Research Article

Mutations in AtPS1 (Arabidopsis thaliana Parallel Spindle 1) Lead to the Production of Diploid Pollen Grains

  • Isabelle d'Erfurth,

    Affiliation: French National Institute for Agricultural Research (INRA), UR254, Versailles, France

  • Sylvie Jolivet,

    Affiliation: French National Institute for Agricultural Research (INRA), UR254, Versailles, France

  • Nicole Froger,

    Affiliation: French National Institute for Agricultural Research (INRA), UR254, Versailles, France

  • Olivier Catrice,

    Affiliation: National Center for Scientific Research (CNRS), UPR2355, Gif sur Yvette, France

  • Maria Novatchkova,

    Affiliation: Research Institute of Molecular Pathology (IMP), Vienna, Austria

  • Mathieu Simon,

    Affiliation: French National Institute for Agricultural Research (INRA), UR254, Versailles, France

  • Eric Jenczewski,

    Affiliation: French National Institute for Agricultural Research (INRA), UR254, Versailles, France

  • Raphaël Mercier mail

    Affiliation: French National Institute for Agricultural Research (INRA), UR254, Versailles, France

  • Published: November 28, 2008
  • DOI: 10.1371/journal.pgen.1000274


Polyploidy has had a considerable impact on the evolution of many eukaryotes, especially angiosperms. Indeed, most—if not all—angiosperms have experienced at least one round of polyploidy during the course of their evolution, and many important crop plants are current polyploids. The occurrence of 2n gametes (diplogametes) in diploid populations is widely recognised as the major source of polyploid formation. However, limited information is available on the genetic control of diplogamete production. Here, we describe the isolation and characterisation of the first gene, AtPS1 (Arabidopsis thaliana Parallel Spindle 1), implicated in the formation of a high frequency of diplogametes in plants. Atps1 mutants produce diploid male spores, diploid pollen grains, and spontaneous triploid plants in the next generation. Female meiosis is not affected in the mutant. We demonstrated that abnormal spindle orientation at male meiosis II leads to diplogamete formation. Most of the parent's heterozygosity is therefore conserved in the Atps1 diploid gametes, which is a key issue for plant breeding. The AtPS1 protein is conserved throughout the plant kingdom and carries domains suggestive of a regulatory function. The isolation of a gene involved in diplogamete production opens the way for new strategies in plant breeding programmes and progress in evolutionary studies.

Author Summary

In the life cycle of sexual organisms, meiosis reduces the number of chromosomes from two sets (2n) to one set (n), while fertilization restores the original chromosome number. However, in case of failure of meiosis to reduce the chromosome number, the fecundation involving the obtained 2n gametes can lead to the formation of an organism with more than two sets of chromosomes (polyploid). Polyploidization occurred widely in the course of evolution of eukaryotes, especially of plants. Besides, many crops are current polyploids, and 2n gametes have been useful for their genetic improvement by allowing crosses between 2n and 4n species. 2n gametes formation is known to be under genetic control but none of the genes involved were identified. We have isolated and characterised a gene (AtPS1) involved in controlling diploid (2n) gamete formation in A. thaliana. In the Atps1 mutant, the second division of meiosis is disturbed, leading to the gathering of chromosomes that had been separated at the first division. Consequently, Atps1 mutants produce 2n male gametes and spontaneous triploid plants in the next generation. The isolation of a gene involved in diplogamete production opens the way for new strategies in plant breeding programmes and progress in evolutionary studies.


Polyploidy, the condition of organisms having more than two sets of chromosomes, has had a considerable impact on the evolution of many fungi, invertebrate, and vertebrate lineages and is particularly prominent in plants [1],[2]. It is estimated that 95% of ferns are polyploids [3] and that almost all angiosperms have experienced at least one round of whole genome duplication during the course of their evolution [4]. Many important crop plants are currently polyploids or retain the vestiges of ancient polyploid events [5][7]. Even plants with small genomes, such as Arabidopsis thaliana, have been affected by polyploidy [8],[9]. However, the mechanisms involved in polyploid formation are still poorly understood. For a long time, polyploids were thought to originate from somatic chromosome doubling[10]. The realisation that gametes with somatic chromosome numbers (2n gametes or diplogametes) widely occur in diploid populations as a result of meiotic failure, led to a change of paradigm[11]; it is now believed that 2n gametes are the major route for polyploidy formation, in particular by leading to the formation of triploids, which then may serve as a bridge/step towards even ploidy levels [2], [12][15]. 2n gametes are also instrumental in the genetic improvement of several polyploid crops, where useful genes from diploid relatives are incorporated into cultivated genotypes [16],[17].

Given their importance in evolution and crop improvement, 2n gametes have been the focus of a considerable amount of research [12],[18]. The best documented and described meiotic abnormalities leading to 2n gamete formation include abnormal cytokinesis, the omission of the first or second division and abnormal spindle geometry. Co-orientation of 2nd division spindles (parallel spindles or fused spindles) is perhaps the most common mechanism resulting in 2n spore formation [12],[18], most notably in potato [14], and was first described more than eight decades ago [19],[20].

Environmental factors, notably temperature and chemical agents, were shown to affect the frequency of 2n gametes [12],[13],[21]. However, 2n gamete production is under strong genetic control [13]. The genetic determination of 2n pollen production was studied in several species [12] and usually fits the segregation pattern expected for a major locus in a background of polygenic variation. To date, however, none of the genes contributing to high frequency 2n gametes production were identified and characterised at the molecular level [22],[23]. This lack of information has slowed down our understanding of the origins of diplogametes, and limited the potential of diplogametes in crop breeding programmes.

In this paper we describe the isolation and characterisation of the first gene, called AtPS1 (Arabidopsis thaliana Parallel Spindle 1), implicated in the formation of a high frequency of diplogametes in plants. We show that meiosis in Atps1 mutants generates diploid male spores, giving rise to viable diploid pollen grains and spontaneous triploid plants in the next generation. Analysis of male meiosis showed that during meiosis II spindles are abnormally orientated, with frequent parallel or fused spindles, leading to the production of two sets of chromosomes instead of four at the end of anaphase II. Genetic analyses of the diploid gametes and epistasis experiments demonstrated that diplogamete formation in Atps1 results from these defects in spindle organisation.


Identification of AtPS1

AtPS1 was identified in a screen for genes potentially involved in meiosis using the Expression Angler tool [24], which selects co-regulated genes, in combination with the AtGenExpress tissue data set [25]. We first chose a subset of known meiotic genes (AtMER3 [26], AtDMC1 [27], SDS [28], AtMND1 [29],[30], AtHOP2 [31], AtMSH5 [32] and AtSPO11-1 [33]) for which the expression data appeared to be relevant: when one of these genes was used as the query in the Expression Angler tool (with default parameters [24]), other known meiotic genes appeared among the first hits. We thus selected a list of genes that appeared among the first 60 hits in at least one query, and in the first 100 hits in at least two independent queries, using one of these seven genes as bait. Following an additional manual selection, including elimination of genes with known function or essential character, we ended up with a list of 138 candidate genes. We examined the phenotype of one to three lines carrying an insertion in each of these genes (218 lines in total) [34][38]. Thirteen genes were not tested because corresponding mutant lines were not found in the databases. We visually screened ~50 plants of each line obtained from stock centers [35],[39] for reduced fruit length, without genotyping. The meiotic products of plants with reduced fertility were then examined. In two independent lines carrying an insertion in the AT1G34355 gene, plants were found to have slightly reduced fertility and unbalanced meiotic products. Chromosome spreads revealed that these plants were polyploid, prompting us to analyze these lines further. Functional characterization of this gene led us to name it AtPS1 (see below). Two other genes with meiotic function were identified in the same screen (R. M., unpublished data).

We amplified the AtPS1 cDNA (EU839993) by RT-PCR on bud cDNA and sequencing confirmed that it is identical to that predicted in the databases (NM_103158). The AtPS1 gene contains 7 exons and 6 introns (Figure 1A) and encodes a protein of 1477 amino acids. BLASTp and PSI-Blast [40] analyses showed that the AtPS1 protein is conserved throughout the plant kingdom and contains two highly conserved regions. An FHA domain (forkhead associated domain) was predicted at the N-terminus (CD-search: 65–140 aa, E-value 2e-11) [41], while the C-terminal conserved region shows similarity to a PINc domain as identified using the SMART outlier homologue search (BLAST: PINc, 1237–1389 aa, E-value 1.00e-84) [42], the InterPro superfamily search (SSF88723: PIN domain-like, 1235–1412 aa, E-value 8.8e-09) [43] as well as borderline similarities in CD-search (smart00670: PINc, 1237–1305 aa, E-value 0.21) (Figure 1B and 1C). No close homologs of AtPS1 containing both the FHA and PINc domain were found outside of the plant kingdom. An FHA domain is a phosphopeptide recognition motif implicated in protein-protein interactions and is found in a diverse range of proteins involved in numerous processes including intracellular signal transduction, cell cycle control, transcription, DNA repair and protein degradation [44]. The PINc domain has been predicted to have RNA-binding properties often associated with RNAse activity [45], and this has now been experimentally confirmed [46]. Accordingly, several PINc domain containing proteins are involved in RNAi, RNA maturation, or RNA decay. The highest level of sequence similarity to the AtPS1 PINc domain in eukaryotes was found among others with S. cerevisiae Swt1[47], mammalian C1orf26, Drosophila CG7206 and SMG6 protein families [45],[48] (Figure 1D).


Figure 1. AtPS1 gene and protein.

A) The intron/exon structure of the AtPS1 gene and location of the three different T-DNA insertions (triangles). The primers used are indicated below the gene diagram. B) Domain architecture of Arabidopsis thaliana AtPS1. In addition to the conserved FHA (IPR000253, 64–132aa) and PIN (IPR006596, 1237–1389aa) protein domains, Arabidopsis AtPS1 contains three repeat regions (rep-reg) which were further analyzed using HHpredid [67] and REPRO [68]. The CAST program detected a serine bias for the 182–1251 amino acid segment, and a glutamic acid-bias from amino acids 559 to 708 [69]. C) Multiple sequence alignment showing regions of highest sequence conservation among plant AtPS1 proteins. Based on sequence similarity, phylogenetic analysis and domain architecture, the listed plant proteins are likely orthologs of the Arabidopsis AtPS1 protein. Full-length plant AtPS1 proteins were aligned and segments of highest conservation were identified using plotcon (EMBOSS package). Non-conserved sequence segments were removed from the alignment. The length of these regions is indicated in box brackets in the corresponding position of the alignment. Domain hits based on a comparison against the Interpro domain database [43] are indicated by gray boxes above the alignment, and the hit-indicators are extended to include adjacent segments which showed sequence and structural conservation. In addition, white boxes highlight a short motif identified as a C repeat element in Arabidopsis thaliana AtPS1. Plant AtPS1 proteins typically contain a highly conserved N-terminal FHA domain and C-terminal PINc domain, which are separated by a compositionally biased sequence of variable length. While a single AtPS1 representative is usually found per species, gene duplication events appear to have occurred in some lineages such as Physcomitrella patens and Glycine max. The predicted Physcomitrella patens co-orthologous sequences do not contain a detectable FHA N-terminal domain, present in all other identified plant AtPS1sequences. This could be due to gain of the FHA domain in the Tracheophyta lineage, or due to loss or degeneration of the FHA domain in the Physcomitrella lineage. D) Multiple sequence alignment of the plant AtPS1 C-terminal PINc domain and representatives of closely related PINc domain homologs. The closest non-plant AtPS1 PINc domain homologs were identified using profile-based similarity searches such as PSI-BLAST and HHpred [70],[71]. Based on these analyses the AtPS1 PINc domain belongs to a PINc subfamily including human C1orf26, human SMG6, and bacterial PhoH proteins. Conservation of functionally important regions between AtPS1 and human SMG6 PINc domains suggest that the AtPS1 PINc domain might confer nuclease activity/function. Asterisks indicate the residues contributing to the negative charged cavity and potential active site of the human SMG6 PINc domain [46],[72]. Protein sequences are listed using a 2 letter species code followed by a reference to the sequence source. Species codes: At Arabidopsis thaliana, Rs Raphanus sativus, Bn Brassica napus, AfxAp Aquilegia formosa x Aquilegia pubescens, Pt Populus trichocarpa,Vv Vitis vinifera, Gm Glycine max, Os Oryza sativa, Sb Sorghum bicolor, Zm Zea mays, Sm Selaginella moellendorffii, Pp Physcomitrella patens, Hs Homo sapiens, Dm Drosophila melanogaster, Vc Vibrio cholerae. Sequences obtained from the NCBI non-redundant protein database are assigned their NCBI accession number, sequence start/end positions and gene name if available. All other sequences were derived from predicted translations of EST contigs or using preliminary gene predictions made available by the Department of Energy's Joint Genome Institute (JGI).


We investigated the role of the AtPS1 gene by isolating and characterizing a series of allelic mutants, identified in several public T-DNA insertion line collections [34],[35],[37]. The Atps1-1 (SALK_078818) and Atps1-2 (WiscDsLox342F09) insertions are in a Columbia (Col-0) background and are in the fourth exon and first intron, respectively. The Atps1-3 (FLAG_456A09) insertion is in a Wassilewskija (Ws-4) background and is located in the second exon (Figure 1A). RT-PCR was carried out using the pAtpsF/pAtpsR primers (Figure 1A) on RNA from the Atps1-3 and Atps1-1 mutants and no detectable levels of the AtPS1 transcript were amplified, indicating that these two alleles are null. When the same primers were used on RNA from the Atps1-2 mutant normal expression levels of this region of the AtPS1 transcript were observed (data not shown). Nevertheless, the phenotype analysis described below strongly suggests that this third allele is also null.

Diploid Spores and Pollen Grains in Atps1

In A. thaliana, male meiosis produces a group of four spores, organised in a tetrahedron, called a tetrad. As expected, male meiotic products in wild type were almost exclusively tetrads (Figure 2). Rarely, (13/304) groups of three spores were also seen but these were most certainly the result of occasional spore superposition. In contrast, the meiotic products in the three independent Atps1 mutants were characterized by a high frequency of dyads and triads (Figure 2). Atps1 mutants did not show any other developmental defects. The Atps1-1 and Atps1-2 mutants produced a majority of dyads (~65%). The Atps1-3 mutant phenotype appeared to be weaker and only 8% of its meiotic products were dyads. Complementation tests between Atps1-1 and Atps1-2 and Atps1-3 and Atps1-1 showed that these mutations are allelic, and thus demonstrated that the dyads observed in this series of mutants are due to disruption of the AtPS1 gene.


Figure 2. Atps1 mutants produce dyads of spores instead of tetrads.

A) Photos of meiotic products in wild type and Atps1 mutants. Scale bar = 10 µm. B) Quantification of meiotic products in Ws-4 (n = 92), Col-0 (n = 212), Atps1-3 (n = 436), Atps1-1 (n = 1125), Atps1-2 (n = 554), the Atps1-1/Atps1-3 F1 (n = 283) and Atps1-1/Atps1-2 F1 (n = 252).


The Atps1-3 mutant exhibited a weaker phenotype than the two other alleles, whereas expression analysis suggested that this allele is also null. As this allele was in a different genetic background (Ws-4) to the two others (Col-0), we tested if this difference could be influencing the strength of the phenotype by introducing the Col-0 mutation into the Ws-4 background and vice versa. As expected for a background effect, the frequency of dyads increased with successive backcrosses when Atps1-3 was introduced into Col-0 (from 8% to 58% after four backcrosses) and decreased when Atps1-1 was introduced into the Ws-4 background (from 64% to 13% after four backcrosses). These results clearly indicate that the frequency of diploid gametes is influenced by multiple genes, with AtPS1 acting as a major gene.

Pollen grain viability was examined by Alexander staining [49] and showed that in the majority of cases the dyads and triads produced by the mutants result in viable pollen grains (more than 95% in the different Atps1 mutants : Col: 0 dead pollen grains out of 181 ; Atps1-1: 44 dead pollen grains out of 948 ; Atps1-2: 3 dead pollen grains out of 363). We did observe however that the pollen grains in mutant plants varied in size (data not shown). We then assessed the ploidy level of Atps1-1 and Atps1-2 pollen grains by quantifying spermatic nuclei DNA. Both mutants exhibited two different populations of pollen grains, one corresponding to viable haploid pollen grains (~40% estimated by maximum likelihood) and another to viable diploid pollen grains (~60% estimated) (data not shown). These proportions are compatible with the proportion of dyads, triads and tetrads observed in the mutants. In summary, the Atps1-1 and Atps1-2 mutants produce a high frequency of viable diploid pollen grains.

Spontaneous Triploids among Atps1 Progeny

Next, we measured the ploidy level of the offspring of diploid Atps1 mutants by flow cytometry. Diploid and triploid plants (30%), but no tetraploid plants, were found among the progenies of Atps1-1 and Atps1-2 mutants (Atps1-1: 38 triploids out of 130 plants; Atps1-2: 30 triploids out of 103 plants). Flow cytometry results were confirmed by karyotyping a subset of 29 plants which were all confirmed to be triploid. This demonstrated that the diploid gametes produced in the Atps1 mutants are involved in fertilisation and produce viable triploid plants. The appearance of triploids, but not tetraploids, suggests that the Atps1 mutations only affect male meiosis. As expected for the absence of a female meiotic defect we never isolated triploid plants when ovules from plants with the Atps1 mutation were fertilised with wild type pollen grains (0 triploids out of 182 plants). When mutant pollen was used for the cross we again observed that 30% of the progeny were triploids (20 triploids out of 56 plants).

The observed frequency of triploid plants (30%) among Atps1-1 and Atps1-2 mutant progeny is lower than expected from the frequency of diploid pollen grains produced by these mutants (~60%). In parallel, more than 50% of seeds obtained by selfing the Atps1-1 and Atps1-2 mutants were thinner than wild type, abnormally colored and shaped, and germinated at a rate of 57%, compared to 99.8% in wild type. We do not believe, however, that this seed mortality phenotype infers a possibly essential role for AtPS1 in embryo development, for the following two reasons: 1) 25% (56/210) of the progeny of selfed heterozygotes were mutant and no dead seed was obtained, showing that the Atps1 mutation does not impair embryo development. 2) The same seed defect (59% of germination) is observed when Atps1 is crossed as male with wild type as female, which shows that a seed with one functional AtPS1 allele may show developmental defects. Thus, a likely explanation for the discrepancy between the frequency of diploid pollen grains and triploids in the progeny is abnormal development of triploid seed, which is commonly observed during crosses between plant species with different ploidy levels. These problems are related to the paternal to maternal ratio, which is very important for normal endosperm development [50]. Using C24 and Ler accessions, Scott et al showed that triploid seeds obtained in diploid X tetraploid crosses germinated at a rate of 90%. We obtained stronger germination defects with Col0, suggesting a background effect on the susceptibility to the paternal/maternal ratio. Another, non-exclusive, explanation for the discrepancy could be that haploid pollen grains out-competed diploid pollen grains, which were shown in some cases to germinate more slowly [51],[52]. Nevertheless, approximately 25% of the triploid embryos appear to be able to overcome these constraints.

The Mechanism Leading to Dyads Production

To unravel the mechanisms leading to dyad production in Atps1-1, we investigated chromosome behaviour during meiosis (Figure 3). Chromosome spreads showed that the meiosis in the Atps1-1 mutant progresses normally and is indistinguishable from the wild type until the end of the telophase I. Synapsis was complete, chiasmata formed (the cytological manifestation of crossovers) and bivalents were seen (compare Figure 3G–I with Figure 3 A–C, for example). At metaphase II, however, differences were seen compared to wild type with the 10 chromosomes aligned in a same plane, causing abnormal looking figures, rather than two well separated metaphase II plates containing five chromosomes each (Compare Figure 3J–K with Figure 3D). In rare cases, metaphase II in Atps1 did appear normal however (Figure 3L). At telophase II, we observed dyads (two sets of 10 chromosomes, Figure 3M), triads (2 sets of five chromosomes and one set of 10, Figure 3N) and normal tetrads (4 sets of 5 chromosomes, Figure 3O). These observations are consistent with the previous finding that Atps1 meiotic products are a mixture of dyads, triads and tetrads.


Figure 3. Meiosis I is normal but meiosis II is defective in Atps1 mutants.

A–F) Wild type meiotic chromosome spreads. A) pachytene. B) metaphase I. C) anaphase I. D) metaphase II. E) Anaphase II F) telophase II. G–O Atps1-1 meiosis. G) pachytene. H) metaphase I. I) anaphase I. J–L) metaphase II. M–O) Anaphase II. M) dyad. N) triad. O) tetrad. Scale bar = 10 µm.


These results and specifically the alignment of the 10 chromosomes at metaphase II suggested that the meiotic spindles in Atps1 mutants are defective at this stage. We thus examined spindle organisation by immunolocalisation with an alpha-tubulin antibody (Figure 4). In wild type plants the majority of metaphase II spindles were roughly perpendicular to each other (Figure 4A), leading to four well separated poles at anaphase II (Figure 4B) and the formation of tetrads (Figure 4C). In the Atps1 mutant, while individual metaphase II / anaphase II spindles appeared regular in most cases their respective orientation was aberrant. The majority of cells had parallel spindles (Figure 4D to 4G), fused spindles (Figure 4H and 4I) or tripolar spindles (Figure 4J and 4K). This defect in spindle orientation explains the appearance of triads and dyads. These conformations cause chromatids, that had been separated at meiosis I, to gather at anaphase II. Occasionally, three to four sets of chromosomes encompassed by a spindle were dispersed in the cell at metaphase II (Figure 4L). This type of defect is probably the cause of the few unbalanced meiotic products observed in the Atps1 mutants.


Figure 4. The relative orientation of spindles at metaphase II is disrupted in Atps1.

A, B, C) Wild type spindles at metaphase II, anaphase II and telophase II, respectively. D–L) Atps1-1 meiocyte at metaphase II / anaphase II. Chromosomes were stained by propidium iodide or DAPI (red, false color) and microtubules by immunolocalisation (green, false color). Scale bar = 5 µm.


The name AtPS1 for Arabidopsis thaliana Parallel Spindle 1 was chosen due to the high percentage of parallel spindles produced by the corresponding mutants.

Thus, parallel spindles at metaphase II in the Atps1 mutants appear to be leading to the formation of dyads. This proposed mechanism implies that unbalanced chromosome segregation at meiosis I would have no impact on the final distribution of chromosomes in the resulting dyad. To test this hypothesis we constructed a double Atspo11-1/Atps1 mutant. The Atspo11-1 mutant (N646172, Atspo11-1-3) [53] displays an absence of bivalents at meiosis [33] (Figure 5A) leading to frequent unbalanced first divisions (Figure 5B) that can be associated with lagging chromosomes (Figure 5C). At metaphase II, unbalanced plates are seen (Figure 5D), leading to unbalanced tetrads (Figure 5E). Lagging chromosomes at anaphase II, lead to multiple metaphase II plates and then polyads with more than four nuclei (Figure 5F).


Figure 5. Unbalanced chromosome distribution at meiosis I does not affect dyad formation in Atps1.

A–F) Atspo11-1 meiotic chromosome spreads. A) metaphase I. B, C) anaphase I. D) metaphase II. E) Unbalanced Tetrad. F) Polyad. G to R) Atps1-1/Atspo11-1 meiosis. G) metaphase I. H, I) anaphase I. J) metaphase II. K) Anaphase II. L, M) Balanced dyad II. N. O) Triad. P, Q) Unbalanced tetrad. R) Polyad. Scale bar = 10 µm.


In the Atspo11-1/Atps1 background the first division was identical to the single Atspo11-1 phenotype. We observed 10 univalents at metaphase I (Figure 5G), leading to missegregation at anaphase I, with two sets of unbalanced chromosomes (Figure 5H) or three sets because of lagging chromosomes (Figure 5I). At metaphase II, we regularly observed two unbalanced metaphase plates, which had a tendency to be parallel instead of perpendicular (Figure 5J). This led to the formation of dyads which were always balanced (Figure 5K to 5L, n = 44). We also observed triads with one set of 10 chromosomes caused by an unbalanced first division followed by the fusion of two of the four second division products (Figure 5N), which is highly consistent with our proposed mechanism. We also observed unbalanced tetrads (Figure 5P and 5Q), expected since the Atps1 mutation is not fully penetrant, and polyads due to lagging chromosomes at the first division (Figure 5R).

Another prediction of the proposed mechanism is that centromere distribution should resemble that seen during mitosis, e.g., any heterozygosity at the centromeres should be retained in the diploid gametes. Indeed, in Atps1, the first division is identical to wild type, with the co-segregation of sister chromatids and separation of homologous chromatids. Thus, in the case of a heterozygous genotype, A/a, at the centromere, following the first division the two A alleles will end up at one pole, and the two a alleles at the opposite pole. In wild type, the second division separates the two sisters leading to four spores with one chromatid. In Atps1, the second division would regroup the products of the first division, thus grouping the a and A allele in each cell, leading to systematic heterozygosis at the centromere. Because of recombination, loci unlinked to centromeres should segregate randomly. We tested this prediction by taking advantage of the two genetic backgrounds of the Atps1-1 (Col-0) and Atps1-3 mutants (Ws-4). F1 plants bearing the two mutations – thus mutant for AtPS1 and heterozygous for any Col-0/Ws-4 polymorphisms – were crossed as male to a third genetic background Landsberg erecta (Ler). Karyotyping and genotyping of the obtained plants for trimorphic molecular markers provided direct information regarding the genetic make up of the pollen grain produced by the mutant (Figure 6). All the diploid pollen grains tested had the predicted genetic characteristics. They were systematically heterozygous at centromeres and segregating–because of recombination–at other loci. These results confirm that the “parallel spindle” defect is indeed the cause of at least the vast majority of 2n pollen in Atps1.


Figure 6. Diploid gametes are heterozygous at centromeres.

Diploid and triploid offspring of the Atps1-1(Col-0) / Atps1-3(Ws-4) ♂×Ler ♀ cross was genotyped for several genetic markers. For each marker plants bearing only the Col-0 allele are in green, plants bearing only the Ws-4 allele are in yellow and plants bearing both the Col-0 and Ws-4 alleles are in red. The Ler alleles are present in all the plants because it was used as the female parent in the cross. The position of each marker (red) and the centromeres (dark blue) are indicated along the chromosomes.



In this study, we identified and described the AtPS1 gene and a corresponding set of mutants that produce pollen grains which are up to 65% diploid and give rise to numerous triploid plants in the next generation. Another Arabidopsis mutant that leads to severe meiotic defects and almost sterility [54],[55] was recently described and reported to produce diploid female gametes [56], but at a frequency of several orders of magnitude lower than the frequency of 2n gametes induced by the Atps1 mutation. By combining cytological and genetic analyses, we carried out a detailed investigation of the mechanism responsible for these 2n pollen grains in Atps1, and established that they result from abnormal orientation of spindles at meiosis II. Interestingly, defects in meiosis II spindles are the most common known mechanisms responsible for the formation of 2n spores, [12],[18] and are the main source of the 2n pollen which is extensively used in potato breeding programmes [14]. In potato, a major locus called ps was shown to be responsible for the parallel spindle phenotype more than 30 years ago [57], but the corresponding gene is still to be identified. As was observed in different ps potato lines, Atps1 mutations only affect male meiosis and the frequency of dyads formed depends on the genetic background. The AtPS1 gene is conserved in higher plants (Figure 1C) and is therefore a good candidate for the gene behind the major ps locus of potato [14].

The fact that Atps1 mutations only affect male meiosis points to a difference in regulation between male and female 2n gametes production. This phenomenon was previously described for mutations that had a specific impact on either male or female meiosis [12],[23],[55],[56],[58]. In the case of parallel spindles, it may stem from the 3-dimension organization of the spores (e.g. tetrahedron in male vs linear or multiplanar arrays in female [59]).

The AtPS1 protein has two domains, a FHA (ForkHead Associated) domain, a phosphopeptide recognition domain found in many regulatory proteins and a PINc domain, which is found in proteins involved in RNA processing [48]. In fungi/metazoa, the AtPS1 PINc domain shows highest similarity with the PINc domains of the Swt1/ C1orf26/ CG7206 and SMG6 protein families followed by SMG5, Dis3 and others. The mammalian C1orf26 and Drosophila CG7206 genes encode related proteins of unknown function, but Interestingly both are overexpressed in testis and ovaries, which is consistent with a putative meiotic role [60],[61]. SMG6 is an essential component of the Nonsense Mediated RNA Decay (NMD) machinery that degrades mRNAs containing premature translation termination codons. SMG6 also plays a role in RNAi [45],[48]. The SMG6 PINc domain has RNA degradation activity [46]. These features suggest that AtPS1 plays a regulatory function, perhaps via RNA decay, which may directly control the orientation of metaphase plates/spindles or be related to meiotic cell cycle control. There is growing evidence that NMD and its components have important functions in various cellular processes, including the cell-cycle [62]. A link between RNA decay and the control of meiosis progression was recently suggested because SMG7, which is a NMD essential component, is involved in progression through meiotic anaphase II in Arabidopsis [63].

Further studies involving AtPS1 should shed light on the poorly understood process of meiosis II. The isolation of a gene involved in 2n gamete production has important implications for deciphering meiosis mechanisms, as well as potentially fundamental applications in evolution studies and plant breeding programmes.

Materials and Methods

Growth Conditions

Arabidopsis plants were cultivated as described in [64]. For germination assays and cytometry experiments Arabidopsis were cultivated in vitro on Arabidopsis medium [65] at 21°C with a 16h day/8h night photoperiod and 70% hygrometry.

Genetic Analysis

The Atps1-1 (SALK_078818) and Atps1-2 (WiscDsLox342F09) lines were obtained from the European Arabidopsis stock centre [39]. The Atps1-3 (FLAG_456A09) insertion is from the Versailles T-DNA collection[35]. Plants were genotyped by PCR (30 cycles of 30 s at 94°C, 30 s at 56°C and 1 min at 72°C) using two primer pairs. For each line the first pair designated is specific to the wild type allele and the second pair is specific to the T-DNA insertion.


Genetic markers used to genotype Atps1-1/Atps1-3×Ler F1 triploid and diploid plants (40 cycles of 20 s at 94°C, 20 s at Tm and 30 s at 72°C): Microsatellite msat1.29450 (located on chromosome I at position 29450001) was amplified (Tm = 57°C) using 5′TCCTTTCATCTTAATATGC3′ and 5′TCTGTCCACGAATTATTTA3′ primers. Microsatellite Msat4.35 (Tm = 58°C) (located on chromosome 4 at position 7549125) was amplified using 5′CCCATGTCTCCGATGA3′ and 5′GGCGTTTAATTTGCATTCT3′ primers. Microsatellite NGA151 (Tm = 58°C) (located on chromosome 5 at position 4669932) was amplified using 5′GTTTTGGGAAGTTTTGCTGG3′ and 5′CAGTCTAAAAGCGAGAGTATGATG3′ primers. The 2 primer pairs specific for the Atps1-1 and Atps1-3 TDNA borders were used as a centromeric marker of the chromosome 1. CAPS markers Seqf16k23 (physical position: 14481813) and CAPSK4 51 (physical position: 5078201) were used as centromeric markers for chromosome 1 and 4, respectively. CAPS Seqf16k23 was amplified (Tm = 60°C) using 5′GAGGATACCTCTTGCTGATTC3′ and 5′CCTGGCCTTAGGAACTTACTC3′ primers and observed after TaqI digestion. CAPS CAPSK4 51 was amplified (Tm = 60°C) using 5′CAATTTGTTACCAGTTTTGCAG3′ and 5′TGAGTTTGGTTTTTTGTTATTAGC3′ primers and observed after MnlI digestion.

Cytology and Flow Cytometry

Final meiotic products were observed as describe in [28] and viewed with a conventional light microscope with a 40× dry objective. Chromosomes spreads and observations were carried out using the technique described in [33]. The DNA fluorescence of spermatic pollen nuclei was quantified using open LAB 4.0.4 software. For each nucleus the surrounding background was calculated and subtracted from the global fluorescence of the nucleus. Meiotic spindles were observed according to the protocol described in [55] except that the DNA was counter-stained with DAPI. Observations were made using an SP2 Leica confocal microscope. Images were acquired with a 63× water objective in xyz and 3D reconstructions were made using Leica software. Projections are shown. Cells were imaged at excitation 488 nm and 405 nm with AlexaFluor488 and DAPI respectively. Arabidopsis genome sizes were measured as described in [66] using tomato Lycopersicon esculentum cv “Montfavet” as the standard. (2C = 1.99 pg, %GC = 40.0%).


Arabidopsis total RNA was extracted using the QUIAGEN RNA kit.

Reverse transcription was done on 5 µg of total RNA using oligo(dT)18 as primer. The RevertAid M-MuLV Reverse Transcriptase enzyme (Fermentas) was used according to the manufacturer's instruction. RT-PCR was carried out on 1 µl of cDNA using the pAtpsF and pAtpsR primers and the following PCR conditions: 30 cycles of 30 s at 94°C, 30 s at 56°C and 1 min at 72°C.


We would like to thank Mathilde Grelon and Christine Mézard for their useful comments on the manuscript. We also thank Olivier Grandjean for his technical assistance. LKG scientific editing and translation improved the English in the manuscript.

Author Contributions

Conceived and designed the experiments: IdE EJ RM. Performed the experiments: IdE SJ NF OC MN RM. Analyzed the data: IdE MN EJ RM. Contributed reagents/materials/analysis tools: MS. Wrote the paper: IdE EJ RM.


  1. 1. Otto SP (2007) The evolutionary consequences of polyploidy. Cell 131: 452–462.
  2. 2. Otto SP, Whitton J (2000) Polyploid incidence and evolution. Annu Rev Genet 34: 401–437.
  3. 3. Masterson J (1994) Stomatal Size in Fossil Plants: Evidence for Polyploidy in Majority of Angiosperms. Science 264: 421–424.
  4. 4. Cui L, Wall PK, Leebens-Mack JH, Lindsay BG, Soltis DE, et al. (2006) Widespread genome duplications throughout the history of flowering plants. Genome Res 16: 738–749.
  5. 5. Gaut BS, Doebley JF (1997) DNA sequence evidence for the segmental allotetraploid origin of maize. Proc Natl Acad Sci U S A 94: 6809–6814.
  6. 6. Lysak MA, Koch MA, Pecinka A, Schubert I (2005) Chromosome triplication found across the tribe Brassiceae. Genome Res 15: 516–525.
  7. 7. Schlueter JA, Lin JY, Schlueter SD, Vasylenko-Sanders IF, Deshpande S, et al. (2007) Gene duplication and paleopolyploidy in soybean and the implications for whole genome sequencing. BMC Genomics 8: 330.
  8. 8. Blanc G, Barakat A, Guyot R, Cooke R, Delseny M (2000) Extensive duplication and reshuffling in the Arabidopsis genome. Plant Cell 12: 1093–1101.
  9. 9. Adams KL, Wendel JF (2005) Polyploidy and genome evolution in plants. Curr Opin Plant Biol 8: 135–141.
  10. 10. Winge O (1917) The chromosomes. Their numbers and general importance. Carlsberg Lab Copenhagen CR trav 13: 131–175.
  11. 11. Harlan JR, De Wet JMJ (1975) On O. Winge and a prayer: the origins of polyploids. The Botanical review 41: 361–690.
  12. 12. Bretagnolle F, Thompson JD (1995) Gametes with the somatic chromosome number: mechanisms of their formation and role in the evolution of autopolyploid plants. New Phytologist 129: 1–22.
  13. 13. Ramsey J, Schemske DW (1998) Pathways, mechanisms and rates of polyploid formation in flowering plants. Annual Reviews of Ecology and Systematics 29: 467–501.
  14. 14. Carputo D, Frusciante L, Peloquin SJ (2003) The role of 2n gametes and endosperm balance number in the origin and evolution of polyploids in the tuber-bearing Solanums. Genetics 163: 287–294.
  15. 15. De Wet JMJ (1968) Diploid-Tetraploid-Haploid Cycles and the Origin of Variability in Dichanthium Agamospecies. Evolution 22: 394–397.
  16. 16. Peloquin SJ, Boiteux LS, Carputo D (1999) Meiotic mutants in potato. Valuable variants. Genetics 153: 1493–1499.
  17. 17. Ramana MS, Jacobsen E (2003) Relevance of sexual polyploidization for crop improvment-A review. Euphytica 133: 3–18.
  18. 18. Veilleux R (1985) Diploid and polyploid gametes in crop plants: mechanisms of formation and utilization in plant breeding. Plant Breeding Reviews 3: 252–288.
  19. 19. Fukuda y (1927) Cytological studies on the development of pollen grains in different races of solanum. Bot Mag Tokyo 41: 459–474.
  20. 20. Ramana MS (1979) A re-examination of the mechanisms of 2n gamete formation in potato and its implications for breeding. Euphytica 28: 537–561.
  21. 21. Kato A, Birchler JA (2006) Induction of tetraploid derivatives of maize inbred lines by nitrous oxide gas treatment. J Hered 97: 39–44.
  22. 22. De storme N, Van Labele M, Geelen D (2007) Formation of unreduced pollen in Arabidopsis thaliana. Commun Agric Appl Biol Sci 72: 159–163.
  23. 23. Barrell PJ, Grossniklaus U (2005) Confocal microscopy of whole ovules for analysis of reproductive development: the elongate1 mutant affects meiosis II. Plant J 43: 309–320.
  24. 24. Toufighi K, Brady SM, Austin R, Ly E, Provart NJ (2005) The Botany Array Resource: e-Northerns, Expression Angling, and promoter analyses. Plant J 43: 153–163.
  25. 25. Schmid M, Davison TS, Henz SR, Pape UJ, Demar M, et al. (2005) A gene expression map of Arabidopsis thaliana development. Nat Genet 37: 501–506.
  26. 26. Mercier R, Jolivet S, Vezon D, Huppe E, Chelysheva L, et al. (2005) Two meiotic crossover classes cohabit in Arabidopsis: one is dependent on MER3,whereas the other one is not. Curr Biol 15: 692–701.
  27. 27. Couteau F, Belzile F, Horlow C, Grandjean O, Vezon D, et al. (1999) Random chromosome segregation without meiotic arrest in both male and female meiocytes of a dmc1 mutant of Arabidopsis. Plant Cell 11: 1623–1634.
  28. 28. Azumi Y, Liu D, Zhao D, Li W, Wang G, et al. (2002) Homolog interaction during meiotic prophase I in Arabidopsis requires the SOLO DANCERS gene encoding a novel cyclin-like protein. Embo J 21: 3081–3095.
  29. 29. Kerzendorfer C, Vignard J, Pedrosa-Harand A, Siwiec T, Akimcheva S, et al. (2006) The Arabidopsis thaliana MND1 homologue plays a key role in meiotic homologous pairing, synapsis and recombination. J Cell Sci 119: 2486–2496.
  30. 30. Panoli AP, Ravi M, Sebastian J, Nishal B, Reddy TV, et al. (2006) AtMND1 is required for homologous pairing during meiosis in Arabidopsis. BMC Mol Biol 7: 24.
  31. 31. Schommer C, Beven A, Lawrenson T, Shaw P, Sablowski R (2003) AHP2 is required for bivalent formation and for segregation of homologous chromosomes in Arabidopsis meiosis. Plant J 36: 1–11.
  32. 32. Higgins JD, Vignard J, Mercier R, Pugh AG, Chris F, et al. (2008) AtMSH5 partners AtMSH4 in the class I meiotic crossover pathway in Arabidopsis thaliana, but is not required for synapsis. Plant J.
  33. 33. Grelon M, Vezon D, Gendrot G, Pelletier G (2001) AtSPO11-1 is necessary for efficient meiotic recombination in plants. Embo J 20: 589–600.
  34. 34. Alonso JM, Stepanova AN, Leisse TJ, Kim CJ, Chen H, et al. (2003) Genome-wide insertional mutagenesis of Arabidopsis thaliana. Science 301: 653–657.
  35. 35. Samson F, Brunaud V, Balzergue S, Dubreucq B, Lepiniec L, et al. (2002) FLAGdb/FST: a database of mapped flanking insertion sites (FSTs) of Arabidopsis thaliana T-DNA transformants. Nucleic Acids Res 30: 94–97.
  36. 36. Sessions A, Burke E, Presting G, Aux G, McElver J, et al. (2002) A high-throughput Arabidopsis reverse genetics system. Plant Cell 14: 2985–2994.
  37. 37. Woody ST, Austin-Phillips S, Amasino RM, Krysan PJ (2007) The WiscDsLox T-DNA collection: an arabidopsis community resource generated by using an improved high-throughput T-DNA sequencing pipeline. J Plant Res 120: 157–165.
  38. 38. Takahashi N, Kuroda H, Kuromori T, Hirayama T, Seki M, et al. (2004) Expression and interaction analysis of Arabidopsis Skp1-related genes. Plant Cell Physiol 45: 83–91.
  39. 39. Scholl RL, May ST, Ware DH (2000) Seed and molecular resources for Arabidopsis. Plant Physiol 124: 1477–1480.
  40. 40. Altschul SF, Madden TL, Schaffer AA, Zhang J, Zhang Z, et al. (1997) Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25: 3389–3402.
  41. 41. Marchler-Bauer A, Bryant SH (2004) CD-Search: protein domain annotations on the fly. Nucleic Acids Res 32: W327–331.
  42. 42. Ponting CP, Schultz J, Milpetz F, Bork P (1999) SMART: identification and annotation of domains from signalling and extracellular protein sequences. Nucleic Acids Res 27: 229–232.
  43. 43. Mulder NJ, Apweiler R (2008) The InterPro database and tools for protein domain analysis. Curr Protoc Bioinformatics Chapter 2: Unit 2 7:
  44. 44. Durocher D, Jackson SP (2002) The FHA domain. FEBS Lett 513: 58–66.
  45. 45. Clissold PM, Ponting CP (2000) PIN domains in nonsense-mediated mRNA decay and RNAi. Curr Biol 10: R888–890.
  46. 46. Glavan F, Behm-Ansmant I, Izaurralde E, Conti E (2006) Structures of the PIN domains of SMG6 and SMG5 reveal a nuclease within the mRNA surveillance complex. Embo J 25: 5117–5125.
  47. 47. Rother S, Clausing E, Kieser A, Strasser K (2006) Swt1, a novel yeast protein, functions in transcription. J Biol Chem 281: 36518–36525.
  48. 48. Domeier ME, Morse DP, Knight SW, Portereiko M, Bass BL, et al. (2000) A link between RNA interference and nonsense-mediated decay in Caenorhabditis elegans. Science 289: 1928–1931.
  49. 49. Alexander MP (1969) Differential staining of aborted and nonaborted pollen. Stain Technol 44: 117–122.
  50. 50. Scott RJ, Spielman M, Bailey J, Dickinson HG (1998) Parent-of-origin effects on seed development in Arabidopsis thaliana. Development 125: 3329–3341.
  51. 51. Ahloowalia (1972) Germination in vitro of ryegrass pollen grains. Euphytica 22: 575–581.
  52. 52. Susin I, Alvarez M (1996) Fertility and pollen tube growth in polyploid melons (Cucumis melo L.). Euphytica 93: 369–373.
  53. 53. Stacey NJ, Kuromori T, Azumi Y, Roberts G, Breuer C, et al. (2006) Arabidopsis SPO11-2 functions with SPO11-1 in meiotic recombination. Plant Journal 48: 206–216.
  54. 54. Agashe B, Prasad CK, Siddiqi I (2002) Identification and analysis of DYAD: a gene required for meiotic chromosome organisation and female meiotic progression in Arabidopsis. Development 129: 3935–3943.
  55. 55. Mercier R, Vezon D, Bullier E, Motamayor JC, Sellier A, et al. (2001) SWITCH1 (SWI1): a novel protein required for the establishment of sister chromatid cohesion and for bivalent formation at meiosis. Genes Dev 15: 1859–1871.
  56. 56. Ravi M, Marimuthu MP, Siddiqi I (2008) Gamete formation without meiosis in Arabidopsis. Nature 451: 1121–1124.
  57. 57. Mok DWS, Peloquin SJ (1975) The inheritance of three mechanisms of diplandroid (2n pollen) formation in diploid potatoes. Heredity 35: 295–302.
  58. 58. Glover J, Grelon M, Craig S, Chaudhury A, Dennis E (1998) Cloning and characterization of MS5 from Arabidopsis: a gene critical in male meiosis. Plant J 15: 345–356.
  59. 59. Schneitz K, Hülskamp M, Pruitt R (1995) Wild-type ovule development in Arabidopsis thaliana: a light microscope study of cleared whole-mount tissue. Plant Journal 7: 731–749.
  60. 60. Su AI, Cooke MP, Ching KA, Hakak Y, Walker JR, et al. (2002) Large-scale analysis of the human and mouse transcriptomes. Proc Natl Acad Sci U S A 99: 4465–4470.
  61. 61. Chintapalli VR, Wang J, Dow JA (2007) Using FlyAtlas to identify better Drosophila melanogaster models of human disease. Nat Genet 39: 715–720.
  62. 62. Isken O, Maquat LE (2008) The multiple lives of NMD factors: balancing roles in gene and genome regulation. Nat Rev Genet.
  63. 63. Riehs N, Akimcheva S, Puizina J, Bulankova P, Idol RA, et al. (2008) Arabidopsis SMG7 protein is required for exit from meiosis. J Cell Sci 121: 2208–2216.
  64. 64. Vignard J, Siwiec T, Chelysheva L, Vrielynck N, Gonord F, et al. (2007) The interplay of RecA-related proteins and the MND1-HOP2 complex during meiosis in Arabidopsis thaliana. PLoS Genet 3: 1894–1906.
  65. 65. Estelle MA, Somerville C (1987) Auxin-resistant mutants of Arabidopsis thaliana with an altered morphology. Mol Gen Genet 206: 200–206.
  66. 66. Marie D, Brown SC (1993) A cytometric exercise in plant DNA histograms, with 2C values for 70 species. Biol Cell 78: 41–51.
  67. 67. Biegert A, Soding J (2008) De novo identification of highly diverged protein repeats by probabilistic consistency. Bioinformatics 24: 807–814.
  68. 68. George RA, Heringa J (2000) The REPRO server: finding protein internal sequence repeats through the Web. Trends Biochem Sci 25: 515–517.
  69. 69. Promponas VJ, Enright AJ, Tsoka S, Kreil DP, Leroy C, et al. (2000) CAST: an iterative algorithm for the complexity analysis of sequence tracts. Complexity analysis of sequence tracts. Bioinformatics 16: 915–922.
  70. 70. Altschul SF, Koonin EV (1998) Iterated profile searches with PSI-BLAST–a tool for discovery in protein databases. Trends Biochem Sci 23: 444–447.
  71. 71. Soding J, Biegert A, Lupas AN (2005) The HHpred interactive server for protein homology detection and structure prediction. Nucleic Acids Res 33: W244–248.
  72. 72. Takeshita D, Zenno S, Lee WC, Saigo K, Tanokura M (2007) Crystal structure of the PIN domain of human telomerase-associated protein EST1A. Proteins 68: 980–989.