Abstract
Auxin plays pleiotropic roles in plant development via gene regulation upon perception by the receptors TRANSPORT INHIBITOR RESPONSE 1/AUXIN SIGNALING F-BOX (TIR1/AFBs). This nuclear auxin signaling (NAS) originated in the common ancestor of land plants. Although complete loss of TIR1/AFBs causes embryonic lethality in Arabidopsis thaliana, it is unclear whether the requirement for TIR1-mediated auxin perception in cell viability can be generalized. The model liverwort Marchanita polymorpha has a minimal NAS system with only a single TIR1/AFB, MpTIR1. Here we show by genetic, biochemical, and transcriptomic analyses that MpTIR1 functions as an evolutionarily conserved auxin receptor. Null mutants and conditionally knocked-out mutants of MpTIR1 were viable but incapable of forming any organs and grew as cell masses. Principal component analysis performed using transcriptomes at various developmental stages indicated that MpTIR1 is involved in the developmental transition from spores to organized thalli, during which apical notches containing stem cells are established. In Mptir1 cells, stem-cell- and differentiation-related genes were up- and down-regulated, respectively. Our findings suggest that, in M. polymorpha, NAS is dispensable for cell division but essential for three-dimensional body plans by establishing pluripotent stem cells for organogenesis, a derived trait of land plants.
Introduction
The common ancestors of land plants diverged from algal sisters about 4.5 million years ago and acquired three-dimensional (3D) bodies with parenchymal cells which contributed to their survival in the terrestrial environment (Delwiche and Cooper, 2015). The common ancestor also established several intercellular communication mechanisms mediated by plant hormones (Bowman et al., 2019). Among them, auxin is proposed to act as a morphogen, whose localization and gradient are critical for developmental aspects such as embryonic patterning (Verma et al., 2021; Liao et al., 2015), organ orientation (Galvan-Ampudia et al., 2020; Guan and Jiao, 2020), and gravitropism (Herud-Sikimic et al., 2021; Su et al., 2017) in plants.
Auxin signal is mainly transduced via the nuclear auxin signaling (NAS) pathway, major players of which are: TRANSPORT INHIBITOR RESPONSE 1/AUXIN SIGNALING F-BOX (TIR1/AFB) subunits of Skp1-Cullin-F-box (SCF)-type E3 ubiquitin ligase complex, which act as auxin receptors, AUXIN/INDOLE-3-ACETIC ACID (AUX/IAA) proteins as transcriptional repressors, and AUXIN RESPONSE FACTOR (ARF) proteins as transcription factors. In the NAS pathway, auxin facilitates the interaction between the leucine-rich-repeat (LRR) domain of TIR1/AFBs and domain II (DII) of AUX/IAAs by filling a hydrophobic cavity in the LRR domain, which in turn promotes the ubiquitination of AUX/IAAs (Gray et al., 1999; 2001; Dharmasiri et al., 2005; Kepinski and Leyser, 2005; Tan et al., 2007). In the absence of auxin, AUX/IAAs interact with ARFs and repress transcription by recruiting TOPLESS (TPL) co- repressors to the target loci of ARF (Kim et al., 1997; Ulmasov et al., 1997; 1999; Tiwari et al., 2003). In other words, in the presence of auxin, TIR1/AFBs promote degradation of AUX/IAA with its DII domain acting as a degron, which in turn enables ARFs to exert transcriptional regulation.
Comprehensive phylogenetic analyses of TIR1/AFBs, AUX/IAAs, and ARFs among land plants and green algae have shown that the NAS pathway was established in the common ancestor of land plants through the acquisition of the TIR1/AFB-AUX/IAA co-receptor mechanism in order to regulate transcriptional regulation by ARFs (Bowman et al., 2017; Flores- Sandoval et al., 2018; Mutte et al., 2018). The loss of all six TIR1/AFB homologs in Arabidopsis thaliana mutant lines showed disturbed division patterns and delayed cell divisions which led to inhibition of embryogenesis (Prigge et al., 2020). Transmission of tir1/afb sextuple mutations through gametes was unaffected, indicating that TIR1/AFBs do not play an essential role in the development of gametophyte, where only a few cell divisions occur (Prigge et al., 2020).
The moss Physcomitrium (Physcomitrella) patens and the liverwort Marchantia polymorpha have been studied as models for gametophyte-dominant species (Rensing et al., 2020; Kohchi et al., 2021). Auxin-dependent interaction between TIR1/AFB and AUX/IAA homologs was demonstrated in P. patens (Prigge et al., 2010). Disturbed auxin perception suppresses the differentiation of caulonema in P. patens (Prigge et al. 2010), whereas in M. polymorpha it leads to the formation of undifferentiated cell masses (Kato et al. 2015). In these studies, auxin perception was inhibited either by knockdown of TIR1/AFBs (Prigge et al. 2010) or by the expression of dominant-negative AUX/IAAs with mutations in DII (Kato et al., 2015), where leaky signal transduction cannot be avoided. For this reason, it is difficult to assess the role of auxin in terms of gametophytic cell survival through the studies described above. Instead, knocking out the TIR1/AFBs is expected to shut down NAS altogether. We chose M. polymorpha for our study, as it encodes a minimal set of the NAS components, including the sole TIR1/AFB and AUX/IAA homologs, MpTIR1 and MpIAA respectively (Bowman et al., 2017; Flores-Sandoval et al., 2015; Kato et al., 2015).
Germinated M. polymorpha spores divide vigorously to produce unorganized tissues, sporelings, and once single-celled apical stem cells are established, they initiate 3D morphogenesis into thalli which differentiate specialized organs such as the gemma cup (Kohchi et al., 2021; Shimamura 2016). Within the dorsal organ gemma cup, multicellular asexual reproductive propagules, gemmae, are produced from single initial cells, which involves apical stem cell formation (Kato et al., 2020). Previous studies have shown that auxin acts as a mobile signal in the thalli (Gaal et al., 1982; Solly et al., 2017) and that NAS regulates the establishment of body axis in gemma development (Kato et al., 2017; Kato et al., 2018). Here, to understand necessity and roles of NAS in plant development, we analyzed molecular functions of MpTIR1 as an auxin receptor and performed knockout-study of MpTIR1 in the liverwort M. polymorpha.
Results
MpTIR1 positively regulates auxin response
To determine whether MpTIR1 is involved in auxin response, wild-type (WT), MpTIR1- overexpressing plants under the MpEF1A promoter (Figure 1A) as well as Mptir1 mutants were treated exogenously with varying concentrations of natural (indole-3-acetic acid (IAA)) and synthetic (1-naphthaleneacetic acid (NAA) and 2,4-dichlorophenoxyacetic acid (2,4-D)) auxins. WT plants showed ectopic rhizoid formation and arrested thallus development in the presence of high levels of auxin (Figure 1B, C; Supplemental Figure 1A–D). Consistent with the assumption that MpTIR1 is an auxin receptor, MpTIR1-overexpressing plants exhibited hypersensitivity to auxins; the responses were observed at lower concentrations (Figure 1B, C; Supplemental Figure 1).
Mptir1-1ko mutants grew as cell masses with no obvious organ development (Supplemental Figure2; see “MpTIR1 is critical for organ development but dispensable for cell survival” section for details); exogenous auxins neither caused morphological changes nor enhanced rhizoid development in Mptir1-1ko cells, although NAA and 2,4-D, but not IAA, slightly affected the growth increment (Figure 1D; Supplemental Figure 2D, E). Mptir1-1komutants harboring a transgene of an MpTIR1 genomic fragment showed rescued morphology with normal auxin responsiveness, as shown by ectopic rhizoid formation and arrested growth in the presence of high concentrations of auxin (Figure 1D). This was also the case with Mptir1-1ko mutants harboring an A. thaliana TIR1 (AtTIR1) transgene driven by the MpTIR1 promoter (Figure 1D). These results suggest that MpTIR1 positively regulates auxin response and that its function is comparable with that of AtTIR1.
MpTIR1 acts as an auxin receptor
Interaction between TIR1/AFBs and AUX/IAAs, and the subsequent AUX/IAA degradation are essential for NAS (Gray et al., 1999; 2001; Dharmasiri et al., 2005; Kepinski and Leyser, 2005). To investigate whether MpTIR1 requires auxin for it to interact with MpIAA, we performed an in vitro pull-down assay. We used the Escherichia coli-expressed fusion protein of glutathione S-transferase (GST) with MpIAA truncated from the 627th amino acid till the C-terminus including DII (GST-MpIAA(627C)) as the bait for M. polymorpha-expressed MpTIR1-3xFLAG. The MpTIR1-3xFLAG proteins showed interaction with MpIAA only when auxin was added to the mixture (Figure 2A). IAA facilitated the MpTIR1-MpIAA interaction in a dose-dependent manner (Figure 2A). NAA and 2,4-D also mediated MpTIR1-MpIAA interaction to the same or a slightly weaker degree as IAA (Figure 2A). GST-MpIAAmutDII(627C), having a mutated sequence in its DII that had been shown to inhibit auxin signaling in M. polymorpha (Kato et al., 2015), did not interact with MpTIR1 regardless of the presence of auxin (Figure 2B). These data suggest that MpTIR1 interacts with the DII of MpIAA in an auxin-dependent manner.
We also investigated the involvement of MpTIR1 in the degradation of MpIAA in vivo using conditional knockout (CKO) mutants generated by transforming Mptir1-1ko cells with a vector (Nishihama et al. 2016) that, under normal conditions, expressed a floxed MpTIR1 coding region under MpEF1A promoter for complementation. After induction by heat shock and dexamethasone (DEX) treatment, Cre recombinase excised the floxed MpTIR1 to express nuclear localization signal (NLS)-fused Citrine for labeling MpTIR1-KO cells (Supplemental Figure 3A–C). Hereafter, we refer to this as Mptir1-1CKO>CitN plants. Then, accumulation of a DII degron peptide probed by mTurquoise2-NLS was assessed in Mptir1-1CKO>CitN plants. If MpTIR1 led its target to degradation, accumulation of DII-mTurquoise2-NLS would be facilitated in the absence of MpTIR1. Higher mTurquoise2-fluorescence signals were detected under MpTIR1 KO-induced conditions than uninduced conditions (Figure 2C, D). By contrast, the mutated DII peptide (see above) failed to enhance signals of mTurquoise2 fluorescence under MpTIR1 KO-induced conditions (Figure 2C, D). These results suggest that MpTIR1 promotes degradation of MpIAA in a DII-dependent manner in vivo, which is a direct indication that MpTIR1 acts as an auxin receptor.
MpTIR1 is essential for auxin-mediated transcriptional regulation
In order to analyze MpTIR1-mediated auxin-dependent transcriptional regulation, WT (5-day- old sporelings) and Mptir1-1kocells were treated with or without 10 μM IAA as well as 10 μM NAA for 4 h and then subjected to RNA-seq analyses to compare transcriptome profiles. Differentially expressed genes (DEGs) were examined in comparison to mock-treated (without auxin) samples for each experiment (Supplemental Data Set 1).
In the WT samples, IAA treatment caused up- and down-regulation of 83 and 76 genes, respectively (Figure 3A). NAA treatment yielded >15-fold larger numbers of DEGs than did IAA treatment (Figure 3A). Nevertheless, the majority of DEGs resulting due to IAA treatment were also differentially expressed in response to NAA with a high correlation (Supplemental Figure 4A, B). These results suggest that although NAA acts more strongly than IAA, both the auxins essentially induce the same transcriptional changes.
In Mptir1-1ko cells, consistent with MpTIR1 being an auxin receptor, DEGs detected upon IAA treatment were almost nil (up in 1 gene, and down in 0 genes; Figure 3A). DEGs upon NAA treatment (up in 112 genes, and down in 253 genes) were also dramatically fewer than that with WT (Figure 3A). Decreased- or non-responsiveness of Mptir1-1ko cells to auxin was further confirmed by real-time PCR on known auxin-responsive genes, MpC2HDZ and MpWIP (Figure 3B; Kato et al., 2017; Kato et al., 2020; Mutte et al., 2018). These results further confirm the role of MpTIR1 in auxin-mediated transcriptional regulation.
MpTIR1 is critical for organ development but dispensable for cell survival
To understand the role of NAS in 3D morphogenesis, MpTIR1 was knocked-out in WT sporelings. The resultant five independent Mptir1-1ko mutants failed to develop thalli and slowly proliferated as cell masses (Figure 4A; Supplemental Figures 2A, B, and 5). We could not observe any phenotypic differences between male and female mutants (Supplemental Figure 2C). The mutants displayed unorderly cell division in the cell masses and failed to differentiate into multicellular organs and rhizoids (Figure 4B). To verify these phenotypes, we also genome- edited MpTIR1 in WT sporelings (Supplemental Figure 6). Four independent mutants (Mptir1- 2ld, Mptir1-3ld, Mptir1-4ld, and Mptir1-5ld), completely lacking the MpTIR1 locus, displayed similar phenotypes to those of the Mptir1-1ko mutants (Supplemental Figure 7). The developmental defects of Mptir1-1ko mutants, besides the auxin response defects described above, were rescued by MpTIR1 or AtTIR1 transgenes driven by the MpTIR1 promoter (Figure 1D), confirming that the impaired organogenesis of Mptir1-1komutants was due to loss of evolutionarily conserved functions of MpTIR1. For a more detailed understanding, we compared the cell compositions of Mptir1-1kocell clumps with that of proliferating WT sporelings. Ten- day-old sporelings had meristematic region(s) consisting of small undifferentiated cells and other regions containing large vacuolated cells (Figure 4C, D). Mptir1-1ko cell masses were not composed of a uniform cell type but a mixture of small cells at outer regions and large vacuolated cells at inner regions (Figure 4, E, F). Taken together, these results suggest that MpTIR1 is dispensable for cell survival but is essential for orderly organogenesis and development.
MpTIR1 is critical for proper patterning and organ differentiation
In order to determine the physiological roles of MpTIR1 in other developmental stages, Mptir1- 1CKO>CitNplants (see “MpTIR1 acts as an auxin receptor” section) and Mptir1-1CKO>tdTNplants were generated. In the latter plants, Mptir1-1ko cells could be visualized using tdTomato-NLS after excision of the floxed MpTIR1 genomic fragment (Figure 5A; Sugano et al. 2018; Suzuki et al., 2020). Mptir1CKO>tdTNgemmae essentially showed proper patterning with a trace of stalk at the bottom and two apical notches at its lateral tips (Figure 5B). When we excised the floxed MpTIR1 transgene during gemma development, the gemma patterning appeared disordered and failed to establish ellipse-shaped bodies and notches (Figure 5C). We then examined the loss-of- MpTIR1 phenotypes after germination of gemmae. Mptir1-1CKO>CitN and Mptir1-1CKO>tdTN gemmae grew up into thalli under mock conditions, whereas the excision of the floxed MpTIR1 transgene in mature gemmae or 1-day-old gemmalings resulted in impaired thallus development and caused cell mass formation (Figure 5D–H, Supplemental Figure 3B, C). These results suggest that MpTIR1 is critical for proper patterning and organ differentiation in M. polymorpha.
Auxin hypo-responsiveness, a common feature between Mptir1-1kocells and sporelings
In order to clarify the differentiation status of Mptir1-1ko cells, we assessed the gene expression patterns using principal component analysis (PCA) of Mptir1-1ko cells, sporelings, and available organ-specific transcriptome data (Bowman et al., 2017; Frank and Scanlon, 2015; Higo et al., 2016; Yasui et al., 2019). In the two-dimensional plot of the first and second principal components (PC1 and PC2, respectively), biological replicates from the same sources were clustered (Figure 6A). WT tissues were grouped into an approximate order of developmental stage (Figure 6A). Spores and sporelings were grouped age-wise along PC2, and these developmentally early tissues were grouped separately from the thalli, reproductive organs, and sporophytes along PC1 (Figure 6A). Vegetative thalli and gemma cups were comparable to older sporelings in PC2, while reproductive organs and sporophytes were grouped along PC2 (Figure 6A). Mptir1-1ko cells were grouped separately from all of these organs; Mptir1-1ko cells were between older sporelings and thalli in PC1 but were similar values to them in PC2 (Figure 6A). These data indicate that Mptir1-1ko mutants failed to differentiate from sporelings into mature thalli due to defects in auxin perception.
We performed another PCA to confirm this using a subset including Mptir1-1ko cells, 5- day-old sporelings, and thalli (Figure 6B). Evidently, PC1 scores were high in Mptir1-1ko and sporelings, but low in thalli, resulting in a large separation along PC1 (Figure 6B). In order to find whether auxin contributed to this particular separation, we examined the relationships between auxin responsiveness and factor loadings for PC1 and PC2. Genes that were downregulated upon IAA treatment were dense in positive factor loading of PC1 while those upregulated were dense in the negative factor loading of the PC1 (Figure 6C). These relationships suggest the involvement of IAA in the regulation of gene expression which contributed to high PC1 values in the case of sporelings and Mptir1-1ko cells, and low PC1 values in the case of thalli (Figure 6B). In contrast, we did not find any correlation between IAA responsiveness and factor loading of the PC2 (Figure 6D). Similar relationships were observed with respect to NAA treatment (Supplemental Figure 8A, B). These observations suggest that auxin hypo-responsiveness is a characteristic feature of Mptir1-1ko cells and sporelings.
MpTIR1 promotes gene expression for organ differentiation
In order to further characterize the differentiation features of Mptir1-1ko cells, we performed transcriptomic pairwise comparisons of Mptir1-1ko cells with sporelings and thalli (Supplemental Data Set 2). With respect to regulatory genes (Supplemental Data Set 3), Mptir1-1ko cells showed significantly lower expression of known transcription factors involved in rhizoid and organ differentiation, such as LOTUS JAPONICUS ROOTHAIRLESS1-LIKE homolog (MpLRL; Breuninger et al. 2016), ROOT HAIR DEFECTIVE SIX-LIKE1 homolog (MpRSL; Proust et al. 2016), and WIP homolog (MpWIP; Jones and Dolan, 2017), when compared with that of sporelings and thalli (Figure 6E, F). Conversely, Mptir1-1kocells highly expressed LOW-AUXIN RESPONSIVE (MpLAXR), which triggers cellular reprogramming to generate undifferentiated cells (Figure 6E, F; Ishida et al., 2022). Auxin appears to regulate an MpTIR1-dependent expression of these transcription factors (Supplemental Figure 8C–F). Our analyses indicate that MpTIR1 plays a critical role in organogenesis by promoting gene expression for differentiation.
Discussion
MpTIR1 is an evolutionarily conserved auxin receptor
In this study, we observed a positive correlation between MpTIR1 expression levels and auxin responsiveness (Figure 1; Supplemental Figures 1; 2D, E). We observed an auxin-dependent direct interaction of MpTIR1 with DII of MpIAA (Figure 2A, B) which promoted the degradation of DII-tagged proteins (Figure 2C, D). Mptir1-1ko mutants were rescued by AtTIR1 (Figure 1D), which further established the role of MpTIR1 as auxin receptor. In M. polymorpha, MpTIR1 transmits auxin signal by capturing and leading MpIAA towards degradation, which allows transcriptional activation as well as competitive repression mediated by the sole class-A and class-B ARFs, MpARF1 and MpARF2, respectively (Flores-Sandoval et al. 2015; Kato et al., 2015; Kato et al. 2017; Kato et al. 2020).
Mptir1-1ko cells did not show any physiological and transcriptional responses to IAA (Figures 1D; 3; Supplemental Figure 2D, E). This could be due to hyper-accumulation of MpIAA regardless of auxin, which indicates that the transcription of MpARF1/2-target genes are strongly repressed in Mptir1 KO cells. Such a constitutively repressive status seems to be different from the absence of A-ARFs, because Mparf1 KO mutants showed much milder developmental defects, although the Mparf1 mutants were also insensitive to auxin (Kato et al., 2017). The latter situation could be explained by impaired recruitment of MpIAA to MpARF1/2- target loci, as MpARF2 interacts with MpIAA less efficiently than does MpARF1, and thus neither transcriptional activation nor strong repression occurs regardless of auxin (Kato et al., 2017; Kato et al., 2018; Kato et al., 2020). Auxin insensitivity was also caused by loss of all AUX/IAAs as demonstrated in P. patens (Lavy et al. 2016), where auxin responses are constitutively saturated in contrast to Mptir1 KO cells.
In M. polymorpha, the synthetic auxins NAA and 2,4-D inhibit the growth much more severely than does IAA (Ishizaki et al., 2012). In this study, M. polymorpha exhibited a much higher number of DEGs with NAA treatment than that with IAA treatment (Figure 3A). However, the amplitude of gene activation or repression by IAA and NAA was comparable with respect to the DEGs regulated by both IAA and NAA (Supplemental Figure 4B). Pull-down assay demonstrated that both IAA and NAA facilitated the interaction between MpTIR1 and MpIAA (Figure 2A). These results suggest that although IAA and NAA both promote MpIAA degradation and enhance transcriptional regulation by MpARF1, we cannot conclude that either of these hormones are functionally stronger. NAA possibly induces more gene expression changes than IAA due to differences in cellular processes such as uptake by passive diffusion (Delbarre et al. 1996) or due to non-specific side effects (Paponov et al. 2019).
MpTIR1-mediated NAS is essential for 3D morphogenesis but not for cell survival
The observations made in this study where KO mutants of MpTIR1 were viable (Figure 4A–C, Supplemental Figures 5, 7), are in line with a previous report indicating tir1/afb sextuple mutation did not affect gametophyte viability in A. thaliana (Prigge et al., 2020). These results seem to indicate that gametophyte viability is not related to TIR1/AFB-mediated NAS.
In M. polymorpha, as is the case for vascular plants, IAA is biosynthesized from tryptophan in a two-step reaction catalyzed first by TRYPTOPHAN AMINOTRANSFERASE OF ARABIDOPSIS (TAA) and then YUCCA homologs (Eklund et al., 2015). Low auxin levels due to KO of the sole TAA homolog MpTAA in M. polymorpha or overexpression of IAA conjugation enzyme resulted in cell masses in sporelings (Eklund et al., 2015; Flores-Sandoval et al., 2015). Besides auxin biosynthesis, block of its signaling by dominant suppression of MpARF-mediated gene regulation due to expression of co-repressor-fused MpARFs (Flores- Sandoval et al., 2015) or induction of stabilized MpIAA (Kato et al., 2015) was shown to cause cell mass phenotype. These phenomena are most likely reflected by the cell mass phenotype resulting from MpTIR1 KO (Figures 4, 5, Supplemental Figures 3, 7). NAS is proposed to control body axis formation in gemma development as KO of MpARF1 disrupts patternings in this process (Kato et al., 2017). In this study, CKO of MpTIR1 in immature gemmae caused structures without notches (Figure 5B, C), probably due to impaired axis formation. These are reminiscent of AtTIR1/AFBs and other downstream elements that control proper patterning in embryogenesis by regulating division orientation in A. thaliana (Prigge et al., 2020; Yoshida et al., 2015). These results support that MpTIR1-mediated NAS is essential for 3D body plan establishment and organogenesis during early development.
As cell migration is restricted by cell walls, cell supply from stem cells towards appropriate directions is essential for orderly plant development. The control of cell supply is attributed to the regular division of apical cells in bryophytes (Harrison, 2017; Moody, 2020). In M. polymorpha, wedge-shaped apical cells, which produce daughter cells toward the dorsal, ventral, and lateral sides, are established during sporelings and gemma development (Shimamura, 2016).
In the cell mass of Mptir1 mutants, establishment of properly shaped apical cells and/or control of division planes may be impaired (Figure 4A, B; Supplemental Figure 7). CKO of MpTIR1 in gemmalings where the apical cells are already established also resulted in cell masses (Figure 5D, E, Supplemental Figure 3B, C), suggesting the disruption of apical cell functions in the absence of MpTIR1. PCA based on transcriptomes revealed clear separation of Mptir1-1ko cells and sporelings from thalli (Figure 6B) and positive correlation of auxin responsive genes to this separation (Figure 6C, D; Supplemental Figure 8A, B). Mptir1-1ko cells showed lower expression of the differentiation-related transcription factors, MpLRL, MpRSL, and MpWIP (Breuninger et al., 2016; Jones and Dolan, 2017; Proust et al., 2016), than sporelings and thalli (Fig6E, F). Judging from this, even though swelled cells were observed (Figure 4F, G), Mptir1- 1ko cells were not assumed to be properly differentiated. A relatively mild knockdown of MpTAA results in thalli with impaired organogenesis (Eklund et al., 2015), supporting that auxin response is required for organ differentiation. Mptir1-1ko cells showed greater expression of a dedifferentiation-related transcription factor, MpLAXR (Ishida et al., 2022), when compared with that of sporelings and thalli (Figure 6E, F). Although molecular functions have not yet been characterized in M. polymorpha, MpR2R3-MYB20, a paralogous gene to GEMMA CUP- ASSOCIATED MYB1 whose overexpression causes undifferentiated cell clumps (Yasui et al., 2019), and MpNAC1, an orthologous gene to CUP-SHAPED COTYLEDONs that act as shoot apical meristem-related boundary genes in angiosperms (Verma et al. 2021), were up-regulated in Mptir1-1ko cells. In the moss P. patens, auxin signaling is proposed to be low in undifferentiated tissues while it is high in differentiating tissues (Thelander et al., 2019), highlighting the conserved roles of auxin to regulate proper differentiation in gametophyte- dominant species. It could be said that the lack of auxin responsiveness of mutants results in the disruption of apical cell functions, which in turn affects organ differentiation, eventually leading to the formation of undifferentiated cell masses in M. polymorpha.
In conclusion, MpTIR1-mediated NAS contributes to establishing 3D body axes through the regulation of apical stem cell functions, including division plane determination and cell differentiation. The findings of this study of MpTIR1, in combination with previous reports of other NAS components in M. polymorpha, would help us understand organogenesis through temporal and spatial regulation of NAS in land plants.
Materials and Methods
Plant materials and growth conditions
Male accession Takaragaike-1 (Tak-1), female accession Takaragaike-2 (Tak-2), and a female accession of their third backcross generation, BC3-38, were used as wild-type (WT) M. polymorpha subsp. ruderalis. Tak-1 and BC3-38 were used for phenotypic analysis. Tak-1, BC3- 38, and BC4 spores which were obtained by crossing Tak-1 and BC3-38 were used to generate MpTIR1-overexpressing (proMpEF1A:MpTIR1-3xFLAG) plants. F1 spores, which were obtained by crossing Tak-1 and Tak-2, were used to generate Mptir1-1ko and Mptir1ld mutants.
M. polymorpha was cultured on half-strength Gamborg’s B5 medium (Gamborg et al., 1968) containing 1% agar under 50–60 μmol photons m-2 s-1 continuous white light at 22°C unless otherwise defined. For crossing, M. polymorpha was grown on soil under far-red irradiated conditions to induce gametangiophore formation as described previously (Chiyoda et al., 2008).
Preparation of plasmid constructs
Oligos used in this study are listed in Supplemental Table 1.
For proMpEF1A:MpTIR1-3xFLAG plants
A coding sequence (CDS) of MpTIR1 without stop codon was amplified from pENTR_MpTIR1 using the primer pair, MpTIR1_entry/MpTIR1_nonstop, and cloned into pENTR/D-TOPO vector (Thermo Fisher Scientific, Massachusetts, U.S.A.) to generate pENTR_MpTIR1_nonstop. The MpTIR1 CDS was then transferred into pMpGWB110 (Ishizaki et al., 2015) by using LR Clonase II (Thermo Fisher Scientific) to generate pMpGWB110_MpTIR1, which was then used for MpTIR1-overexpression experiments and pull- down assays.
For pull-down assay
MpIAA or MpIAAmutDII CDS spanning from the 627th codon till the stop codon was amplified from vectors containing the respective sequences (Kato et al., 2015) using the primer pair, EcoRI-MpIAA_DII/MpIAA-NotI. The PCR products and pGEX6P-1 vector were digested with EcoRI (Takara Bio, Shiga, Japan) and NotI (Takara Bio) and then ligated to generate pGEX6P-1_MpIAA and pGEX6P-1_MpIAAmutDII. Each of these vectors was introduced into E. coli Rosetta2(DE3) strain for induction of recombinant proteins.
For homologous recombination of the MpTIR1 locus
5′- and 3′- homologous arms (3,462 and 3,367 bp, respectively) were amplified from a PAC clone including the MpTIR1 locus (pMM23-241G5; Okada et al., 2000) using the primer pairs, MpTIR1_KO_F1/MpTIR1_KO_R1 and MpTIR1_KO_F2/MpTIR1_KO_R2, respectively. The resultant 5′- and 3′- homologous arms were cloned into pJHY-TMp1 (Ishizaki et al., 2013), using the In-Fusion HD cloning kit (Clontech, Mountain View, CA) to generate pJHY- TMp1_MpTIR1, which was then used for homologous recombination of the MpTIR1 locus.
For the complementation cassette for MpTIR1
A genomic fragment spanning from 5,618-bp upstream of the start codon to 1,081-bp downstream of the stop codon was amplified from pMM23-241G5 (Okada et al., 2000) using the primer pair, MpTIR1_usEntry/MpTIR1_R15, and cloned into pENTR/D-TOPO vector (Thermo Fisher Scientific) to generate pENTR_gMpTIR1. The genomic fragment was then transferred into pMpGWB301 (Ishizaki et al., 2015) by using LR Clonase II (Thermo Fisher Scientific) to generate pMpGWB301_gMpTIR1, which was then used for complementation experiments.
For the expression of AtTIR1 under the MpTIR1 promoter
An MpTIR1 promoter sequence spanning from 5,618-bp upstream of the start codon to the start codon was amplified from pMM23-241G5 (Okada et al. 2000) using the primer pair, MpTIR1_usEntry/MpTIR1_R6, and cloned into pENTR/D-TOPO vector (Thermo Fisher Scientific) to generate pENTR_proMpTIR1. An N-terminal 3xFLAG-tagged AtTIR1 CDS was amplified from a vector containing the sequences, pAN19_TIR1 (a sincere gift from Keiko U. Torii and Naoyuki Uchida), using the primer pair, AscI-Flag_F/AtTIR1_AscI_R. pENTR_proMpTIR1 and the 3xFLAG-AtTIR1 PCR products were digested with AscI (New England Biolabs, Massachusetts, U.S.A.) and then ligated into pENTR_proMpTIR1 to generate pENTR_proMpTIR1:3xFLAG-AtTIR1. The resultant proMpTIR1:3xFLAG-AtTIR1 fragment was then transferred into pMpGWB301 (Ishizaki et al., 2015) by using LR Clonase II (Thermo Fisher Scientific) to generate pMpGWB301_proMpTIR1:3xFLAG-AtTIR1, which was then used for expression studies of AtTIR1 under the MpTIR1 promoter.
To generate Mptir1-1CKO>tdTN plants
A NOS terminator sequence was amplified from a vector containing the sequence, pMpGWB302 (Ishizaki et al., 2015), using the primer pair, NotI-NosT_F/NotI-NosT_R. The resultant NOS terminator fragment and pENTR_gMpTIR1 were digested with NotI (Takara Bio) and then ligated to generate pENTR_NosT-gMpTIR1. The resultant NosT-gMpTIR1 fragment was then transferred into pMpGWB337tdTN (Sugano et al., 2018) by using LR Clonase II (Thermo Fisher Scientific) to generate pMpGWB337tdTN_NosT:gMpTIR1, which was then used for MpTIR1 CKO experiments.
To generate Mptir1-1CKO>CitN plants
The CDS of MpTIR1 was amplified from RNA of WT plants by reverse transcription (RT)-PCR using a primer pair, MpTIR1_entry/MpTIR1_stop, and cloned into pENTR/D-TOPO vector (Thermo Fisher Scientific) to generate pENTR_MpTIR1. The MpTIR1 fragment was then transferred into pMpGWB337 (Nishihama et al., 2016) by using LR Clonase II (Thermo Fisher Scientific) to generate pMpGWB337_MpTIR1, which was then used for MpTIR1 conditional knockout experiments.
To generate proMpEF1A:DII-mTurquoise2-NLS/Mptir1-1CKO>CitN plants and proMpEF1A:mutDII-mTurquoise2-NLS/Mptir1-1cko>CitN plants
DII and mutDII sequences of MpIAA were amplified from vectors containing the respective sequences (see Kato et al., 2015) using the primer pair, MpIAA_dN3/MpIAA DII_R1, and then cloned into pENTR/D-TOPO vector (Thermo Fisher Scientific) to generate pENTR_DII and pENTR_mDII. An mTurquoise2-NLS fragment was amplified from a vector containing the sequence, pMpGWB337mT2N, using the phosphorylated primer pairs, Aor51HI- mT2_F/NOSt_head_SacI_NLS_mTurq_R, digested with SacI (Takara Bio), and then ligated with Aor51HI- (Takara Bio) and SacI- (Takara Bio) digested pMpGWB203 (Ishizaki et al., 2015) to generate pMpGWB203_Gateway:mT2N. The DII- and mutDII- fragments were transferred into pMpGWB203_Gateway:mT2N by using LR Clonase II (Thermo Fisher Scientific) to generate pMpGWB203-DII-mT2N and pMpGWB203-mutDII-mT2N, respectively. These vectors were then used for degradation assays of DII-tagged protein.
For locus deletion of MpTIR1 using CRISPR/Cas9 genome editing
Oligos encoding each guide RNA (gRNA) sequence were annealed with their corresponding antisense oligos (see Supplemental Table 1). The annealed oligos were ligated with BsaI (New England Biolabs) digested vectors with the following combinations: 5′gRNA1 and 5′gRNA3 were ligated with pMpGE_En_04, 5′gRNA2 and 5′gRNA4 were ligated with pBC-GE12, 3′gRNA1 and 3′gRNA3 were ligated with pBC-GE23, and 3′gRNA2 and 4′gRNA4 were ligated with pBC-GE34. 5′gRNA1/2-, 5′gRNA3/4-, 3′gRNA1/2-, and 3′gRNA3/4-pairs are active guide RNA-pairs for a nickase version of the CRISPR/Cas9 genome-editing system (Hisanaga et al., 2019; Koide et al. 2020). Four vectors including one of the active 5′gRNA-pairs and one of the active 3′gRNA-pairs were digested by BglI (New England Biolabs) and ligated at once. The resultant gRNA expression cassettes were transferred into pMpGE017 by using LR clonase II (Thermo Fisher Scientific) to generate pMpGE017_MpTIR1_5′gRNA1/2_3′gRNA3/4, pMpGE017_MpTIR1_5′gRNA3/4_3′gRNA1/2, and pMpGE017_MpTIR1_5′gRNA3/4_3′gRNA3/4. These vectors were then used for genome editing of the MpTIR1 locus. Mptir1-2ld mutation was caused by pMpGE017_MpTIR1_5′gRNA1/2_3′gRNA3/4. Mptir1-3ld and Mptir1-4ldmutations were caused by pMpGE017_MpTIR1_5′gRNA3/4_3′gRNA1/2. Mptir1-5ld mutation was caused by pMpGE017_MpTIR1_5′gRNA3/4_3′gRNA3/4. pMpGE_En04, pBC-GE12, pBC-GE23, pBC-GE34 and pMpGE017 were developed by Keisuke Inoue in Kyoto University.
Plant transformation
Binary vectors were transformed into WT plants as previously described (Ishizaki et al., 2008 and Kubota et al., 2013). To transform Mptir1-1ko mutants, Mptir1-1ko cell masses were cocultured with Agrobacterium GV2260 harboring binary vectors in liquid 0M51C medium under continuous white light. After 2 or 3 days of co-cultivation, the Mptir1-1ko cells were washed with sterilized water and then cultured on half-strength Gamborg’s B5 medium (Gamborg et al., 1968) containing 1% agar, 100 mg/L cefotaxime, and 0.5 μM chlorsulfuron for selection.
Plant genotyping
For genotyping, small plant fragments were crushed in 100 μL of buffer (100 mM Tris-HCl (pH 9.5), 1 M KCl, and 10 mM EDTA), diluted with 400 µL of sterilized water, and then used as templates for PCR.
Mptir1-1ko candidate plants were genotyped by PCR using crude DNA extracts and primer pairs A (MpTIR1_L21/MpTIR1_R12), B (MpTIR1_L14/MpEF_GT_R1), and C (EnSpm_L2/MpTIR1_R13; Supplemental Figure 2A), as described previously (Ishizaki et al., 2013).
Mptir1ld candidate plants were genotyped by amplifying the MpTIR1 genomic locus from crude DNA extracts using the primer pair, MpTIR1_L30/MpTIR1_R21. The resultant DNA fragments were treated with Exonuclease I and Shrimp Alkaline Phosphatase (New England Biolabs), purified with Fast GeneTM Gel/PCR Extraction Kit (NIPPON Genetics Co., Tokyo, Japan), and then sequenced with the primers, MpTIR1_L45 or MpTIR1_R20.
Molecular determination of sex of the plants was performed by amplifying U- and V- chromosome markers from crude DNA extract, as described previously (Fujisawa et al., 2001) using modified primer pairs, rhf-73F_new/rhf-73R_new and rbm-27F_new/rbm-27R_new, respectively.
CKO analysis
In order to induce KO of MpTIR1 in young plants, 1-day-old Mptir1-1CKO>CitN, proMpEF1A:DII- mTurquoise2-NLS/Mptir1-1CKO>CitN, and proMpEF1A:mutDII-mTurquoise2-NLS/Mptir1- 1CKO>CitN gemmalings were treated with approximately 3 μL of 10 μM dexamethasone (DEX) solution to enable nuclear localization of glucocorticoid receptor (GR)-fused Cre proteins, dried for several minutes, and then incubated at 37°C for 1 h. Each gemmaling was treated once more with the same procedure at 4-h intervals.
In order to induce KO of MpTIR1 in gemmae, Mptir1-1CKO>tdTN plant gemmae were treated with approximately 3 μL of 10 μM DEX solution, dried for several minutes, and then incubated at 37°C for 1 h. Each gemma was treated once more with the same procedure at 4-h intervals.
In order to induce KO of MpTIR1 in immature gemmae, 14-day-old Mptir1-1CKO>tdTN plant thalli were vacuum-infiltrated with 5 μM DEX solution, dried for approximately 20 min, and then incubated at 37°C for 1 h. Each thallus was treated twice with the same procedure at 4- to 5-h intervals. The treated thalli were further cultivated under normal condition for a few weeks. Then, gemmae were taken from gemma cups and used for microscopic analysis.
Sectioning of plant tissues
In order to observe cell compositions, 10-day-old sporelings and Mptir1-1ko cell masses grown on the half-strength Gamborg’s B5 agar media were pre-fixed in 2.5% (v/v) glutaraldehyde and 2% (w/v) paraformaldehyde in 50 mM phosphate buffer (pH 7.2) at 4°C overnight. The samples were then washed with 50 mM phosphate buffer, and treated with 2% (w/v) OsO4 solution at room temperature for 2 h, and then washed with 8% (w/v) sucrose. The samples were then dehydrated by 25, 50, and 80% (v/v) ethanol solutions before they were finally dehydrated in 100% (v/v) ethanol, and then embedded into the resin, Quetol 812 (Nissin EM Co., Kyoto, Japan). The resin-embedded samples were sectioned at 1 µm with a diamond knife using Ultracut-UCT (Leica, Wetzlar, Germany). The sections were stained with toluidine blue solution and used for microscopic analyses.
Microscopic analyses
Thalli, sporelings, and Mptir1 cell masses were observed using microscopes SZX16 (Olympus, Tokyo, Japan), M205C (Leica, Wetzlar, Germany), Axiophot (Zeiss, Oberkochen, Germany), or BZ-X710 (Keyence, Osaka, Japan). Plate cultures were photographed using EOS Kiss X3 (Canon, Tokyo, Japan). Z-series images of 2-day-old proMpEF1A:mutDII-mTurquoise2- NLS/Mptir1-1CKO>CitN plant notches were taken using a confocal microscope FLUOVIEW FV1000 (Olympus). SEM images of Mptir1 cell masses and Mptir1-1CKO>tdTN plants were taken by TM3000 (Hitachi High Technologies, Tokyo, Japan), as described previously (Nishihama et al. 2015).
Image manipulations
Bright field (BF) and fluorescence images taken with the M205C were merged using an image analysis software, Fiji (http://fiji.sc/; Schindelin et al., 2012). Z-series confocal images were two- dimensionally projected with max intensity and then merged by using Fiji. BF and fluorescence images taken with BZ-X710 were merged by using BZ-X Analyzer (Keyence).
Measurement of plant areas
Plant area measurements were performed using Fiji. Bright field images were split into RGB channels by the “Split Channels” function. B-channel images were converted into binary images by the “Auto Threshold” function with the “Default” method. Plant areas were then measured by the “Find Edges” and subsequent “Analyze Particles” functions.
Quantification of nuclear fluorescence intensities
Images of 2-day-old proMpEF1A:DII-mTurquoise2-NLS/Mptir1-1CKO>CitN or proMpEF1A:mutDII- mTurquoise2-NLS/Mptir1-1CKO>CitNplant notches, which were cultured for 1 day after KO induction, were quantified using Fiji by measuring mTurquoise2 fluorescence intensities in the nuclei. Five to six biological replicates were prepared for each condition. A series of 45 confocal images at 5-μm intervals were two-dimensionally projected with sum intensity of the slices. From each image, nucleus and background regions were manually selected as 8-μm circles in diameter at 25 locations each. An average intensity value of each nucleus region was subtracted by mean values of background regions from the same image. The data generated were plotted and was analyzed for statistical significance (see “Statistics and graphics” section).
Pull-down assay
E. coli Rosetta2(DE3) strain harboring the GST-MpIAA(627C) or GST-MpIAAmutDII(627C) vectors were precultured in 5 mL of LB liquid medium at 37°C overnight. The overnight grown cultures were then inoculated into 300 mL of fresh LB medium and incubated at 37°C until the OD600 reached 0.5. Isopropyl β-D-thiogalactopyranoside (IPTG) was then added to the culture at the final concentration of 0.1 mM for the induction of protein expression; the cultures were incubated at 37°C for 5 h. After 5h, cells were harvested by centrifugation for 10 minutes at 4,000 xg at 4°C, resuspended in ice-cold sonication buffer (PBS and 1 mM dithiothreitol), and subjected to lysis by sonication. The cell lysates were then centrifuged for 30 min at 12,000 xg at 4°C. The supernatants were purified with PierceTM Disposable Plastic Columns (Thermo Fisher Scientific) of Glutathione Sepharose 4B (GE Healthcare Life Sciences, Massachusetts, U.S.A.).
Fourteen-day-old proMpEF1A:MpTIR1-3xFLAG plants were harvested and immediately frozen in liquid nitrogen. The frozen samples were homogenized with three-fourth volume per g of tissue of extraction buffer (150 mM NaCl, 100 mM Tris-HCl pH 7.5, 0.5% Nonidet P-40, 10 μM dithiothreitol, 1 mM phenylmethanesulfonyl fluoride, 1 μg/mL pepstatin A, and 10 μM MG132) and then melted on ice. Debris were removed by centrifugation at 16,000 xg at 4°C for 15 min after which the supernatant was further filtered through a 0.45-μm pore syringe filter. Protein concentration of the samples was measured using Bradford Protein Assay Kit (Bio-Rad Laboratories Inc., California, U.S.A.).
2.5 mg each of the protein samples were incubated with 10 μL of the GST- MpIAA(627C)- or GST-MpIAA(627C)-conjugated Glutathione Sepharose beads and auxin at 4°C for 30 min. After three times of washing with ice-cold extraction buffer, the beads were mixed with 2x Laemmli sample buffer (100 mM Tris-HCl, pH 6.8, 4% [w/v] SDS, 10% [v/v] 2- mercaptoethanol, and 20% [v/v] glycerol), and boiled at 95°C for 5 min. Samples were separated by SDS-PAGE on a 10% acrylamide gel, and transferred onto polyvinylidene fluoride membranes (Bio-Rad Laboratories, Inc.). Membranes were incubated with anti-FLAG (1:5,000; Sigma-Aldrich, Missouri, U.S.A.) or anti-GST (1:2,000; Nacalai tesque, Kyoto, Japan) for 1 h, respectively, washed with PBST (PBS and 0.1% Tween-20), and then incubated with anti-mouse IgG (1:10,000; GE Healthcare) for 1 h. Bands were visualized with ECL Prime reagent (GE Healthcare) and ImageQuant LAS 4010 (GE Healthcare).
Real-time PCR
For real-time PCR of MpTIR1, 10-day-old Tak-1 and proMpEF1A:MpTIR1-3xFLAG plants were harvested and immediately frozen in liquid nitrogen. For real-time PCR of auxin-responsive genes, the F1 spores and Mptir1-1ko cell masses were precultured in half-strength Gabmorg’s B5 liquid medium for 5 days, treated with 10 μM NAA or solvent control for 4 h, then harvested and immediately frozen in liquid nitrogen. RNA was extracted from the frozen samples using TRIzol (Thermo Fisher Scientific) as described previously (Kubota et al., 2014). Reverse transcription to cDNA and subsequent quantitative PCR were performed as described previously (Kato et al., 2017). Primer pairs, MpTIR1-qPCR_F2/MpTIR1-qPCR_R2, MpC2HDZ-qPCR_F1/MpC2HDZ- qPCR_R1, MpWIP-qPCR_F1/MpWIP-qPCR_R1, and MpEF-qPCR_F/MpEF-qPCR_R were used to quantify MpTIR1, MpC2HDZ, MpWIP, and MpEF1A transcripts, respectively (Supplemental Table1). MpEF1A was used as an internal control. Relative expression levels were calculated by Pfaffl’s method (Pfaffl, 2001).
RNA-sequencing
The F1 spores and Mptir1-1kocell masses were precultured in half-strength Gamborg’s B5 liquid medium for 5 days, and then treated with 10 μM IAA or solvent control for 4 h. Plants were then harvested and immediately frozen in liquid nitrogen. RNA extraction from frozen samples was performed using RNeasy Plant Mini Kit (QIAGEN, Venlo, the Netherland). RNA Libraries were prepared using a NEBNext Ultra II Directional RNA Library Prep Kit for Illumina (New England Biolabs) and sequenced as single end reads using the NextSeq500 platform (Illumina, California, U.S.A.). Total RNA was extracted from the F1 spores and Mptir1-1ko cell masses treated with 10 μM NAA or solvent control in the same way. Library preparation and subsequent paired-end RNA-sequencing was performed by Macrogen Japan (Tokyo, Japan) using the NovaSeq6000 platform (Illumina).
RNA-seq data analysis
For quality control, raw read data were pre-filtered using fastp (version 0.20.1; Chen et al., 2018) with default settings for SE- and PE-sequence data, respectively. The filtered reads were then mapped onto the M. polymorpha genome (v5.1r1 + U-chromosomal genes of v3.1) using STAR (version 2.6.1c; Dobin et al., 2013) with default settings for SE- and PE-sequence data, respectively. Following analyses were performed in R (version 4.0.0; R Core Team, 2020). Reads mapped on exons were counted using the “featureCounts” function of the Rsubread package (version 2.2.2; Liao et al., 2019). Pairwise comparisons were performed by Wald test using the DESeq2 package (version 1.28.1; Love et al., 2014). In the pairwise comparisons between Mptir1-1ko cells and WT tissues, U chromosomal genes were excluded. All the four combinations of comparisons between Mptir1-1kocells (mock samples for IAA or those for NAA) and public thalli data (9-day-old thalli from Higo et al., 2016 or 7-day-old thalli from Yasui et al., 2019) were performed. The shared DEGs among all comparisons were chosen, since all data sets were derived from different experiments, batch effects were not taken into consideration. PCA was performed by the “prcomp” function of the stats package (version 4.0.0; R Core Team, 2020) with log2 transformed read counts of all genes. Factor loadings were calculated as √l ∗ ℎi/ui, where l, hi, and ui represent the eigenvalues of the covariance, the eigenvectors of each gene, and the square root of variance of each gene, respectively.
Statistics and graphics
Statistical tests were performed by R (version 4.0.0; R Core Team, 2020). The stats package (version 4.0.0; R Core Team, 2020) was used for Welch’s t-test (Figure 1A) and Pearson’s correlation test (Supplemental Figure 4B). The NSM3 package (version 1.15; Schneider et al., 2020) was used for Steel-Dwass test (Figure 1C; Supplemental Figure 1B, D). The lawstat package (version3.4; Gastwirth et al., 2020) was used for Brunner-Munzel test (Figure 2D). The multcomp package (version 1.4.13; Hothorn et al., 2008) was used for ANOVA and subsequent Tukey-Kramer test (Figure 3B), and Dunnett test (Supplemental Figure 2E). The DESeq2 package (version 1.28.1; Love et al., 2014) was used for Wald test (Supplemental Data Set 1, 2). Graphs were drawn by R using the ggplot2 package (version 3.3.1; Wickham, 2016), the ggsignif package (version 0.6.0; Ahlmann-Eltze, 2019), and the UpSetR package (version 1.4.0; Conway et al. 2017; Gehlenborg, 2019).
Accession Numbers
Sequence data from this article can be found in the GenBank libraries (http://www.ncbi.nlm.nih.gov) or the MarpolBase (https://marchantia.info) under the following accession numbers: AtTIR1 (AT3G62980); MpTIR1 (Mp6g02750 / Mapoly0035s0062); MpIAA (Mp6g05000 / Mapoly0034s0017); MpARF1 (Mp1g12750 / Mapoly0019s0045); MpARF2 (Mp4g11820 / Mapoly0011s0167); MpC2HDZ (Mp2g24200 / Mapoly0069s0069); MpLRL (Mpzg01410 / Mapoly0502s0001); MpWIP (Mp1g09500 / Mapoly0096s0050); MpTAA (Mp5g14320 / Mapoly0032s0124). Other M. polymorpha transcription factors used in RNA-seq analysis are listed in Supplemental Data Set 3.
Transcriptome data obtained in this study are stored at DNA Data Bank of Japan Sequence Read Archive (https://www.ddbj.nig.ac.jp/dra) under project number DRA013690. Other public transcriptome data obtained from the Sequence Read Archive (https://www.ncbi.nlm.nih.gov/sra) are listed in Supplemental Table 2.
Supplemental Data
Supplemental Figure 1. Genetic evidence for MpTIR1 involved in auxin response.
Supplemental Figure 2. Genotyping and auxin responses of Mptir1-1ko mutants.
Supplemental Figure 3. Verification of induced MpTIR1 KO.
Supplemental Figure 4. Significant overlap between IAA- and NAA-responsive genes.
Supplemental Figure 5. Growth of Mptir1-1ko mutants.
Supplemental Figure 6. Generation and genotyping of MpTIR1-locus deletion mutants.
Supplemental Figure 7. Reproducibility of Mptir1 defects in Mptir1ld mutants.
Supplemental Figure 8. Contribution of auxin-responsive genes to transcriptional properties of Mptir1-1kocells.
Supplemental Table 1. Oligos used in this study.
Supplemental Table 2. Public RNA-seq data used in this study.
Supplemental Data Set 1. Pairwise comparisons between auxin- and mock-treated samples in WT or Mptir1-1ko cells.
Supplemental Data Set 2. Pairwise comparisons between Mptir1-1ko cells and WT samples.
Supplemental Data Set 3. All M. polymorpha transcription factors tested in RNA-seq data analysis.
Funding
This work was supported by MEXT/JSPS KAKENHI (grant numbers: JP18J12698 and 21K20649 to H.S., 12J07037, 19K23751 and 21K15125 to H.K., JP16K07398 and JP20H04884 to R.N., and 25113009, JP17H07424 and JP19H05675 to T.K.) and SPIRITS 2017 of Kyoto University to R.N.
Author contributions
H.S., H.K., R.N. and T.K. designed the research and wrote the paper; H.S., H.K., and M.I. performed research and analyzed the data.
Supplemental Figures and Tables
Acknowledgements
We thank Keiko U Torii and Naoyuki Uchida for kindly providing the vector, pAN19_TIR1. We thank Keisuke Inoue for kindly providing the vectors, pMpGE_En04, pBC-GE12, pBC-GE23, pBC-GE34 and pMpGE017. We thank Takefumi Kondo and Yukari Sando for sequencing RNA. We would like to thank Editage (www.editage.com) for English language editing.
Footnotes
Material distribution footnote The author 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) is Takayuki Kohchi (tkohchi{at}lif.kyoto-u.ac.jp).