, the NPC has been recognized as a site of transcriptional activation (Gerber et al., 2004; Akhtar and Gasser, 2007). In yeast, a connection between the chromatin-bound Spt-Ada-Gcn5 acetyltransferase (SAGA) transcriptional coactivator complex, the nuclear pore protein Mlp1, and the RNA export complex TREX-2 (also known as the Thp1-Sac3-Cdc31-Sus1 complex) is implied in this activation. The SAGA histone acetyl transferase component Gcn5, the plant Mlp1 homolog NUA, and subunits of TREX-2 have all been identified in Arabidopsis, making it worthwhile to test if a similar connection might be involved in regulating plant gene expression (Stockinger et al., 2001; Xu et al., 2007b; Lu et al., 2010; Yelina et al., 2010). Nucleoporins are bound to hundreds of genomic sites, as identified by chromatin immunoprecipitation experiments and fusion of Nups to micrococcal nuclease (Schmid et al., 2006; Capelson et al., 2010; Vaquerizas et al., 2010; Wälde and Kehlenbach, 2010). Genes associated with Nups are typically highly to moderately expressed, in contrast to the LAD-located genes. Nups also contact chromatin away from the NPC, and interactions with the most highly active genes actually occur in the nucleoplasm (Kalverda and Fornerod, 2010; Kalverda et al., 2010). The rich and growing evidence on the regulation of gene expression by both NE and NPC components should encourage the plant community to also investigate this so far untouched question in plant model systems. Specifically, addressing whether the putative lamin-like plant proteins affect gene expression, investigating the spatial distribution of histone marks and of gene-rich and gene-poor areas of the genome, and testing Nup-chromatin interactions could open up a new area of investigation into the spatial organization of gene expression in plants. DUAL ROLES OF NE COMPONENTS DURING MITOSIS Plants, like all higher eukaryotes, undergo open mitosis when the NE breaks down and the separation of the nucleoplasm from the cytosol vanishes, until the NE reforms after a cell completes division. A cell needs to accurately segregate not only the genetic material and all the organelles but also the NE membranes with its specific protein components. According to the ER-retention model (Collas and Courvalin, 2000), some NE components are retained in the mitotic ER network during cell division, but numerous other ones localize to diverse mitotic structures and play crucial roles in consecutive stages of the division process (Rabut and Ellenberg, 2001; Griffis et al., 2004; Xu et al., 2008; Lee et al., 2009). Both the localization patterns and a variety of developmental phenotypes point to these functions. Preprophase/Prophase One of the canonical mitotic functions of the plant NE is to act as a microtubule (MT) organizing center (MTOC; Stoppin et al., 1994; Canaday et al., 2000). Plant cells undergo drastic MT array rearrangements during cell division, forming cortical and radial MTs, the preprophase band (PPB), the spindle, and phragmoplast structures. At the onset of mitosis, the cortical MTs depolymerize and rearrange into the PPB surrounding the nucleus. This initial cytoskeletal change is crucial for the fate of a dividing cell, since this transient MT array demarcates the future cortical division site, where a cell will separate into two daughter cells (Van Damme and Geelen, 2008; Müller et al., 2009). RanGAP1 is a NE-associated protein that is delivered to the PPB in an MT-dependent manner, and it remains associated with the cortical division site during mitosis and cytokinesis, constituting a continuous positive marker of the plant division plane (Xu et al., 2008). RanGAP1 is thus a molecular landmark left behind by the PPB, which later guides the phragmoplast and the forming cell plate, since the silencing of RanGAP1 in Arabidopsis roots leads to mispositioned cell walls similar to other mutants with division plane defects (Smith et al., 2001; Xu et al., 2008). At this stage, another NE-associated protein, Rae1, is targeted to the PPB (Lee et al., 2009; Fig. 2). This localization of Rae1 reflects its association with mitotic MTs throughout mitosis as well as at least partial involvement of the PPB in spindle assembly, since the RNA interference inhibition of Nicotiana benthamiana Rae1 (NbRae1) in BY-2 cells led to the formation of disorganized or multipolar spindles and defects in chromosome segregation (Lee et al., 2009). Indeed, in plants, the PPB marks the plane perpendicular to the axis of symmetry, the spindle (Lloyd and Chan, 2006). The PPB is linked to and cross-communicates with the nucleus through bridging MTs, which partly mediates the establishment of the bipolarity of a cell and the central positioning of the nucleus (Granger and Cyr, 2001; Ambrose and Wasteneys, 2008). This arrangement facilitates the formation of the prophase spindle perpendicular to the PPB. Figure 2. Open in new tabDownload slide Mitotic locations of NE-associated proteins. See text for details. Figure 2. Open in new tabDownload slide Mitotic locations of NE-associated proteins. See text for details. At this stage, the NE, acting as an MTOC, promotes the nucleation of MTs on its surface (Stoppin et al., 1994, 1996; Canaday et al., 2000). An essential factor of the MT-nucleating complex is the γ-tubulin ring complex, which is conserved among the kingdoms (Schmit, 2002). In mammals, the minimal complex functioning as an MTOC is composed of γ-tubulin, γ-TUBULIN COMPLEX PROTEIN2 (GCP2), and GCP3, which all have orthologs in the Arabidopsis genome (Canaday et al., 2004). Besides their sequence similarity, γ-tubulin, AtGCP2, and AtGCP3 were detected in the same complex in vivo, localized at the NE and the cell cortex, and were required for MT nucleation in Arabidopsis, corroborating the conserved function of the plant γ-tubulin ring complex (Erhardt et al., 2002; Seltzer et al., 2007). Interestingly, a nuclear rim-associated fraction of histone H1 was shown to have MT-organizing activity in BY-2 cells and to promote MT nucleation through the formation of complexes with tubulin and the elongation of radial MTs (Hotta et al., 2007; Nakayama et al., 2008; Fig. 2). Recently, a biophysical interaction between Ran and histone H1 and their colocalization at the nuclear rim have indicated a possible role for histone H1 in the organization of MTs adjacent to the NE in Leishmania donovani (Smirlis et al., 2009). Prior to the disappearance of the PPB in plant prophase, a rapid NE breakdown occurs (Dixit and Cyr, 2002). Both processes seem to require phosphorylation events carried out by a cyclin-dependent kinase (CDK) and its regulatory protein, cyclin B (CYCB). The CDK/CYCB complex promotes PPB disassembly in plants (Hush et al., 1996), the depolymerization of nuclear lamins in vertebrates, Caenorhabditis elegans, and yeast (Nigg, 1992; Daigle et al., 2001; Galy et al., 2008), and the disassembly of nucleoporins in animal cells (Macaulay et al., 1995; Favreau et al., 1996). This mitotic phosphorylation releases lamins and some nuclear membrane and nuclear pore proteins, enabling progression through the NE breakdown. Among plant nuclear pore proteins with dynamic mitotic relocalization, there is, for instance, NUA (the Arabidopsis homolog of Tpr/Mlp1/Mlp2/Megator; Jacob et al., 2007; Xu et al., 2007b) and Rae1 (Lee et al., 2009; Fig. 2). Metaphase The Ran gradient controls the spindle assembly in animal cells. High concentrations of RanGTP around chromosomes (and high RanGDP concentration at the cell periphery) attract importins and release nuclear localization signal-containing cargo proteins (Dasso, 2001; Weis, 2003). These cargos are, for instance, spindle assembly factors, such as targeting protein for Xklp2 (TPX2), Rae1, and NuMA (for Nucleus and Mitotic Apparatus; Carazo-Salas et al., 1999; Kaláb et al., 1999, 2006; Ohba et al., 1999; Wilde and Zheng, 1999; Wiese et al., 2001; Caudron et al., 2005). Arabidopsis TPX2 is nuclear in interphase, but it is actively exported in prophase, enriched around the NE, and then accumulates in the vicinity of the prospindle (Vos et al., 2008; Fig. 2). After its release from importin-dependent inhibition, TPX2 promotes spindle formation around chromosomes through MT nucleation (Gruss and Vernos, 2004; Vos et al., 2008). Simultaneously, human TPX2 targets Aurora A to the spindle and activates it (Bayliss et al., 2003; Gruss and Vernos, 2004; Kawabe et al., 2005; Vos et al., 2008). In plant and animal cells, the coordination of chromosomal and cytoskeletal events in mitosis is partly mediated by the chromosomal passenger complex. Aurora kinases (in Arabidopsis, Aurora1 and -2) are thought to play this role through mediating the positioning information of the PPB to the formation of the bipolar prophase spindle (Carmena and Earnshaw, 2003; Vagnarelli and Earnshaw, 2004; Demidov et al., 2005). At the onset of prophase, AtAurora1 and AtAurora2 are associated with the ONM and then gradually migrate to the poles of the prospindle as mitosis progresses (Demidov et al., 2005; Fig. 2). Tobacco NbRae1, a homolog of Rae1/mrnp41 in metazoans, Gle2p (for GLFG lethal 2p) in Saccharomyces cerevisiae, and Rae1 in Schizosaccharomyces pombe, exhibits a mitotic function besides its role as an mRNA export factor associated with the NPC (Whalen et al., 1997; Pritchard et al., 1999; Griffis et al., 2004; Lee et al., 2009). Mammalian Rae1 is a mitotic spindle checkpoint component in conjunction with Bub3 and forms a complex with Nup98 and the Cdh1-activated anaphase-promoting complex, preventing the degradation of Securin before anaphase (Whalen et al., 1997; Babu et al., 2003; Jeganathan et al., 2005). NbRae1 associates with the spindle and was shown to function in the proper spindle organization and chromosome segregation (Lee et al., 2009; Fig. 2). NbRae1 silencing resulted in delayed progression of mitosis, which led to plant growth arrest, reduced cell division activities in the shoot apex and the vascular cambium, and increased ploidy levels in mature leaves. Together, these results suggest a conserved function of the Rae1 proteins in spindle organization among eukaryotes, which is distinct from their roles at the interphase NE. In metaphase, while histone H1 relocalizes along the condensed chromosomes (Nakayama et al., 2008), Aurora3 and -1 are associated with centromeric regions of chromosomes (Demidov et al., 2005; Kawabe et al., 2005) and RanGAP1 localizes to kinetochores and the spindle (Joseph et al., 2002; Xu et al., 2008). Mammalian RanGAP1 is targeted to kinetochores in a SUMO-dependent manner (Joseph et al., 2002, 2004). Thus, it remains enigmatic how Arabidopsis RanGAP1, which lacks the SUMOylation domain, is targeted to kinetochores. In view of human RanGAP1, found only on the attached sister chromatids (Joseph et al., 2004), the exact timing of kinetochore association and the function of plant RanGAP1 at this cellular location remains to be verified. Recently, the cell cycle dynamics of Apium graveolens NMCP1 and NMCP2 (AgNMCP1 and AgNMCP2) were investigated (Kimura et al., 2010). Both proteins associate with the NE in interphase, disassemble simultaneously during prometaphase, and reaccumulate around the reforming nuclei (Fig. 2). However, while AgNMCP1 was mainly localized to the spindle and accumulated on segregating chromosomes, AgNMCP2 dispersed in the mitotic cytoplasm in vesicular structures that could be distinguished from the bulk endomembrane system. This vesicular signal might represent the NE membranes absorbed into the ER network upon NE breakdown. Two Arabidopsis homologs of the spindle pole body protein Sad1 were initially discovered in a survey for cytokinesis-related genes (Hagan and Yanagida, 1995; Van Damme et al., 2004). These Arabidopsis SUN domain proteins are NE markers in plants (Graumann et al., 2010). Oda and Fukuda (2011) and Graumann and Evans (2011) carefully followed the localization dynamics of both proteins through the cell cycle using transgenic Arabidopsis plants and stably transformed BY-2 cells, respectively. Both groups reported the localization of SUNs in mitotic ER membranes and an asymmetric reassociation with the decondensing telophase chromatin, with an envelope-like structure first appearing at the surface next to the spindle poles and a delayed reappearance of the envelope at the surface close to the phragmoplast (Fig. 2). This might indicate that NE assembly lags behind at the phragmoplast-proximal surface of the daughter nuclei, and potentially this area remains open longer to nonrestricted exchange between nucleus and cytoplasm. Alternatively, because SUN1/2 are nuclear proteins, it might indicate that nuclear pores at the phragmoplast-proximal surface lag behind in regaining full import capacity. These scenarios can be distinguished by also following ONM and NPC proteins as well as generic markers for active nuclear import. Anaphase/Telophase As chromosomes migrate to opposing spindle poles, a plant-specific MT structure, the phragmoplast, is formed to allow the completion of cell division through the assembly of a new cell wall between the separating sister nuclei (Verma, 2001; Jürgens, 2005). Besides the proteins involved in vesicular trafficking and fusion (for review, see Van Damme and Geelen, 2008), some NE-associated proteins have been found to mark the phragmoplast and/or the cell plate as well. The localization of Rae1 and SUN1/2 at the cell plate (and the phragmoplast for Rae1; Fig. 2) suggests a tight linkage between the NE components and the cytoskeleton during mitosis. Thus, it would be of utmost interest to identify plant interactors of SUN proteins both at the NE and at the cell plate. Such data would shed more light on molecular bridges across the perinuclear space, linking the nucleoskeleton to the cytoskeleton, as well as on functions of NE proteins in cell division. Apart from Rae1, other nuclear rim-associated proteins colocalize with SUNs at the cell plate as well. For instance, Arabidopsis ONM proteins, WIP1, WIP2, WIT1, and WIT2, are redistributed to the cell plate during cytokinesis (Patel et al., 2004; Xu et al., 2007a; Zhao et al., 2008; Fig. 2). Both WITs and WIPs are required for RanGAP1 anchoring to the NE in the root meristem, but only one of the protein families, either WIPs or WITs, is sufficient to target RanGAP1 to the NE in differentiated cells (Zhao et al., 2008). The cell plate localization of RanGAP1 (as well as its PPB and cortical division site association), on the other hand, is independent on both WIPs and WITs, suggesting that interphase and mitotic targeting of RanGAP1 require different mechanisms. Therefore, identification of the molecular players involved in RanGAP1 localization and function(s) during plant cell division would be of great importance. OUTLOOK Over the past years, much progress has been made in unraveling the molecular players residing at the nuclear periphery in animal, yeast, and plant cells. Numerous INM, ONM, as well as nuclear lamina and nuclear pore proteins have been brought to the stage via homology-based reverse genetics, forward genetics, or proteomics approaches. The NE components have been shown not only to separate the nucleoplasm from the cytosol and to constitute a selective barrier for nucleocytoplasmic transport but are also involved in nuclear mobility, signal transduction, chromatin attachment, and transcriptional activation and repression. Subcellular localization as well as thorough phenotypic analyses have delivered additional spatiotemporal information regarding NE-associated proteins. Namely, in plants, these molecular players have been implicated in such mitotic events as spindle assembly, chromosome segregation, MTOC-like function, cortical division site demarcation, and NE reformation upon cytokinesis. The concept of NE components having additional roles throughout cell division is fascinating but very challenging to dissect experimentally. Therefore, certain biological questions remain to be addressed. In vivo “fishing expeditions” using NE molecules as baits would possibly elucidate the protein interactors involved in particular processes of cell division as well as targeting mechanisms of these molecules to diverse cellular addresses. 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Plant Cell 20 : 1639 – 1651 Google Scholar Crossref Search ADS PubMed WorldCat Zhao Q Meier I ( 2011 ) Identification and characterization of the Arabidopsis FG-repeat nucleoporin Nup62 . Plant Signal Behav 6 : 330 – 334 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by the National Science Foundation (grant nos. MCB–0641271 and MCB–0919880 to I.M.). * Corresponding author; e-mail [email protected]. www.plantphysiol.org/cgi/doi/10.1104/pp.111.185256 © 2012 American Society of Plant Biologists. All rights reserved. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)