The role of floral organs in carpels, an Arabidopsis loss-of-function mutation in MicroRNA160a, in organogenesis and the mechanism regulating its expression

Authors


For correspondence (fax +1 414 229 3926; e-mail dzhao@uwm.edu).

Summary

MicroRNAs (miRNAs) have emerged as key regulators of gene expression at the post-transcriptional level in both plants and animals. However, the specific functions of MIRNAs (MIRs) and the mechanisms regulating their expression are not fully understood. Previous studies showed that miR160 negatively regulates three genes that encode AUXIN RESPONSE FACTORs (ARF10, -16, and -17). Here, we characterized floral organs in carpels (foc), an Arabidopsis mutant with a Ds transposon insertion in the 3′ regulatory region of MIR160a. foc plants exhibit a variety of intriguing phenotypes, including serrated rosette leaves, irregular flowers, floral organs inside siliques, reduced fertility, aberrant seeds, and viviparous seedlings. Detailed phenotypic analysis showed that abnormal cell divisions in the basal embryo domain and suspensor led to diverse defects during embryogenesis in foc plants. Further analysis showed that the 3′ region was required for the expression of MIR160a. The accumulation of mature miR160 was greatly reduced in foc inflorescences. In addition, the expression pattern of ARF16 and -17 was altered during embryo development in foc plants. foc plants were also deficient in auxin responses. Moreover, auxin was involved in regulating the expression of MIR160a through its 3′ regulatory region. Our study not only provides insight into the molecular mechanism of embryo development via MIR160a-regulated ARFs, but also reveals the mechanism regulating MIR160a expression.

Introduction

MicroRNAs (miRNAs) are small (∼21 nucleotides) non-coding RNAs that are key post-transcriptional controllers of gene expression in both plants and animals. MicroRNAs regulate gene expression by guiding the cleavage of target mRNAs or attenuating the translation of their target genes (Bartel and Bartel, 2003; Carrington and Ambros, 2003; Jones-Rhoades et al., 2006; Chen, 2008; Bartel, 2009; Voinnet, 2009). In plants, primary miRNAs (pri-miRNAs) are transcribed from MIRNA (MIR) genes. Stem–loop segments derived from pri-miRNAs are cleaved by RNase III-type endonucleases (Dicer) to produce paired miRNAs with approximately 21 nucleotides, including two-nucleotide 3′ overhangs. After liberation of the miRNA duplexes, mature miRNAs are loaded into an RNA-induced silencing complex (RISC), where they interact with target mRNAs by complementary matching for cleavage or translational attenuation. The general functions of miRNAs have mainly been revealed by analyzing mutants that are impaired in miRNA biogenesis, such as dcl1 (Park et al., 2002; Kurihara and Watanabe, 2004), ago (Kidner and Martienssen, 2004; Baumberger and Baulcombe, 2005), hen1 (Boutet et al., 2003), hyl1 (Song et al., 2007), se (Grigg et al., 2005; Yang et al., 2006), and ddl (Yu et al., 2008) mutants. The specific functions of miRNAs are primarily established through gain-of-function approaches, including ectopic expression of miRNAs and expression of miRNA-resistant versions of their targets. So far, only a handful of loss-of-function mutants with detectable phenotypes have been identified in MIRs in plants and other organisms (Miska et al., 2007; Voinnet, 2009). In addition, the temporal and spatial actions of miRNAs have not been extensively studied. Therefore, it is of great importance to investigate the function of individual miRNAs and the mechanisms controlling MIR gene expression.

Auxin is essential for plant vegetative growth and reproductive development. AUXIN RESPONSE FACTORs (ARFs), which are released by the auxin receptor E3 ubiquitin ligase complex SCFTIR1, regulate the expression of a large set of auxin-responsive genes by binding to auxin response elements (AuxREs) in their promoters (Guilfoyle and Hagen, 2007; Mockaitis and Estelle, 2008). Among 23 ARFs in Arabidopsis, recent studies showed that miRNAs and trans-acting-small interfering RNAs (ta-siRNAs) play important roles in controlling transcript abundance of ARF2, -3, -4, -6, -8, -10, -16, and -17 (Okushima et al., 2005; Guilfoyle and Hagen, 2007). The trans-acting small interfering RNA ARFs (tasiR-ARFs) target ARF2, -3, and -4 (Allen et al., 2005; Williams et al., 2005; Fahlgren et al., 2006). The gradient of tasiR-ARFs established by small RNA movement is critical for patterning ARF3 in leaf development (Chitwood et al., 2009). The expression of ARF6 and -8 is regulated by miR167, which is essential for anther and ovule development (Wu et al., 2006). Furthermore, gain-of-function analyses showed that miR160 is important for plant development by negatively regulating the expression of ARF10, -16, and -17 (Mallory et al., 2005; Wang et al., 2005; Liu et al., 2007). Based on transient expression experiments and protein structures, ARFs 5–8, and 19 may function as transcriptional activators, while ARFs 1–4, 9–18, and 20–22 possibly act as repressors (Ulmasov et al., 1999; Tiwari et al., 2003; Okushima et al., 2005). So far, only ARF5 (MONOPTEROS) and ARF7 (NON-PHOTOTROPIC HYPOCOTYL4) have been shown to play direct roles in embryogenesis (Hardtke and Berleth, 1998; Harper et al., 2000; Hamann et al., 2002; Hardtke et al., 2004). Plants expressing a miR160-resistant version of ARF17 produced aberrant seedlings, suggesting that miR160 may control early embryo development (Mallory et al., 2005). miR160 is derived from three genes: MIR160a, -b, and -c. However, it is not clear how a specific MIR160 regulates plant development, particularly embryo development, by post-transcriptional regulation of ARF10, -16, and -17.

Arabidopsis has about 190 MIR genes (http://www.mirbase.org). To date, the function of only a few miRNAs has been studied by loss-of-function analysis (Vaucheret et al., 2004; Baker et al., 2005; Gasciolli et al., 2005; Guo et al., 2005; Allen et al., 2007; Cartolano et al., 2007; Sieber et al., 2007; Nag et al., 2009; Wu et al., 2009). For example, mir159a or mir159b single mutants have no phenotypes, while the mir159ab double mutant exhibited diverse abnormalities, including reduced apical dominance, curled leaves, reduced fertility, and small seeds (Allen et al., 2007). eep1, which is impaired in MIR164c function, produced extra petals in early flowers (Baker et al., 2005). Both mir164a and mir164b single mutants formed more lateral roots (Guo et al., 2005). Analysis of the mir164abc triple mutant uncovered functional redundancy and specialization among three MIR164 genes (Sieber et al., 2007). Here, we characterize a miRNA loss-of-function mutant, floral organs in carpels (foc), in which the MIR160a gene is disrupted by a Ds transposon insertion in its 3′ regulatory region. foc plants exhibited a wide range of intriguing phenotypes in leaf, flower, embryo, and seed development. We demonstrate that the 3′ region of MIR160a is required for its expression pattern. During embryogenesis in foc plants, abnormal cell divisions in the basal embryo domain and suspensor cause various embryonic defects mainly due to the altered expression pattern of ARF16 and -17. foc plants are also deficient in auxin responses. Moreover, auxin regulates the expression of MIR160a, possibly through potential auxin response elements in its 3′ regulatory region. Our results suggest that MIR160a is required for the development of multiple organs, particularly the embryo, through a mechanism involving auxin signaling.

Results

foc has defective flower development

To identify those genes important for reproductive development using the loss-of-function approach, we screened Arabidopsis Ds transposon insertion lines (Sundaresan et al., 1995; Zhao et al., 2002; Jia et al., 2008). One mutant showed interesting phenotypes in both vegetative and reproductive development. Besides producing serrated rosette leaves (Figure S1 in Supporting Information), the mutant plants exhibited diverse abnormalities in reproductive development, including flower development. The mutant plants produced abnormal flowers with long pedicles (Figures 1a,b and S2a,b). Wild-type buds are enclosed by sepals and open flowers have four sepals, four petals, six stamens, and two fused carpels (Figure 1a,c,g). However, the mutant plants frequently formed buds with unfurled sepals (Figure 1b,h). Seventy per cent of the mutant flowers had narrow sepals and petals (Figure 1d), and flowers produced in late stages had variable numbers of floral organs (Figures 1e and S2e–h). In short-day conditions, 30% of the first 10 mutant siliques became swollen (Figure 1f). More strikingly, floral organs formed inside these swollen carpels (Figures 1i and S2i–k). We therefore named this mutant floral organs in carpels (foc). Eventually, inflorescences emerged from siliques (Figures 1j and S2i). Our results suggest that FOC plays important roles in controlling floral organ identity and formation.

Figure 1.

foc has defective flower development.
(a) A wild-type inflorescence. (b) A foc inflorescence showing flowers with unfurled sepals. (c) A wild-type flower. (d) A foc flower exhibiting narrow sepals and petals. (e) A foc flower showing one sepal-like, two petal-like, and one stamen-like structures. (f) Developing wild-type silique (left) and two foc swollen siliques (middle and right). (g) Scanning electron microscope (SEM) image showing a wild-type stage 7 bud enclosed by sepals. (h) A SEM image showing an unfurled foc stage 7 bud, indicated by stalks on stamen primordia and the slotted tube of the gynoecium (arrowheads, sepal primordia; arrow, a filamentous structure). (i) A SEM image showing floral organs generated inside a foc silique (arrows, stamen-like structures; arrowheads, ovules). (j) An emerged inflorescence from a foc silique.
Scale bars: 1 mm in (a–f), (j); 25 μm in (g, h); 50 μm in (i).

foc has abnormal seed and embryo development

foc plants are defective in seed development and exhibit reduced fertility (Figure S3a–e). Compared with wild-type seeds, foc seeds are variable in size (Figure S3f,g). Moreover, foc plants show a viviparous phenotype (Figure S3c–i). Semi-thin sectioning revealed various abnormalities in late seed development (Figure S3j–m), which cause diverse defects in young seedlings (Figure S4).

The seed and seedling phenotypes suggest that foc plants are defective in early embryo development. Therefore, we analyzed embryo development by preparing whole-mount squashes of embryos. Wild-type embryo development is initiated from a zygote. Lineages with different developmental fates are initially established by producing a small, spherical apical (or terminal) cell and a large elongated basal cell (West and Harada, 1993; Laux et al., 2004). The first round of cell division in the apical and basal cells results in the formation of a two-celled embryo consisting of an embryo proper and a suspensor (Figure 2a). Subsequent cell divisions give rise to embryos at the octant (eight cells, Figure 2b), dermatogen (16-cells, Figure 2c), globular (Figure 2d), triangular (or transition, Figure 2e), heart (Figure 2f), torpedo (Figure S3j), and bent cotyledon stages (Figure S3l). However, abnormal embryo development in foc occurred at the very beginning of embryogenesis. In wild-type embryos, the suspensor contains a single file of cells that are derived from a series of transverse divisions of the basal cell (Figure 2a–e). In foc embryos, at the two- to four-cell stage, suspensor cells appeared to divide both transversely and longitudinally, resulting in the formation of a double-filed suspensor (Figure 2g). Furthermore, the mutant embryo proper failed to differentiate normally, leading to a non-spherical and asymmetric embryo at the octant stage (Figure 2h). foc embryos did not form normal hypophyses due to abnormal cell divisions in both the suspensor and embryo proper (Figure 2i,j). These early defects caused aberrant embryos in later stages (Figure 2k,l).

Figure 2.

foc has defective early embryo development.
(a, g) A wild-type embryo at the two to four cell stage, showing a longitudinal cell division in the apical cell and transverse cell division in suspensor cells (a) while a foc embryo shows aberrant longitudinal cell divisions in suspensor cells, resulting in a double-filed suspensor (g, arrowheads).
(b, h) A wild-type embryo at the octant (eight cell) stage exhibiting an embryo proper resulting from both transverse and longitudinal cell divisions, and a single-filed suspensor resulting exclusively from transverse cell divisions (b), while a foc embryo shows abnormal cell divisions in central and basal embryo domains (h, arrow).
(c, i) A wild-type embryo at the dermatogen stage (16 cell, c); while a foc embryo at the dermatogen-like stage shows abnormal cell divisions in the embryo proper (i).
(d, j) A wild-type embryo at the globular stage showing hypophysis (d, arrowheads), while a foc embryo at the globular-like stage shows a triple-filed suspensor (j, arrow), but no typical hypophysis.
(e, k) A wild-type embryo at the triangular (transition) stage, showing cotyledon buttresses (e, arrows), while a foc embryo at the triangular-like stage has no suspensor (k).
(f, l) A wild-type embryo at the heart stage, showing enlarged cotyledon lobes (f, arrows), while a foc embryo at the heart-like stage shows two asymmetric cotyledon lobes (l, arrows).
Scale bars: 25 μm in (a–l).

By analyzing embryos at specific times after fertilization, we found that the distribution of foc embryos at different stages was significantly different from that of wild type (Table 1). By 2 days after pollination (DAP), most wild-type embryos (93% or 119/128) had reached the dermatogen and globular stages, while only 7% (9/128) of embryos were at the quadrant and octant stages (Table 1). In contrast, by 2 DAP, 51% (45/88) of foc embryos were still at the quadrant or octant stage (Table 1). After 2 DAP, besides morphological defects, the development of foc embryos was more severely retarded and developmental stages were widely distributed (Table 1). In summary, abnormal cell divisions in the basal embryo domain and uppermost part of the suspensor caused various defects during embryogenesis in foc plants. Furthermore, embryo development in foc plants was severely retarded and asynchronous.

Table 1.   Comparison of embryogenesis between wild-type and foc plants. Siliques from wild-type and foc plants were dissected. Whole-mount preparations of embryos were cleared and examined and the numbers of embryos at each developmental stage were recorded
DAPGenotypeQuadrant or octantDermatogenGlobularTriangularHeartTorpedoBent cotyledonMature or desiccation
  1. DAP, days after pollination.

2wt96257     
 foc45376     
3wt  55670   
 foc82254177   
4wt    19958 
 foc 113319163 6
5wt     3499 
 foc  4121042244
7wt       145
 foc    2 3102

FOC is MIR160a

To identify FOC, we first performed genetic analyses. Wild-type plants were crossed to foc plants. All F1 plants exhibited the wild-type phenotype, suggesting that foc is recessive. In the F2 generation, approximately one quarter (55 mutant: 171 wild type) of the plants displayed mutant phenotypes, indicating that these diverse phenotypes may be caused by a single gene mutation. However, thermal asymmetric interlaced (TAIL)-PCR (Liu et al., 1995; Zhao et al., 2002) revealed two Ds insertion sites within 6198 bp of each other (Figure S5a). One Ds (designated Ds1) was located 835 bp upstream of the ATG start codon of At2g39170 (Figure S5a), which encodes an unknown protein. The other Ds (Ds2) was located downstream of MIR160a (At2g39175) (Figure S5a). We cloned the 967-bp full-length transcript of MIR160a by 5′ and 3′ rapid amplification of cDNA ends (RACE)-PCR (Figure S6) and determined that the Ds2 insertion was 1635 bp from the 3′ region of MIR160a.

Our RT-PCR results demonstrated that MIR160a was expressed in seedlings, stems, roots, inflorescences, and mature leaves in wild-type plants (Figure S5b). However, in foc, the MIR160a transcript was not detected in stems, inflorescence, or mature leaves. In young seedlings, MIR160a expression was decreased, while its expression in roots appeared normal. Our results indicate that Ds2 severely disrupts the expression of MIR160a.

Due to the linkage of two Ds insertions, we performed complementation experiments. Two constructs, ETA and ETB, harboring a 3820 bp genomic DNA fragment for At2g39170 and a 4921 bp fragment for MIR160a (At2g39175) respectively, were generated to create transgenic plants. Thirty PCR-verified transgenic plants for each construct were chosen to cross with +/foc plants. In 23 F2 populations, plants with the ETB transgene exhibited a wild-type phenotype in the foc background, after PCR verifications. Conversely, every plant carrying the ETA transgene still had foc phenotypes in the foc background. Taken together, our results strongly suggest that Ds2 inserted in the 3′ region of MIR160a is responsible for the foc phenotypes.

The 3′ region of FOC is required for its expression pattern

To further determine whether the 3′ region is required for the FOC expression, we generated three constructs using the GUS reporter gene (Figure 3a). By analyzing plants expressing a ProFOC:GUS:3′FOC (Figure 3a, #1, wild-type version) transgene, we found that FOC was expressed in cotyledons, roots, and rosette leaves of young seedlings (Figure 3b top, c). FOC was also strongly expressed in root tips (Figure 3e), lateral roots (Figure 3g), stems (Figure 3i), vascular tissues of mature leaves (Figure 3k), and in young buds and open flowers (Figure 3m,p). FOC was primarily expressed in anthers and carpels (Figure 3s) as well as in developing embryos (Figure 3v, leftmost three, w). Our results indicate that FOC is expressed in all organs, especially in reproductive organs.

Figure 3.

 The 3′ region of FOC is required for its expression pattern.
(a) A diagram showing three constructs: #1, ProFOC:GUS:3FOC; #2, ProFOC:GUS:Δ3FOC; and #3, ProFOC:GUS (open arrowhead indicates the site of the Ds insertion in the 3′ region of foc, while solid arrowheads indicate the positions of three potential auxin response elements in the 3′ region of FOC). (b) A ProFOC:GUS:3FOC young seedling (top) expressing GUS at similar level in root, but at a higher level in cotyledons, relative to expression levels in a ProFOC:GUS plant (bottom). (c, d) GUS activity was higher in cotyledons and young true leaves of ProFOC:GUS:3FOC seedlings (c) than that of ProFOC:GUS (d). (e–h) ProFOC:GUS:3FOC plants with similar GUS expression patterns in root tip (e) and lateral root (g) as in ProFOC:GUS plants (f, h). (i, j) GUS staining in stem vascular bundles of ProFOC:GUS:3FOC plants (i), but not of ProFOC:GUS plants (j). (k, l) GUS staining in mature true leaves of ProFOC:GUS:3FOC (k), but almost undetectable in ProFOC:GUS leaves (l). (m–r) GUS staining of inflorescences showing strong expression in ProFOC:GUS:3FOC young buds and open flowers (m, p), with GUS signal but greatly reduced expression in ProFOC:GUS:Δ3′FOC (n, q). In ProFOC:GUS plants, GUS activity was almost undetectable in young buds and markedly reduced in open flowers (o, r). (s–u) Relative to expression in ProFOC:GUS:3′FOC flower (s), GUS activity in carpels was reduced in ProFOC:GUS:Δ3′FOC (t) and ProFOC:GUS flowers (u). (v–x) GUS signal in ProFOC:GUS:3′FOC embryos (v, left three and w), but not in ProFOC:GUS embryos (v, right three and x). (y) Quantification of GUS activity (nm 4-methylumbelliferone (4-MU)/min/mg).
Abbreviation: Inf., inflorescence. Scale bars: 1 mm in (b–d, m–o), (v); 50 μm in (e–h), (w, x); 0.5 mm in (i, j), (p–u); 0.5 cm in (k, l).

Transgenic plants harboring ProFOC:GUS (Figure 3a, #3, lacking the 3′ region) and ProFOC:GUS:Δ3FOC (Figure 3a, #2, with a partial 3′ region) showed GUS staining patterns different from plants expressing ProFOC:GUS:3FOC. In ProFOC:GUS plants, GUS staining in the young seedling roots (Figure 3b bottom), root tips (Figure 3f), and lateral roots (Figure 3h) was similar to that of ProFOC:GUS:3FOC (Figure 3b top, e,g). However, GUS activity was reduced in cotyledons and young rosette leaves (Figure 3b bottom, d). Almost no GUS activity was detected in stems (Figure 3j), mature leaves (Figure 3l), young buds (Figure 3o,r), carpels (Figure 3u), and developing embryos (Figure 3v right three, x). ProFOC:GUS:Δ3FOC plants showed reduced GUS staining intensities when compared with ProFOC:GUS:3FOC (Figure 3n,q,t) plants. The quantification of GUS activity confirmed the results from GUS staining (Figure 3y). Our results indicate that the 3′ regulatory region of FOC is required for its expression in aerial organs, particularly in reproductive organs, including flowers and embryos.

The accumulation of mature miR160 and expression of ARF10, -16, and -17 are altered in foc plants

To examine the effect of the foc mutation on overall accumulation of mature miR160, we performed small RNA Northern blot assays. In the wild type, miR160 was readily detected in both leaves and inflorescences, with higher levels in the inflorescences (Figure 4a). However, miR160 was substantially reduced in foc leaves and inflorescences (Figure 4a). Accumulation of miR160 showed a 72% reduction in foc inflorescences, relative to levels in wild type, while a 21% reduction was found in leaves (Figure 4b), suggesting that the foc mutation has a more profound effect on miR160 homeostasis in the inflorescence. The effect of the foc mutation on accumulation of miRNA appeared to be specific to miR160, since the accumulation of miR168 was not affected in either leaves or inflorescences (Figure 4c,d). The accumulation of miR171 was also found to be unaffected by the foc mutation (data not shown).

Figure 4.

 RNA blot assays showed that accumulation of mature miR160 was decreased in foc plants.
(a) A representative RNA gel blot image (top) showing that accumulation of mature miR160 was considerably reduced in foc leaves and inflorescences. Ethidium bromide staining of 5S rRNA and tRNAs are shown as loading controls (bottom).
(b) A diagram showing that accumulation of miR160 is more significantly reduced in foc inflorescences than in foc leaves. Signals from wild-type leaves were set as 100%. Bars indicate standard deviation.
(c), (d) The accumulation of miR168 was not significantly changed in foc leaves or inflorescences.
Abbreviations: Inflor., inflorescence; wt, wild type.

To test whether the altered expression of MIR160a (FOC) and the accumulation of mature miR160 affected the miR160 target genes ARF10, -16, and -17, we carried out RT-PCR and quantitative real time RT-PCR experiments. Compared with wild-type expression levels, the RT-PCR results showed that expression of ARF10, -16, and -17 was increased in most examined foc organs, particularly in young seedlings, inflorescences and mature leaves (Figure S5b). In addition, the quantitative real time RT-PCR results demonstrated that ARF10, -16, and -17 were expressed at significantly higher levels in young foc seedlings and inflorescences, relative to their levels in the wild type (Figure S5c). Our results suggest that the impaired function of MIR160a results in increased expression of ARF10, -16, and -17 in foc plants.

We further examined expression of MIR160a and ARF10, -16, and -17 during embryonic development in wild-type and foc plants using in situ hybridization. In the wild type, the expression of MIR160a was uniformly detected using the MIR160a antisense probe in the embryo proper at the octant and dermatogen stages (Figure 5a,b), while the expression of MIR160a was stronger in whole embryos at the globular and triangular stages (Figure 5c,d). At the heart stage, MIR160a was primarily expressed in cotyledon primordia and in the vascular primordium (Figure 5e). At the torpedo stage, the MIR160a expression was decreased and was mostly in the cotyledons and hypocotyl epidermal cells (Figure 5f). In foc embryos, MIR160a was almost undetectable (Figure 5g–i).

Figure 5.

 Expression of ARF10, -16, and -17 is altered during embryo development in foc.
(a, b) Expression of MIR160a (160a in a–j) was detected in the embryo proper at the octant (a) and dermatogen (b) stages in wild type. (c, d) MIR160a was strongly expressed in whole embryos at the globular (c) and triangular (d) stages in the wild type. (e) MIR160a expression was primarily expressed in cotyledon primordia and in the vascular primordium at the heart stage in the wild type. (f) MIR160a was primarily expressed in cotyledons and epidermal cells in hypocotyl (arrows) at the early torpedo stage in the wild type. (g–i) MIR160a was almost undetectable at the octant (g), triangular (h), and late heart (i) stages in foc. (j) A wild-type embryo at the heart stage hybridized with the MIR160a sense probe, showing no signal. Similar results were observed using sense probes for ARF10, -16, and -17 genes (data not shown). (k, l) ARF10 was expressed at a higher level in foc cotyledons (l) relative to the expression in the wild type (k) at the late heart stage. (m, n) The expression of ARF16 was greatly increased in the vascular primordium in the foc embryo (n) relative to expression in the wild type at the globular stage (m). (o, p) ARF16 was weakly expressed in cotyledon primordia and the vascular primordium at the heart stage in the wild type (o), while ARF16 was strongly expressed in the whole embryo in foc (p). (q, r) ARF16 expression was greatly increased in cotyledons and procambium (arrows) in foc embryos (r) relative to the expression levels in the wild type at the late heart stage (q). (s, t) The foc embryo had higher ARF17 expression (t) than did the wild type at the octant stage (s). (u, v) ARF17 was mainly detected in cotyledons and the central embryo domain at the heart stage in the wild type (u), while in foc ARF17 was strongly expressed in the basal embryo domain (arrow, v). (w, x) ARF17 expression became more predominant in foc hypocotyls (arrow, x) than in the wild type (w) at the torpedo stage.

Expression levels and domains of ARF16 and -17 were markedly changed in foc plants. Although the expression pattern of ARF10 remained similar during early stages, after the late heart stage it was expressed at higher levels in foc cotyledons than in the wild type (Figure 5k,l). In the wild type, ARF16 was expressed in the vascular primordium at the globular stage (Figure 5m), in cotyledon primordia and the vascular primordium at the heart stage (Figure 5o), and in cotyledons and the procambium at the late heart stage (Figure 5q). However, in foc ARF16 expression was greatly increased at the globular stage (Figure 5n), and strongly expressed in embryos at the heart stage (Figure 5p). In addition, the expression of ARF16 was substantially increased at the late heart stage, particularly in the procambium (Figure 5r). Relative to expression in the wild type, ARF17 expression was much stronger at the octant stage in foc (Figure 5s,t). ARF17 was principally expressed in cotyledons and the central embryo domain at the heart stage in the wild type (Figure 5u). However, in foc ARF17 expression was not only increased in cotyledons, but was also strong in the basal embryo domain (Figure 5v). In the wild type, ARF17 was weakly expressed in the hypocotyl at the torpedo stage (Figure 5w); however, in foc it was strongly expressed in the hypocotyl, particularly in the root meristem (Figure 5x). No hybridization was detected when MIRI60a, ARF10, -16, and -17 sense probes were used (Figure 5; data not shown). Our results indicate that the expression levels and domains of ARF16 and -17 are considerably altered in foc embryos, which may cause pleiotropic phenotypes in embryo development.

foc is deficient in auxin responses

Auxin is known to play major roles in anther and embryo development (Jenik and Barton, 2005; Cecchetti et al., 2008). The miR160 target genes ARF10, -16, -17 are involved in auxin signaling (Mallory et al., 2005; Okushima et al., 2005; Wang et al., 2005; Guilfoyle and Hagen, 2007). Furthermore, expression of MIR160a and ARF10, -16, -17 was markedly affected in foc aerial organs, particularly in reproductive organs. Therefore, we examined the auxin response using DR5::GUS as a proxy for auxin levels (Ulmasov et al., 1997). In DR5::GUS plants, GUS activity was strong in cotyledons and moderate in the margin and veins of mature leaves (data not shown). However, in foc DR5::GUS plants, GUS expression was greatly reduced in those organs (data not shown).

In DR5::GUS plants, GUS was strongly expressed in anthers of young buds and carpels of pollinated flowers (Figure 6a,c,e). However, in foc DR5::GUS plants GUS expression was greatly reduced in anthers of young buds, and was not detectable in carpels of old flowers (Figure 6b,d,f). Furthermore, compared with DR5::GUS expression in the wild type, GUS staining was greatly reduced in foc embryos (Figure 6i,j). After treatment with 50 μm 1-naphthalene acetic acid (NAA), the DR5:GUS inflorescences showed greatly enhanced GUS expression, indicated by strong staining in both young buds and old flowers, particularly in sepals and carpels (Figure 6g). In contrast, no enhancement of GUS staining was observed in the foc background (Figure 6h). Our results indicate that foc is defective in auxin signal responses.

Figure 6.

foc is defective in the auxin signaling response.
(a, b) Inflorescences showing GUS expression in young buds and open flowers in wild type (a), but little or none in foc (b). (c, d) Stage 10 buds showing strong GUS expression in wild-type anthers (c), but much weaker expression in foc anthers (d). (e, f) Pollinated flowers showing GUS staining in wild-type carpels (e), but no staining in foc carpels (f). (g, h) Inflorescences after treatment with 50 μm NAA, showing enhanced and expanded GUS expression in young and old flowers, particularly in sepals and carpels in wild type (g), but no enhancement in foc (h). (i) Carpels showing GUS staining in wild-type embryos (left three), but less or no staining in foc embryos (right three). (j) The quantification of GUS activity (nm 4-methylumbelliferone (4-MU)/min/mg).
Abbreviations: wild type (WT), DR5:GUS plants; foc, foc DR5:GUS plants; Inf., inflorescence. Scale bars: 1 mm in (a, b), (g–i); 250 μm in (c–f).

Auxin positively regulates expression of MIR160a

Three potential AuxREs were found in the 3′ regulatory region of MIR160a by searching the PLANTCARE database (http://bioinformatics.psb.ugent.be/webtools/plantcare/html/; Figure 3a), leading to the hypothesis that expression of MIR160a might be regulated by auxin. To test this idea, we examined whether exogenous auxin could alter MIR160a expression through its 3′ regulatory region. Taking advantage of our ProFOC:GUS:3FOC and ProFOC:GUS transgenic plants, we first treated young seedlings at 5 days after germination (DAG) with 10 μm indole-3-acetic acid (IAA). In ProFOC:GUS:3FOC seedlings, IAA treatments caused a large increase in GUS expression in cotyledons after 2, 6 or 12 h (Figure 7a–d), whereas without IAA treatment, GUS expression levels were only slightly increased in cotyledons (Figure 7e–h). No significant changes in GUS expression were detected with IAA treatments in ProFOC:GUS seedlings (Figure 7i–p). We obtained similar results using seedlings with true leaves (Figure S7).

Figure 7.

 Exogenous auxin regulates expression of MIR160a through its 3′ regulatory region.
(a)–(d) ProFOC:GUS:3FOC seedlings treated with 10 μm of indole-3-acetic acid (IAA) in 1/2 × MS solution for 0 (a), 2 (b), 6 (c), and 12 (d) h showing greatly increased GUS activity in cotyledons.
(e–h) ProFOC:GUS:3FOC seedlings treated with 1/2 × MS solution for 0 (e), 2 (f), 6 (g), and 12 (h) h showing slightly increased GUS activity over time in cotyledons.
(i–p) ProFOC:GUS seedlings treated with 1/2 × MS solution (i–l) or 10 μm IAA (m–p) for 0 (i,m), 2 (j,n), 6 (k,o), and 12 (l,p) h, showing no changes in GUS activity.
Scale bar: 0.5 cm in all panels.

To further test whether IAA treatment increased MIR160a expression, we performed quantitative real time RT-PCR using young seedlings. Without IAA treatment, MIR160a expression was slightly elevated after 0.5 and 1 h, but greatly increased after 2 h (Figure 8a). However, with IAA treatments, the levels of MIR160a expression were significantly increased after 0.5, 1.0, and 2.0 h (Figure 8a). In wild-type seedlings, the expression levels of ARF10, -16, and -17 did not change greatly after IAA treatment (Figure 8c–e). However, in foc seedlings, the expression of ARF10 was greatly increased after 1.0 h of IAA treatment (Figure 8c), while the expression of ARF16 and -17 was significantly increased after 0.5, 1.0, and 2.0 h of IAA treatments (Figure 8d,e). Thus, the steady expression patterns of ARF10, -16, and -17 after IAA treatments of wild-type seedlings might be mainly down-regulated by the increased expression of MIR160a. In summary, our results indicate that auxin up-regulates expression of MIR160a in seedlings, possibly through the AuxREs in its 3′ regulatory region.

Figure 8.

 Quantitative real time RT-PCR results show that the expression of MIR160a in seedlings is up-regulated by auxin.
Transcripts of seedlings without indole-3-acetic acid (IAA) treatments were used as standards for normalization. *Indicates the difference is significant (P-values <0.01 or 0.05). Abbreviations: wt, wild type; h, hour.
(a) The expression of MIR160a was significantly increased after 0.5, 1.0, and 2.0 h of IAA treatment.
(b) Expression of IAA5 was used to confirm that the IAA treatments was effective.
(c–e) Expression of ARF10, -16, and -17 remained similar after IAA treatment in wild-type seedlings, while their expression was significantly increased in foc seedlings in most of time courses.

Discussion

Regulation of ARF10, -16, and -17 by MIR160a in embryo development

We characterized the function of MIR160a in regulating plant development, particularly embryogenesis, by analyzing the novel phenotypes of foc, the MIR160a loss-of-function mutant in Arabidopsis. Moreover, we demonstrated that the 3′ regulatory region of MIR160a is required for its expression pattern and regulation by auxin. The dynamic and differential distribution of auxin activity during embryo development suggests that both ARF activators and repressors are required for embryonic cell patterning in Arabidopsis (Willemsen and Scheres, 2004; Jenik and Barton, 2005; Bowman and Floyd, 2008). Among the available loss-of-function mutants (for at least 18 ARF genes), only single mutants of arf2, arf3, arf5, arf7, arf8, and arf19 have phenotypes in growth or development, indicating that most ARFs function redundantly (Sessions et al., 1997; Hardtke and Berleth, 1998; Harper et al., 2000; Ellis et al., 2005; Nagpal et al., 2005; Wilmoth et al., 2005; Goetz et al., 2006; Schruff et al., 2006; Okushima et al., 2007). So far, only ARF5 and -7 have been found to play direct roles in controlling embryogenesis. monopteros/arf5 is defective in establishing cell division patterns of the embryo proper and of the uppermost cell of the suspensor (Hardtke and Berleth, 1998; Hamann et al., 2002). ARF5 is expressed in the apical cell and its daughter cells until the 16-cell stage. ARF5 expression then becomes restricted to provascular cells. A mutation in NON-PHOTOTROPIC HYPOCOTYL4 (ARF7), the closest homolog of ARF5, strongly enhances the arf5 phenotype (Harper et al., 2000; Hardtke et al., 2004). The extensively overlapping expression patterns of ARF5 and -7 further indicate that they function redundantly during embryogenesis. Therefore, ARF5 and -7, serving as potential transcriptional activators, are essential for embryo development.

Our study showed that ARF10, -16, and -17, known to be regulated by MIR160a, play important roles in controlling embryogenesis. The MIR160 gene family consists of three members (MIR160a, -b, and -c). Ectopic expression of MIR160a, -b, or -c causes disorganized root caps and more lateral roots, whereas expression of a miR160 resistant-version of ARF16 (mARF16) results in reduced fertility and fewer lateral roots (Wang et al., 2005). Plants expressing an mARF17 version exhibited diverse abnormalities in vegetative and reproductive development (Mallory et al., 2005). The aberrant seedlings produced in mARF17 plants indicate that ARF17 may control early embryonic development. Analyses using similar approaches revealed that the negative regulation of ARF10 by miR160 is essential for plant development and ABA signaling (Liu et al., 2007). foc plants produce embryos with pleiotropic defects, including abnormal suspensors, no hypophysis, and irregular cell patterning in the central embryo domain. Abnormal cell differentiation in the boundary between the embryo proper and the uppermost suspensor cell may cause some suspensor cells to eventually incorporate into the embryo. It is also possible that mir160a embryos are defective in cell patterning in central embryo domains. Our in situ hybridization results demonstrate that MIR160a is expressed throughout early embryogenesis. Besides greatly increased levels of expression of ARF16 and -17, the expression domain of ARF17 is shifted in foc embryos. Our results show that auxin activity is decreased in foc, suggesting that ARF10, -16, and -17 may repress auxin signaling. At the triangular and heart stages, auxin activity is mainly detected in cotyledon apices as well as in the hypophysis and its derivatives. Therefore, ARF10, -16, and -17 might contribute as repressors in establishing and maintaining auxin signals in embryogenesis. In summary, our results provide insight into the molecular mechanism of embryo development involving ARFs, which are regulated by MIR160a.

Regulation of MIR160a expression by auxin

Transcriptional and post-transcriptional controls of MIR expression have not been extensively studied, although the spatial and temporal actions of miRNAs are critical for their function. The levels of mature miRNAs are determined by their transcription, processing, and incorporation into the RISC. Previous experiments show that the level of miR164 increased after treatment with 10 μm NAA (Guo et al., 2005), while changes in miR160, miR164, or miR167 were not detected after treatment with 10 μm IAA (Mallory et al., 2005). The level of ARF17 mRNA also remained unchanged (Mallory et al., 2005). In seedlings, exogenous IAA induces a slow increase in ARF16 expression, followed by a substantial increase after 5 h (Wang et al., 2005). Microarray analyses showed that expression of ARF10, -16, and -17 remain at similar levels before and after 1 and 3 h of IAA treatment (Goda et al., 2004, 2008; Paponov et al., 2008). Our quantitative RT-PCR results agree with previous findings when the seedlings were treated with 10 μm IAA. However, the overall expression of ARF10, -16, and -17 was substantially increased after treatments with 10 μm IAA in foc seedlings, suggesting that the increased expression of MIR160a induced by IAA treatment down-regulates expression of ARF10, -16, and -17. We also found that MIR160a expression in inflorescences is up-regulated by IAA (data not shown). Three potential AuxREs in the 3′ regulatory region of MIR160a might be required for regulating the expression of MIR160a by auxin. Taken together, the spatial and temporal expression of MIR160a may be sophisticatedly regulated by the 5′ promoter and 3′ regulatory region, as well as by the local concentration of auxin.

Functional analysis of MIR160 genes using the loss-of-function approach

Our study demonstrated a specific function for MIR160a using a loss-of-function approach. MIR160a (At2g39175), -b (At4g17788), and -c (At5g46845) may play different roles in regulating the miR160 target genes ARF10, -16, and -17. In the MIR164 family, single loss-of-function mutants in mir164a, -b, and -c showed subtle but different phenotypes in flower development or lateral root formation (Baker et al., 2005; Guo et al., 2005). However, the mir164abc triple mutant exhibited enhanced and novel phenotypes (Sieber et al., 2007). Although gene numbers vary with species, MIR160 is deeply conserved in the plant kingdom from moss (Physcomitrella patens) to higher plants (Axtell et al., 2007). Our phylogenetic analysis using sequences in the stem–loop regions of MIR160 from different plant species showed that Arabidopsis MIR160b and -c were very similar, but distant from MIR160a (data not shown). That the loss-of-function mutation of MIR160a alone results in severe phenotypes indicates that MIR160a may play a primary role in regulating plant development. Resistant versions of ARF10, -16, and -17 interfere with the recognition of miR160s derived from all three MIR160 genes (Mallory et al., 2005; Wang et al., 2005; Liu et al., 2007). In the loss-of-function foc mutant, the altered expression of ARF10, -16, and -17 was affected only by miR160 derived from MIR160a. In addition, expression of ARF10, -16, and -17 was affected simultaneously in foc, which resulted in severe and novel phenotypes. It will be interesting to study the specific function of MIR160b and -c using the loss-of-function approach, and to test the possibilities of specialization and redundancy among the three MIR160 genes in Arabidopsis.

Experimental procedures

Plant materials and growth conditions

The Arabidopsis thaliana foc mutant is in the Landsberg erecta background. foc was also backcrossed to the Columbia (Col-0) ecotype. The DR5::GUS line was kindly provided by Dr Thomas Guilfoyle (University of Missouri, Columbia, Missouri). Plants were grown on Metro-Mix 360 soil at 22°C under 16 h of light/8 h of dark or 8 h of light/16 h of dark.

Identification of FOC

To identify FOC, TAIL-PCR was performed using foc genomic DNA (Table S1) (Liu et al., 1995; Zhao et al., 2002). The full-length cDNA of MIRNA160a was obtained by RACE-PCR. The RACE-PCR products were cloned into the pGEM-T Easy vector (Promega, http://www.promega.com/) and sequenced.

Phenotypic analyses and microscopy

Micrographs were taken with an Olympus DP70 digital camera through stereo (Olympus SZX-RFL) and compound (Olympus BX51) microscopes (http://www.olympus.com/). Embryo development was analyzed using a whole-mount squash method essentially as previously described (Schneitz et al., 1995; Blilou et al., 2005; Zhou et al., 2009). Siliques between 1 and 7 DAP were fixed overnight in FAA [50% (w/v) ethanol, 5% (w/v) acetic acid, and 3.7% (w/v) formaldehyde] at 4°C. After rehydration in an ethanol series, siliques were transferred onto a slide and ovules were dissected in Hoyer’s buffer (70% chloral hydrate and 4% glycerol). Samples for scanning electron microscopy (SEM) were prepared and examined as previously described using a Hitachi S-570 scanning electron microscope (http://www.hitachi.com/) (Bowman et al., 1989; Zhao et al., 2001).

Vector construction and plant transformation

All DNA and cDNA fragments were PCR-amplified by Phusion High-fidelity DNA polymerase (New England Biolabs, http://www.neb.com/) (Table S1), then cloned into the pENTR™ TOPO® vector (Invitrogen, http://www.invitrogen.com/), and finally introduced into Gateway binary vectors using the Gateway® LR recombinase II enzyme mix (Invitrogen). Plant transformation was performed using Agrobacterium tumefaciens strain GV3101 (Clough and Bent, 1998). Transformants were screened on 1/2 × MS agar plates containing kanamycin (50 mg/L) and hygromycin (50 mg/L).

For complementation experiments, two constructs were generated by cloning a 4921-bp genomic fragment of FOC (At2g39175, ETB) and a 3820-bp genomic fragment of At2g39170 (ETA). To examine regulation of FOC expression, three constructs were generated: ProFOC:GUS:3FOC, ProFOC:GUS:Δ3FOC, and ProFOC:GUS. Based on the sequence of the FOC transcript and the Ds location, a 1502-bp fragment was used as the FOC promoter, while 2488-bp and 1456-bp fragments were used as the FOC 3′ regions for ProFOC:GUS:3FOC and ProFOC:GUS:Δ3′FOC constructs, respectively. The FOC promoter was introduced into the Gateway pGWB3 binary vector with the GUS gene, then the FOC 3′ fragments were cloned into a SacI site downstream of the GUS gene.

Hormone Treatments

Five-day-old ProFOC:GUS:3FOC and ProFOC:GUS T2 transgenic seedlings were transferred onto Whatman filter paper soaked with 1/2 × MS liquid media containing 10 μm IAA and 0.1% ethanol in a Petri dish. Mock treatments were performed using the same solution but without IAA. Seedlings were collected at 0, 2, 6, and 12 h for GUS staining. For quantitative real time RT-PCR, 5-day-old wild-type and foc seedlings were treated similarly, and then collected at 0, 0.5, 1, and 2 h for total RNA extraction. For NAA treatment, DR5:GUS and foc DR5:GUS inflorescences were incubated with 50 μm NAA in 0.01% Silwet L-77 and 0.1% ethanol in the morning, and collected for GUS staining after 6 h. Mock treatments were performed with the same solution lacking NAA.

GUS staining and GUS activity assay

Histochemical GUS staining (Willemsen et al., 1998) and GUS activity assays (Jefferson et al., 1987) were performed essentially as described previously. For the GUS activity assay, fluorescence was determined using a Synergy HT multi-mode microplate reader at 360 nm (excitation) and 460 nm (emission) (BioTek, http://www.biotek.com). Protein concentration was measured using the Bradford method and GUS activity was calculated as nmol 4-methylumbelliferone (4-MU)/min/mg protein.

Small RNA northern blots

Small RNA Northern blot assays were carried out essentially as described (Xie, 2010). Briefly, total RNA extracts (15 μg) were resolved in a 17% polyacrylamide gel containing 7 m urea, transferred to Nytran SuPerCharge nylon membrane (Whatman, http://www.whatman.com/), and then probed with 32P-labeled oligonucleotides with a sequence complementary to the target small RNAs. To enhance detection sensitivity, locked nucleic acid (LNA)-modified (Valoczi et al., 2004) oligonucleotides (Exiqon, http://www.exiqon.com/) specific for miR160 (a gift from James Carrington, Oregon State University, Corvallis, OR, USA), miR168, and miR171, respectively, were used as probes. Two RNA oligos of 21 and 24 nucleotides were used as size markers. Before exposure of the blots to X-ray films, radioactive signals from blots were captured in a phosphorimager (Storm, Amersham Biosciences, http://www1.gelifesciences.com) and analyzed using ImageQuant software.

RNA in situ hybridization

RNA in situ hybridizations were performed on wild-type and foc silique sections essentially as described (Long and Barton, 1998; Zhao et al., 2002). The 5′ region upstream stem loop of MIR160a, as well as specific cDNAs of ARF10, -16 and -17 were PCR-amplified (Table S1) and then cloned into the pGEM®-T Easy Vector (Promega). Antisense and sense probes were synthesized using the SP6/T7 DIG RNA Labeling Kit (Roche, http://www.roche.com/).

RT-PCR and quantitative real time RT-PCR

Total RNAs were extracted from different plant tissues using the RNeasy Plant Mini Kit (Qiagen, http://www.qiagen.com/). The RNA concentrations were measured with a NanoDrop ND-1000 spectrophotometer (Thermo Scientific, http://www.nanodrop.com). Reverse transcription reactions were carried out using a QuantiTect Reverse Transcription Kit (Qiagen). The PCR and quantitative real time PCR (DNA Engine Opticon 2 system) were performed using primers as noted for specific experiments (Table S1). ACTIN2 was used as an internal control. Fast SYBR Green PCR Master Mix (Applied Biosystems, http://www3.appliedbiosystems.com/) was used for quantitative real-time PCRs. The quantitative RT-PCR results were analyzed as described previously (Pfaffl et al., 2002). Three independent experiments were carried out. Each value indicates the average and a standard error.

Acknowledgements

We thank S. Forst, D. Heathcote, M. Rawluk, C. Seubert, C. Starrett, and Y. Zhao for technical assistance and critical comments on this manuscript, S. McCormick for editing, the ABRC for providing Ds lines, T. Schuck for plant care, and T. Nakagawa for providing Gateway binary vectors. We also thank J. Carrington for sharing the miR160 LNA probe. This work was supported by a grant from the University of Wisconsin-Milwaukee Research Growth Initiative (RGI) Program (to DZ), an NSF grant IOS-0721192 (to DZ), and the Shaw Scientist Award from the Greater Milwaukee Foundation (to DZ).

Ancillary