Adventitious shoot organogenesis contributes to the fitness of diverse plant species, and control of this process is a vital step in plant transformation and in vitro propagation. New shoot meristems (SMs) can be induced by the conversion of lateral root primorida/meristems (LRP/LRMs) or callus expressing markers for this identity. To study this important and fascinating process we developed a high-throughput methodology for the synchronous initiation of LRP by auxin, and subsequent cytokinin-induced conversion of these LRP to SMs. Cytokinin treatment induces the expression of the shoot meristematic gene WUSCHEL (WUS) in converting LRP (cLRP) within 24–30 h, and WUS is required for LRP → SM conversion. Subsequently, a transcriptional reporter for CLAVATA3 (CLV3) appeared 32–48 h after transfer to cytokinin, marking presumptive shoot stem cells at the apex of cLRP. Thus the spatial expression of these two components (WUS and CLV3) of a regulatory network maintaining SM stem cells already resembles that seen in a vegetative shoot apical meristem (SAM), suggesting the very rapid initiation and establishment of the new SMs. Our high-throughput methodology enabled us to successfully apply a systems approach to the study of plant regeneration. Herein we characterize transcriptional reporter expression and global gene expression changes during LRP → SM conversion, elaborate the role of WUS and WUS-responsive genes in the conversion process, identify and test putative functional targets, perform a comparative analysis of domain-specific expression in cLRP and SM tissue, and develop a bioinformatic tool for examining gene expression in diverse regeneration systems.
Plants are able respond to environmental challenges with impressive metabolic and developmental flexibility. Adaptable development is achieved by controlling the activities of stem cell niches (meristems) in root and shoot, and regulating the growth of tissues produced from them. In addition to primary root and shoot apical meristems (RAMs and SAMs), reiterative development from lateral root primordia (LRP) in the root and axillary meristems in the shoot is regulated in response to diverse cues and signaling inputs (reviewed in McSteen and Leyser, 2005; Scheres, 2007). Remarkably, many plant species also initiate adventitious root and shoot meristems (SMs) de novo (reviewed in Steeves and Sussex, 1989; Kerstetter and Hake, 1997) for clonal propagation (e.g. poplar), or to recover from injury. In the root system, adventitious shoots can be initiated from pericycle-derived cells, LRP and RAMs (Bonnet and Torrey, 1966; Peterson, 1970; Spencer-Barreto and Duhoux, 1994).
Artificial induction of adventitious shoots using auxin and cytokinin was first developed over 60 years ago (Skoog, 1950; Skoog and Miller, 1957), and remains a vital step in micropropagation and transformation protocols. Shoot organogenesis is generally induced from callus by cytokinin treatments. Callus was long considered an essential step in in vitro organogenesis, providing a mass of apparently disorganized and undifferentiated cells amenable to ‘redifferentiation’. However, recent studies have shown that callus has much in common with LRP, in its proliferation from xylem-pole pericycle cells and gene expression patterns (Che et al., 2007; Atta et al., 2009; Sugimoto et al., 2010). Furthermore, shoot organogenesis has also been induced from LRP/LRP-like organs without an intervening callus stage (Atta et al., 2009).
Shoot organogenesis can also be induced by genetic elevation of endogenous cytokinin levels (Zuo et al., 2002), or misexpression of specific SAM-associated genes, such as DORNRÖSCHEN (DRN) (Banno et al., 2001) and WUSCHEL (WUS) (Gallois et al., 2004). In loss-of-function wus mutants, stem cells of the SAM are consumed shortly after germination (Laux et al., 1996), and WUS is functionally required for shoot organogenesis from callus (Gordon et al., 2007). Within the SAM, WUS is expressed in the organizing centre and the protein migrates to overlying cells of the central zone, CZ (Yadav et al., 2011), where it specifies stem cell fate (Mayer et al., 1998). Within the CZ WUS activates CLAVATA 3 (CLV3), which negatively regulates WUS expression via the CLAVATA pathway, a feedback loop that helps maintain a constant population of stem cells (Fletcher et al., 1999; Brand et al., 2000). When WUS is misexpressed in the root, or induced with cytokinin, it also stimulates the expression of CLV3 in adjacent cells (Gallois et al., 2004; Gordon et al., 2007).
Recent studies have enhanced our understanding of the relationship between WUS and cytokinin in SAM function and shoot organogenesis. Type-A ARABIDOPSIS RESPONSE REGULATORS (ARRs) are negative regulators of cytokinin signaling (To et al., 2004), and are transcriptionally repressed by WUS (Leibfried et al., 2005; Busch et al., 2010). In characterizing cytokinin-induced WUS expression, Gordon et al. (2009) found CLV-dependent and -independent mechanisms were involved, primarily mediated through cytokinin receptors ARABIDOPSIS HISTIDINE KINASE 2 (AHK2) and AHK4.
In addition to cytokinin-signaling genes, WUS binds at least two different motifs to regulate the transcription of numerous targets (Busch et al., 2010). Furthermore, WUS stimulates transcription of TOPLESS (TPL) (Busch et al., 2010), which modulates auxin signaling and represses root-specifying genes (Long et al., 2002; Szemenyei et al., 2008; Smith and Long, 2010). Thus WUS appears to modulate aspects of cytokinin and auxin signaling, two plant hormones that play key antagonistic roles in root and shoot organogenesis.
In this study we adapt a method for synchronous LRP induction and couple it to cytokinin-mediated WUS upregulation to provide a high-throughput system for studying shoot organogenesis via LRP → SM conversion. Expression of transcriptional reporters for CLV3 and WUS identified key stages in LRP → SM conversion, guiding transcriptome analysis and revealing potential targets mediating the process. Furthermore, because there is a functional requirement for WUS in SM initiation from callus/LRP, we analyzed the transcriptomes of wus loss-of-function mutants, and WUS expression domains within cLRP, identifying genes transcriptionally responsive to WUS activity (WUS responsive) involved in this process. In addition to enhancing our understanding of shoot organogensis, and the role of WUS, it is hoped that our work on this intriguing developmental phenomenon will also inform regeneration in natural systems.
Incipient stem cell niche conversion
To study the cytokinin-induced conversion of LRP/LRMs to shoots we first generated synchronous LRP initiation by exposing 3- or 4-day-old seedlings to 10 μm 1-naphthaleneacetic acid (1-NAA), a synthetic auxin used in transcriptomic studies of LRP (Himanen et al., 2004). Seedlings have few LRP prior to NAA treatment, which initiates LRP at every available position along the primary root xylem-pole pericycle. Transfer to cytokinin-enriched media induces the synchronous conversion of induced LRP into SMs, and emerging shoots become visible within 5 days (Figure 1a–c). This simple methodology appears robust, and we have used it to rapidly induce shoots from roots of other brassicas and poplar (Figure 1d).
A high-throughput methodology, adapted from Birnbaum et al. (2005), permitted the transfer of hundreds of plants between hormone treatments and rapid sampling (Figure 1a,b). In agreement with previous studies, 24 h after auxin treatment the LRP induced by the treatment reached between three and five cell layers, a stage that precedes commitment to self-sustaining LRMs (Sussex et al., 1995).
WUSCHEL reporters are expressed in cLRP within 30 h
In our experiments we used well-characterized transcriptional reporters for WUS and CLV3, with expression patterns reflecting mRNA in situ hybridization experiments (Reddy and Meyerowitz, 2005; Gordon et al., 2007; Yadav et al., 2009). Within the treated roots, WUS reporters were first visible 19 h after transfer from NAA to isopentenyladenine (2iP), and were weakly and sporadically expressed outside the LRP (Figure 2a). WUS reporter expression was first seen within cLRP 24–36 h after transfer to 2iP (Figure 2b), but rarely in the outermost layer of cells (2C–E). Expression of this transcriptional reporter for the organizing center of the SAM within former LRP shows that an important change in cell identity, towards that associated with SMs, is already underway just a day after exposure to cytokinin.
A pCLV3::GFP-ER reporter appeared 32–48 h after transfer to 2iP marking between two and four cell layers at the apex of cLRP, above and overlapping with populations of small cells expressing pWUS::DsRED-N7 (Figure 2c,d), and was never seen in the absence of the WUS reporter, consistent with previous studies that have found CLV3 expression to be dependent upon the WUS gene product (Laux et al., 1996; Brand et al., 2002). Thus, transcriptional reporter expression of these key regulators of the SAM stem cell population assumed a spatial relationship within cLRP, reflecting that in SAMs. Expression of these two components of a regulatory network responsible for maintaining a shoot meristematic stem cell population constitutes another significant step in the establishment of a new SM.
Transcriptome analysis of LRP–SM conversion reveals changes related to hormone signaling, meristem identity, cell cycle and photosynthesis
By enriching samples with synchronously developing LRP, we hoped to focus on gene expression pertaining to LRP–SM conversion. For transcriptome analysis, key time points in LRP → SM conversion were selected based on the aforementioned reporter analysis. Tissues were sampled after 24 h of exposure to NAA (0 h 2iP), and subsequently after 19, 30 and 48 h of 2iP treatment. The first time point corresponds to saturated LRP development within the primary roots, and 19 h of 2iP treatment corresponds with the initial expression of pWUS::DsRED-NLS. After 30 h of treatment with 2iP there is consistent expression of this marker within cLRP prior to the detection of pCLV3::mGFP-ER. Treatment with 2iP for 19 h precedes the detectable expression of SAM markers within cLRP, perhaps corresponding to an intermediate state between LRP and initiating SMs. We reasoned that a 30 h time point would reveal gene expression events associated with early SM initiation. After 48 h of treatment with 2iP the CLV3 marker is expressed within 10–25% of cLRP, reflecting gene expression representing the initial establishment of organized SMs.
Exogenous auxin and cytokinin were expected to generate marked changes in gene expression. Because expression of key developmental regulators could be relatively small when ranked against this background, lenient criteria were used for the initial selection of differentially expressed genes (DEGs). Using MAS5 processed data, each time point was compared pairwise with every other time point, applying a fold-change threshold of 1.75, a P-value cut-off of <0.05 and the rejection of mean expression values <50 units.
Figure 3 provides an overview of DEGs identified in each comparison (also Table S1). As expected, large numbers of DEGs (2399–3472) were identified in comparisons between 0 h and 2iP time points. There was also a marked overlap in the DEGs from each of these comparisons, with 1700 DEGs identified in all three (Figure 3). For comparisons between cytokinin treatment periods, 340 and 642 DEGs were identified between 19 and 30 h, and between 19 and 48 h, respectively. Interestingly, although the 30 and 48 h time points of 2iP treatment separate important changes in reporter gene expression, no DEGs were found in this comparison.
The expression of positive cell-cycle regulators and RM-associated genes is decreased by cytokinin, whereas the expression of photosynthetic and SAM-associated genes is increased
Consistent with the presumed changes in identity occurring as 2iP promotes LRP → SM conversion, reduced expression of many RM/LRP-associated genes and increased expression of SM-associated genes were observed (Table S2). However, several key SAM-associated genes, including CLV3, did not pass the expression value cut-off. Misexpression of CLV3 has been shown to precipitate consumption of the RAM (Fiers et al., 2005), but CLV3 reporter expression, and low expression values of CLV3, suggest it does not play a key role in the initial loss of LRM identity.
Unsurprisingly, transferring seedlings from high-auxin to high-cytokinin media is reflected in the increased expression of many cytokinin-responsive signaling genes, and in the reduced expression of auxin-induced signaling and metabolic genes (Tables S3 and S4, and over-represented gene ontology, GO, categories in Table S1), suggestive of an involvement in maintaining hormone signaling or metabolic homeostasis.
Cytokinins and certain cytokinin-signaling components promote differentiation of chloroplasts and expression of photosynthetic genes (Schmulling et al., 1997; Argyros et al., 2008), and all photosynthesis-related plant-encoded DEGs were found to be increased in steady-state transcript levels by exogenous 2iP in our study (over-represented GO categories in Table S1).
The reduced expression of type-A and -B cyclins and cyclin-dependent kinases, and the increased expression of three cyclin-dependent kinase inhibitors (Table S5), suggests a reduction in cell division/numbers of dividing cells on 2iP. Reduced expression of other positive regulators of cell division, such as EF2a, and decreased expression of many histones (e.g. S-phase marker HIS4; Table S5) support this interpretation. These observations are consistent with studies showing that cytokinins inhibit cell division in RMs and LRP founder cells (Werner et al., 2003; Li et al., 2006; Dello Ioio et al., 2007).
Gene ontology (GO) enrichment analysis revealed transcription factors were over-represented amongst DEGs in most pairwise comparisons (Table S1). This category contained 61 differentially expressed homeobox genes, some of which have known roles in meristem and organ initiation. For example, WOX13 is dynamically expressed during RAM/LRP initiation, and showed decreased expression on 2iP. Conversely, ARABIDOPSIS THALIANA HOMEOBOX 1 (ATH1) and PENNYWISE (PNY) interact with SHOOT MERISTEMLESS 1 (STM1) in the SAM, and their expression was increased.
Comparison with callus-based regeneration reveals an overlap with DEGs identified in LRP→SM conversion
To identify potential key regulators of shoot regeneration common to different in vitro systems, we compared targets identified in studies of shoot organogenesis from callus with LRP → SM conversion. Che et al. (2006) analysed transcriptome changes during root or shoot organogenesis from callus, and described the ‘top-20’ DEGs with increased/decreased expression during callus induction, or subsequent shoot or root induction. Of the top-20 DEGs identified during presumed commitment to shoot organogenesis, 11 and 12 of the genes with increased and decreased expression, respectively, also appeared amongst the DEGs identified in our study, with similar patterns of expression (Table S6).
We then surveyed genes previously identified as affecting shoot organogenesis for differential expression during LRP → SM conversion (Table S7). One of these, HLS1/COP3, is a negative regulator of callus-based shoot regeneration (Chatfield and Raizada, 2008), and exhibits >14-fold decrease in expression levels on 2iP in our study (Table S7). Interestingly, RSM1 overexpression phenocopies loss of HLS1 function (Hamaguchi et al., 2008), and RSM1 expression was increased more than eightfold on 2iP.
WUS is required for LRP → SM conversion in the high-throughput system
Misexpression of WUS is sufficient to generate ectopic shoots in root tissue (Gallois et al., 2004), and is functionally required for shoot organogenesis from root-derived callus (Gordon et al., 2007). We have tested the effect of three wus loss-of-function mutant alleles (Methods S1) in the high-throughput LRP–SM system, and found an absolute requirement for a functional copy of the gene (Figure 4). Our data also suggest a positive gene dosage effect of WUS on the numbers of SMs generated (Figure 4).
Transcriptome analysis of wus mutants identifies WUS-responsive DEGs
To further examine the role of WUS in LRP → SM conversion, we compared transcriptomes of loss-of-function wus mutants with the wild type (WT). We first compared a reported wus null allele, SAIL_150_G06 (McElver et al., 2001; Sonoda et al., 2007), with WT using the aforementioned 2iP treatment periods. Three biological replicates were recorded with Affymetrix ATH1 microarrays, and mas5 processed data analysed using the limma (linear models for microarrays) package (Smyth, 2005). A significance cut-off of P < 0.05, a minimum fold change >1.5 and a minimum expression value of 50 yielded 543 DEGs in total.
Figure 5 shows the distribution and overlap of the DEGs identified, and a summary of over-represented GO categories (details Tables S8 and S9). Using our lenient selection criteria, between 63 and 121 genes were found to have increased or decreased steady-state transcript levels in the mutant at each time point. So, although the developmental consequences of wus loss of function are dramatic, the perturbation of gene expression was small compared with that associated with the hormone treatments used to induce LRP and conversion. Additionally, unlike our WT time course, very little overlap was found between time points in terms of DEGs identified, suggesting that examining downstream consequences of wus loss of function successfully focused on transcriptome changes relevant to discrete stages in WUS-dependent development.
After 30–48 h of 2iP treatment, the WUS reporter was consistently expressed in WT cLRP and the most relevant transcriptome differences were anticipated at these times. Amongst the DEGs identified in these comparisons, the most over-represented GO categories included the biological processes of post-embryonic development, apotosis, post-translational protein modification, glucoside biosynthesis, phenylpropanoid biosynthesis and responses to light. This output resembles the groups of GO categories described by Busch et al. (2010) in a genomic study identifying WUS-responsive genes: namely, the regulation of development (including meristem and cell death), metabolism (including glucosinolate) and response to stimuli. Furthermore, the distribution of WUS-repressed and -induced genes described by Busch et al. (2010) amongst the DEGs we identified (Figure 6) suggests that our approach has been successful in identifying a subset of WUS-responsive DEGs during LRP → SM conversion. After 19 h of 2iP treatment, WUS reporter expression is weak and sporadic in tissues around cLRP, and the ratio of WUS-repressed and -induced genes amongst DEGs suggests WUS loss of function has not yet directly affected the transcription of targets (Figure 6). Conversely, after 30 h of 2iP treatment the proportion of WUS-repressed genes amongst the DEGs with higher expression in wus increases to more than fivefold that of WUS-induced genes, suggesting that the loss of WUS has permitted elevated expression of these genes. Conversely, although the number of WUS-repressed genes has decreased amongst DEGs with reduced expression in wus at this time, they outnumber WUS-induced genes. However, after a further 18 h, WUS expression within WT cLRP indicates that >11% of the DEGs with reduced expression in wus belong to the WUS-induced group, and none to the WUS-repressed group (Figure 6). Furthermore, after 48 h of 2iP treatment the proportion of WUS-repressed genes amongst DEGs with increased expression in wus remained higher than WUS-induced genes. In addition, we surveyed promoter and intron sequences of DEGs for cis-element sequences bound by WUS (Lohmann et al., 2001; Busch et al., 2010), and found them to be over-represented amongst DEGs after 48 h of 2iP. Two or more instances of the 6-bp sequence CACGTG (Busch et al., 2010) were found within 500 bp upstream of 5.4% (1.7% expected, P = 0.003) of the DEGs with increased expression in wus, and two or more instances of the sequence TTAATSS (Lohmann et al., 2001) were found within the introns of 7.94% (3.45% expected) of DEGs with decreased expression in wus, although the latter observation was not deemed significant (P = 0.055).
Thus, for a subset of genes the effect of WUS upregulation within cLRP seems broadly consistent with published findings on the regulation of gene expression by WUS. In addition, we have identified many targets not previously identified as WUS responsive that are differentially regulated in the loss-of-function mutant under the specific conditions associated with LRP → SM conversion (Table S8, and GO analysis in Table S9).
As might be expected, DEGs included those associated with developmental processes in SMs (Table S8), including CUP-SHAPED COTYLEDONS 1 and 3 (CUC1 and CUC3) and BLADE ON PETIOLE 2 (BOP2). However, many meristem-associated genes were expressed at low levels, and despite enrichment of samples with cLRP, SAM-associated genes positively regulated by WUS (e.g. CLV3; Brand et al., 2002; Yadav et al., 2011), or repressed by WUS (e.g. CLV1; Busch et al., 2010), were not identified as DEGs. It may be that the relevant cell types still represent an insubstantial fraction of samples, or that our sampling precedes the significant upregulation of many meristem-associated genes.
An important role of WUS in meristem function is believed to be the regulation of cytokinin-inducible ARRs. Leibfried et al. (2005) used inducible misexpression to isolate WUS-responsive genes, and identified four type-A ARR genes (ARR5, ARR6, ARR7 and ARR15) as WUS-repressed. In contrast, no ARR genes were amongst DEGs with higher expression in wus, and few cytokinin-related targets were identified as differentially expressed after 30–48 h of treatment with 2iP (Table S8), from which it is difficult to infer an outcome upon cytokinin-signaling output. This discrepancy may reflect different tissues sampled, and cytokinin treatments masking the impact of WUS on ARR expression in our study.
To identify associations and putative functional relationships between DEGs, cluster analysis was performed and each cluster analyzed for over-represented GO terms. The DEGs were grouped into 18 clusters by k-means clustering, seven of which yielded statistically over-represented GO terms (P < 0.05 Hochberg false discovery rate; Figure S1; Table S10). Although many GO categories overlapped with those identified in our time-point comparison (Table S9), several interesting new groups were highlighted. These included: lipase activity (cluster 5), nuclear protein import (cluster 6) and, endo-1,4-β-xylanase activity (cluster 7). Endo-1,4-β-xylanases are associated with cell expansion and shape changes, and inclusion of three (of five on the ATH1 microarray) within cluster 7 suggests WUS, or dependent processes, reduce these activities.
After 30 h of treatment with 2iP, we hoped to identify early events in WUS-dependent LRP → SM conversion. To provide greater resolution of WUS-related DEGs at this time point, we examined the transcriptome of another wus mutant allele (GABI_870H12). GABI-KAT constructs were designed for activation tagging, but in this line an intragenic insertion appears to drive elevated expression of a truncated non-functional product (Methods S1). Heterozygotes yield loss-of-function wus phenotypes in approximately 25% of progeny, and these homozygous mutants are unable to undergo LRP → SM conversion. For comparison of the two alleles with the WT, processed data were filtered to remove genes absent in one wus mutant allele, but not the other. Using the limma package of BioConductor, a P-value threshold of <0.05 and a minimum fold change of 1.5 in one genotype, 144 DEGs similarly regulated in both wus mutant alleles were identified (Table S11). Of these initial DEGs, 21% overlapped with those identified in our wus SAIL/WT comparison. Applying the fold change cut-off to both alleles increased overlap to 37%, comprising 25 and 48% of DEGs up- or downregulated in a wus mutant background, respectively. The observed differences in overlap between genes with increased or decreased levels of expression could reflect differences in the function of the mutant gene products, but as both mutant alleles seem functionally similar in terms of LRP → SM conversion, the subset of mutual DEGs appears to offer stronger candidates for genes mediating WUS-dependent LRP → SM development. Consistent with this view, the proportion of WUS-induced to WUS-repressed genes amongst DEGs with reduced expression in wus increased in this two-allele comparison (Figure 6).
Insertional knock-outs in WUS-responsive candidates affect LRP → SM conversion
To explore the roles of putative WUS-responsive targets identified by transcriptome analysis, we refined selection criteria to test insertional knock-outs of promising targets. To enrich for potential direct targets of WUS, we surveyed <1 kb upstream of the wus mutant DEGs for two or more instances of sequences corresponding to putative WUS-binding cis-elements: CACGTG and TTAATSS. As functional TTAATSS sequences were originally identified within an intron, we included DEGs with two or more instances within introns. These candidates were then surveyed with the Arabidopsis eFP Browser (Winter et al., 2007) for genes displaying differential expression in the SAM (Yadav et al., 2009) or embryo (Casson et al., 2005). Initially, 32 wus mutant DEGs were selected, of which 28 possessed corresponding T-DNA insert lines. Homozygous lines were assayed for LRP → SM conversion by scoring the numbers of shoots initiated on seedling roots treated with 4.4 or 2.2 μm 2iP for 5–7 days. The lower cytokinin concentration was included to screen for enhanced shoot-induction rates. A total of 39 homozygous insertion lines, corresponding to 27 genes, were tested in at least two replicates (Figure 7; Table S12). Four of these lines, corresponding to four different DEGs with reduced expression in wus during LRP → SM conversion, showed a consistent reduction in shoot initiation rates (Figure 7). All mutants appear phenotypically normal, apart from their deficit in shoot initiation. However, further work will be required to determine the relevance of the mutations to WUS-dependent processes and LRP → SM conversion.
Cell-specific profiling of the WUSCHEL domain of cLRP
Cell-specific expression profiling has been used to explore gene expression within specific domains of RMs (Birnbaum et al., 2003; Brady et al., 2007; Gifford et al., 2008) and SMs (Yadav et al., 2009), with improved sensitivity and resolution in relating transcriptome changes to development and responses to stimuli. Although our samples were enriched with cLRP, the relative contribution by key cell types/domains may be insufficient to resolve key genes, illustrated by low WUS expression values. We therefore isolated and profiled pWUS::mGFP5-ER cells to compare expression in the WUS domain of cLRP with the WUS domain of established SMs.
Tissues harvested after 30 h of treatment with 2iP were protoplasted rapidly (1 h) and cells expressing pWUS::mGFP5-ER were isolated with a fluorescence-activated cell sorter (FACS). Isolated RNA underwent two cycles of amplification and transcriptomes were recorded with Affymetrix ATH1 arrays. Previous studies, using RM and SM cells, have identified genes that respond to protoplasting with changes in expression (Birnbaum et al., 2003; Yadav et al., 2009), and these targets were removed from comparisons.
The mean expression value of WUS in pWUS::mGFP5-ER cells was about sevenfold higher than the whole root, suggesting successful enrichment for WUS-expressing cells, but was lower than that obtained for the WUSp domain of SAMs (Yadav et al., 2009). This latter observation was expected because the expression of transcriptional reporters for WUS only began in cLRP at this time.
Pearson correlation coefficients of 0.97–0.99 for comparisons indicate a high level of reproducibility between our pWUS::mGFP5-ER replicates (Table S13). Lower correlation coefficients (0.513–0.596) were found between pWUS::mGFP5-ER cells in our experiments and those isolated by Yadav et al. (2009). In addition to the likely differences in expression arising from harvesting WUS-expressing cells from different organs, low correlation values probably reflect differing culture conditions, particularly the high concentrations of hormones used in our experiments.
Next we identified genes differentially expressed within pWUS::mGFP5-ER cells from cLRP compared with whole root samples, and examined how these genes were distributed amongst those assigned to SAM domains (Yadav et al., 2009). Figure 8 shows that of the genes previously assigned to each SAM domain, the highest proportion of overlap with the DEGs from pWUS::mGFP5-ER cells is the WUSp domain of the SAM at 60.3%, compared with 35.7 and 45.5% for CLV3p and FILAMENTOUS FLOWER (FILp) domains, respectively. Moreover, a higher proportion of the overlapping genes in the WUS SAM domain are DEGs, with higher expression in pWUS::mGFP5-ER cells from cLRP: 42.0%, compared with 14.8 and 14.1% in CLV3 and FIL domains, respectively. This suggests that within hours of initiating WUS reporter expression in cLRP, the transcriptome of these cells began to resemble theWUSp domain of an SAM.
Examining the distribution of sequences associated with WUS cis-elements we found the 6-bp sequence CACGTG was significantly over-represented 500–1000 bp upstream of DEGs from pWUS::mGFP5-ER cells in cLRP (Figure 8). Two or more TAATTSS sequences within introns were also observed at a higher frequency than expected, but the cut-off of P < 0.05 was not met (P = 0.056). Enrichment of DEGs from this domain with elements mediating transcriptional responses to WUS is consistent with rapid transcriptional responses to WUS expression within cLRP.
We then examined the distribution WUS-responsive genes identified in our transcriptome analysis of wus mutants, and those identified by Busch et al. (2010), amongst DEGs from pWUS::mGFP5-ER cells in cLRP. Interestingly, genes found to have increased expression in pWUS::mGFP5-ER cells were enriched amongst those found to have increased expression in wus (Figure 9a). Conversely, those with decreased expression in pWUS::mGFP5-ER cells were enriched amongst those with decreased expression in wus (Figure 9a). Similarly, WUS-induced and WUS-repressed genes (Busch et al., 2010) were found to be enriched amongst those with decreased and increased expression, respectively, in pWUS::mGFP5-ER cells (Figure 9b). These findings mirror those of Busch et al. (2010), who found WUS-repressed genes were enriched within the WUS domain of the SAM, whereas WUS-induced genes were enriched among transcripts expressed in the combined CLV3 and WUS domains, and combined CLV3 and FIL domains. It was suggested that this finding reflected a contribution by both direct and indirect targets amongst WUS-responsive genes, and linked the prevalence of transcripts with reduced expression in the WUS domain to evidence that WUS acts primarily as a transcriptional repressor (Leibfried et al., 2005; Ikeda et al., 2009), modulating target gene expression. Our findings are in line with these hypotheses, and may also reflect the duration of WUS expression in cLRP and WUS non-cell autonomous activity (Mayer et al., 1998; Gallois et al., 2004; Yadav et al., 2011).
To characterize transcriptome differences between WUS domains within established SAMs and within cLRP we compared pWUS::mGFP5-ER cells in our experiment with WUSp SAM cells (Yadav et al., 2009). pWUS::mGFP5-ER-associated DEGs with more than twofold difference in expression comprised 1224 DEGs with elevated expression in cLRP, compared with WUSp SAM, and 1452 DEGs with lower expression (Table S14). GO analysis (Tables 1 and S15) revealed the enrichment of various ‘response to stimuli’ terms, including the responses to hormones (ABA, ethylene, cytokinin and auxin), which may reflect the exogenous hormones applied in our study. Differential over-representation of root versus shoot developmental terms (Table 1) amongst DEGs from cLRP is consistent with continuing conversion of the treated root tissues. Over-representation of the GO categories for meristem initiation, and meristem structural organization amongst DEGs with lower expression in the WUS domain of cLRP, may also reflect the transitional state of these organs.
Table 1. Selected enriched gene ontology (GO) terms amongst differentially expressed genes (DEGs) identified in the transcriptome of fluorescence-activated cell sorted (FACS) cells expressing pWUS::mGFP-ER from lateral root primordia (LRP) undergoing conversion to shoot meristems (SMs; after 30 h of treatment with 2iP), compared with the WUSp expression domain of apetala1/cauliflower double mutant SMs (Yadav et al., 2009). GO analysis was performed using agriGO (http://bioinfo.cau.edu.cn/agriGO)
Singular enrichment analysis was used with Fisher's statistical test, Yekutieli multi-test adjustment and a minimum significance level of P < 0.05.
Selected enriched GO terms. DEG >2 fold UP in pWUS::GFP 30 h 2iP versus SAM WUSp (Yadav et al., 2009)
Response to chemical stimulus
Response to stress
Toxin catabolic process
Response to hormone stimulus
Glycoside metabolic process
Response to reactive oxygen species
Selected enriched GO terms. DEG >2 fold DOWN in pWUS::GFP 30 h 2iP versus SAM WUSp (Yadav et al., 2009)
ncRNA metabolic process
Meristem structural organization
Photosynthesis, light harvesting
Veg. to reprod. phase transition of meristem
Meristem initiation/organization includes targets interacting with WUS, or transcriptionally modulated by it. TOPLESS (TPL) and TOPLESS-RELATED 4 (TPR4) encode for transcriptional co-repressors that interact with WUS (Kieffer et al., 2006). Expression of TPL is enhanced by WUS (Busch et al., 2010), and directly targets PLETHORA 1 and 2 (PLT1 and PLT2) genes (Smith and Long, 2010), which are master regulators promoting basal/root fate (Aida et al., 2004; Galinha et al., 2007). Two other WUS-induced meristematic genes, STM1 and a CYCLIN-DEPENDENT KINASE B2;1 (CDKB2;1) were also found to have lower expression in the newly initiated WUS-reporter domain. CLV1 is a direct target of WUS, which represses CLV1 expression (Busch et al., 2010). If factors that positively regulate the expression of CLV1 within the SAM are absent in cLRP this might account for the lower expression of CLV1 in the WUS domain of these organs. As a shoot stem cell population, marked by CLV3 reporter expression, has not developed at this time, it is unsurprising that expression of other elements of the WUS–CLV pathway have yet to be established, including CLV1 and POLTERGEIST (POL) (Song et al., 2006).
Other interesting enriched biological processes amongst DEGs with lower expression in cLRP included: non-coding RNA processing, RNA methylation and histone modification. The establishment of a functional shoot stem cell niche can be expected to involve coordinated regulation of transcription, and the stability and functional output of numerous interacting targets. The categories histone modification, RNA methylation and ncRNA categories may include genes mediating these processes in initiating or established SAMs that have yet to be upregulated in LRP.
In vitro shoot regeneration and adventitious shooting in diverse natural systems can occur through the conversion of LRP or RMs (Bonnet and Torrey, 1966; Peterson, 1970; Spencer-Barreto and Duhoux, 1994; Atta et al., 2009; Sugimoto et al., 2010). To investigate this phenomenon we used a high-throughput methodology for synchronous LRP → SM conversion, enabling us to apply a systems approach to further understanding the process.
Characterizing reporter expression for WUS and CLV3 identified important stages in the conversion of LRP into SMs, permitting transcriptome analysis to elaborate the critical role played by WUS in the process. The appearance of WUS reporter expression, prior to the initiation of a shoot stem cell population marked by pCLV3::GFP-ER, suggests that the upregulation of CLV3 does not play a primary role in inhibiting LRP development and initial stages of conversion. Cell-specific profiling and comparison of WUSp domains in cLRP and SMs indicated that the expression of many shoot meristematic genes, including other elements of the CLV pathway, are not yet induced when WUS reporter expression is initiated in cLRP. However, the over-representation of sequences associated with cis-elements bound by WUS amongst DEGs suggests that upregulation of WUS within cLRP drives rapid changes in the transcriptional activity during conversion. Similarly rapid changes in gene expression associated with the re-specification of cell identity have been shown during regeneration of RAMs (Sena et al., 2009).
Transcriptome analysis of wus loss-of-function mutants and cell-specific profiling permitted us to remove the background of gene expression perturbed by exogenous hormone treatments, and focus upon the DEGs reflecting important developmental transitions and the role of WUS in them. These studies revealed differential regulation of known WUS-responsive genes and identified new putative WUS-responsive targets associated with LRP → SM conversion. In addition to reflecting our specific culture conditions and ectopic expression of WUS in root tissues, it is hoped that some of these new targets may be uniquely associated with the WUS-dependent conversion process.
A scan for candidates with a functional role in LRP → SM conversion was enriched for potential WUS-interacting genes by cross-referencing our putative WUS-responsive targets with available embryo and SAM-related transcriptome data, and for enrichment with WUS-binding cis-element sequences. This approach identified four homozygous mutant lines that negatively impacted shoot induction rates via LRP → SM conversion. Further work will be required to determine whether the associated genes do indeed make a functional contribution to LRP → SM conversion downstream of WUS. Because high-throughput LRP → SM conversion is likely to be dependent upon appropriate responses to auxin and cytokinin, and other response networks affecting growth in culture, a relatively high proportion of mutations may affect the process.
Overall, we have laid important groundwork in characterizing key transitions early in the conversion of incipient root stem cell niches to SMs, and have identified targets affecting the process. The approach offers a tractable system to investigate questions pertaining to pluripotency, cell fate reprogramming and stem cell niche patterning. Historically, in vitro organogenesis has provided an invaluable tool for research and biotechnology. To facilitate further comparative analysis of regeneration systems, and provide a resource for the community, we have constructed a ‘Regeneration’ eFP browser (http://bar.utoronto.ca/efp_arabidopsis/cgi-bin/efpWeb.cgi?dataSource=Regeneration; see Figure S2 for an example visualization) for visualizing gene expression in our data sets alongside those obtained in studies of callus-based systems (Che et al., 2006; Sugimoto et al., 2010), cell-specific profiling (Yadav et al., 2009) and RAM regeneration (Sena et al., 2009).
Plant materials and growth conditions
Arabidopsis thaliana ecotypes Columbia (CS1092), Landsberg erecta (CS20), line WUSpro:GFP-ER (CS23897; Jonsson et al., 2005), line WUSpro:DsRed-N7 CLV3pro:GFP-ER (CS23895; Gordon et al., 2007), wus-1 (CS15) and wus null allele SAIL_150_G06 (CS807292), and all SALK and SAIL T-DNA mutant lines (Figure 7; Table S12), were provided by the Arabidopsis Biological Resource Center (ABRC, http://abrc.osu.edu), and wus GABI-KAT allele (GABI_870H12) was provided by the Nottingham Arabidopsis Stock Centre (NASC, http://arabidopsis.info).
Seedlings were grown aseptically on 200-μm Nitex mesh sheets (Sefar, http://www.sefar.com) upon phytogel (Sigma-Aldrich, http://www.sigmaaldrich.com) solidified media (2 g L−1) containing half-strength MS salts, 4.5 mm 2-(N-morpholine)-ethanesulphonic acid (MES) and 1% sucrose, pH 5.7. Hormone induction media contained full-strength Gamborg's B5 with vitamins (PhytoTechnology Laboratories, http://www.phytotechlab.com), 20 g L−1 glucose, 0.5 g L−1 MES and 2 g L−1 Phytagel, pH 5.8, amended with 1000 × stocks of either 1-NAA or 2iP.
Seedlings were transferred on Nitex from half-strength MS plates at 3–4 days after germination to plates containing 10 μm NAA for 24 h to induce LRP, then transferred to 4.4 or 2.2 μm 2iP to promote LRP → SM conversion. For screening mutants and testing wus alleles, the numbers of shoots were scored after 5 and 7 days of 2iP treatment. Shoots were defined as three or more leaves initiated in a radial pattern around a presumed SM.
Images were acquired with a Leica upright DM 6000CS microscope connected to a TCS SP5 system (Leica, http://www.leica.com). Argon (50 mW) and GreenHeNe (1.2 mW) lasers were used for excitation. Maximal projections of z-stacks are shown.
Protoplasting and fluorescence-activated cell sorting (FACS)
Protoplasting and sorting were performed according to the methodology described by Birnbaum et al. (2005) using the enzyme mix described by Yadav et al. (2009). Approximately 250 seedling roots were harvested within 5 min, sliced and placed in protoplasting solution for 1 h. Between 1000 and 25 000 protoplasts were sorted into 350 μl of reverse transcriptase (RT) buffer (RNA Easy kit; Qiagen, http://www.qiagen.com) within 10 min, and flash frozen.
Microarray experiments and analysis
The WT, mutant analyses and cell-specific experiments were performed in triplicate, and RNA was extracted from 250 treated seedling roots pooled for each sample. Roots were removed with a scalpel blade across the Nitex mesh and flash frozen. Time points sampled included: 0, 19, 30 and 48 h 2ip treatment. RNA was extracted from seedling roots or sorted protoplasts using an RNA Easy kit (Qiagen). RNA extracted from sorted protoplasts underwent amplification using the GeneChip® IVT Express Kit (Affymetrix, http://www.affymetrix.com). For each sample, 5 μg of total RNA was reverse transcribed (SuperScript II; Invitrogen, http://www.invitrogen.com), labelled and hybridised to the Arabidopsis ATH1 Genome Array (Affymetrix). Data were pre-processed using mas5/gcos (Hubbell et al., 2002) implemented in r (R Development Core Team, 2011) and BioConductor (Gentleman et al., 2004), with a TGT value of 100. Expression data were filtered to remove probe sets reporting low transcript abundances (mean expression value <50, approximately 2.5-fold higher than the background). Differentially expressed genes were identified by raw P value and fold change, or by contrasts made using linear models for microarrays (limma) (Smyth, 2005) implemented in r/BioConductor (Gentleman et al., 2004). See Methods S1 for the quantitative real-time PCR (qRT-PCR) validation of arrays.
GO enrichment analysis
Singular enrichment analysis of DEGs was performed using agriGO (http://bioinfo.cau.edu.cn/agriGO; Zhou et al., 2010), using Fisher as the statistical test method, Hochberg (false-discovery rate) or Yekutieli (false-discovery rate under dependency) as the multi-test adjustment method, and minimum mapping entries were set at three or five, and ATH1 was used as the background.
We offer special thanks to Thanh Nguyen for her invaluable contribution, and extend our gratitude to Ryan Austin, Shu Hiu, Connor Chatfield and Henry Hong for technical assistance. We thank Siobhan Brady, Venu Reddy and Daphne Goring for expert advice and seed resources. We are grateful to Bruce Hall and Andrew Petrie for plant care. This research was funded by the National Science and Engineering Research Council, the Centre for the Analysis of Genome Evolution and Function at the University of Toronto, and Genome Canada.