Stem cells in the root and shoot apical meristem provide the descendant cells required for growth and development throughout the lifecycle of a plant. We found that mutations in the Arabidopsis MAINTENANCE OF MERISTEMS (MAIN) gene led to plants with distorted stem cell niches in which stem cells are not maintained and undergo premature differentiation or cell death. The malfunction of main meristems leads to short roots, mis-shaped leaves, reduced fertility and partial fasciation of stems. MAIN encodes a nuclear-localized protein and is a member of a so far uncharacterized plant-specific gene family. As main mutant plants are hypersensitive to DNA-damaging agents, expression of genes involved in DNA repair is induced and dead cells with damaged DNA accumulate in the mutant meristems, we propose that MAIN is required for meristem maintenance by sustaining genome integrity in stem cells and their descendants cells.
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In higher plants, stem cells are positioned in the shoot apical meristem (SAM) at the tip of the shoot and in the root apical meristem (RAM) at the tip of the root. They constitute a pool of undifferentiated cells that continually provides new cells for post-embryonic growth and development throughout the lifecycle of a plant. Stem cells are maintained in a specific cellular micro-environment, the stem cell niche, in which organizing centres provide signals and physical support to maintain the stem cells in their undifferentiated state (Scheres, 2007).
The organizing centre in the SAM is located beneath three layers of stem cells and characterized by expression of the homeodomain transcription factor WUSCHEL (WUS) (Mayer et al., 1998). In the RAM, a unicellular layer of stem cells surrounds the rarely dividing quiescent centre (QC) cells, which act as an organizing centre in the root and are characterized by expression of the transcription factor encoded by WUSCHEL-RELATED HOMEBOX 5 (WOX5) (Dolan et al., 1993; Haecker et al., 2004; Sarkar et al., 2007). Stem cell daughters that are located proximal to the QC retain the capacity to divide and form the epidermis, ground tissue and vasculature. In contrast, the distal stem cell daughters differentiate into starch-containing columella cells, without further cell divisions.
Correct positioning of the root stem cell niche, stem cell maintenance and function and the mitotic activity of stem cell daughters are controlled by a number of largely plant-specific transcription factors (Iyer-Pascuzzi and Benfey, 2009; Bennett and Scheres, 2010). The AP2 transcription factors PLETHORA1 (PLT1) and PLETHORA2 (PLT2) provide an apical–basal patterning signal (Blilou et al., 2005), the GRAS transcription factors SHORTROOT (SHR) (Benfey et al., 1993; Helariutta et al., 2000) and SCARECROW (SCR) (DiLaurenzio et al., 1996; Wysocka-Diller et al., 2000) generate a radial patterning signal. It is well established that the phytohormone auxin plays a crucial role in positioning of the stem cell niche of the RAM, and that an auxin response maximum in the distal stem cell region regulates and maintains the function of the QC (Friml et al., 2003). As PLT expression is regulated by auxin, an apical–basal auxin gradient in the root results in a gradient of PLT expression. The overlapping maxima of apical–basal (auxin, PLT1, PLT2) and radial (SHR, SCR) patterning signals thereby specify the stem cell niche in the root (Sabatini et al., 2003; Blilou et al., 2005; Scheres, 2007).
In addition, several reports have shown that meristem organization and maintenance are also severely affected in mutants for genes that are implicated in chromatin organization, DNA replication or cell-cycle regulation. For example, mutants of the cyclin-dependent kinase gene cdka;1 display cell cycle-specific defects but have also strong developmental defects, including disturbed organization and maintenance of stem cell fate in the RAM (Nowack et al., 2012). Further prominent examples are the fasciata mutants (fas1 and fas2), which represent loss-of-function mutants for the large subunits of CHROMATIN ASSEMBLY FACTOR–1 (CAF–1). The CAF–1 complex is important for nucleosome assembly on synthesized DNA, and fas1 and fas2 mutants show severe defects in organization and maintenance of the stem cell niche in both the SAM and the RAM (Kaya et al., 2001; Ono et al., 2006). Similar defects were observed in mutants for the nuclear protein BRUSHY1 (BRU1), which has a putative function in maintaining the structural and functional stability of chromatin (Takeda et al., 2004). In bru1 mutants, the defective organization of the meristem was accompanied by constitutive activation of the DNA damage pathway. Likewise, mutants for the chromatin remodelling factors MINISCULE1 and 2 (MINU1 and MINU2) are unable to initiate and maintain stem cell populations in the RAM and SAM, and show severe defects during embryogenesis (Sang et al., 2012).
Moreover, mutants for the topoisomerase MGOUN1, which functions during replication and transcription (Laufs et al., 1998; Graf et al., 2010), or for the TEBICHI (TEB) gene, which encodes a protein with putative DNA helicase and polymerase activity, show defects in cell-cycle progression and meristem maintenance and constitutive activation of the DNA damage response (Inagaki et al., 2006, 2009). Response to DNA damage is mediated by the highly conserved ATAXIA-TELANGIECTASIA MUTATED (ATM) and ATM/RAD53-RELATED (ATR) protein kinases. They induce transcription of genes involved in cell-cycle control, chromatin structure and DNA damage repair (Culligan et al., 2006; Mannuss et al., 2012). A recent study has shown that stem cells are hypersensitive to DNA damage and undergo ATM/ATR-dependent non-apoptotic programmed cell death upon mild treatments with genotoxic agents. It was proposed that this pathway provides a mechanism to safeguard genome stability in stem cells and thus to prevent accumulation of mutations in stem cells and their descendant cells (Fulcher and Sablowski, 2009).
This paper describes identification of the uncharacterized mutant maintenance of meristems (main), which exhibits defects in the morphology of shoot organs as well as retarded root growth. The developmental abnormalities in main are associated with disorganized root and shoot meristems and loss of stem cell activity. A high number of cells with damaged DNA, increased expression of genes that are implicated in DNA repair, and the presence of dead cells in the RAM of main led to the hypothesis that the plant-specific MAIN gene product is required for protection of genome stability in plant meristems.
Identification of main mutants
The main–1 mutant was identified in a genetic screen for altered GFP distribution in Arabidopsis root tips. T–DNA mutagenesis was performed using an Arabidopsis line (ecotype C24) that expressed GFP from the companion cell-specific Arabidopsis SUCROSE TRANSPORTER2 promoter (pSUC2::GFP), in which GFP moves into sieve elements, is translocated to sinks and is unloaded symplastically from the phloem ends (Imlau et al., 1999; Hoth et al., 2005; Stadler et al., 2005a,b).
In roots of pSUC2::GFP-expressing plants, strong GFP fluorescence is seen in the vascular strands that end approximately 200 μm above the tip and as a cloud of fluorescence from symplastically unloaded GFP in the meristem (Figure 1a, see also Figure 3a). After T–DNA insertion mutagenesis, eight of 8000 microscopically screened T2 lines were unable to unload GFP from the root vasculature (arrow in Figure 1b, see also Figure 3b). In addition to the reduced GFP unloading phenotype, one mutant line exhibited severe defects in the RAM and SAM, and was therefore named main for maintenance of meristems. Here, we describe characterization of the main-1 and main-2 mutants.
In main–1, the T–DNA insertion is located in exon 3 of At1 g17930, 746 bp downstream of the start ATG (Figure 1e). A second insertion mutant line for this gene was obtained from the GABI-Kat collection (Kleinboelting et al., 2012), which we named main–2 (GABI-Kat ID 728H05; Li et al., 2003). In main–2, the T–DNA is located in intron 5, 1523 bp downstream of the start ATG (Figure 1e). Both insertions resulted in complete loss of full-length MAIN mRNA (Figure 1f). However, residual expression of a truncated mRNA covering the regions before and after the insertion was detected in both mutant lines (Figure S1).
We analysed the unloading phenotype of main–2 by crossing main–2 plants (ecotype Columbia, Col–0) with wild-type (WT) plants (Col–0) that expressed GFP under the control of the SUC2 promoter (Figure 1c) (Schneidereit et al., 2008). In this background, main–2 showed an identical phenotype (Figure 1d) to main–1 (Figure 1b).
In addition to the GFP unloading phenotype, both mutants displayed several developmental abnormalities. Soil-grown main–1 and main–2 mutants were smaller than WT (Figure 1g–j), and most of their rosette leaves had a lanceolate shape (arrows in Figure 1h,j) with altered vein patterning (Figure 1k). When grown on MS medium, the developmental defects were even more obvious, revealing dramatically reduced growth of the primary root (Figure 1l and Figure S2). Moreover, both main mutants had short siliques with a large number of aborted seeds (mean 60%) (Figure 1m–o).
The phenotype of both lines segregated with the T–DNA insertion and was restricted to homozygous mutants. In addition, we prepared a complementation construct with the MAIN coding sequence flanked by its 1462 bp promoter region and the 3′ UTR (pMAIN::MAIN-CDS-UTR), which was introduced into main–1 and main–2 plants. Four independent complemented lines were grown on plates and on soil, and altogether 67 plants were analysed. In each of the plants harbouring the complementation construct, the main phenotype was completely rescued (Figure 1p–r). These results demonstrate that the insertion in MAIN is responsible for the observed phenotype, and that both mutants are loss-of-function mutants.
MAIN is a nuclear protein and is mainly expressed in meristems
To investigate the subcellular localization of MAIN, we fused GFP to the MAIN cDNA under the control of the CaMV 35S promoter (p35S::MAIN-GFP). This construct was used for particle bombardment of tobacco (Nicotiana tabacum) epidermis cells, and the results showed that the fusion protein localized exclusively to the nucleus (Figure 2a). A construct in which GFP was fused to the 5′ end of MAIN and particle-bombarded into epidermis cells of Arabidopsis, tobacco and onion (Allium cepa) confirmed the localization to the nucleus (Figure S3). As MAIN may be targeted to the nucleus by a predicted nuclear localization sequence (NLS) containing the amino acid sequence KRKRR (Brameier et al., 2007; Tsugeki et al., 2009), a fusion construct was generated in which a stop codon was introduced before this sequence. This fusion was no longer targeted to the nucleus, but remained in the cytoplasm (Figure S3). After modifying the NLS from KRKRR to KLNQR, the fusion protein also localized to the cytoplasm (Figure 2b), indicating that the KRKRR sequence is a functional NLS.
To study the tissue-specific expression pattern of MAIN, a translational fusion was generated comprising the complete MAIN genomic region, including the promoter region that was used for the complementation construct, fused to the uidA (GUS) gene of Escherichia coli (pMAIN::MAIN-GUS) or GFP (pMAIN::MAIN-GFP). Both constructs were used for stable transformation of Arabidopsis WT plants and revealed essentially the same expression pattern. A detailed analysis with the GFP construct revealed that MAIN was expressed in all cells of the root meristem, including the stem cell niche (Figure 2c,d), but its expression was confined to the vasculature in the differentiation zone (Figure 2e,f). This pattern was confirmed in histological cross-sections of pMAIN::MAIN-GUS seedling roots 3 days after germination (DAG) (Figure 2g,h). In general, MAIN was most strongly expressed in all meristematic tissues, i.e. the SAM (arrows in Figure 2i,j), the RAM (Figure 2e), the lateral root meristems (Figure 2f) and the adventitious root meristem (arrowhead in Figure 2j).
In the aerial parts, pMAIN::MAIN-GUS staining was generally weaker than in the root, and, apart from the SAM, was mainly detected in the vasculature of leaves at all developmental stages (Figure 2i–k); this became more pronounced after prolonged staining (24 h; Figure 2k). Expression of MAIN was also seen in the nuclei of pollen tubes (Figure 2l) and in the ovary of young and mature flowers (Figure 2m). A more detailed analysis in flowers of pMAIN::MAIN-GFP lines revealed GFP signals in the ovules (arrows in Figure 2n), as well as the carpels (arrow in Figure 2o).
main mutants have a disorganized RAM
The specific expression pattern of MAIN, together with the drastically reduced primary root growth of main mutant seedlings, suggested that MAIN may function specifically in meristems. Therefore, we next analysed the root meristem of main-1 and main-2 mutants in more detail. Confocal images of propidium iodide (PI)-stained roots revealed severe defects in the RAM of main roots, with an abnormal organization of the stem cell niche and an irregular arrangement of cells in the proximal meristem and the columella (Figure 3b). The QC cells (arrow in Figure 3b) lost their specificity in main mutant roots and became morphologically indistinguishable from the surrounding cells at later developmental stages (Figures S2 and S4). Moreover, the clearly defined cell division pattern observed in WT RAM was disturbed in main mutants, leading to distorted division planes, such as anticlinal instead of periclinal divisions in the endodermal layer (Figure 3b, see also arrows in Figure 6f,g).
A short root phenotype may either be caused by a reduced length of the cells or a reduced number of cells. To distinguish between these two possibilities, we measured the size of the meristematic region by quantifying the number of meristematic cells in the cortical cell file at 3, 5 and 7 DAG. In main-2 mutants, the number of meristematic cells was reduced by approximately one third compared to WT, and increased only very slowly over time (Figure 3c). Next, we crossed the main-2 mutant lines with a cyclin–GUS marker line, in which cyclin–GUS is specifically expressed in the late G2 and M phases of the cell cycle (Colon-Carmona et al., 1999). In the main–2 background, fewer cells expressed the cyclin–GUS marker, confirming that the number of dividing cells was reduced (Figure 3d,e). In addition, the cells of the main-1 and main-2 root meristem exhibited premature elongation (arrowhead in Figure 3b) and differentiation, as indicated by root hair formation close to the root tip (Figure 3g, see also Figure 1b,d). The reduced meristem and premature differentiation were confirmed by analysing the main phenotype in the background of an AtSUC2p::tmGFP9 marker line (Figure 3f), in which a membrane-bound version of GFP is expressed from the phloem-specific AtSUC2 promoter (Stadler et al., 2005b). In this marker line, the GFP fluorescence is confined to differentiated companion cells, which are seen approximately 500 μm from the tip in WT roots (arrowhead in Figure 3f). In main-2 mutants, the distance of AtSUC2p::tmGFP9-expressing cells from the tip was reduced to approximately 250 μm (arrowhead in Figure 3g). At later developmental stages (18 DAG), vacuolized cells were detected close to the QC, indicating that no meristematic cells were present in the main mutant (Figure S2).
The modified pseudo-Schiff propidium iodide (mPS–PI) technique (Truernit et al., 2008) was used for confocal visualization of cell-wall and storage polysaccharides. This method revealed that main-1 mutants contained fewer starch granules in the columella compared to WT (Figure 3h,i) and the presence of starch granules in the columella initial cells (red arrowhead in Figure 3i), indicating that stem cells are differentiated and not maintained in main mutants (Bureau et al., 2010).
Another very prominent feature of PI-stained main mutant roots was the presence of strong intracellular staining in meristematic cells proximal to the QC and in the stem cell niche itself (asterisks in Figure 3b,g), indicating that these cells were dead (Truernit and Haseloff, 2008; Fulcher and Sablowski, 2009; Cools et al., 2011). Dead cells were not observed in WT roots grown under exactly the same conditions, and the number of dead cells was variable among individual main seedlings.
In summary, these results show that loss of MAIN function leads to loss of meristem cell identity, reduced cell division activity and increased cell death of meristematic cells in the root.
Expression and function of MAIN starts during embryo development
As the severe defects during root development were already observed as early as 2 DAG, we examined the expression of MAIN during embryo development. Analysis of pMAIN::MAIN-GFP lines revealed that the MAIN–GFP fusion protein accumulated in all cells of the embryo at all developmental stages (Figure 4a–d). Next we analysed the phenotype of main–2 mutants during embryo development. While most embryos from homozygous main mutant plants showed normal development, several abnormal cell division patterns were observed in approximately 18% of the investigated embryos (total number of investigated embryos n =88) (Figure 4e–k). These involved anticlinal rather than periclinal divisions of the hypophysis in globular embryos (Figure 4e,f) or periclinal rather than anticlinal divisions of protodermal cells in transition-stage embryos (arrows in Figure 4i–k). In addition to these defects in cell-plate orientation, individual main mutant embryos showed additional unusual cell divisions that completely destroyed embryonic patterning (Figure 4g, arrow in Figure 4h). These results indicate that embryo development is affected in main mutants to a variable degree, ranging from no or only slight defects to severe defects, ultimately leading to 60% reduced seed set.
main mutants have a disorganized SAM and show a fasciation phenotype
Strong pMAIN::MAIN-GUS expression was also observed in the SAM. The effect of the main mutation on SAM development was thus analysed in more detail. Longitudinal sections prepared from the SAM of plants at 3 DAG revealed a disturbed cellular organization of the meristem. The cells of the L1 and L2 layers remain clonally distinct and undergo exclusively anticlinal divisions in WT plants, but frequently showed irregular or periclinal divisions in main-2 mutants (Figure 5a,b).
In addition, 9.8 ± 0.8% of mature main-2 plants (n =103) showed a fasciation phenotype. Fasciation was seen in the stems of both main mutants (arrow in Figure 5d, cross-section shown in Figure 5f) but never in WT control plants (Figure 5c, cross-section shown in Figure 5e). Some main flowers contained more than one ovary (arrowheads in Figure 5h), showed lost organ identity (asterisks in Figure 5h,i, indicating sepals carrying papillae) and had fused flowers and pedicels (Figure 5i).
In summary, main mutants have cell-patterning defects in those cells in which MAIN is mainly expressed, i.e. in cells of the RAM and the SAM. This leads to phenotypic alterations in the organs derived from them, such as roots, leaves, stems and flowers.
main mutants show altered expression of root patterning genes
To further investigate the function of MAIN for correct cell patterning in the RAM, we used well-characterized reporter lines and crossed them into the main-2 mutant background. The transcription factor SCARECROW (SCR) is required for the maintenance of proximal stem cells and for correct radial patterning of the root (DiLaurenzio et al., 1996; Wysocka-Diller et al., 2000). In WT, pSCR::erGFP expression is found in the endodermis, the endodermis/cortex initials and the QC (Figure 6a), but SCR expression in main mutants was patchy, and was sometimes confined to individual cells within the endodermal layer (Figure 6b,c). This low or absent pSCR activity correlated with additional cell divisions in the adjacent cortex layer (arrow in Figure 6b) and the formation of supernumerary cell files (arrows in Figure 6c).
The QC is the central organizer of the root stem cell niche, and the pWOX5::erGFP marker is specifically expressed in QC cells throughout the development of WT roots (Figure 6d) (Sarkar et al., 2007; Ditengou et al., 2008). However, in main mutants, pWOX5::erGFP was ectopically expressed in the endodermis/cortex initials and their daughter cells (arrows in Figure 6e). This additional expression of pWOX5::erGFP increased during development and was even more pronounced in roots of older seedlings (Figure S4).
Auxin plays a central role in the organization and maintenance of the root stem cell niche (Sabatini et al., 1999; Friml et al., 2003). We therefore analysed the distribution of this phytohormone in main mutants. The auxin-responsive reporter pDR5::GFP (Ottenschlager et al., 2003) shows maximum expression in the QC and the columella cell files in WT roots (Figure 6f). In the main–2 mutant, pDR5::GFP showed a similar expression pattern, but the maximum in the QC was lost (arrow in Figure 6g), indicating a change in auxin distribution. Altered auxin flux may result from altered expression of auxin efflux carriers such as those encoded by the PIN genes (Blilou et al., 2005). We analysed GFP lines for PIN1, PIN2 and PIN7 expression in the main mutant background, and did not find any major changes compared to WT. An exception was the PIN1 reporter, which showed a shift of expression to the columella cell files in main mutants (arrow in Figure 6i) that was never observed in WT (Figure 6h) (Friml et al., 2002). To further analyse this phenotype, we performed a quantitative real time–PCR analysis for PIN1, PIN2, PIN3, PIN4 and PIN7 on RNA isolated from RAMs of seedlings at 3 DAG. However, no major changes in expression between WT and the main-2 mutant were observed (Figure S5).
To investigate whether MAIN is involved in auxin signalling, we monitored the expression of primary auxin response genes such as INDOLE-3–ACETIC ACID INDUCIBLE1 (IAA1) and IAA19 in seedlings at 3 DAG that had been incubated for 2 h on medium containing 10 μm auxin. The main-2 mutants behaved like WT seedlings, and both genes were readily induced after auxin treatment. Moreover, the expression of MAIN was not changed in WT seedlings upon addition of auxin. We also monitored the expression pattern of the DR5::GFP reporter after incubation of seedlings with auxin or with auxin inhibitors, and obtained similar results in the main mutant background and in WT (Figure S5).
The results indicate that MAIN is not directly involved in auxin-related processes, and therefore that the main phenotype is most likely not caused by altered auxin signalling or distribution.
main mutants exhibit increased DNA damage and are hypersensitive to DNA-damaging agents
Meristematic cells have been shown to be particularly sensitive to DNA damage (Fulcher and Sablowski, 2009). We observed an accumulation of dead cells in the main RAM as revealed by PI staining of the roots (Figure 7a,c). Quantification of the mean area of dead cells in the meristem of PI-stained roots revealed that this was significantly increased (25-fold) in main-2 mutants compared to WT seedlings at 3 DAG (Figure 7e). Dead cells were almost exclusively observed in the proximal meristem, and provascular cells were affected most frequently, resulting in areas of dead cells that covered 10–12 cells of the stelar region. Only individual dead cells were detected in the endodermis or cortex layer. These observations showed that MAIN may be involved in the control of genome integrity. To test this hypothesis, we first analysed PI-stained roots of WT seedlings that had been treated for 24 h with zeocin, which induces DNA double-strand breaks. After incubation with zeocin, WT seedlings displayed essentially the same phenotype as untreated main seedlings, with a comparable amount of dead cells in the meristem (Figure 7e) and premature elongation of meristematic cells (arrowhead in Figure 7b). In addition, analysis of pDR5::GFP and pPIN1::PIN1-GFP expression in zeocin-treated WT plants revealed an altered expression pattern similar to that in main–2 mutants (Figure S6). This indicates that the observed ectopic pPIN1::PIN1-GFP expression and reduced pDR5::GFP expression in the main–2 mutant background may be a secondary effect of the accumulation of dead cells in the RAM.
Second, we tested the sensitivity of main-2 mutant seedlings to DNA-damaging agents by incubating them with zeocin, which resulted in an even higher accumulation of dead cells (Figure 7d,e). This result shows that main mutants are still able to respond to DNA damage, and indicates a hypersensitivity of main mutants to DNA-damaging agents. To confirm this finding, WT and main-2 mutant seedlings at 3 DAG were transferred to medium containing zeocin, the DNA-alkylating agent methyl methanesulfonate (MMS) or no DNA-damaging agent, and further grown for 12 days (Figure 7f). WT plants that were transferred to zeocin- or MMS-containing medium had shorter roots and smaller rosettes compared to plants transferred to MS medium. However, the main mutants were much more strongly affected by this treatment, and exhibited a hypersensitive reaction to the DNA-damaging substances (Table S1).
Third, we analysed the expression of DNA repair genes, which are known to be induced upon DNA damage caused by double-strand breaks (Deveaux et al., 2000; Lafarge and Montane, 2003; Zhu et al., 2011). Quantitative real time-PCR analysis on RNA extracted from isolated RAMs of seedlings at 3 DAG revealed that ATGR1, RAD51, BRCA1, PARP2, KU70 and XRCC4 were all significantly up-regulated in main-2 mutants (Figure 7g) compared to WT. Double-strand breaks are repaired via the non-homologous end-joining pathway and the homologous recombination pathway (Puchta, 2005; Waterworth et al., 2011). Our analysis revealed that both the non-homologous end-joining pathway (KU70 and XRCC4) and the homologous recombination pathway (ATGR, RAD51, BRCA1 and PARP2) are constantly activated in main mutants.
Finally, we analysed whether the constitutive activation of the DNA damage response pathway in main mutants was due to an actual increased level of DNA damage. To this end, we tested DNA integrity in main-2 mutants and WT using the comet assay (Menke et al., 2001). In this assay, the percentage of DNA in the comet tail, which represents the degree of damaged DNA, was more than twofold higher in the main mutants compared with WT, and was similar to the value for WT plants treated with 40 μg ml−1 zeocin for 7 days (Figure 7h,i). Taken together, these results strongly suggest that loss of main function leads to an increased level of DNA damage specifically in dividing cells.
MAIN (At1g17930) is a member of a uncharacterized family of 14 DUF1723 proteins in Arabidopsis (Figure S7 and Table S2). DUF1723 domains are plant-specific and were initially described in a study on DNA-binding domains in eukaryote-specific WRKY and GCM1 transcription factors (Babu et al., 2006). In line with the identification of DUF1723 domains in transcription factor-related proteins, we show that MAIN carries a NLS in its C–terminus and that it localizes to the nucleus.
MAIN is mainly expressed in meristematic cells throughout development, and our detailed phenotypic analysis revealed that, in main mutant meristems, stem cells are gradually lost by premature differentiation or cell death. We propose that MAIN is a plant-specific factor that is essential for the maintenance and stability of stem cells and their immediate descendant cells.
Loss of MAIN function leads to irregular cell divisions, embryo abortion and premature differentiation of stem cells
MAIN is expressed in all cells of embryos (Figure 4a–d) and in meristematic tissues of adult Arabidopsis plants (Figure 2c–o). Loss-of-function mutations in main lead to variable embryonic phenotypes, ranging from mis-shaped embryos that are eventually aborted (Figure 4g,h), through embryos with individual abnormal cell divisions (Figure 4f,j) to WT embryos that produce viable seeds and allow analyses of main effects on root development. This phenotypic variability in embryos may be due to the presence of a factor with (partially) redundant function, i.e. another uncharacterized but closely related member of the DUF1723 family (Figure S7).
Moreover, the occurrence and intensity of DNA damage is likely to be induced randomly in main mutant cells. This may also explain the phenotypic variability observed in roots of individual main plants as seen in Figure 6, in which a mutant with almost no dead cells is shown (Figure 6b) next to mutants with many damaged cells (Figure 6e,g).
The main–2 mutant appeared to be somewhat more strongly affected than main–1 with regard to rosette diameter and embryo lethality (Figure 1o,r). We tested whether this could be explained by residual expression of truncated mRNAs that are at least partially functional (Figure S1). However, truncated mRNAs covering the regions before and after the insertion were detected and expressed at similar low levels in both mutants. As both insertions are located in the central region of the gene and lead to complete loss of full-length mRNA, it is unlikely that these truncated mRNAs are functionally relevant. Another possibility to explain the difference in the phenotypic strength may be that main–1 has a C24 background while main–2 has a Col–0 background, and that these two ecotypes are differently affected by loss of MAIN function.
Drastically reduced growth of the primary root was observed in each of the main homozygous mutant progeny. The QC cells that keep the surrounding stem cells in an undifferentiated state (Scheres, 2007) are gradually lost in main mutants, and the stem cells as well as their daughter cells lost their undifferentiated state and underwent premature differentiation. In mutants that are unable to specify a QC, the root meristem is eventually lost, as stem cells and their descendant cells stop dividing (Sabatini et al., 2003; Aida et al., 2004). MAIN is expressed in all meristematic cells, and therefore MAIN is most likely not a factor that is specifically required to maintain the QC. Instead, we propose that main mutants are unable to maintain a stable pool of undifferentiated and cell division-competent cells in the meristem, and that the QC is lost in order to replace damaged stem cells.
Genome stability is defective in dividing cells of main mutants
It has been shown that stem cells of animals as well as stem cells of plants are specifically susceptible to genotoxic stress and readily undergo differentiation or cell death after treatment with DNA-damaging agents (Rich et al., 2000; Fulcher and Sablowski, 2009; Sherman et al., 2011). Several lines of evidence suggest that the severe defects in the main root meristem are caused by DNA damage-induced cell death of dividing cells. main seedlings not only had an increased number of dead cells among stem cells and their immediate descendant cells, but also showed constitutive expression of typical DNA damage-induced genes (Figure 7g). Moreover, the comet assay revealed a significant increase in the amount of nuclei with damaged DNA in main mutants compared with WT (Figure 7i). The accumulation of dead cells and cells with damaged DNA in the main meristem is likely to interfere with positional information like auxin and may contribute to false divisions and premature differentiation. WT plants suffering from DNA damage show an altered auxin distribution and premature cell elongation and differentiation in the meristematic region, similar to main mutants (Figure S6). These results strongly suggest that the auxin-related defects observed in main mutants are a secondary effect resulting from cell death in the meristem.
The phenotypic characteristics of main mutants such as accumulation of dead cells in the RAM, patchy SCR expression and ectopic WOX5 expression (Figure 6b,c,e), as well as fasciated stems and short roots, have also been described in mutants for genes that are involved in DNA replication or DNA damage repair. For example, mutants for TEBICHI (TEB), whose product is involved in DNA replication, accumulate dead cells, show patchy SCR expression, slightly altered DR5 activity in the root tip, constitutively activated DNA damage response and hypersensitivity to DNA-damaging agents (Inagaki et al., 2006). However, teb mutants show more frequent and severe defects during embryo development than main mutants. Another example are fasciata mutants, which have defects in chromatin assembly during DNA replication and repair. fasciata mutants are similar to main mutants as they show an accumulation of dead cells in the RAM, starch-containing initials and patchy SCR expression, as well as stochastically occurring defects during embryo development and altered leaf shape resulting from a disturbed balance between cell division and differentiation (Kaya et al., 2001). Moreover, mutants for BRU1, whose product is thought to control chromatin stability (Takeda et al., 2004) and MGNOUN1, encoding a DNA topoisomerase, are similar to main mutants, showing increased expression of DNA repair genes, accumulation of cells with damaged DNA in the RAM, and partially fasciated stems. However, mgnoun1 mutants are more affected in the stem cell niche of the SAM, while main mutants have stronger defects in the RAM (Graf et al., 2010). MINU1 and MINU2 encode chromatin-remodelling ATPases, and their loss of function leads to defects in the RAM similar to main mutants, but minu mutants also show severe embryonic defects and loss of SAM activity (Sang et al., 2012). Taken together, these phenotypic similarities, together with the nuclear localization and expression of MAIN in meristematic tissues, suggest that MAIN may also be involved in chromatin assembly during DNA replication and repair, or may be involved in the DNA replication machinery itself. Recently, a new factor for stem cell maintenance named MERISTEM DISORGANIZATION 1 (MDO1) was identified, which is involved in protecting genome integrity (Hashimura and Ueguchi, 2011). Like MAIN, MDO1 encodes an unknown plant-specific protein with a similar loss-of-function phenotype that was also associated with increased DNA damage in meristematic cells. This indicates that plants, which maintain active stem cells throughout their lifecycle, have evolved plant-specific factors that ensure genome integrity during cell division.
In summary, our results show that MAIN is a factor that regulates genome integrity in dividing cells and is therefore essential for a stable maintenance of plant meristems. Whether MAIN as an exclusively nuclear-localized protein acts as a transcriptional regulator, or is involved in chromatin remodelling or DNA replication, will be addressed in future experiments.
Strains and growth conditions
Arabidopsis thaliana was grown on soil in growth chambers (16 h light\8 h dark, 22°C), greenhouses (ambient conditions) or on Petri plates (16 h light\8 h dark, 22°C).
Generation of mutant lines and screening
The pGPTV-BAR (Becker et al., 1992) derivative pBAR-35S (AJ251014; Guillermo Cardon and Peter Huijser, Max Planck Institute for Plant Breeding Research, Department of Comparative Development and Genetics, Cologne, Germany, unpublished results) was used for T–DNA mutagenesis of pSUC2::GFP plants. Seeds of 1500 transformed T0 plants were selected for BASTA (http://agrar.bayer.de) resistance. Seedlings of 8000 BASTA-resistant T1 plants were screened on Petri plates for reduced unloading of GFP after 12–16 days growth (16 h light\8 h dark, 22°C).
Identification of T–DNA insertion sites and transcript levels
T–DNA insertion sites were determined by TAIL-PCR (Liu and Whittier, 1995) in main–1 using primers LBT0b, LBT0c, LBT0 and AD3, or by sequencing a PCR-derived fragment in main–2 obtained using GabiKat-LB1 and At1 g17930-PIGr. MAIN mRNA levels in WT and mutant lines were determined by RT–PCR using primers MAINc+1f and MAINc+1437r (full-length transcript), MAINc+1f and At1 g17930+421rev (truncated transcript before the T–DNA insertion), At1g17930c+1270f and At1g1793UTR+1704rev (truncated transcript after the T–DNA insertion). ACT2 mRNA levels in WT and mutant lines were determined by RT–PCR using primers AtACT2 g+846f and AtACT2 g+1295r. Primer sequences are provided in Table S3.
For complementation, a 1462 bp promoter fragment and a 2393 bp genomic fragment were amplified using primers At1 g17930 g-1465f + At1 g17930 g + 3r and At1 g17930 g-PciIf + At1 g17930 g+2393r, respectively, which add PciI sites to the 3′ end of pMAIN and the 5′ end of the coding sequence. Fragments were cloned into pGEM–T Easy (Promega, http://www.promega.com), yielding pRG108 for the gene and pRG109 for the promoter, and combined to yield pRG110. A 3911 bp ApaI/SpeI fragment of pRG110 was cloned into pER8 (Zuo et al., 2000), yielding pUW114 and transformed into plants (Clough and Bent, 1998).
MAIN reporter gene constructs
Addition of NcoI sites to the MAIN cDNA using primers MAIN-cDNA–N and MAIN-cDNA–C replaced the stop codon by a codon for glycine. This cDNA was cloned into pSO35e (Klepek et al., 2005) upstream of the GFP ORF to create pSF1. For the N–terminal GFP fusion, the MAIN coding sequence was amplified using primers MAIN-gene–N and MAIN-gene–C and cloned into pSO35em, in which the GFP stop codon has been replaced by a BglII site, to create pDW10. The pSF1 and pDW10 plasmids were particle-bombarded into epidermis cells of Nicotiana tabacum (pSF1, pDW10), Arabidopsis thaliana (pDW10) and Allium cepa (pDW10) (Klepek et al., 2005). MAIN sequences with modified or deleted NLS were amplified from genomic DNA using the MAIN-gene–N primer and the primer At1g17930-KLNQR (changing KRKRR into KLNQR to create pSF1b), or At1g17930c1389r (introducing a stop codon in front of the NLS to create pDW10a.
For construction of pMAIN::MAIN-GFP and pMAIN::MAIN-GUS, a 3254 bp fragment was amplified from genomic DNA usig primers At1g17930 g-1465f and At1g17930g+1792r, and cloned into pMDC107 and pMDC163 (Curtis and Grossniklaus, 2003), yielding pUW16 and pUW17.
GUS staining and histological analysis
GUS staining was performed for at least 1 h as described previously (Weingartner et al., 2011). For histological analyses, roots and SAMs of seedlings at 3 DAG and stems of mature plants were fixed and embedded as described previously (Scheres et al., 1994), and 20 μm sections were cut using a Leica RM 2135 rotary microtome (Leica Microsystems, http://www.leica-microsystems.com). SAMs were additionally stained with Astra Blue as described previously (Scheres et al., 1995).
RNA isolation and quantitative real time-PCR
RNA was isolated from root tips of agar-grown WT and mutant seedlings at 3 DAG using an RNeasy plant mini kit (Qiagen, http://www.qiagen.com), and reverse-transcribed using a QuantiTect reverse transcription kit (Qiagen). Quantitative real time-PCR was performed using Brilliant III Ultra-Fast SYBR Green QPCR Master Mix (Agilent Technologies, http://www.agilent.com) and a Rotor-Gene Q real-time cycler (Qiagen). Expression levels were calculated relative to ACT2. Primers used for quantitative real time-PCR are listed in Table S3.
Microscopy and fixation
Images were obtained using a Zeiss Axioskop (Carl Zeiss, http://zeiss.com) or Leica MZFLIII (Leica Microsystems). Cell walls were stained with PI as described by Stadler et al. (2005a) or with mPS–PI as described by Truernit et al. (2008). Confocal images were obtained using a Leica TCS SP5 confocal microscope (Leica Microsystems). The areas of dead cells were quantified using Leica LAS AF software. Embryos were fixed in chloral hydrate solution (chloral hydrate/water/glycerol 8:3:1) for 20 min, and examined using the Leica TCS SP5 confocal microscope with a differential interference contrast filter.
The comet assay was performed using the A/N protocol as described previously (Menke et al., 2001; Olive and Banath, 2006).
We thank Rebecca Günther, Ingrid Schiessl, Gudrun Steingräber (University of Erlangen-Nürnberg, Molekulare Pflanzenphysiologie, Germany) for experimental help, Philip N. Benfey (Duke University, Department of Biology, Durham, NC) for pSCR::erGFP seeds, Thomas Laux (University of Freiburg, Faculty of Biology, Germany) for pWOX5::erGFP seeds, Klaus Palme (University of Freiburg, Germany) for pPIN1::PIN1-GFP seeds, Markus Geißler (University of Fribourg, Department of Biology-Plant Biology, Switzerland) for pDR5::GFP seeds, László Bögre (Royal Holloway University of London, School of Biological Sciences, UK) for the pCYCB::GUS construct, and Ben Scheres (Utrecht University, Department of Biology, The Netherlands) for pPLT1::CFP seeds. This work was supported by grants from the Deutsche Forschungsgemeinschaft to N.S. (SA 382/8 and SA 382/22).