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Plant Cell. 2005 Nov; 17(11): 3035–3050.
PMCID: PMC1276028

Cell Cycle Progression in the Pericycle Is Not Sufficient for SOLITARY ROOT/IAA14-Mediated Lateral Root Initiation in Arabidopsis thalianaW in Box


To study the mechanisms behind auxin-induced cell division, lateral root initiation was used as a model system. By means of microarray analysis, genome-wide transcriptional changes were monitored during the early steps of lateral root initiation. Inclusion of the dominant auxin signaling mutant solitary root1 (slr1) identified genes involved in lateral root initiation that act downstream of the auxin/indole-3-acetic acid (AUX/IAA) signaling pathway. Interestingly, key components of the cell cycle machinery were strongly defective in slr1, suggesting a direct link between AUX/IAA signaling and core cell cycle regulation. However, induction of the cell cycle in the mutant background by overexpression of the D-type cyclin (CYCD3;1) was able to trigger complete rounds of cell division in the pericycle that did not result in lateral root formation. Therefore, lateral root initiation can only take place when cell cycle activation is accompanied by cell fate respecification of pericycle cells. The microarray data also yielded evidence for the existence of both negative and positive feedback mechanisms that regulate auxin homeostasis and signal transduction in the pericycle, thereby fine-tuning the process of lateral root initiation.


Auxins have long been put forward as potent stimulators of cell division (Gautheret, 1939), but although considerable progress has been made in our understanding of both auxin signaling (Weijers and Jürgens, 2004) and cell cycle progression (Inzé, 2005), the molecular mechanisms by which these processes are connected remain poorly understood (Vanneste et al., 2005). The initiation of lateral roots, marked by specific cell divisions in the pericycle (Casimiro et al., 2001), is triggered by auxin (Torrey, 1950) and is, therefore, an ideal model system for studying how auxin signaling activates cell cycle progression.

The onset of lateral root formation coincides with the occurrence of a series of anticlinal, asymmetric divisions in the xylem pole pericycle (Malamy and Benfey, 1997); hence, cell cycle activation is inherently connected with lateral root initiation. Activation and progression through the major phases of the cell cycle (G1, S, G2, and M) are governed by the control of cyclin-dependent kinases (CDKs). The activity of these CDKs can be modulated through interacting regulatory components (cyclins [Roudier et al., 2000; Dewitte et al., 2003; Lee et al., 2003] and CDK subunits [De Veylder et al., 1997]), inhibitory components (inhibitors of CDKs and Kip-related proteins [Wang et al., 1997; De Veylder et al., 2001]), or through stimulatory (Shimotohno et al., 2004) and inhibitory phosphorylation (Sun et al., 1999). Beeckman et al. (2001) showed that in Arabidopsis thaliana, pericycle cells leaving the root apical meristem remain in G1 phase. Under normal conditions, only the pericycle cells at the xylem pole retain the capability to respond to an inductive signal, such as auxin, to initiate lateral roots. Interestingly, the expression of the cell cycle inhibitor KRP2 is strongly downregulated in xylem pole pericycle cells when roots are subjected to auxin treatment (Himanen et al., 2002; Casimiro et al., 2003). However, it can be anticipated that numerous regulatory proteins will be involved in lateral root initiation.

For global studies of developmental processes at the molecular level, relatively large quantities of material are required but are almost impossible to obtain for lateral root initiation because of the small subsets of pericycle cells involved (Laskowski et al., 1995; Kurup et al., 2005). Based on the observation that inhibition of polar auxin transport prevents lateral root initiation (Casimiro et al., 2001), a lateral root-inducible system (LRIS) was developed and characterized (Himanen et al., 2002, 2004). Germination on media with sufficiently high levels of 1-N-naphthylphthalamic acid (NPA) results in roots without lateral root initiation sites, which can be induced synchronously throughout the pericycle by subsequent transfer to media with high auxin concentrations (Himanen et al., 2002). This system allows the monitoring of the sequential signaling and cell cycle progression during lateral root initiation (Himanen et al., 2004).

Prior to inducing cell division, auxins have to be perceived and transmitted to turn on the cell cycle machinery. The current understanding of early auxin signal transduction is focused on the 26S proteasome-dependent proteolysis of specific small short-lived nuclear proteins, designated auxin/indole-3-acetic acid (AUX/IAA) (reviewed in Dharmasiri and Estelle, 2004). AUX/IAA proteins belong to a family of 29 members (Liscum and Reed, 2002) and act as negative regulators by repressing auxin response factors (ARFs) (Tiwari et al., 2004). When the auxin concentration increases, the interaction between the SCFTIR1 E3-ligase and AUX/IAA proteins is stimulated by direct binding of auxin to a family of auxin binding F-boxes (Dharmasiri et al., 2005a, 2005b; Kepinski and Leyser, 2005), resulting in oligo-ubiquitination of AUX/IAA proteins (Gray et al., 2001). Such ubiquitinated proteins are usually targeted for 26S proteasome-mediated proteolysis (Hershko and Ciechanover, 1998). Thus, high auxin concentration derepresses ARF activity, allowing the primary auxin response to take place. When AUX/IAA proteins are mutated in domain II, essential for the interaction with the SCFTIR1 complex (Ramos et al., 2001), their stability increases dramatically, resulting in auxin-resistant phenotypes (Ouellet et al., 2001; Zenser et al., 2001). Over the years, an increasing number of dominant and semidominant aux/iaa mutants have been described that exhibit point mutations in domain II (Liscum and Reed, 2002). When mutated in domain II, solitary root1 (slr1) mutants develop a primary root without any sign of lateral root initiation (Fukaki et al., 2002). Therefore, the SLR/IAA14 protein is probably a central regulator of lateral root initiation.

Here, lateral root initiation was used as an in planta model to study the interaction between the auxin signaling pathway and cell cycle activation. For this purpose, the lateral rootless phenotype of the early auxin signaling mutant slr1 was utilized to compare on a genome-wide level the transcriptional changes that occur in root segments of the wild type and slr1 during auxin-induced lateral root initiation. Complementation by overexpression of CYCD3;1 in the slr1 pericycle was able to trigger cell divisions, but no lateral root initiation, suggesting that more is involved than the activation of cell cycle progression in the pericycle. The transcriptional data suggest that during lateral root initiation, counteracting feedback regulation acts on auxin homeostasis and the signal transduction machinery. Furthermore, the analysis of the transcriptional changes in the wild type versus the slr1 mutant allowed some cell cycle regulators to be pinpointed as potential targets of the primary auxin signaling pathway.


slr1 as a Tool to Link Auxin Signaling to Lateral Root Initiation

The dominant mutation in SLR/IAA14 results in complete absence of detectable lateral root initiation sites, formation of aberrant root hairs, and agravitropic root growth (Fukaki et al., 2002). For a better notion of the tissues and cell types that become the primary targets of the reduced auxin response in the mutant, we analyzed the expression pattern of SLR/IAA14 at the cellular level by anatomical sections of 3-d-old roots of transgenic plants harboring a promoter IAA14/β-glucuronidase (PIAA14:GUS) construct. Expression of mutant slr/iaa14 under control of this promoter sequence has been shown to be sufficient to phenocopy the root phenotype of slr1 (Fukaki et al., 2002). Furthermore, tissue-specific expression of stabilized SLR/IAA14 in the xylem pole pericycle conferred the lateral rootless phenotype of slr1 (Fukaki et al., 2005). The expression of PIAA14:GUS was strongest in the xylem pericycle cells, in the xylem-associated cells of mature root tissues, and almost absent from phloem pole pericycle cells (Figures 1A and 1B). This stronger expression at the xylem pole, the site of lateral root initiation in Arabidopsis, suggests a causal link between the reduced auxin responsiveness and the lack of lateral roots in the mutant. During lateral root development, GUS activity was strongest in the youngest lateral root primordia, whereas it was nearly absent in mature lateral root meristems (see Supplemental Figure 1 online). In epidermal cells of mature root tissues, the expression was also low (Figure 1A). In the elongation zone, expression was strongest in epidermal cells (Figure 1C), whereas more distally, it was restricted to the lateral root cap (Figure 1D). Plants harboring the PIAA14:GUS construct were crossed into the slr1 background, resulting in F1 plants exhibiting a reduced level of expression compared with that of the wild type (Figures 1E and 1F). Anatomical sections showed that the tissue specificity of PIAA14:GUS expression was maintained in slr1 (data not shown). When the primary root length was compared between 2-week-old wild-type and mutant seedlings, no significant differences were observed (data not shown), suggesting that the meristematic activity of the slr1 primary roots was not affected. To verify whether slr1 mutants still normally specified the pericycle, the enhancer trap line, J0121, a marker for xylem pericycle cell identity (Casimiro et al., 2001), was crossed into the slr1 mutant. The green fluorescent protein expression pattern of J0121 did not change significantly when compared with the wild-type situation (Figures 1G and 1H). Because CDKA;1 is a central regulator of cell cycle progression and its expression is associated with competence to divide (Hemerly et al., 1993), the PCDKA;1:GUS fusion was analyzed. Its expression was equally strong in the pericycle of the wild type and slr1 (Figures 1I and 1J). Also, expression of PALF4:GUS, fused with ALF4, a protein with unknown function that is required for maintaining pericycle cells competent to form lateral roots (DiDonato et al., 2004), was unaltered in slr1 (data not shown). These observations indicate that the changes in auxin response and cell cycle activation in the mutant are probably not due to an altered pericycle identity or a reduced competence to divide.

Figure 1.
Expression Analysis in the Wild Type and slr1.

To compare the transcriptional changes associated with auxin induction in the wild-type and the slr1 situations, we used the recently developed LRIS that allows the synchronization of lateral root initiation in roots devoid of lateral roots (Himanen et al., 2002). The possibility that this LRIS might induce lateral root initiation in the mutant was first addressed before initiating large-scale experiments. Therefore, specific GUS reporter lines were crossed into the mutant background, and their response to a 12-h auxin treatment following a 72-h growth on NPA was characterized. In 72-h NPA-treated seedlings, no expression of the auxin-responsive PDR5:GUS construct could be detected in either genotype (data not shown). When seedlings had been transferred to α-naphthaleneacetic acid (NAA) for 12 h, PDR5:GUS activity was strongly induced throughout the root in the wild type and to a lesser extent in slr1 (Figures 1K and 1L), suggesting that the slr1 mutation also results in reduced auxin responsiveness within the LRIS. To assay cell cycle progression within the LRIS, plants harboring GUS reporter constructs for the G2-to-M transition-associated genes CYCB1;1 and CDKB1;1 were used. The expression of CYCB1;1 is strong in pericycle cells undergoing the first divisions of lateral root initiation (Beeckman et al., 2001; Himanen et al., 2002), whereas that of CDKB1;1 starts from the S phase into the G2-to-M transition (Porceddu et al., 2001; Menges and Murray, 2002). With these two GUS reporter lines, no expression was detected in the pericycle in either genotype when germinated on NPA (data not shown). When transferred to auxin for 12 h, wild-type pericycle cells had very strong GUS activity for both markers, hinting that these pericycle cells divide actively (Figures 1M and 1O). The destruction box fused to GUS in PCYCB1;1:GUS results in proteolysis during the late mitosis. Therefore, because of its patchy GUS pattern, the GUS-stained pericycle cells might have undergone at least one cycle of cell division during the 12-h NAA treatment. On the other hand, no GUS activity could be induced in slr1 pericycle cells (Figures 1N and 1P), reflecting the complete absence of mitotic activity, following 12 h of auxin treatment. These data legitimize the use of the slr1 mutant in a genome-wide transcriptional analysis of lateral root initiation.

Microarray Setup and Statistical Analysis

Within the LRIS, pericycle cells have been shown to be blocked in the G1 phase when germinated on NPA (Himanen et al., 2002, 2004). Subsequent transfer to auxin media was sufficient to trigger the primary auxin response within 2 h, resulting in a synchronous induction of lateral root initiation over the entire pericycle (visualized by PCYCB1;1:GUS expression in Figure 2). To gain insight into the early events of lateral root initiation, samples for microarray analysis were taken at the time points 0 h NAA (= 72 h NPA), 2 h NAA, and 6 h NAA for both the wild type and slr1 (highlighted stages in Figure 2). Per time point, ~1000 root segments were sampled and two independent biological replicates were performed. To minimize contamination with nonrelevant tissues and dividing cells, root segments were cut above the root apical meristem and below the root-hypocotyl junction. These segments were further subjected to the required steps for microarray analysis with the ATH1 Affymetrix chips harboring 22,746 probe sets (see Methods). Analysis of variance on the normalized gene expression data assessed the significance of three major sources of variability affecting the expression level: the duration of the auxin treatment (time), the genotype, and the interaction between these two. Correction for multiple comparisons was performed by controlling the false discovery rate, and q-values were calculated according to Storey and Tibshirani (2003). At a stringency level of P < 0.001, 3110 genes had a significantly modulated expression profile during the experiment. Furthermore, none of them could be rejected based on the calculated q-values (q < 0.05) (see Supplemental Table 1 online).

Figure 2.
Schematic Representation of the LRIS.

Cross-Table Clustering to Identify Lateral Root Initiation Genes

The generally used approaches of hierarchical (Eisen et al., 1998) and K means (Tavazoie et al., 1999; Soukas et al., 2000) clustering allow the classification of gene expression patterns of single-series microarray time courses. However, for the comparison of multiple-series microarray time courses, no standard algorithms are established. Most reports on such issues restrict themselves to fold-change comparisons (Tian et al., 2002; Ullah et al., 2003) or basic diagrams (Puthoff et al., 2003; Taji et al., 2004). Less frequently, clustering is used to identify differences between expression profiles in different genotypes (De Paepe et al., 2004). Here, a new approach was developed based on the commonly used single-series time course K means clustering algorithm (Figure 3A). By combining the data irrespective of the genetic background (0, 2, and 6 h) with the K means clustering algorithm, the gene expression profiles could be classified into 14 clusters (Figure 3B). Subsequently, the clusters were plotted at the headings of rows and columns of a cross-table. In this cross-table format, the expression profile of each gene is summarized by a uniquely identifiable cluster combination corresponding to the expression pattern in the wild type and in slr1. Per cluster combination, the gene frequency was calculated and included into the cross-table; a color code was implemented to assess easily the differential dynamics of expression between both genotypes (Figure 3C; see Supplemental Table 1 online). Hereafter, we will refer to this methodology as cross-table clustering. This approach allows a high-resolution representation of gene expression patterns across multiple genotypes and provides a tool to compare the transcriptional changes in the wild type versus mutant within the framework of a unified set of expression profiles.

Figure 3.
Cluster Analysis of the Expression Data.

In order to gain further insight into the data set, the functional relevance of all types of cluster combinations was evaluated. The wild-type expression profiles were subdivided into three major patterns: upregulated (Figure 3B, clusters 1 to 6), constitutive (Figure 3B, clusters 7 to 11), and downregulated (Figure 3B, clusters 12 to 14). These expression patterns were compared with the corresponding expression profiles in slr1. The genes represented by cluster combinations on the diagonal of the cross-table (gray) exhibit expression profiles that are similar in both genotypes, whereas above (red and orange) and below (green and blue) the diagonal, the induction rates in the wild type are higher and lower than those in the mutant, respectively. The Gene Ontology classification for biological function (Berardini et al., 2004) was applied on all annotated genes of each subgroup and subsequently compared with all annotated significant genes within our experiment with the EASE 2.1 software (Hosack et al., 2003). The 913 genes that were upregulated dependent on wild-type SLR/IAA14 stability (selection in Figure 3C; see Supplemental Table 2 online) were significantly enriched (PBonferoni < 0.01) in proliferation-related biological functions, such as cell cycle, nucleic acid metabolism, and protein metabolism (Figure 4; see Supplemental Table 3A online), but restricting this selection to the earliest induced genes (Figure 3B, cluster 1 to 3) resulted in a significant enrichment (PBonferoni < 0.01) in the biological functions response to endogenous stimulus and response to auxin stimulus (see Supplemental Table 3B online). These observations strongly correlate with the previous observations of Himanen et al. (2004), who found that within the LRIS, genes related to signaling were upregulated early prior to cell cycle activation.

Figure 4.
Overrepresented Functional Categories within the SLR/IAA14-Mediated Upregulated Genes.

In conclusion, the cross-table clustering methodology on our data set reliably identified 913 genes that depend on rapid SLR/IAA14 degradation for normal auxin responsiveness. Because slr1 does not initiate lateral roots, even upon auxin treatment, such genes might encode potential regulatory proteins required for lateral root initiation and will be designated as lateral root initiation (LRI) genes hereafter.

Validation of LRI Genes by Database Comparison

With the same LRIS, we had performed previously a microarray study based on cDNA microarrays representing ~4600 genes (Himanen et al., 2004). At a stringency level of 0.005, 906 genes had been found to be significantly expressed. The profiles of 343 genes were clustered into three upregulated clusters. From the selected 913 LRI genes, defined as auxin inducible in an SLR/IAA14-dependent manner, 122 genes were also represented in the earlier data set (see Supplemental Table 4 online). This result suggests that, despite the differences in type of array (cDNA array versus Affymetrix gene chip) and the differences in statistical analysis, a high level of reproducibility was obtained by the LRIS and implies also that a high level of confidence can be attributed to the profiles associated with the LRI genes, as suggested by the q-values.

The arf7 arf19 double mutants have been shown to phenocopy the lateral rootless phenotype of slr1 to a large extent (Okushima et al., 2005; Wilmoth et al., 2005), indicating that SLR/IAA14 inhibits the activity of these ARFs to block lateral root initiation. Recently, a yeast two-hybrid assay confirmed that SLR/IAA14 interacts with ARF7 and ARF19 (Fukaki et al., 2005). Therefore, both slr1 and arf7 arf19 mutants can be expected to have similar target genes. The auxin inducibility in arf7 arf19 has been assessed by Okushima et al. (2005), who assayed the effects of a 2-h treatment with 5 μM IAA on 5-d-old seedlings of wild type, arf7, arf19, and arf7 arf19 in a microarray that gave a robust overview of early auxin-induced genes. The respective ARF7- and ARF19-dependent auxin inducibility of the 913 LRI genes was extracted from the published data set (see Supplemental Table 5 online). Of the 913 LRI genes, 99 were induced more than twofold in the wild type, whereas the auxin inducibility of these genes was abolished in the arf7 arf19 double mutants (see Supplemental Table 6 online). Because in this experimental setup only a 2-h IAA treatment was used, a large percentage of these genes belong to the early induced LRI genes (83 out of 365 genes in wild-type clusters 1 to 3). Due to large experimental differences, we cannot exclude that more LRI genes act downstream of ARF7 and ARF19. These results indicate that the use of the LRIS provides a high reproducibility among divergent experiments, giving a high-resolution image on differential expression during lateral root initiation.

Cell Cycle Progression during Lateral Root Initiation

Within these 913 LRI genes, cell division–related genes were identified, such as APC8/CDC23, PCNA1, RNR1, MCM3, MCM4, MCM7/PROLIFERA, RPS18A/PFL1, RPS13A/PFL2, FtsZ1-1, FtsZ2-1, CYCD3;2, CYCA2;4, CYCB2;5, CDKB2;1, and CKL3 (Table 1). APC8/CDC23 is part of the anaphase-promoting complex (APC) that is involved in targeting A- and B-type cyclins for degradation during mitosis (Capron et al., 2003). Proliferating cell nuclear antigen (PCNA1) belongs to the DNA replication machinery, and its expression is associated with the S phase (Hübscher et al., 2002). Minichromosome maintenance (MCM) proteins are conserved eukaryotic replication factors involved in the initiation of DNA replication (Stevens et al., 2002), and ribonucleotide reductase large subunit (RNR1) is part of a rate-limiting enzyme in the synthesis of nucleotides (Elledge et al., 1993). The ribosomal proteins encoded by POINTED FIRST LEAVES1 (PFL1) and PFL2 are produced during lateral root formation (Van Lijsebettens et al., 1994; Ito et al., 2000). FtsZ is involved in plastid divisions (Osteryoung et al., 1998). On the other hand, we recovered very few core cell cycle genes as defined by Vandepoele et al. (2002). The represented genes consisted of one G1-to-S (CYCD3;2), an S phase–related (CYCA2;4) and two G2-to-M–related genes (CYCB2;5 and CDKB2;1), and a recently identified CDK-like protein-encoding gene (CKL3) (Menges et al., 2005). However, in the root part concerned, most core cell cycle genes are expressed in low abundance at this high stringency level (P < 0.001). When the stringency level was reduced to P < 0.01, the number of significantly modulating cell cycle genes increased to 28 (q < 0.05) (see Supplemental Table 7 online).

Table 1.
Known Auxin- and Cell Cycle–Related Genes within the LRI Genes

Three cell cycle genes of the stringent selection, CYCD3;2, CYCA2;4, and CDKB2;1, contain at least one auxin-responsive element (ARE; TGTGTC or GAGACA) within a sequence 1000 bp upstream of their 5′ untranslated region (Figure 5A), suggesting that these genes may be part of the primary auxin response with their expression controlled by unstable AUX/IAA proteins (Ulmasov et al., 1997). Consistently, treating 5-d-old roots with the protein synthesis inhibitor cycloheximide (CHX) for 2 and 4 h resulted in a progressively strong induction of CYCA2;4 and CDKB2;1, respectively (Figure 5B). However, for CYCD3;2, only a weak (nonsignificant) CHX-mediated induction was observed, suggesting that the AREs in its promoter may not be functional.

Figure 5.
Analysis of Primary Auxin Responsiveness of CYCD3;2, CYCA2;4, and CDKB2;1.

Consistent with the observation that their induction is strongly dependent on normal SLR/IAA14 degradation, CYCA2;4 and CDKB2;1 are probably directly regulated through a labile repressor, such as SLR/IAA14. Taken together, these results argue for a direct link between auxin signaling and cell cycle activation during lateral root initiation both at the S phase and at the G2-to-M transition.

Cell Cycle Progression in the Pericycle Is Not Sufficient to Complement the Lack of Lateral Roots in slr1

Overexpression of G1/S regulators, such as CYCD3;1 (Dewitte et al., 2003) or the transcription factor complex E2Fa/DPa (De Veylder et al., 2002), is sufficient to trigger several rounds of cell division in cell types that normally remain quiescent. Can overexpression of these cell cycle regulators induce pericycle cell proliferation and possibly even lateral roots in the absence of a normal SLR/IAA14-dependent auxin response? To address this question, slr1 was crossed into the transgenic lines and their respective wild types. The root phenotypes of the F1 seedlings were analyzed in detail. All the wild-type (Columbia [Col-0] and Landsberg erecta [Ler]) and CYCD3;1OE (in Ler background) plants formed normal roots (Figures 6A, 6E, and 6F). However, E2Fa/DPaOE (in Col-0 background) showed a clear reduction in primary root length and in visible lateral roots (Figure 6B). The reduced lateral root density of E2Fa/DPaOE was also observed in cleared 10-d-old roots and can be rescued by auxin treatment (I. De Smet, unpublished results). When slr1 was crossed into Col-0, E2Fa/DPaOE, Ler, or CYCD3;1OE, no lateral roots could be observed (Figures 6C, 6D, 6G, and 6H). Surprisingly, after closer microscopic inspection of the root, only in CYCD3;1OE × slr1 were regions seen in which the pericycle cells were shorter (Figure 6L) than those of mature Ler and Ler × slr1 (Figures 6I and 6K). These series of short cells were found exclusively in the pericycle cell layer and are most probably the result of one or more extra rounds of cell division. In E2Fa/DPaOE × slr1 and Col-0 × slr1 roots, no such regions with shortened pericycle cells could be observed (data not shown). Unlike in a stage I primordium, which is normally composed of small radially swollen cells in the midst of larger elongated cells (Figure 6J; Malamy and Benfey, 1997), the short pericycle cells in CYCD3;1OE × slr1 roots had a uniform cell size and were not radially swollen (Figure 6L). Also absent were stage II lateral root primordia, which are the result of a periclinal division of stage I primordium cells. To verify whether these regions of shortened cells were the consequence of extra rounds of cell division after leaving the meristem, the expression levels of the S phase–associated CYCA2;4 gene (Figure 6M) and G2/M-specific CYCB1;1 (Figure 6N) and CYCB2;5 (Figure 6O) were analyzed via real-time RT-PCR in the CYCD3;1OE × slr1 background. In slr1 background, the expression of these cell cycle genes was strongly reduced, whereas that of both genes appeared to be restored by CYCD3;1 overexpression. To assess whether this cell division activity in the pericycle of the mutant was associated with a recovered capacity to initiate a new organ, the expression of the PLETHORA1 (PLT1) gene was analyzed in CYCD3;1OE × slr1 roots. PLT1 encodes a transcription factor that has been shown to be associated with quiescent center specification downstream of the auxin signal (Aida et al., 2004). Because a new quiescent center has to be specified during lateral root formation, PLT1 expression can be used as a marker for early stages of lateral root organogenesis (see Supplemental Figure 2 online). Nevertheless, PLT1 expression remained low in CYCD3;1OE × slr1 roots (Figure 6P). These results indicate that the requirements for organogenetic processes, such as lateral root initiation, are more complex than simple activation of the cell cycle and that additional SLR/IAA14-dependent signaling is needed to develop a new organ.

Figure 6.
Complementation of Cell Cycle Defect in slr1.

Complex Auxin Signaling-Dependent Mechanisms Regulate Lateral Root Initiation

As expected, besides cell cycle–related genes, several central regulators of auxin signaling were recovered among the LRI genes (Table 1). Nine AUX/IAA genes were identified within this selection. For two of them, gain-of-function mutants with defects in lateral root formation have been described before (axr5/iaa1 [Park et al., 2002; Yang et al., 2004] and msg2-1/iaa19 [Tatematsu et al., 2004]). The remaining seven AUX/IAA genes have not been characterized functionally. In addition to this type of negative feedback regulation, several potential AUX/IAA targets, such as ARF4, ARF16, and ARF19, were also strongly upregulated in a SLR/IAA14-dependent fashion, especially ARF19, which seems to play an important role in regulating lateral root formation because arf7 arf19 double mutants exhibit an slr1-like phenotype (Okushima et al., 2005; Wilmoth et al., 2005). Furthermore, both ARF7 and ARF19 were shown to interact with SLR/IAA14 in a yeast two-hybrid assay (Fukaki et al., 2005). In conclusion, our microarray data show SOLITARY ROOT/IAA14-dependent expression of both AUX/IAA and ARF genes, suggesting that at the level of auxin signaling, lateral root initiation is tightly regulated through both negative and positive feedback.

Attenuating and Maintaining Auxin Levels during Lateral Root Initiation

Auxin conjugation, transport, and biosynthesis contribute to local auxin homeostasis. The data set presented (Table 1) indicates that these three processes are a part of lateral root initiation.

The GH3 gene family consists of the first identified auxin-responsive genes (Hagen et al., 1991) and has been divided into three major groups, according to their substrate specificity and sequence similarities (Staswick et al., 2002). Six group II GH3 proteins have been shown to act as IAA-amido synthetases, providing a mechanism to reduce the level of active auxin by conjugation to amino acids (Staswick et al., 2005). Interestingly, all five GH3 genes identified as LRI genes belong to the group II GH3 family, and besides ARF8 (Tian et al., 2004), they also seem to be regulated through ARF7 and ARF19 transcription factors (Okushima et al., 2005). Also, UGT84B1, coding for an enzyme involved in the conjugation of glucose to IAA (Jackson et al., 2002), was induced in the wild type, but not in slr1. By contrast, IAR1, which codes for a putative ZIP family transporter protein and is necessary for the production of IAA out of IAA–amino acid conjugates (Lasswell et al., 2000), was strongly repressed in the wild type and to a lesser extent in slr1. A negative feedback mechanism interfering with the level of free auxin is therefore likely to be active during lateral root initiation.

On the other hand, many components of the auxin transport machinery were recovered within the LRI genes. Not only did the putative auxin influx carriers AUX1 and Like-AUX1 3 (LAX3) (Parry et al., 2001) emerge, but so did the auxin efflux facilitators, such as PIN1, PIN3, and PIN7 (Paponov et al., 2005), and some proteins involved in their correct localization, such as PINOID/PID (Friml et al., 2004), PINOID-like, and a PID-interacting protein, TCH3 (Benjamins et al., 2003). Moreover, the multidrug resistance protein involved in polar auxin transport encoded by PGP1 (Lin and Wang, 2005) was found among the LRI genes. The corresponding mutants display defects in lateral root initiation (Bennett et al., 1996; Benková et al., 2003). The auxin inducibility of polar auxin transport genes (influx and efflux) was also observed within the vascular cambium of hybrid aspen (Schrader et al., 2003). Interestingly, PIN1, PIN3, and PIN7 were also shown to depend on ARF7 and ARF19 for their auxin inducibility (Okushima et al., 2005), and their AUX/IAA-dependent inducibility has been implicated in their functional redundancy (Vieten et al., 2005).

Recently, auxin has been shown to negatively regulate its own biosynthesis (Ljung et al., 2005). Two genes encoding enzymes involved in the Trp-dependent IAA biosynthesis pathway (CYP79B2 and CYP79B3) (Zhao et al., 2002) and their transcriptional regulator ATR1/MYB34 (Celenza et al., 2005) were downregulated upon auxin treatment in the wild type, but to a lesser extent in slr1. Interestingly, YUCCA, a gene encoding another rate-limiting enzyme of the Trp-dependent IAA biosynthesis pathway (Zhao et al., 2001), was more induced in slr1 than in the wild type.

Taken together, these data suggest that auxin homeostasis is disturbed in slr1. Therefore, the auxin content of seedling roots of the wild type and slr1 was determined by gas chromatography–selected reaction monitoring–mass spectrometry (Ljung et al., 2005). Indeed, a significantly higher auxin concentration was found in the most apical 3 mm of slr1 primary roots than those of the wild type (P < 0.005; Figure 7A). Correspondingly, the increased auxin concentration in slr1 could be visualized with PDR5:GUS (Figures 7B to 7E).

Figure 7.
Analysis of Auxin Content in Col-0 and slr1.


The results of our genome-wide transcriptome analysis allow us to present a model describing a regulatory network that integrates some of the most likely occurring processes upon auxin perception during the onset of lateral root initiation (Figure 8). The auxin signaling cascade starts off by the auxin-stimulated targeting of AUX/IAA proteins for degradation, thereby derepressing the transcriptional activity of ARFs (reviewed in Dharmasiri and Estelle, 2004). Therefore, the auxin-mediated degradation of the SLR/IAA14 protein might be the first crucial event in the preamble toward lateral root initiation (Figure 8A).

Figure 8.
Model of SLR/IAA14-Dependent Lateral Root Initiation.

The Importance of Cell Cycle Progression during Lateral Root Initiation

When the auxin-induced transcriptional changes in root segments of the wild type and slr1 are compared, the inhibition of auxin signal transduction in the mutant is correlated with the failure to induce cell cycle–related genes. Although the role of auxin in cell cycle progression is generally accepted, we found evidence that this stimulation may occur via an AUX/IAA-ARF–dependent pathway. Indeed, CYCA2;4 and CDKB2;1 both contain at least one ARE in their promoter sequence, and their auxin inducibility has been shown to depend on the normal degradation of SLR/IAA14. Furthermore, both genes are induced by the inhibition of protein synthesis, suggesting that a labile repressor is controlling their transcription. Because AUX/IAA proteins are regarded as labile repressors (Ouellet et al., 2001), it is tempting to consider CYCA2;4 and CDKB2;1 to be primary targets of the SLR/IAA14 signaling. These results are indicative for a potential molecular mechanism by which auxin signaling may feed into the cell cycle. Nevertheless, the functionality of the respective AREs remains to be proven.

However, stimulation of the cell cycle was not sufficient to trigger the formation of lateral roots in the slr1 background. Only CYCD3;1OE was capable of initiating a round of cell divisions in slr1 pericycle cells, giving rise to a single layered proliferation zone (Figure 8A) that did not progress further into lateral root developmental stages, let alone lateral root primordia.

Cell Fate Respecification of the Pericycle

The inability to form lateral roots after cell cycle stimulation of the pericycle suggests that besides cell cycle progression, slr1 is unable to respecify the identity of pericycle cells into that of lateral root primordia. Analogous to the situation found in early embryogenesis, the formation of lateral roots also seems to depend on cell fate alteration through the occurrence of asymmetric divisions. In case of embryogenesis and primary root development, polar auxin transport installs an auxin maximum, visualized by the auxin reporter construct PDR5:GUS (Sabatini et al., 1999), restricting PLT expression that specifies stem cell identity (Aida et al., 2004; Blilou et al., 2005). Likewise, polar auxin transport is required to form lateral roots because mutants defective in polar auxin transport fail to produce lateral roots (Benková et al., 2003; Geldner et al., 2004). Transcripts of the polar auxin transport machinery appear to be unable to accumulate normally in auxin-treated slr1 roots, preventing the installation of an auxin maximum. Therefore, it might be argued that fine-tuned polar auxin transport could be the missing factor to specify stem cell identity for lateral root initiation in slr1. Other mutants defective in polar auxin transport were unable to produce individual lateral root primordia upon auxin supplementation; instead, multilayered proliferation zones (Figure 8A) developed (Benková et al., 2003; Geldner et al., 2004). Nevertheless, none were detected in the slr1 root that has increased auxin levels, even when cell division was stimulated by CYCD3;1OE. However, PLT expression was not restored by CYCD3;1OE in slr1, suggesting that the induced pericycle cell divisions do not correspond to lateral initiation sites. Thus, the impaired auxin transport in slr1 would probably not be the only element responsible for its inability to form lateral roots. Polar auxin transport is necessary for providing auxin to the pericycle and for organizing the auxin gradient in the developing lateral root primordium; induction of other factors are probably also needed to turn into organogenesis.

Such factors, which are indispensable for induction of asymmetric divisions, acquisition of different cell fates, and organogenesis, might be represented by members of the WUSCHEL-related homeobox (WOX) family. During embryogenesis, WOX genes are expressed in restricted areas of the embryo and are believed to be involved in cell fate specification (Haecker et al., 2004). In this respect, it is worth mentioning that one member of the WOX family, WOX13, was retrieved among the LRI genes. The SLR/IAA14-mediated auxin inducibility (cluster combination 6.8) of WOX13 suggests that it is involved in the respecification of pericycle cells during lateral root initiation. Moreover, the functional analysis of other proteins encoded by the LRI genes will probably result in the identification of potential cell fate respecification factors and will provide new insights into this intriguing developmental process.

A Binary Switch Mechanism Is Involved in Lateral Root Initiation

Our results suggest that lateral root initiation is regulated through positive as well as negative feedback loops (Figure 8B, green and red arrows). Two types of regulation with negative effects on auxin concentration and/or signal transduction can be distinguished from the data set (Figure 8B, red arrows). One type is brought about by the strong upregulation of genes encoding enzymes involved in auxin conjugation. Conjugation of IAA to amino acids or sugars can reduce the free auxin level, although some conjugates of IAA and amino acids are reversible and can contribute to the pool of free IAA (Rampey et al., 2004). However, the contribution of IAA oxidation is believed to be much more important for auxin homeostasis than that of conjugation (Kowalczyk and Sandberg, 2001). Secondly, in contrast with the auxin-induced instability of AUX/IAA proteins (Gray et al., 2001), their expression was found to be strongly induced within the LRIS. Because these proteins encode potent inhibitors of ARF-mediated transcription (Tiwari et al., 2004), their induction provides a direct negative feedback onto the auxin signal transduction.

An example of a positive feedback loop (Figure 8B, green arrows) is the active auxin transport that has been shown to be auxin inducible in a SLR/IAA14-dependent manner. The installation of its own transport machinery matches the self-organizing nature of auxin transport suggested in the canalization hypothesis proposed by Sachs (1988). This supposition implies that high auxin levels in the pericycle would be actively reinforced by the locally increased auxin transport potential. Furthermore, the downregulation of IAA biosynthesis genes and the upregulation of the IAA conjugation potential in early stages of lateral root initiation suggest that young primordia depend on auxin import for their development. Later on, in emerged lateral root primordia, a regained capacity for auxin biosynthesis has been observed (Ljung et al., 2005). Indeed, only excised lateral root primordia at later stages of development can grow in culture without added hormones (Laskowski et al., 1995). Furthermore, expression of several ARFs is also induced during lateral root initiation, reducing the AUX/IAA versus ARF ratio with an increased sensitivity toward auxin as a consequence and amplification of the auxin signal to stimulate the onset of lateral root initiation.

In summary, in the proposed model, lateral root initiation is subjected to two counteracting regulatory circuits that result in a binary switch mechanism. When the auxin concentration is low, the auxin response acts to dampen small fluctuations in auxin concentration and no lateral roots are initiated. When this negative regulation is overcome, auxin concentration and responsiveness are actively reinforced by a positive feedback mechanism. This positive spiral activates the developmental program of lateral root initiation. When SLR/IAA14 is mutated, no lateral roots can be produced. Also, auxin homeostasis is misregulated in slr1, leading to auxin accumulation. Thus, our model is consistent with the experimental data and provides a basis to understand the complex self-regulatory mechanisms involved in lateral root initiation.


Plant Material and Growth Conditions

In this study, we analyzed the Arabidopsis thaliana Heynh. ecotypes Col-0 and Ler, the mutant slr1 (Fukaki et al., 2002), the promoter fusions PIAA14:GUS (Fukaki et al., 2002), PCDKA;1(cdc2a):GUS (Hemerly et al., 1993), PCDKB1;1:GUS (de Almeida Engler et al., 1999), PCYCB1;1:GUS (Colón-Carmona et al., 1999), PDR5:GUS (Ulmasov et al., 1997), PALF4:GUS (DiDonato et al., 2004), E2Fa/DPaOE (De Veylder et al., 2002), CYCD3;1OE (Dewitte et al., 2003), and the xylem pole pericycle-specific GAL4 enhancer trap line J0121 (http://www.plantsci.cam.ac.uk/Haseloff/geneControl/catalogues/Jlines/record/record_0.html). Seeds were always germinated on medium derived from standard Murashige and Skoog medium on vertically oriented square plates (Greiner Labortechnik) in a growth chamber under continuous light (110 μE·m−2·s−1 photosynthetically active radiation, supplied by cool-white fluorescent tungsten tubes; Osram) at 22°C (Himanen et al., 2002). Supplements consisted of 10 μM NPA (Duchefa), 10 μM NAA (Sigma-Aldrich), or 30 μM CHX (Sigma-Aldrich).

Histochemical and Histological Analysis

The GUS assays were performed as described by Beeckman and Engler (1994). For microscopic analysis, samples were cleared by mounting in 90% lactic acid (Acros Organics) (analysis of GUS stainings) or using the clearing method described by Malamy and Benfey (1997) (analysis of pericycle cell length). All samples were analyzed by differential interference contrast microscopy (Leica DMLB).

For anatomical sections, GUS-stained samples were fixed overnight in 1% glutaraldehyde and 4% paraformaldehyde in 50 mM phosphate buffer, pH 7. Samples were dehydrated and embedded in Technovit 7100 resin (Heraeus Kulzer) according to the manufacturer's protocol. For proper orientation of the samples, we used a two-step embedding methodology, based on a pre-embedding step to facilitate orientation in 0.5-mL Eppendorf tubes (De Smet et al., 2004). Sections of 5 μm were cut with a microtome (Minot 1212; Leitz), dried on Vectabond-coated object glasses, counterstained for cell walls with 0.05% ruthenium red for 8 min (Fluka Chemica), and rinsed in tap water for 30 s. After drying, the sections were mounted in DePex medium (British Drug House) and covered with cover slips.

Photographs were taken with a CAMEDIA C-3040 zoom digital camera (Olympus) and processed with Photoshop 7.0 (Adobe Systems).

Microarray Analysis and Data Processing

Col-0 and slr1 seeds were germinated on medium containing 10 μM NPA and transferred 3 d after germination under continuous light to 10 μM NAA, according to the points of the time course (0, 2, and 6 h). All sampling points were performed in duplicate. For each sampling, ~1000 root segments between root apical meristem and root-hypocotyl junction were pooled. From the pooled LRIS, RNA of root segments was extracted with the RNeasy kit (Qiagen). Out of 5.8 μg of total RNA, biotinylated copy RNA was produced (Hennig et al., 2003), of which 20 μg was fragmented and hybridized to the ATH1 arrays (Affymetrix). Washing, detection, and scanning were performed as described by Hennig et al. (2003). Raw data were processed with the statistical algorithm of the Affymetrix Microarray Suite 5.0 (Liu et al., 2002). Box plots of log2-transformed normalized value distributions of all arrays show that most array-to-array effects were taken care of by the normalization procedure (see Supplemental Figure 3 online). The normalized data were subjected to two-factor analysis of variance with Microsoft Excel. The false positives were controlled by measuring the false discovery rate (q-value) (Storey and Tibshirani, 2003) with the freely available software QVALUE (http://genomine.org/qvalue/). Significant profiles were preprocessed (Figure 3A) prior to clustering. The optimal number of clusters was estimated with the figure of merit calculations (Yeung et al., 2001) and K means clustered (Soukas et al., 2000) in the Multiple Experiment Viewer 2.2 of The Institute for Genome Research (Saeed et al., 2003). The cross-table was constructed with Photoshop 7.0.

Real-Time PCR

RNA was extracted with the RNeasy kit. Poly(dT) cDNA was prepared from 1 μg of total RNA with Superscript III reverse transcriptase (Invitrogen) and quantified on an iCycler apparatus (Bio-Rad) with the qPCR core kit for SYBR green I (Eurogentec). PCR was performed in 96-well optical reaction plates heated for 10 min to 95°C to activate hot start Taq DNA polymerase, followed by 50 cycles of denaturation for 60 s at 95°C and annealing extension for 60 s at 58°C. Target quantifications were performed with specific primer pairs designed with the Beacon Designer 4.0 (Premier Biosoft International). All PCRs were performed in triplicate. Expression levels were first normalized to ACTIN2 expression levels that did not show clear systematic changes in Ct value and then to the respective expression levels in the wild type (Ler). The primers used to quantify gene expression levels were At1g42970/GAPDH, 5′-TCTTTCCCTGCTCAATGCTCCTC-3′ and 5′-TTTCGCCACTGTCTCTCCTCTAAC-3′; At3g18780/ACTIN2, 5′-TTGACTACGAGCAGGAGATGG-3′ and 5′-ACAAACGAGGGCTGGAACAAG-3′; At1g80370/CYCA2;4, 5′-GCTCCAGATCGCCTCCAAG-3′ and 5′-CACGCAGGTTGTAGTAGATG-3′; At1g20590/CYCB2;5, 5′-GAGTTTCACACAGGCTAC-3′ and 5′-AGATGTGTTGTACTTCCTG-3′; and At3g20840/PLT1, 5′-ACGATATGCCTTCCAGTGATG-3′ and 5′-TTCAGACCCATTCCTTGTGC-3′.

Quantification of IAA

Wild-type and mutant slr1 seedlings were grown on vertical plates (1× Murashige and Skoog medium, 1% sucrose, and 1% agar, pH 5.7) under long-day conditions (16 h light, 8 h darkness). Samples were collected, extracted, and purified as described by Ljung et al. (2005). Of the primary root, the apical 3 mm was collected from 7-d-old seedlings and cut in 1-mm sections. For each sample, sections from 50 seedlings were pooled. Endogenous IAA content was analyzed by gas chromatography–selected reaction monitoring–mass spectrometry (Edlund et al., 1995). Isotopic dilution was calculated based on the addition of 100 pg 13C6-IAA per sample. Four replicates were analyzed for each sample.

Accession Numbers and Data Deposition

All microarray data will be available in the Genevestigator database (Zimmermann et al., 2004; https://www.genevestigator.ethz.ch) and in the public repository Gene Expression Omnibus upon publication (http://www.ncbi.nlm.nih.gov/geo/) under accession number GSE3350. The accession numbers of the genes used in this study are At4g14550 (SLR/IAA14), At3g54180 (CDKB1;1), At3g48750 (CDKA;1), At4g37490 (CYCB1;1), At4g34160 (CYCD3;1), At2g36010 (E2Fa), At5g02470 (DPa), At5g20730 (ARF7), At5g37020 (ARF8), At5G11030 (ALF4), At3G20840 (PLT1), and At4g35550 (WOX13). Accession numbers of other genes discussed in the text are listed in Table 1.

Supplemental Data

The following materials are available in the online version of this article.

  • Supplemental Table 1. List of the 3110 Significant Genes.
  • Supplemental Table 2. List of the 913 Lateral Root Initiation Genes.
  • Supplemental Table 3. Overrepresented Gene Ontology Functional Categories of LRI Genes versus All Significant Genes and of the Early Induced LRI Genes versus All Significant Genes.
  • Supplemental Table 4. Overlapping Upregulated Genes between This Study and That of Himanen et al. (2004) with the Cluster Combination and the Cluster Number and Profile, Respectively.
  • Supplemental Table 5. Expression Profiles of the LRI Genes in the arf7 arf19 Data Set from Okushima et al. (2005).
  • Supplemental Table 6. LRI Genes with arf7 arf19–Dependent Auxin Inducibility.
  • Supplemental Table 7. Cell Cycle Genes Differentially Regulated at a Stringency Level of P < 0.01.
  • Supplemental Figure 1. PIAA14:GUS Activity throughout Different Stages of Lateral Root Development.
  • Supplemental Figure 2. Histochemical Localization of plt1-1::GUS Activity Using a Promoter-Trap Line during Different Stages of Lateral Root Formation.
  • Supplemental Figure 3. Box Plots of Log2-Transformed Normalized Value Distributions of All Arrays.

Supplementary Material

[Supplemental Data]


We thank Satoshi Tameda for GUS staining, Gun Löfdahl and Marzanna Gontarczyk for skillful technical assistance, the Nottingham Arabidopsis Stock Centre, the ABRC, Peter Doerner, John Celenza, Tom Guilfoyle, and Jim Murray for sharing material, Ben Scheres and Jiří Friml for critical reading of the manuscript, and Martine De Cock for help in preparing it. This work was supported by a grant from the Interuniversity Poles of Attraction Program–Belgian Science Policy (P5/13), in part by Grant-in-Aid for Scientific Research on Priority Areas (Molecular Basis of Axis and Signals in Plant Development) and for Scientific Research for Young Scientists from the Ministry of Education, Culture, Sports, Science, and Technology of Japan to H.F., and a grant from the Research for the Future program of the Japan Society for the Promotion of Science to M.T. S.V. and I.D.S. are indebted to the Institute for the Promotion of Innovation through Science and Technology in Flanders for predoctoral fellowships.


The authors responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) are: Hidehiro Fukaki (h-fukaki@bs.naist.jp) and Tom Beeckman (eb.tnegu.bsp@namkceeb.mot).

W in BoxOnline version contains Web-only data.

Article, publication date, and citation information can be found at www.plantcell.org/cgi/doi/10.1105/tpc.105.035493.


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