Functional characterization of the CKRC1/TAA1 gene and dissection of hormonal actions in the Arabidopsis root

Authors


(fax +86 931 8912565; e-mail rchen@noble.org or +1 580 224 6692; gqguo@lzu.edu.cn).

Summary

Cytokinin (CK) influences many aspects of plant growth and development, and its function often involves intricate interactions with other phytohormones such as auxin and ethylene. However, the molecular mechanisms underlying the role of CK and its interactions with other growth regulators are still poorly understood. Here we describe the isolation and characterization of the Arabidopsis CK-induced root curling 1 (ckrc1) mutant. CKRC1 encodes a previously identified tryptophan aminotransferase (TAA1) involved in the indole-3-pyruvic acid (IPA) pathway of indole-3-acetic acid (IAA) biosynthesis. The ckrc1 mutant exhibits a defective root gravitropic response (GR) and an increased resistance to CK in primary root growth. These defects can be rescued by exogenous auxin or IPA. Furthermore, we show that CK up-regulates CKRC1/TAA1 expression but inhibits polar auxin transport in roots in an AHK3/ARR1/12-dependent and ethylene-independent manner. Our results suggest that CK regulates root growth and development not only by down-regulating polar auxin transport, but also by stimulating local auxin biosynthesis.

Introduction

Virtually every aspect of plant development is controlled by phytohormones, with extensive interactions between their signal transduction pathways (Gray, 2004; Weiss and Ori, 2007). Auxins and cytokinins (CKs) are two such hormones involved in diverse processes including embryogenesis (Müller and Sheen, 2008), post-embryonic root and shoot development (Werner and Schmülling, 2009), and in vitro organogenesis (Skoog and Miller, 1957). Added to the complexity of hormonal regulation is the fact that their de novo biosynthesis, transport and growth responses are often tissue-specific and highly sensitive to developmental and environmental cues (Coenen and Lomax, 1997).

Root growth is critical for plant survival and productivity as roots not only anchor plants in soil but also are the primary organ for uptake of water and mineral nutrients. Root growth is controlled by the rate of cell division in the root apical meristem and cell differentiation at the transition zone immediately after the cell division zone. When the number of cells from cell division equals the number of cells that differentiate and leave the cell division zone, the apical meristem has reached a stable size. Several hormones, including auxin, CK and ethylene, have been shown to control root growth by regulating cell division and differentiation (Ioio et al., 2007; Laplaze et al., 2007; Stepanova et al., 2007). Interestingly, all three hormones inhibit primary root elongation but differ in their regulation of cell division and differentiation. For example, auxin increases, but CK decreases, the root meristem zone (MZ) (Ioio et al., 2007), whereas ethylene has no apparent effect on the root MZ (Růžička et al., 2007). Central to the hormonal regulation of root growth are the auxin maximum at the root tip and auxin concentration gradients generated and maintained by de novo auxin biosynthesis (Zhao et al., 2001; Stepanova et al., 2008; Tao et al., 2008; Zhao, 2008) and carrier-mediated intercellular transport (Ioio et al., 2008; Pernisováet al., 2009; Růžička et al., 2009).

It has been shown that ethylene inhibits cellular elongation at the elongation zone by increasing the auxin level through promotion of auxin biosynthesis and basipetal auxin transport (bPAT) at the root tip (Růžička et al., 2007; Swarup et al., 2007). The effect of CK on root growth is more complex. On the one hand, CK inhibits cellular elongation at the elongation zone, partly due to promotion of ethylene biosynthesis and responses (Cary et al., 1995; Vogel et al., 1998; Rashotte et al., 2005; Kuderova et al., 2008). On the other hand, CK promotes cell differentiation at the transition zone and reduces the root MZ, and this is ethylene-independent (Kuderova et al., 2008; Růžička et al., 2009). The effects of hormones on root growth are dosage-dependent and frequently involve interactions amongst signaling pathways. Recently, CK has been shown to negatively regulate the expression of some PIN genes that encode auxin efflux carriers, indicating that one aspect of the effects of CK on root growth is through the regulation of polar auxin transport (PAT) (Ioio et al., 2008; Pernisováet al., 2009; Růžička et al., 2009). However, how various signaling pathways are integrated to regulate root growth is still largely unknown.

Here we report identification and characterization of the Arabidopsis CK-induced root curling 1 (ckrc1) mutant, a new allele of taa1. Our results indicate that CK not only inhibits PAT but also positively regulate the expression of CKRC1/TAA1 and other auxin biosynthesis genes to increase local auxin levels in the root.

Results

ckrc1 mutants exhibit reduced primary root growth, insensitivity to CK, and a defective root gravitropic response

The roles of auxin and CK in post-embryonic root development are well established. To isolate genes potentially involved in CK and auxin interactions, we initially screened and isolated mutants that were resistant to CK in terms of primary root growth. A dozen such putative CK-resistant mutants were isolated by screening approximately 40 000 T-DNA insertion lines (Alonso et al., 2003), one of which also exhibited strong root curling when grown on a vertical plate in the presence of 0.1 μmtrans-zeatin (ZT) (Figure 1b), 1.0 μm 6-benzylaminopurine (6-BA) (Figure S1c) or thidiazuron (TDZ) (Thomas and Vatteran, 1986) (Figure S1d; Figure 1a). We named this mutant CK-induced root curling 1-1 (ckrc1-1). When grown on a vertical plate in the absence of CK, the ckrc1-1 mutant exhibited reduced primary root and hypocotyl growth (Figure 1a,c,d), and a defective root gravitropic response (GR) compared to wild-type Col-0 (WT) plants (Figure 1e). However, the apical hooks in etiolated seedlings and the shoot GR appeared to be normal, and flowering time under long-day conditions was only slightly delayed in the ckrc1-1 mutant (Figure S2a–d).

Figure 1.

ckrc1-1 exhibits altered responses to CK, root and hypocotyl elongation and an altered root GR.
(a, b) WT and ckrc1-1 seedlings were grown on mock medium (a) and 0.1 μm ZT (b) for 7 days.
(c, d) The elongation rate of primary roots (c) and hypocotyls (d) was reduced in the ckrc1-1 mutant compared with WT plants (= 28–56; ***< 0.001, Student’s t-test).
(e) The root GR after 90° reorientation was altered in the ckrc1-1 mutant compared with WT plants (= 45–83).

To determine whether the reduced primary root growth in ckrc1-1 is a result of decreased root meristematic activity, reduced cellular elongation or both (Beemster and Baskin, 2000; Kuderova et al., 2008), we compared the root structures of ckrc1-1 and WT plants. Figure 2(a–c) shows that the root MZ was significantly reduced in 7-day-old ckrc1-1 seedlings (159.0 ± 34.4 μm) compared with that in WT (235.0 ± 24.3 μm). The difference in the root MZ (32% shorter) is much more pronounced than the difference in the mean mature epidermal cell length (14%) between ckrc1-1 (114.0 ± 20.6 μm) and WT (132.0 ± 17.5 μm) (Figure 2c), suggesting that the reduced primary root growth in the ckrc1-1 mutant was mainly due to reduced meristematic activity. Consistent with this, real-time quantitative RT-PCR data showed that expression of cyclin B1 genes was reduced in ckrc1-1 roots (Figure S3).

Figure 2.

 Root morphology of ckrc1-1.
(a, b) DIC (a) and laser confocal (b) images of representative roots show reduced root MZ in ckrc1-1 compared with WT plants.
(c) The reduction in MZ (= 48–50; **< 0.01, Student’s t-test) is much greater than the reduction in mature epidermal cell length (= 137–140; *< 0. 05).
(d) Starch staining of root cap columella cells of WT and ckrc1-1.

Potassium iodide staining indicated that the appearance of starch granule-filled statolith, a structure that is believed to be a part of the gravitropic sensing apparatus (Chen et al., 1999), was normal in the ckrc1-1 root cap (Figure 2d), suggesting that the defective GR is not caused by a defect in the gravity sensing process. However, only five layers of columella cells were present in the mutant compared to the six layers of columella cells that are typically present in WT roots under the growth conditions tested (Figure 2b). Recently, Ding and Friml (2010) reported that high levels of auxin promote the differentiation of distal stem cells into columella cells in the Arabidopsis root cap, and low levels of auxin in the auxin biosynthesis mutants wei8-1 (allelic to ckrc1-1, see below) and yuc result in fewer tiers of columella cells but extra tiers of distal stem cells. Taken together, these results suggest that CKRC1 plays a key role in multiple developmental processes including primary root growth and the root GR.

ckrc1-1 roots exhibit altered responses to CK, auxin and ethylene

The ckrc1-1 mutant was isolated from a screen of mutants that showed reduced sensitivity of primary root elongation to a low concentration of CK. We further tested the dosage response of its primary root growth to three CKs: ZT, TDZ and 6-BA. ckrc1-1 roots exhibited insensitivity to all three CKs at most concentrations tested. This is in contrast with WT roots, which exhibited increased sensitivity to higher concentrations of CKs (Figures 3a and S4a,b). An exception was that 6-BA at 0.01 μm slightly increased the primary root growth of both WT and the ckrc1-1 mutant (Figure S4a).

Figure 3.

 Primary root elongation response of the ckrc1-1 mutant.
WT and the ckrc1-1 mutant were grown in the presence of various concentrations of ZT (a), IAA (b) and ACC (c), and primary root elongation was measured on the 7th day. All experiments were repeated three times. Values are means ± SD (= 35–60 for each repeat).

Auxin is known to interact with CK and plays an important role in root development and GR. We next tested whether ckrc1-1 roots exhibit altered response to auxin. Application of the auxins indole-3-acetic acid (IAA), 1-naphthyleneacetic acid (1-NAA) and 2,4-dichlorophenoxylic acid (2,4-D) at low concentrations of 0.01, 0.1 and 0.01 μm, respectively, restored the primary root growth of ckrc1-1 mutant to the WT level (IAA) or nearly to the WT level (NAA and 2,4-D) (Figures 3b and S4c,d), and rescued the root curling and defective GR phenotype of the mutant on a vertical plate (Figures S5 and S6a). However, application of 2,4-D and IAA at concentrations of 0.05 μm or higher or of 1-NAA at 0.5 μm or higher similarly inhibited the primary root growth of both WT and ckrc1-1, suggesting that ckrc1-1 roots are as sensitive to high concentrations of auxin as those of WT plants (Figures 3b and S4c,d).

Because the negative effect of CK on root growth is partially linked to ethylene (Cary et al., 1995; Vogel et al., 1998; Rashotte et al., 2005; Kuderova et al., 2008), we also tested the sensitivity of ckrc1-1 to 1-aminocyclopropane-1-carboxylic acid (ACC), the immediate precursor of ethylene biosynthesis. The ckrc1-1 mutant exhibited significant resistance to ACC compared with WT plants (Figure 3c). Similar to the effects of low concentrations of the three auxins, application of 0.32 μm ACC rescued the weak root curling phenotype and the gravitropic defect of the mutant (Figures S1f–i and S6a). However, in contrast to low concentrations of the three auxins, ACC treatment could not rescue the strong curling mutant phenotype induced by 0.1 μm ZT (Figure S5).

Molecular cloning of the CKRC1 gene

To clone the CKRC1 gene, we first back-crossed the ckrc1-1 mutant into WT plants for two generations. As F1 lines showed a WT-like phenotype and F2 lines segregated for WT and ckrc1-1 in a 3:1 ratio (141:48), ckrc1-1 is a single recessive mutation. Inverse PCR amplification of T-DNA flanking sequences (Appendix S1) identified a T-DNA insertion in the 4th intron of the annotated gene At1g70560 (Figure 4a). Subsequent co-segregation analysis in the F2 segregating population linked the T-DNA insertion to the mutant phenotype. Additional sequence analysis of regions surrounding the T-DNA insertion site indicated no sequence alterations in the adjacent regions.

Figure 4.

CKRC1 encodes a tryptophan aminotransferase involved in the IPA pathway of IAA biosynthesis (a) CKRC1 gene structure and location of mutations.
(b) Transcription levels of CKRC1 detected by quantitative RT-PCR in the three alleles and the over-expression line (OE). Values are means ± SD of three biological repeats.
(c) Molecular complementation of mutants by the full-length CKRC1 cDNA [ckrc1-1(C)].
(d) CKRC1 mediates tryptophan-dependent IAA biosynthesis through the IPA pathway (Stepanova et al., 2008; Tao et al., 2008; Sugawara et al., 2009).
(e) Comparison of DR5::GUS expression patterns in root tips of WT and ckrc1-1 supplied with 0.8 μm IPA, 0.04 μm IAA or 3 μm l-tryptophan (l-Trp). In the mutant, IAA and IPA but not tryptophan are capable of restoring the WT pattern, with a densely stained area (auxin response maximum) at the quiescent center (QC).

To confirm that At1g70560 is the CKRC1 gene, we identified two additional mutant alleles (N119862 and N118178) (Tissier et al., 1999) named ckrc1-2 and ckrc1-3 (Figure 4a), respectively. Homozygous lines of both alleles exhibited phenotypes resembling that of ckrc1-1 (Figures S7 and S8). Allelic series tests by genetic crossing showed that neither N119862 nor N118178 complemented the ckrc1-1 mutant phenotype in the F1 generation (Figure S7), indicating that N119862 and N118178 are ckrc1 alleles and At1g70560 is the CKRC1 gene. Consistent with these results, introducing the full-length cDNA of At1g70560 into ckrc1-1 under the control of the CaMV 35S promoter fully complemented the mutant phenotype (Figure 4c).

At1g70560 encodes a protein of 391 amino acids, sharing significant sequence similarities with alliinases from Allium species (EC 4.4.1.4) that are annotated as C–S lyases catalyzing cleavage of cysteine-containing compounds (Figure S9). Database searches indicated that At1g70560 belongs to a family of five members in the Arabidopsis genome. Recently, mutant alleles of At1g70560 were identified from independent screenings for alterations in the shade avoidance response (Tao et al., 2008), the ethylene response (Stepanova et al., 2008) and response to the PAT inhibitor NPA (Yamada et al., 2009). Both in vitro biochemical and in vivo feeding studies, together with structural analysis of the encoded protein, confirmed that At1G70560 is actually a tryptophan aminotransferase (TAA1) that is required for IAA biosynthesis via the indole-3-pyruvic acid (IPA) pathway (Figure 4d) (Stepanova et al., 2008; Tao et al., 2008). As expected, IAA and IPA, but not l-tryptophan can both rescue the ckrc1-1 mutant root phenotype (Figures S5 and S10), and restored the WT auxin response maximum pattern in mutant root tips (Figure 4e). Similar rescuing effects by IAA and IPA have also been reported for the tir2 mutant (Yamada et al., 2009). It should be noted that IPA can be broken down in solution to IAA and other compounds within days (Tam and Normanly, 1998). However, we have not tested whether this has contributed to some aspects of the IPA treatment under our assay conditions. Consistent with a defect in auxin biosynthesis, the endogenous IAA level in the ckrc1-1 mutant was significantly reduced (Figure 5a), but responses to exogenous IAA or ZT treatments were normal, as revealed by quantitative RT-PCR analysis of the expression of IAA1/IAA2 and ARR5/ARR15 genes (Ferreira and Kieber, 2005), respectively (Figure S11). Moreover, root bPAT activity in the ckrc1-1 mutant was reduced (Figure 5b), consistent with the observation that auxin can promotes its own efflux in roots (Paciorek et al., 2005).

Figure 5.

 Endogenous IAA and basipetal auxin transport were reduced in the ckrc1-1 mutant.
(a) Quantification of IAA content in 7-day-old WT and ckrc1-1 seedlings by GC/MS. Values are means ± SD of three biological repeats (*< 0.05, Student’s t-test).
(b) Root bPAT activity levels of WT and the ckrc1-1 mutant. Values are means ± SD (= 6–10; ***< 0.001, Student’s t-test).

Tissue-specific and hormonal regulation of CKRC1 expression

To investigate the details of tissue-specific expression of CKRC1/TAA1, we introduced a CKRC1 promoter (2.2 kb)–GUS reporter fusion (CKRC::GUS) into Arabidopsis WT plants. Staining of the GUS activity in independent transgenic lines showed that the CKRC1 promoter is highly active in the shoot apical meristem, vascular tissues of cotyledons, stems, sepals, stamen filaments, and primary and secondary roots (Figure 6a–c,f), consistent with our quantitative RT-PCR results (Figure 6g). The CKRC1 promoter was also active in the margins of leaves and cotyledons (Figure 6a,b), the shoot and root junction (Figure 6a), root cap columella cells (Figure 6f), the stigma (Figure 6c,e) and the base of the silique (Figure 6d). The expression pattern of the CKRC1::GUS reporter is consistent with the expression pattern of a functional translational fusion (TAA1::TAA1:GFP) previously reported by Stepanova et al. (2008) (Figure 6f,j), but differs from that reported by Yamada et al. (2009).

Figure 6.

CKRC1 expression patterns.
Tissue-specific and hormonal regulation of CKRC1 expression was examined using the CKRC1::GUS reporter gene (a–f), quantitative RT-PCR (g–i) and a TAA1::TAA1:GFP fusion (j–o). Quantitative RT-PCR data are means ± SDs of three biological repeats. Bars labeled with different letters in (g) are significantly different (< 0.01, Student’s t-test). For (h) and (i), asterisks indicate significant differences compared with the control (*< 0.05, **< 0.01). GFP fluorescence (j–o) was observed in more than 50 roots for each treatment.

Primary roots of ckrc1-1 are insensitive to CK and ethylene (Figures 3a,c and S4a,b). To determine the underlying mechanisms, we examined whether the two hormones similarly regulate CKRC1 transcription. Quantitative RT-PCR and TAA1::TAA1:GFP imaging analyses showed that both ZT and ACC up-regulated the CKRC1 transcript level and the GFP signal in WT seedlings and root tips (Figure 6h–o). Up-regulation of TAA1 expression by ACC has also been reported by Stepanova et al. (2008), and ethylene is known to promote local auxin biosynthesis in roots (Růžička et al., 2007; Swarup et al., 2007). To more precisely determine the extent of the IAA increase upon CK treatment, we performed GC/MS analysis of the endogenous IAA levels. The IAA level in 7-day-old seedlings significantly increased by 41% in WT, but by only 23% in ckrc1-1 (difference not statistically significant) (Figure 7), indicating that a significant proportion of CK-induced auxin biosynthesis is dependent on CKRC1.

Figure 7.

 Effects of ZT on endogenous IAA levels in WT and ckrc1-1 seedlings determined by GC/MS.
Values are means ± SDs of seven biological repeats (**< 0.01, Student’s t-test).

It has been shown that CK and ethylene interact in the regulation of primary root growth (Cary et al., 1995; Vogel et al., 1998; Rashotte et al., 2005; Kuderova et al., 2008). We tested whether ethylene and CK also interact in up-regulation of CKRC1 transcription, using chemical inhibitors and mutants with compromised ethylene/CK signaling. Both quantitative RT-PCR and GFP fluorescence analyses showed that the ethylene signaling inhibitor Ag+ effectively blocks the effect of ethylene but not of ZT treatment on CKRC1 transcription (Figure 6h,k–o). A positive effect of ZT treatment was also observed in two ethylene signaling mutants, etr1-3 and ein2-5, but not in the CK signaling mutants ahk3-1 and arr1-3/12-1 (Figure 6i). Collectively, these results suggest that ethylene and CK act independently via their respective signaling pathways in up-regulating CKRC1 expression.

We next investigated whether CK and ethylene (ACC) affect the expression of other genes involved in auxin biosynthesis (Barlier et al., 2000; Zhao et al., 2001, 2002; Cheng et al., 2007; Tao et al., 2008), using the AtGenExpress visualization tool (http://jsp.weigelworld.org/expviz/expviz.jsp). The results show that ZT significantly up-regulates the expression of CKRC1/TAA1 and YUCCA 2, 4 and 8 by >1.5-fold, and down-regulates the expression of CYP79B3, YUCCA 5 and 6, and WEI2/ASA1 by <0.75-fold (Figure S12). On the other hand, ACC up-regulates the expression of CKRC1/TAA1 and YUCCA 2 by >1.5-fold, and down-regulates the expression of SUR2, CYP79B2, CYP79B3, and YUCCA 5 and 6 by <0.75-fold (Figure S12). These data indicate that, although CK and ethylene have different target specificities, they act on a common set of genes involved in auxin biosynthesis.

CK up-regulates local auxin synthesis but down-regulates PAT in the root

Although both CK and ethylene up-regulate CKRC1 transcript levels and induce IAA biosynthesis (Figure 7) (Růžička et al., 2007; Swarup et al., 2007), our data show that CK exacerbates root curling (Figure 1b), whereas ethylene rescues both weak root curling on MS medium and the GR defects of ckrc1-1 (Figures S1i and S6a). Because 2,3,5-triiodobenzoic acid (TIBA) could induce root curling (Figure S1e) and auxin rescued the ckrc1-1 mutant phenotype (Figure S1f–h), mimicking the effects of CK and ethylene, respectively, we examined the effects of CK and ethylene on intercellular PAT. [3H]IAA transport assays showed that ZT significantly reduced but ACC increased the rate of both acropetal PAT (aPAT, from shoot to root) and basipetal PAT (bPAT, from root tip to the elongation zone) in WT and ckrc1-1 roots (Figure 8).

Figure 8.

 Effects of ZT and ACC on relative aPAT (a, b) and bPAT (c, d), as determined using the [3H]IAA feeding method (Daniel et al. 2009).
Relative values were normalized by taking activities of all genotypes on MS (controls) as 1, as absolute values are not directly comparable due to large differences in root structure and length between WT and the various mutant combinations. Values are means ± SD (= 10–17; *< 0.05, **< 0.01 and ***< 0.001, Student’s t-test).

We further tested whether the inhibitory effect of CK and the promoting effect of ethylene on PAT depend on their respective signaling pathways and auxin efflux carriers. The negative and positive effects of ZT and ACC were significantly reduced in the CK signaling mutants ahk3-1 and arr1-3/12-1 and the ethylene signaling mutant etr1-3, respectively (Figure 8), indicating involvement of their signaling pathways. Moreover, ZT also had an inhibitory effect on both aPAT and bPAT in the ethylene signaling mutant etr1-3. Mutations in PIN2, whose product is mainly involved in bPAT, attenuated the negative effect of ZT on bPAT (Figure 8c). Similarly, a mutation in MDR1, whose product is reported to be responsible for about 70% of the aPAT (Lewis et al., 2007), also reduced the effect of ZT on aPAT (Figure 8a).

Effects on auxin biosynthesis and PAT can lead to changes in auxin distribution and alter the expression pattern of the auxin response reporter gene DR5::GUS/GFP (Blilou et al., 2005; Růžička et al., 2007). ZT treatment significantly reduced DR5::GUS reporter gene expression in the root tip of ckrc1-1, but not in the WT (Figure S13a), consistent with stimulation of auxin biosynthesis and inhibition of its PAT by CK. On the other hand, expression of the DR5::GUS reporter gene was increased by ACC treatment of WT plants (Figure S13b), consistent with a promotive effect of ACC on CKRC1 expression and PAT. In contrast to ZT, ACC restored DR5rev::GFP expression in the bottom flank of gravistimulated ckrc1-1 roots (Figure S14). However, ACC did not restore the reduced auxin response maximum in the root tip of ckrc1 (Figure S13b), suggesting a critical role for CKRC1/TAA1-mediated local auxin biosynthesis in maintenance of the auxin response maximum in root tips of WT plants, and possibly accounting for the ability of ethylene and the inability of CK to rescue the root GR defects of the ckrc1 mutant.

Although both auxin (IAA) and CK (ZT) inhibit primary root growth (Figure 9a), IAA increased the root apical MZ, but decreased the mature epidermal cell length (Figure 9b,c) (Růžička et al., 2007). On the other hand, ZT decreased both the apical MZ, which is independent of ethylene, and the mature epidermal cell length, which is dependent on ethylene (Figure 9a–c) (Růžička et al., 2009). To investigate the contribution of CKRC1/TAA1-mediated local auxin synthesis to regulation of root growth, we compared the root growth parameters between WT and ckrc1-1 plants grown in the presence of ZT, ethylene (ACC) and IAA at concentrations that result in 50% inhibition of WT root growth. The results indicate that ACC, ZT and IAA did not inhibit, or only weakly inhibited, the primary root growth of ckrc1-1 compared to the 50% inhibition of WT root growth at the concentrations tested (Figure 9a), indicating a key role for CKRC1/TAA1 in auxin-, CK- and ethylene-mediated inhibition of root growth. In ckrc1-1, the ZT treatment further decreased root MZ with no apparent effect on cell elongation (Figure 9b,c). In contrast, the ACC treatment increased MZ more significantly but decreased mature epidermal cell length less significantly in the ckrc1-1 mutant than in the WT roots in an ethylene-dependent manner (Figure 9b,c).

Figure 9.

 Effects of CK, ethylene and auxin on primary root length, MZ and mature epidermal cell length of WT and ckrc1-1.
Treatment with the ethylene signaling inhibitor Ag+ and the double/triple mutant analyses on ckrc1-1 etr1/ahk3/arr1 arr12 indicate that the effects of ZT on MZ are dependent on AHK3/ARR1/12 signaling, but independent of ethylene. Values are means ± SD (> 60 seedlings; *< 0.05, **< 0.01 and ***< 0.001, Student’s t-test).

Discussion

Separate but overlapping functions of CK and ethylene in auxin local biosynthesis and PAT in the root

Several auxin over-production mutants have been identified, but few auxin-deficient mutants have been reported (Woodward and Bartel, 2005; Tao et al., 2008). Here we show that the ckrc1 mutant is deficient in auxin and allelic to other three reported auxin-deficient mutants, namely shade avoidance 3 (sav3) (Tao et al., 2008), weak ethylene insensitive 8 (wei8) (Stepanova et al., 2008) and transport inhibitor response 2 (tir2) (Yamada et al., 2009). Stepanova et al. (2008) and Tao et al. (2008) independently cloned the same gene, which they named TAA1, and showed that TAA1 has tryptophan aminotransferase activity, converting tryptophan into IPA. The IPA pathway is one of multiple pathways proposed for the biosynthesis of IAA (Figure 4d) (Woodward and Bartel, 2005) and is the least characterized (Tao et al., 2008). TAA1/CKRC1 has been shown to be required for plant shade avoidance responses, tissue-specific effects of ethylene, embryo and organ development, and hypocotyl elongation. Here we show that CK up-regulates local auxin biosynthesis through positive regulation of TAA1/CKRC1 transcription and down-regulates PAT in the root. Results from phenotypic analysis of the ckrc1 mutant indicate that ethylene and CK play opposite roles in PAT, and local auxin biosynthesis is required to maintain the correct root apical meristem size.

CK promotes local auxin biosynthesis by up-regulating CKRC1/TAA1 transcription, but inhibits PAT in the root

The regulatory mechanisms underlying CK and auxin interactions have largely remained a mystery. The inhibitory effect of CK on root growth has been previously shown to mainly depend on ethylene (Cary et al., 1995; Vogel et al., 1998; Rashotte et al., 2005; Kuderova et al., 2008), which is known to stimulate local auxin biosynthesis and PAT (Růžička et al., 2007; Swarup et al., 2007; Negi et al., 2008). However, recent evidence has indicated that the negative effect of CK on MZ is ethylene-independent (Kuderova et al., 2008; Růžička et al., 2009). That CK is inhibitory to auxin transport is inferred indirectly from the negative effect of CK on expression of the auxin efflux carrier PIN gene in Arabidopsis and tobacco BY-2 cells (Ioio et al., 2008; Pernisováet al., 2009; Růžička et al., 2009). Ioio et al. (2008) reported that ZT inhibits expression of three PIN genes (PIN1, 3 and 7) through the ARABIDOPSIS HISTIDINE KINASE 3/ARABIDOPSIS RESPONSE REGULATOR 1 (AHK3/ARR1) two-component signaling pathway involving the downstream SHY2 gene, encoding a repressor of auxin signaling. Růžička et al. (2009) reported that benzylaminopurine significantly down-regulates PIN1 and 3 expression, but stimulates PIN7 expression at both transcription and protein levels. These disagreement between studies may be due to the different chemicals used (i.e. ZT versus benzylaminopurine) and/or different materials/conditions, or may be a reflection of the complex and subtle nature of CK regulation of PINs. Using direct [3H]IAA transport assays, our results confirm the overall negative effect of CK on PAT. We further show that the negative effect of CK is partially mediated by its regulation of auxin efflux carrier genes via the AHK3/ARR1/12 signal pathway, but not the ETHYLENE RESPONSE 1 (ETR1)-mediated ethylene signal pathway (Figure 8). Combined with its positive effect on auxin biosynthesis, these results suggest that CK can promote local accumulation of auxin.

Both aPAT and bPAT are similarly inhibited by CK in WT and ckrc1-1 (Figure 8). However, there are significant differences in root growth and MZ size between WT and ckrc1-1 in response to CK. These observations indicate a key role for CKTC1/TAA1-mediated auxin biosynthesis in CK regulation of root growth, which has not been reported previously. The reduced MZ size in ckrc1 roots suggests a role for CKRC1-mediated auxin biosynthesis in the maintenance of MZ size (Figure 2a–c). However, the finding that CK up-regulates CKRC1 in WT roots (Figure 6h–o) appears to contradict its negative effect on MZ size (Beemster and Baskin, 2000; Werner et al., 2003). Our results suggest that CK regulates MZ size through its combined effects on auxin biosynthesis and PAT. Consistent with this view, Blilou et al. (2005) reported that the root MZ size is determined by an ‘auxin reflux’ loop mediated mainly by the auxin efflux carriers PIN2, PIN3, and PIN7. Interestingly, Ioio et al. (2007) reported that CKs derived from the vascular tissue in the transition zone have a negative effect on MZ. Recently, Jones et al. (2010) reported that CK induces auxin biosynthesis and increases the steady-state level of auxin in roots and shoots of Arabidopsis. They showed that CK up-regulates by more than twofold several tryptophan-dependent IAA biosynthesis genes, including CYP79B2, CYP79B3, YUCCA6 and NIT3, in developing root tips. A statistically insignificant increase (at the P < 0.05 level) in TAA1 expression was detected after 24 h of ZT treatment. However, quantitative RT-PCR analysis after short-term ZT treatments (<6 h) revealed no positive effects of CK on TAA1 expression. Our results using various mutants and molecular analysis of Arabidopsis roots clearly indicate that CK negatively regulates root growth by stimulating TAA1/CKRC1-mediated local auxin biosynthesis in an AHK3/ARR1/ARR12-dependent manner. Future work to identify effectors of the AHK3/ARR1/ARR12 signaling pathway and the role of aPAT and bPAT in CK regulation of root growth promises to shed new light on the complex interactions between CK and auxin in regulating plant development.

Experimental procedures

Plant material and growth conditions

All Arabidopsis mutant alleles and lines except ahk3-1 (Ws ecotype) are in the Col background. Germination and plant growth took place at 22°C with a 16 h light/8 h dark cycle. For growth analyses, seedlings were grown on MS medium containing 0.8% w/v agar and 30 g L−1 sucrose. For vertical growth, 1% w/v agar plates were used.

Mutant screening and genetic analysis

Approximately 40 000 Arabidopsis thaliana (Col-0) T-DNA insertion lines (stock numbers CS76502, CS76504, CS76506 and CS76508) were purchased from the Arabidopsis Biological Resource Center (http://abrc.osu.edu/). Seeds were germinated on MS medium containing 0.45 μm TDZ (0.1 mg L−1). Putative mutant seedlings were screened 2 weeks later. They were planted into soil to harvest seeds for further genetic analysis and allelism tests.

To introduce a reporter gene into the mutant, DR5::GUS marker line (Ulmasov et al., 1997) was crossed with ckrc1-1 and double homozygotes were identified in the F3 population. etr1-3 (stock number N3070), eto1-1 (N3072), ahk3-1 (N6562) and arr1-3 arr12-1 (N6981) were obtained from the Arabidopsis Biological Resource Center or the Nottingham Arabidopsis Stock Centre. To generate double mutants, mutants were crossed with ckrc1-1, the F1 progeny of the crosses were propagated, and double homozygotes were identified in the F2 population.

Phenotype characterization

For root inhibition assays, seedlings were germinated and grown vertically on MS medium containing various hormones or compounds for 7 days. All the data were the mean of three separate experiments using at least 20 seedlings.

For the root GR, approximately 80 seedlings were assessed for each genotype or allele. To assess the inflorescence stem GR, 5-week-old bolted plants were placed horizontally in a completely dark place at 23ºC, and the GR of their stems was photographed after 1, 2 and 5 h. The hypocotyl GR was tested by subjecting 3-day-old dark-grown seedlings to a gravitational stimulus for 24 h.

Both DIC and confocal microscopy were performed on 5-day-old vertically grown roots. For DIC, roots were cleared by the chloral hydrate method as described by Inagaki et al. (2006). For confocal microscopy, fresh materials were stained with 10 μg ml−1 propidium iodide for 30 min, and the excitation wavelength was 543 nm. For starch grain observation, roots of 4-day-old seedlings were stained in Lugol solution (1.3% I2 and 2% KI) for 3 min, followed by clearing in chloral hydrate solution.

Gene cloning and sequence analysis

Inverse PCR was used to clone the mutant gene. Genomic DNA was digested completely using HindIII and ligated using T4 DNA ligase. Two rounds of PCR were performed using two sets of nested primers, LBa1/Z4 and LB6313/Z3 (Appendix S2). The PCR fragments were subcloned and sequenced. The downstream flanking sequence was amplified by PCR using LB6313 and CO1404R primers (Appendix S2). Flanking sequences (Appendix S1) were used to design gene-specific primers to determine hetero-/homozygosity and co-segregation.

For sequence alignment, homologs in Arabidopsis and other organisms were obtained using a BLASTP query on A. thaliana UniProt (protein) sequences and the green plant protein database (http://arabidopsis.org/Blast/index.jsp). Multiple sequence alignments were performed using the Clustal W service of the NPS@;ClustalW multiple alignment website (http://npsa-pbil.ibcp.fr/cgi-bin/npsa_automat.pl?page=npsa_clustalw.html) and BoxShaded at its server website (http://www.ch.embnet.org/software/BOX_form.html). The resulting alignments were used to generate a neighbor-joining tree using Mega 2.1 software, with the garlic alliinase (1LK9_A) (Kuettner et al., 2002; Shimon et al., 2007) as the outgroup (Kumar et al., 2001).

Molecular complementation and promoter::GUS transgenic plants

A 2.2 kb promoter sequence was amplified using PL1404/PR1404 primers (Appendix S2) and subcloned into a modified pCAMBIA1300 binary vector which harbors a GUS gene to generate a promoter::GUS reporter gene construct. A 1.3 kb full-length cDNA fragment was amplified by RT-PCR using cD1404L/cD1404R primers (Appendix S2), and placed under the control of the CaMV 35S promoter in a pCAMBIA1300 vector. All amplified DNA fragments were confirmed by sequencing, and the constructed binary vectors were introduced into either WT plants (for promoter::GUS) or ckrc1-1 plants (for 35S-cDNA) by an Agrobacterium tumefaciens-mediated (strain GV3101) floral-dip transformation method (Clough and Bent, 1998). Primary transformants were isolated on MS medium containing 25 mg L−1 hygromycin (Sigma, http://www.sigmaaldrich.com/) and transferred to soil to grow to maturity.

RNA preparation and expression analysis

RNA was isolated using Trizol (Invitrogen, http://www.invitrogen.com/) and reverse-transcribed using a reverse transcription kit (Takara, http://www.takara-bio.com/). Quantitative RT-PCR was performed in an ABI7000 real-time PCR system (http://www.appliedbiosystems.com/) using Power SYBR green chemistry (Takara). Primer sequences used are listed in Appendix S2.

For determination of the regulation of CKRC1 expression by various hormones, 7-day-old seedlings grown on MS medium were treated separately with 1.0 μm ZT, 10 μm ACC or 1 μm IAA for 1 h, 10 μm Ag+ for 3 h, or 10 μm Ag+ for 2 h followed by 1.0 μm ZT or 10 μm ACC for 1 h.

For analysis of cytokinin- and auxin-inducible gene expression, 7-day-old seedlings grown on MS medium were treated in liquid MS medium with 10 μm ZT for 30 min (Laxmi et al., 2006) or 20 μm IAA for 1.5 h (Tian et al., 2002).

Endogenous auxin measurement

Endogenous IAA levels were measured in 7-day-old seedlings as described by Edlund et al. (1995). Seven-day-old seedlings were treated with 10 μm ZT for 24 h in liquid MS medium, and samples were purified along with [13C6]IAA internal standard (Cambridge Isotope Laboratories, http://www.isotope.com/). Approximately 70 mg fresh weight material was used for each sample and the [13C6]IAA internal standard was added before homogenization. After extraction and purification, trimethylsilylation was performed by treatment with 20 μl bis-(trimethylsilyl)trifluoroacetamide and 10 μl pyridine at 60°C for 30 min. Levels of endogenous IAA were determined by GC-TOF MS (Pegasus® IV; LECO, http://www.leco.com/) by measuring the ratio of the abundance of m/z 202 to that of m/z 208.

Histochemical GUS assay

Roots or seedlings of the DR5::GUS marker line and the ckrc1-1/DR5::GUS double homozygous line after various treatments were incubated in 1 mm X-gluc (5-bromo-4-chloro-3-indolyl-β-d-glucuronide) and 50 mm potassium phosphate buffer, pH 7.5, with 0.1% v/v Triton X-100 for GUS staining as described by Jefferson et al. (1987).

PAT assay

The isotope method for the PAT assay has been described previously (Lewis and Muday, 2009). Seedlings were germinated on control medium, and transplanted to control or treatment plates on the 6th day after sowing.

For aPAT, after a 24 h treatment, a 100 nm [3H]IAA agar cylinder was applied at the aligned root/shoot junctions, and the seedlings were incubated in the dark in the inverted position, to prevent [3H]IAA from diffusing along the root, for 18 h. The apical 5 mm of each root tip was excised. Every batch of four segments were placed in 2.5 ml of scintillation liquid in a 3 ml scintillation vial, and radioactivity was measured for 2 min on a Beckman scintillation counter.

For bPAT, after a 24 h treatment, a 100 nm [3H]IAA agar cylinder was applied next to the root tip, and seedlings were incubated in the dark. bPAT was measured after 5 h by first removing the apical 1 mm in contact with the agar line, then excising an apical section of approximately 5 mm from what remains. Every four segments were placed in 2.5 ml of scintillation liquid in a 3 ml scintillation vial, and radioactivity was measured for 2 min on a Beckman scintillation counter.

Acknowledgements

We thank Dr Jane Murfett (Department of Biochemistry, University of Missouri, Columbia, MO) for providing DR5::GUS, Jose M. Alonso (Department of Genetics, North Carolina State University, Raleigh, NC) for TAA1::GFP, the Arabidopsis Biological Resource Center and the Nottingham Arabidopsis Stock Centre for mutant pools and individual lines, and Miss Zhen Xue (Institute of Botany, Chinese Academy of Sciences) for IAA quantification. This work was supported by grants from the Chinese National Science Foundation (90717114) and the Chun Hui project from the Ministry of Education of China.

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