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Keywords:

  • MYB transcription factor;
  • male sterility;
  • dehiscence;
  • ms35;
  • Arabidopsis thaliana;
  • anther development

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

A male sterile mutant with a defect in anther dehiscence was identified in an Arabidopsis thaliana population mutagenized with the Zea mays transposon En-1/Spm. Mutants produce viable pollen that can fertilize when released mechanically from the anthers. Mutant stamens are of normal size and shape, but lack cell wall fortifications in the endothecial cell layer of the anther, which are required for the dehiscence process. The mutant phenotype was shown to be caused by a transposon insertion in AtMYB26, disrupting the putative DNA-binding domain of this R2R3-type MYB transcription factor. RT-PCR revealed that expression of AtMYB26 is restricted to inflorescences. Sterility was shown to be stable under several environmental conditions. The high stability of the sterile phenotype, together with the fact that pollen is functional, makes AtMYB26 and its orthologs a valuable tool for manipulating male fertility in higher plants.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Many different developmental steps are required to achieve male fertility in higher plants. The ontogeny of the male organs, filaments and anthers has to be specified, and the male gametophyte has to develop. Finally, pollen has to be released. Mutants defective in either of these processes provide valuable tools for studying male sterility in higher plants. In Arabidopsis thaliana, mutants have been described that are defective in meiosis or post-meiotic pollen development and therefore do not develop functional pollen grains (Chaudhury et al., 1994; Dawson et al., 1993; He and Mascarenhas, 1998; He et al., 1996; Preuss et al., 1993; Regan and Moffatt, 1990; Sanders et al., 1999; Taylor et al., 1998; Wilson et al., 2001; Yang et al., 1999). Several male sterile mutants affected in the mechanism or timing of the release of pollen from the anthers have been reported (Dawson et al., 1993, 1999; Ishiguro et al., 2001; von Malek et al., 2002; Park et al., 1996, 2002; Sanders et al., 1999, 2000; Stintzi and Browse, 2000). As pollen is at least partially functional in some of these mutants, sterility of such plants can be overcome by mechanically opening the pollen sacs, making these mutants especially attractive for an application in breeding and hybrid seed programmes.

Dehiscence of the anther is a multistage process. Between the locules of the anther, the septum and stomium cells have to differentiate and then undergo a degeneration programme allowing breakage of the anther wall. Furthermore, cells of the endothecium have to enlarge and their walls have to be strengthened to allow proper dehiscence (Dawson et al., 1999; Sanders et al., 1999). In self-pollinating species like A. thaliana, these processes have to occur at the correct time, when the female organ pistil is receptive and in the right place, so that the filament of the stamen is at the proper height for pollen delivery onto the stigma.

To date, all genes affecting anther dehiscence that have been characterized are involved in the jasmonic acid (JA) pathway. Mutants of these genes display defects in filament elongation and timing of anther dehiscence, as well as reduced pollen viability. The mutant coi1 (Xie et al., 1998) is JA insensitive, while the triple mutant fad3/fad7/fad8 (Feys et al., 1994) and the mutants opr3/dde1 (Sanders et al., 2000; Stintzi and Browse, 2000), dad1 (Ishiguro et al., 2001) and aos/dde2-2 (von Malek et al., 2002; Park et al., 2002) are defective in JA synthesis.

Three other mutants with defects in anther dehiscence have been described in A. thaliana, but the cloning of the corresponding genes has not been reported previously. In non-dehiscense1 (Sanders et al., 1999), filaments elongate properly and anthers enter the dehiscence programme, including expansion of the endothecium and degeneration of the septum region. Later, however, the endothecium and connective tissue degenerate completely, resulting in anthers filled with functional pollen surrounded by an anther wall consisting exclusively of the epidermal cell layer. In ms35 (Dawson et al., 1999), formerly referred to as msH (Dawson et al., 1993), filaments also elongate properly, but functional pollen grains are not released from the anthers, although the stomium is cleaved. This defect is associated with a lack of secondary thickening in the endothecium cells. Similarly, in the mutant por1 (Park et al., 1996), anthers fail to release pollen, although the stomium is cleaved. However, in contrast to ms35, stamen filaments show a delayed growth rate and pollen grains are not functional.

Here, we present the isolation of an anther-dehiscence mutant from a population of transposon-tagged A. thaliana insertion lines and the identification of the responsible mutant gene. The mutant phenotype is caused by a transposon insertion in the gene AtMYB26 encoding a putative R2R3-type MYB-transcription factor. As in ms35, the filaments elongate properly but the anthers, although cleaved at the stomium, fail to release the functional pollen grains. The dehiscence defect is associated with a defect in fortification of the endothecium cell walls. Performing crosses with ms35 has shown that myb26 is allelic to ms35.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Identification of a mutant with non-dehiscent anthers

Screening an A. thaliana population (Wisman et al., 1998a,b) mutagenized with the maize transposon En-1/Spm (Schwarz-Sommer et al., 1985) for morphological changes, we have identified a sterile mutant developing only small and empty siliques (Figure 1a). Mutant plants produce normal siliques containing seeds when fertilized with wild-type pollen, indicating that female fertility is not affected. Flowers of mutant plants are of normal size and shape, and the filaments of the stamen elongate as in wild type (Figure 1b). Anthers contain pollen and develop to normal size but fail to dehisce (Figure 1b,c). Scanning electron micrographs revealed that mutant and wild-type pollen grains are indistinguishable (Figure 1d). Furthermore, if the mutant anthers were opened mechanically and the released pollen grains were used for self-pollination of mutant plants, siliques containing seeds were obtained. Pollination with mutant plants was more difficult than with the wild type because pollen grains tended to stick together, but all seeds obtained gave rise to sterile plants, showing that the self-pollination was successful. In addition to the described male sterility phenotype, mutants show delayed apical senescence, a higher number of inflorescences and flowers and the formation of terminal aberrant flowers co-segregating with sterility. Such defects have been observed in various other male sterile mutants before and are thought to be a result of the lack of fertilization rather than the pleiotropic effects of a particular mutant (Chaudhury et al., 1994).

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Figure 1. Description of the phenotype.

Comparison of mutant (myb26, left) and wild-type (wt, right) plants.

(a) Mutants are sterile and thus develop only small siliques without seeds, while wild-type plants of the same age develop swollen siliques containing seeds.

(b) Flowers of mutant plants are of normal size; the filaments of stamen elongate properly, but anthers fail to dehisce.

(c) SEM picture of anthers of open flowers (200-fold magnification). While anthers of mutant plants are still closed, anthers of the wild type are fully dehisced and have released the pollen grains.

(d) SEM of pollen grains (2000-fold magnification). Pollen of the mutant are indistinguishable from wild-type pollen.

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Anther development

The anthers of sterile plants appear as in the wild type and develop to normal size and shape, but directly prior to dehiscence mutant anthers appear less swollen (Figure 2a,b). Cross-sections of mutant inflorescences revealed that anther development is indistinguishable from that in the wild type during phase one and early stages of phase two (as defined by Sanders et al., 1999; Figure 2c,d). Later in phase two of anther development, which includes the dehiscence programme, differences can be seen. While maturation of pollen grains continues and the disappearance of the tapetum and middle layer appears normal, expansion of the endothecium layer, which occurs in wild-type anthers, cannot be observed in the mutant (Figure 2i,j). At later stages, degradation of the septum and breakage of the stomium can be seen in the mutant as in wild type (Figure 2e,f), but the subsequent shrinkage of the anther walls resulting in the release of pollen grains does not take place.

image

Figure 2. Anther development.

Anthers from the largest flower bud of myb26-2(a) appear less swollen than wild-type anthers of the same stage (b). Toluidine-blue-stained cross-sections of anthers of myb26-2(c) and the wild type (d), corresponding to stage 10 according to Sanders et al. (2000), do not reveal differences between myb26-2 and wild type. At stage 13, breakage of the stomium can be seen in myb26-2(e) and in the wild type (f) but the endothecium cells appear less swollen in myb26-2. UV illumination of cleared anthers of myb26-2(g) and the wild type (h) taken from open flowers revealed a net-like structure of lignified material in wild type, which is lacking in myb26-2. The anther of myb26-2 is still closed and contains pollen grains, which are visible as a result of the autofluorescence of sporopollenin, while the wild-type anther is dehisced and has released the pollen grains. Lignified vascular tissue is visible in myb26-2 and wild type. Phloroglucinol-stained cross-sections of anthers corresponding to stage 12 reveal that the endothecium cells of myb26-2(i) are not swollen while the endothecium cells of wild type (j) appear swollen and show rigid cell walls. Staining of cellulosic material with calcofluor white using cross-sections of anthers corresponding to stage 11 reveal bar-like cellulosic thickenings in the walls of wild type (l), which are not visible in myb26-2(k).

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UV-illuminated cleared wild-type anthers usually reveal a net-like structure of autofluorescent material, indicating lignification of the anther wall (Figure 2h), which could be seen first in bud no. 4 (buds were numbered acropetally starting with the oldest bud). These structures could not be found in the mutant (Figure 2g). Cross-sections of flower buds stained with calcofluor white revealed that cellulose deposition on radial cell walls of endothecium cells, as seen in the wild type (Figure 2l), is also missing in sterile plants (Figure 2k).

Cloning of AtMYB26

Segregation analysis of sister plants of the original isolate showed a 3 : 1 ratio of fertile to sterile plants, indicating that the phenotype is caused by a single recessive mutation. As the mutant was isolated from a transposon-mutagenized population, it was expected to be tagged by an En-1 insertion. Southern analyses using the 5′-end of the En-1 transposon as probe were therefore performed with 14 individuals (five mutants, five homozygous wild types and four heterozygous plants according to segregation analysis of progeny), revealing that the individuals carried between six and 10 En-1 insertions, one of which co-segregated with the mutation causing sterility (Figure 3a).

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Figure 3. Identification of the mutant locus.

Individuals from a line segregating for sterility representing five homozygous individuals carrying the wild-type allele (wt/wt), four heterozygous plants (wt/myb26) and five homozygous mutants (myb26/myb26) and a Col-0 wild-type control (C) were investigated by Southern analysis using genomic DNA cleaved with EcoRI.

(a) Segregation analysis: the blot was hybridized with an En-1-left end-specific probe. One band (indicated by an arrow) is only present in the individuals carrying at least one copy of the mutant allele showing co-segregation of this En-1 insertion with the mutant locus.

(b) Co-segregation of an En-1 insertion site with the mutant phenotype. Hybridization was performed with an isolated En-1 flanking region. All individuals not carrying the mutant allele show a single band also present in Col-0 and therefore represent the wild-type allele of the isolated locus, while in the mutants this fragment is barely visible. Instead, a smaller fragment, which also hybridizes with the En-1-probe, gives a signal. In the heterozygous plants, both signals are present. The faint signal corresponding to the wild-type allele, which is visible in the mutants, represents excision events of the active transposon from the insertion site.

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The flanking regions of eight different En-1 insertions were isolated from one individual sterile plant. Using these regions as probes in gel blot experiments, one of them could be shown to correspond to the transposon insertion co-segregating with the mutant phenotype (Figure 3b).

The En-1 used to generate our population is an autonomous element encoding its own transposase. Therefore, the causal relation of the transposon insertion in the isolated locus and the sterile phenotype could be proven by investigating revertants, which could be found easily by screening sterile individuals for the rare appearance of fully developed siliques containing fertile seeds. These seeds gave rise to progeny segregating for sterility. Southern blot analyses of these plants using the isolated flanking DNA as a probe revealed that restoration of fertility was correlated in all plants with a loss of the insertion in at least one allele of the isolated locus (data not shown), indicating that this particular insertion is responsible for the anther dehiscence defect.

The flanking region of the insertion causing the male sterile phenotype was sequenced, revealing that the insertion is located within the coding region of At3g13890, also called AtMYB26 (Romero et al., 1998; Stracke et al., 2001), encoding a putative MYB-transcription factor. The En-1 insertion is located behind the 14th codon of AtMYB26, thus disrupting the putative DNA-binding domain (Figure 4a). Our mutant will therefore be referred to as myb26-2.

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Figure 4. Genomic organization of the AtMYB26-locus and its mutant alleles.

(a) AtMYB26 consists of three exons (I, II, III). In myb26-2, the open reading frame is disrupted by an En-1 insertion within the first exon.

(b) Sequences of the AtMYB26 5′-regions of the En insertion allele myb26-2 and stable myb26-mutants created by imprecise excision of the En-1 element, which were named myb26-3, myb26-4 and myb26-5.

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Isolation of stable mutant alleles

As a result of the fact that the transposon inserted within AtMYB26 is still active, it was necessary to isolate a stable mutant. As excision of the transposon from an integration site is often not precise leaving a footprint behind (Cardon et al., 1993), we screened for such events by crossing a sterile plant with Col-0. The resulting F1 generation was subsequently screened via PCR for the loss of the En-1 insertion within AtMYB26. Of 80 individuals tested, five were shown to have lost the En-1 insertion. The AtMYB26 5′-region of these individuals was amplified by PCR and sequenced revealing overlapping sequences in four cases, which indicate that excision of the En-1 created a deletion or insertion. In one case, the wild-type sequence had been restored.

Progeny of these plants were grown and sterile individuals found among the progeny of three individuals with a putative imprecise excision, while progeny of the individual that had restored the wild-type sequence were fertile, confirming the causal relationship between a mutation in AtMYB26 and the sterile phenotype. The AtMYB26 5′-regions of these sterile plants were amplified by PCR and sequenced, revealing that in one case a 9 bp deletion had occurred which led to the loss of three amino acids in the MYB domain. In the other individuals, insertions of, respectively, four and two nucleotides could be found creating a stop codon in the open reading frame behind the 14th codon (Figure 4b).

Sequence analysis

AtMYB26 encodes a protein of 358 amino acids, which in its N-terminal part shows homology to a MYB DNA-binding domain. This domain generally comprises up to three repeats of about 53 amino acids, each forming a helix-turn-helix structure. AtMYB26, like most plant MYB factors, contains only two of these repeats covering amino acids 12–115 and thus belongs to the R2R3-type of MYB transcription factors. The R2R3-type MYB genes have been classified into 22 groups according to conserved amino acid motifs present C-terminal to the MYB domain. Classification into similar groups often also reflects a functional conservation. AtMYB26 groups with AtMYB103 and AtMYB67 (Stracke et al., 2001), displaying highest overall identity (49.1%) with AtMYB67 at the amino acid level. However, AtMYB26 and AtMYB67 exhibit only 79.8% identity within the MYB domain, whilst 87.3% identity within this domain can be found with a putative transcription factor from rice (gene: P0456F08.4, protein id: BAB39404.1). The overall identity with this protein, however, is only 43.9%. The function of these putative MYB transcription factors is not known.

Expression studies

To determine the expression pattern of AtMYB26, Northern blot analysis was performed. Using as probe the AtMYB26 cDNA lacking the region encoding the conserved MYB domain, no transcript was detected, although RNAs from different organs were used, indicating that AtMYB26 is weakly expressed. Therefore, we performed RT-PCR to examine the expression of AtMYB26. Total RNA was extracted from stem, inflorescences, cauline leaves, young and old rosette leaves, seedlings and roots of A. thaliana Col-0 wild-type plants and reverse transcribed into cDNA. Both conventional RT-PCR and real-time RT-PCR amplified the AtMYB26 transcript only in inflorescences (Figure 5a,b). RT-PCR performed on flower buds of different stages revealed that expression is not detected in young buds (buds prior to pollen mitotic division) and first appears in buds 3–5 of the inflorescence (when buds are numbered acropetally starting with the oldest bud) containing anthers in the stages of pollen mitotic divisions and tapetal degeneration. Transcript can still be found at a lower level in older buds and even open flowers (Figure 5c).

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Figure 5. Relative quantification of the transcript in different organs of Col-0 wild-type plants by PCR.

(a) RT-PCR. Total RNA was isolated from different plant organs and used for a one-step RT-PCR. Ten microlitres of each of the 50 µl PCR assays was loaded on a 1% agarose gel.

(b) Real-time RT-PCR. Real-time PCR was performed with AtMYB26-specific probe and primers in 40 cycles. The relative amounts of transcript, compared to the transcript level in inflorescences, were calculated using the values given in the table. The ct-values of the different plant organs represent the medium of a triplicate (SD of the three values shown). To obtain a relative view of the AtMYB26 expression in the various organs, the results were normalized to the ct-value in inflorescences: Δ = ct-inflorescence − ct-x (x: ct-value in another organ). As the amount of PCR product should duplicate after each cycle, the relative amount of AtMYB26 transcript was then calculated as 2Δ and the result converted into percentages.

(c) RT-PCR using flower buds of different stages (buds were numbered acropetally starting with the oldest bud): young buds correspond to buds 6-centre of inflorescence (buds prior to pollen mitotic division); middle buds represent buds 3–5 of inflorescence (pollen mitotic divisions/tapetal degeneration); old buds represent buds 1 and 2 (unopened petals visible, tricellular pollen). As control, RT-PCR was performed on the same samples using actin-specific primers.

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In order to determine the temporal and spatial expression patterns within inflorescences in more detail, in situ hybridizations were performed on inflorescences, which, however, did not give a signal confirming the weak expression level of AtMYB26.

Stability test

The stability of the sterile phenotype was tested using plants carrying the stable allele myb26-4 (see Figure 4b) by growing plants at 22°C and subjecting them for 4 days to low (10°C) and high (30°C) temperatures. Plants were visually screened for the appearance of siliques containing seeds. Only 10 filled siliques were found on a total of 45 plants with each plant carrying more than 2000 empty siliques. At 22°C, only one silique containing seeds was found (representing 0.003% of siliques). A slightly increased proportion of siliques containing seeds was found after subjecting the plants to 30°C (three siliques, 0.01% of scored siliques) and 10°C (six siliques, 0.02% of scored siliques). Although this might indicate an effect of temperature on the stability of the phenotype, it can still be considered high under different conditions, making AtMYB26 and its homologues valuable tools for manipulating male fertility in higher plants.

Jasmonate insensitivity of myb26

Several mutants affected in anther dehiscence are affected in synthesis or sensing of jasmonate (Ishiguro et al., 2001; von Malek et al., 2002; Park et al., 2002; Sanders et al., 2000; Stintzi and Browse, 2000). Although the phenotypes described for these mutants, which show defects in filament elongation and timing of dehiscence as well as defective pollen, differ in many respects from the phenotype of myb26, we tested if the stable mutant myb26-5 might be rescued by application of jasmonic acid and methyl jasmonate, respectively. As a control, we used the mutant dde2-2 described by von Malek et al. (2002). While dde2-2 mutants regained fertility upon treatment with both jasmonic acid and methyl jasmonate, the sterile phenotype of myb26-5 could not be overcome, showing that synthesis of jasmonic acid is not affected in myb26.

MS35 gene identification

The phenotype of myb26 strongly resembles that described for the male sterile mutant ms35 (Dawson et al., 1999). The region around the ms35 mutations has been mapped using recombinants between hy2-ms35 (breakpoints at approximately 0.18 cm intervals) and ms35-gl1 (every 1.0 cm; Dawson et al., 1999; Sozen, 2000). The MS35 gene was shown to co-segregate with the ATHCHIB and nga162 markers, and to map 0.18 cm away from act11, towards hy2 (Sozen, 2000). Using the recombinant inbred map (Lister and Dean, 1993), ATHCHIB and nga162 markers map to 19.1 and 20.56 cm, respectively. The AtMYB26 gene also maps to 20.56 cm and lies within 31.5 kb of nga162, suggesting that MS35 may be allelic to MYB26. Crosses were therefore performed between myb26-3 and ms35 giving rise to sterile progeny. This shows that myb26-3 is allelic to ms35, which should therefore be referred to as myb26-1.

In order to determine the mutation present in ms35, we have PCR amplified and direct sequenced the AtMYB26 region, from −3174 upstream to 500 bp beyond the transcriptional termination sequence, in Ler and the ms35 mutant. We found the coding region and downstream sequence to be identical; however, aberrations were seen in the upstream region. PCR amplifications were possible throughout this region for the Ler line. However, amplification, including long-PCR, across a 235 bp region that lies between −1288 and −1523 bp upstream of the translation start was not possible in the ms35 mutant. The sequence up to and beyond this point is identical to that seen in Ler. We therefore believe that a major chromosomal rearrangement has occurred in this region, which prevents normal MYB26 gene expression, resulting in a failure of dehiscence in the ms35 mutant. This is supported by RT-PCR data, which shows reduced levels of AtMYB26 expression in ms35 buds as compared to Ler wild type (data not shown). A chromosomal rearrangement could also explain the mapping data for ms35 (Sozen, 2000) that show suppression of recombination within the ATHCHIB-nga162 region, as no recombinants occurred in this region despite the fact that it spans 670 kb/1.46 cm and breakpoints were expected every 0.18 cm.

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

We have isolated a male sterile mutant, myb26-2, from a population of transposon-tagged A. thaliana lines. Sterility is caused by a defect in anther dehiscence, while the pollen itself is shown to be functional. The failure of dehiscence is associated with a defect in the development of the anther walls. During the late stages of anther development, the wild-type endothecium cells appear swollen and show deposition of lignified cellulosic secondary wall thickenings. In myb26-2, these cell wall fortifications are missing and the endothecium cells do not expand. The endothecium has been proposed to contribute to two steps of anther dehiscence. First, an inwardly directed force from the anther wall, which is driven by swelling of the epidermal and endothecial cells, causes the rupture of the stomium. Subsequently, an outward bending force, leading to full opening of the stomium permitting pollen release, is caused by differential shrinkage of thickened and unthickened parts of the endothecium (Keijzer, 1987). While the breakage of the stomium takes place in myb26-2, the retraction of the anther walls does not occur, suggesting that, especially, this latter process requires the endothecium cell wall fortifications as proposed by Dawson et al. (1999).

The phenotype of the myb26 mutants strongly resembles that described for the male sterile mutant ms35 (Dawson et al., 1999). Furthermore, AtMYB26 maps to the top arm of chromosome 3 at about 20.5 cm, while ms35 maps and co-localizes with markers mapping to this region (Sozen, 2000). Crosses between myb26-3 and ms35 resulted in sterile progeny, demonstrating that the two mutants are allelic. PCR amplification and sequence analysis indicate the presence of a major rearrangement in the AtMYB26 5′-region of the ms35 mutant.

The sterile phenotype of our mutant was found to be because of a mutation in AtMYB26 encoding a putative MYB-tanscription factor. MYB-transcription factors are found in all major eukaryotic groups and are characterized by one to three copies of imperfect helix-turn-helix motives designated R1, R2 and R3, which form a DNA-binding domain. In plants, MYB genes form a large family with, e.g. 136 members predicted in the A. thaliana genome by Stracke et al. (2001). One hundred and twenty-five of them, including AtMYB26, belong to the R2R3 class with two repeats, which, according to current knowledge, is plant specific (Riechmann et al., 2000). The importance of the MYB domain for the function of AtMYB26 was shown by the fact that a three-amino acid deletion within the first repeat, as found in myb26-3, is sufficient to confer the mutant phenotype.

The functions of several R2R3-type MYB transcription factors from various plant species have been characterized, and it seems that they mainly regulate plant-specific processes (Stracke et al., 2001). They have been shown to act as either transcriptional activators or repressors controlling development and determination of cell fate and identity and regulating different biosynthetic pathways like tryptophan synthesis and the phenyl propanoid pathway (for review, see Jin and Martin, 1999; Riechmann et al., 2000; Stracke et al., 2001).

AtMYB26 is the first MYB transcription factor that has been shown to be required for male fertility. However, pollen itself is viable in the loss of function mutant. Sterility is caused by anther non-dehiscence associated with a lack of cell wall fortifications consisting of lignin and cellulose material in the endothecium of the pollen sac. Therefore, the main defect seems to be a failure to properly establish the properties of this cellular layer. Other MYB factors have already been shown to control development and determination of cell fate. MIXTA from Antirrhinum majus (Glover et al., 1998; Noda et al., 1994) and GL1 and WER from A. thaliana (Lee and Schiefelbein, 1999; Marks and Feldmann, 1989; Wada et al., 1997) have been shown to confer cell-type specificity. They are required for the determination of the fate of epidermal cells being involved in the decision to form conical cells in petals, trichomes in leaf and stem and root hairs in the roots. AS1 from A. thaliana (Byrne et al., 2000) and its orthologs from other species, which are required for proper meristem function, are other examples for MYB genes involved in cell differentiation.

As, in myb26, lignification has been shown to be missing within the endothecium, it can also be assumed that AtMYB26 is involved in regulating the phenyl propanoid pathway. Several MYB transcription factors have already been shown to regulate this pathway, e.g. in A. thaliana those encoded by AtMYB4 (Jin et al., 2000), PAP1 and PAP2 (Borevitz et al., 2000), TT2 (Nesi et al., 2001) as well as AtMYB21 and AtMYB86 (Shin et al., 2002). Further data come from other plant species where MYB factors have been shown to be involved in regulating the synthesis of anthocyanin, flavonol and phlobaphene (for review, see Jin and Martin, 1999). An effect of MYB transcription factors on lignification has been shown in the PAP1 and PAP2 overexpressing lines, which display an enhanced accumulation of lignin (Borevitz et al., 2000), and by expression of the A. majus genes AmMYB308 and AmMYB330 in tobacco, which results in the repression of phenolic acid metabolism and lignin biosynthesis (Tamagnone et al., 1998). AtMYB26 could therefore be another MYB transcription factor regulating the phenyl propanoid pathway, thus affecting lignification in the endothecium. This is supported by the fact that expression of AtMYB26 was observed at those stages of anther development when secondary cell wall fortifications are formed in the endothecium. However, as other cell wall fortifications like deposition of cellulose are also missing in the endothecium of myb26, a function of AtMYB26 in specifying the identity of this cellular layer seems more likely.

Sequence comparison of AtMYB26 with other MYB factors does not give any hint towards its molecular function. Sequence homology has, in several cases of MYB transcription factors, been shown to correspond to a similar function, e.g. the A. thaliana genes WER and GL, which cluster together and are able to functionally complement each other displaying different biological functions only because of their different spatial expression patterns (Lee and Schiefelbein, 2001). AtMYB26, however, does not cluster with any of the functionally characterized MYB factors (Stracke et al., 2001).

Male sterile plants are useful tools for hybrid seed production. However, the use of many lines, mainly based on cytoplasmic male sterility, require the availability of appropriate restorer lines. Furthermore, the successful application of such plants depends on whether the male-sterile female parent can be multiplied efficiently (Perez-Prat and van Lookeren Campagne, 2002). Fertilization of myb26 mutants does not require the use of a special restorer line to obtain fertile progeny. Furthermore, as myb26 pollen is functional, male sterile lines can be propagated by self-pollination after mechanically opening the anthers. Therefore, male sterility based on a mutation in AtMYB26 and its orthologs in crop plants could offer some advantages over currently available male sterility systems.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Plant material

The myb26-2 mutant was identified in a population of A. thaliana, ecotype Columbia (Col-0), mutagenized with the En-1 transposon of Zea mays (Baumann et al., 1998; Wisman et al., 1998a,b). A. thaliana Col-0 was used as wild type was used for transformation and crossing. The ms35 was obtained by X-ray seed mutagenesis (Dawson et al., 1993), Ler seed was obtained from the Nottingham Arabidopsis Stock Centre. Plants were grown in a cooled greenhouse in 5 cm clay pots filled with soil. The medium temperature was 20°C. If necessary, additional light was provided during 16 h.

To test the sensitivity of myb26 to jasmonic acid and methyl jasmonate, respectively, mutants were subjected to either of the following treatments for 4 days: An aqueous solution of 2 mm jasmonic acid (J-2500, Sigma, St Louis, USA) containing 0.1% Tween 20 was applied as droplets of about 50 µl to flower buds twice per day. Alternatively, mutants were sprayed twice per day with an aqueous solution of 450 µm methyl jasmonate (39,270–7, Aldrich, Steinheim, Germany) containing 0.1% Tween 20.

Molecular biology techniques

Standard molecular biology techniques were performed as described by Sambrook et al. (1989). Isolation of genomic DNA was performed using the Plant DNA Isolation Kit (Roche Diagnostics, Mannheim, Germany). Amplification of flanking regions was carried out according to Steiner-Lange et al. (2001). PCR products were cloned into pGem-Easy (Promega, Mannheim, Germany).

Hybridization was performed with DNA fragments radioactively labelled with α32PdCTP using the Ready-To-Go DNA Labelling Beads (-dCTP; Amersham Biosciences, Freiburg, Germany). Pre-hybridization and hybridization were carried out in 6× SSPE, 30 g l−1 SDS, 200 mg l−1 PVP, 200 mg l−1 Ficoll and 100 mg l−1 salmon sperm DNA at 68°C. Washes were performed in 2× SSC, 0.1% SDS at 68°C. Probed filters were stripped by two successive washes in 0.2 m NaOH, 0.1% SDS at 37°C.

For preparation of an AtMYB26-specific probe lacking the MYB domain, a 669 bp fragment was isolated from plasmid pRD74 consisting of a PCR product representing nucleotides 1272–1941 of the AtMYB26 cDNA (Ralf Stracke, personal communication) cloned into the SmaI site of pCR-ScriptSK+ (Stratagene, La Jolla, USA).

PCR and primers

Polymerase chain reaction was generally carried out in a total volume of 50 µl with 0.4 µm primers and 200 µm dNTPs using the Advantage 2 polymerase (Clontech, Paolo Alto, USA) according to the manufacturers' instructions. PCR conditions were: 94°C for 4 min; 35 cycles of 94°C for 30 sec, 64°C for 1 min and 73°C for 1.5 min followed by a final elongation step at 73°C for 3 min. Primers MTL1 (ACT CGC TGG GAA ATC TGG TAA CGC T) and MTL2 (CAT GCT GCA ACA AGC AAA AGG TGA AG) were used for PCR amplification of the AtMYB26 5′-region. MTL3 (ACC GTG ATG ATG GTG GAC ATG AG) was used as the primer for sequencing the former En-1-integration sites of plants having lost the insertion and in combination with MTL2 for performing RT-PCR. RTL1 (ATG GGT CAT CAC TCA TGC TG) and RTR1060 (GTC CAC AAG AGA TTG GCG ACG A) were also used for RT-PCR. Act2F (TGC TGA CCG TAT GAG CAA AG) and Act2R (CAG CAT CAT CAC AAG CAT CC) served as primers for the RT-PCR actin control. Confirmation of an En-1 insertion within AtMYB26 was performed with primers MTL1 and En91R (TGC AGC AAA ACC CAC ACT TTT ACT TC). RS89 (TTT GCT CAC AAA CCT TCC TTA TCA C) and RS88 (ATG ACG TAC TGT CCA CAA GAG ATT G) were used for the preparation of an AtMYB26-specific probe lacking the MYB domain using an AtMYB26-cDNA clone as template. Preparation of a probe specific for the En-1 5′-end was performed as described by Wisman et al. (1998a).

Real-time PCR and RT-PCR

Total RNA was isolated from different organs (i.e. inflorescence, young and old rosette leaf, cauline leaf, root and seedling) of A. thaliana Col-0 plants, using total RNA Isolation Reagent (biomol GmbH, Hamburg, Germany). The isolated RNA was treated with DNase I (Roche Diagnostics GmbH, Mannheim, Germany) and precipitated in 2.75 m LiCl to eliminate genomic DNA contamination, followed by a final purification using RNeasy columns (Qiagen, Hilden, Germany).

RT-PCR amplifications were performed with 2 µg of total RNA in a Peltier Thermal Cycler (model PTC-225; MJ Research) using the OneStep RT-PCR Kit (Qiagen, Hilden, Germany). The amplification products were visualized on a 1% (w/v) agarose gel via ethidium bromide staining. RT-PCR conditions were: 30 min at 50°C, 15 min at 95°C; 40 cycles of 1 min at 94°C, 1 min at 64°C, 1 min at 72°C and a final extension for 10 min at 72°C. RT-PCR primers were MTL2 and MTL3.

For performing real-time PCR, the purified total RNA was reverse transcribed with the Superscript First-Strand Kit (Gibco Life Technologies, Karlsruhe, Germany) using 250 ng of oligo(dT) (Sigma, St Louis, USA) and 25 ng of random hexamer primer (Amersham Biosciences, Freiburg, Germany) per reaction. Reactions required 0.5 mm dNTP and 5 mm MgCl2. To normalize the input of cDNA, real-time PCR assays were performed in a Taqman ABI 7700 using the Platinum Quantitative PCR Supermix-UDG (Invitrogen Life Technologies, Karlsruhe, Germany) and pre-developed Human 18S Taqman Reagents (primers and Vic-labelled probe), which are recommended by ABI for the normalization in all eukaryotes. According to the results, the input into each of the quantitative assays performed with the AtMYB26-specific probe and primers was normalized to the 18S rRNA genes. The following primers and Taqman probe specific for the AtMYB26 coding sequence were used: FP: CCA TGG ATG TTG GAG CTC TGT T; RP: GCT TCC ACG TTT AAG ATG CAG GTC T and the intron-spanning probe (5′-labelled with 6-Fam dye and 3′-labelled with TAMRA): CAT GCA GGT TTG CAG AGA TGT GGA AAG AG. Conditions for real-time PCR assays were: 2 min at 50°C, 2 min at 95°C, 40 cycles of 15 sec at 95°C, 1 min at 60°C. Assays were performed in triplicates. The presence of residual genomic DNA in the RNA used for this assay had been excluded in advance by performing the real-time PCR assays with the purified total RNA, omitting the reverse transcription step, and the AtMYB26-specific probe and primers.

Microscopical techniques

Light microscopy was performed using a ZEISS Axiphot Fluorescence microscope (Jena, Germany). Pictures were taken using either a digital imaging system (Leica Microsystems, Bensheim, Germany) including a JVC KV-F70 digital camera, PC and imaging software or a digital imaging system (INTAS, Göttingen, Germany) including a Spot Diagnostics Instruments Inc. digital camera, PC and imaging software.

For studying autofluorescence of lignified material, a filter with the following specifications was used: G: 365, FT: 395, LP 420. Prior to microscopy, anthers were cleared in 70% (v/v) lactic acid for 3 days at 60°C.

To determine the cellular structure of myb26-2 and Col-0 anthers, whole inflorescences were embedded in Paraplast Plus (Sherwood, St Louis, USA) and subsequently used for preparing 8 and 5 µm thin cross-sections (Microtome: Jung Autocut 2055, Leica Microsystems, Nussloch, Germany), respectively. After transferring the sections to slides (Superfrost Plus, Menzel), they were incubated for 30 sec at room temperature in an aqueous 0.05% (w/v) toluidine blue solution and subsequently washed twice for 5 min with sterile water. After drying the sections at room temperature, the Paraplast Plus was removed by incubating the slides thrice in 100% Histoclear (National diagnostics, Hessle Hull, UK). Then, the slides were treated with Entellan and subsequently covered with a cover slip.

For studying lignification, a phloroglucinol staining was established. For this purpose, the paraffin-embedded cross-sections were treated with 100% Histoclear and subsequently incubated in 100% ethanol. Staining was then performed with 2% (w/v) phloroglucinol in 100% ethanol for 1 h at room temperature. Subsequently, the sections were mounted with 18.5% (v/v) HCl.

To show the deposition of cellulose in the endothecium, the fifth oldest bud of inflorescences of myb26-2 and wild-type plants was fixed in 2% (w/v) paraformaldehyde/0.25% (v/v) glutaraldehyde for 2 h and subsequently embedded in paraffin. Semi-thin sections were stained with calcofluor white as described by Dawson et al. (1999).

For scanning electron microscopy (SEM), anthers were mounted on aluminium specimen stubs using Tissue-Tek O.C.T. compound (Sakura Finetek, Tokyo, Japan) and immediately shock-frozen in liquid nitrogen. Samples were subsequently transferred to a LEO DSM scanning electron microscope (LEO electron microscopy, Oberkochen, Germany) equipped with a cryo transfer chamber (Gatan GmbH, München, Germany). Samples were sputter-coated with gold and examined at voltages of 2–20 kV.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

The authors would like to thank Eva-Marie Schlösser, Heidrun Häweker and Marc Wolff for technical assistance and Rolf Hirz for help with SEM microscopy. Anna Sorensen and Bernd Weisshaar are acknowledged for helpful discussions and for critically reading the manuscript. We further thank Ralf Stracke for plasmid pRD74 and for sharing unpublished results, Bernadette von Malek for providing us with the seeds of dde2-2 and the ADIS-Unit of our Institute for performing all sequencing. This work was supported by grants from the German Federal Ministry for Education and Research (BMBF) and from the British Biotechnology and Biological Sciences Research Council (BBSRC).

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  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
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