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Conserved regulatory mechanism controls the development of cells with rooting functions in land plants Thomas Ho Yuen Tam, Bruno Catarino, and Liam Dolan1 Department of Plant Sciences, University of Oxford, Oxford OX1 3RB, United Kingdom

Land plants develop filamentous cells—root hairs, rhizoids, and caulonemata—at the interface with the soil. Members of the group XI basic helix–loop–helix (bHLH) transcription factors encoded by LOTUS JAPONICUS ROOTHAIRLESS1-LIKE (LRL) genes positively regulate the development of root hairs in the angiosperms Lotus japonicus, Arabidopsis thaliana, and rice (Oryza sativa). Here we show that auxin promotes rhizoid and caulonema development by positively regulating the expression of PpLRL1 and PpLRL2, the two LRL genes in the Physcomitrella patens genome. Although the group VIII bHLH proteins, AtROOT HAIR DEFECTIVE6 and AtROOT HAIR DEFECTIVE SIX-LIKE1, promote roothair development by positively regulating the expression of AtLRL3 in A. thaliana, LRL genes promote rhizoid development independently of PpROOT HAIR DEFECTIVE SIX-LIKE1 and PpROOT HAIR DEFECITVE SIX-LIKE2 (PpRSL1 and PpRSL2) gene function in P. patens. Together, these data demonstrate that both LRL and RSL genes are components of an ancient auxin-regulated gene network that controls the development of tip-growing cells with rooting functions among most extant land plants. Although this network has diverged in the moss and the angiosperm lineages, our data demonstrate that the core network acted in the last common ancestor of the mosses and angiosperms that existed sometime before 420 million years ago. auxin

| bHLH | evolution | rhizoids | root hairs

T

he evolution of rooting structures was a key morphological innovation that occurred when plants colonized the relatively dry continental surfaces of the planet sometime before 470 million year ago. The rooting structures of the earliest diverging groups of land plants comprised systems of rhizoids. Rhizoids are either unicellular filaments (in liverworts and hornworts) or multicellular (in mosses) and elongate into the growth substrate or air surrounding the plant. Bryophyte (liverwort, moss, and hornwort) rhizoids develop on gametophytes, the multicellular haploid stage in the life cycle. The evolution of vascular plants was accompanied by an increase in the morphological diversity of the sporophyte, the diploid multicellular stage of the plant life cycle (1). Roots, multicellular axes derived from meristems with protective caps, were a key innovation that first appeared in the fossil record ∼380 million years ago and likely evolved at least twice—at least once among the Lycophytes and at least once in the Euphyllophyte clade (2). Roots generally grow into the soil and expand the plant–soil interface into different soil horizons. For example, some root systems extend deep into the soil (taproots), whereas others proliferate in the nutrient-rich horizons near the soil surface. However, unicellular, filamentous protuberances called root hairs, which are morphologically similar to rhizoids, develop on all roots with few exceptions (2–5). Despite the different contexts in which they develop, root hairs and rhizoids carry out similar rooting functions, including nutrient uptake and anchorage (6, 7). Group VIII basic helix–loop–helix (bHLH) transcription factors encoded by ROOT HAIR DEFECTIVE SIX-LIKE (RSL) genes positively regulate root-hair development in the angiosperm Arabidopsis thaliana and both rhizoid and caulonema in

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the moss Physcomitrella patens (8–12). To test whether the conservation of RSL function is unique or indicative of a more general, conserved regulatory mechanism, we determined whether other components of the gene-regulatory network controlling angiosperm root-hair development are conserved among land plants. Group XI bHLH transcription factors encoded by LOTUS JAPONICUS ROOTHAIRLESS1-LIKE (LRL) genes are also key regulators of root-hair development in angiosperms. LRL genes encode several proteins that regulate root-hair development in diverse angiosperms, including ROOTHAIRLESS1 (LjRHL1) in Lotus japonicus, AtLRL1, AtLRL2, AtLRL3 in A. thaliana (13, 14), and ROOTHAIRLESS1 (OsRHL1) in Oryza sativa (15). Other LRL genes include OsPTF1, involved in phosphate starvation tolerance in rice (16), and AtUNE12, involved in female gametophyte development in Arabidopsis (17). Here we define the function of LRL genes in the moss P. patens and conclude that, together, LRL and RSL genes form the core of an ancient network that operated in the common ancestor of the mosses and angiosperms that existed in the Silurian Period 415–435 million years ago. Results Two LRL Genes Are Present in the Genome of the Moss P. patens. The

LRL transcription factors belong to group XI of plant bHLH transcription factors (18) and share a conserved LRL domain of 36 amino acids between the bHLH domain and the C terminus (Fig. 1A). To identify LRL homologs in P. patens, we performed BLAST searches in selected land plant genomes using the AtLRL3 (At5g58010) sequence as a query. Phylogenetic analyses identified two LRL genes in P. patens, designated PpLRL1 Significance This work describes the discovery of an ancient genetic mechanism that was used to build rooting systems when plants colonized the relatively dry continental surfaces >470 million years ago. We demonstrate that a group of basic helix–loop–helix transcription factors—the LOTUS JAPONICUS ROOTHAIRLESS1LIKE proteins—is part of a conserved auxin-regulated gene network that controls the development of tip-growing cells with rooting functions among extant land plants. This result suggests that this mechanism was active in the common ancestor of most land plants and facilitated the development of early land plant filamentous rooting systems, crucial for the successful colonization of the land by plants. Author contributions: T.H.Y.T. and L.D. designed research; T.H.Y.T. and B.C. performed research; T.H.Y.T., B.C., and L.D. analyzed data; and T.H.Y.T., B.C., and L.D. wrote the paper. The authors declare no conflict of interest. This article is a PNAS Direct Submission. Freely available online through the PNAS open access option. 1

To whom correspondence should be addressed. Email: [email protected].

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10. 1073/pnas.1416324112/-/DCSupplemental.

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Edited by Philip N. Benfey, Duke University, Durham, NC, and approved June 11, 2015 (received for review August 22, 2014)

Fig. 1. (A) Schematic representation of gene structure and amino acid alignment of LRL transcription factors. Only the bHLH and LRL domains are shown. (B) Tree of LRL proteins and related groups of bHLH proteins based on ML statistics. This tree resolved PpLRL1, PpLRL2, OsRHL1, AtLRL1, AtLRL2, AtLRL3, AtLRL4, and AtUNE12 within a monophyletic group, which also contains four additional rice and four additional Selaginella moellendorffii genes (Fig. S1). aLRT values are indicated above the nodes. Two other groups of plant bHLHs involved in root-hair and rhizoid development, group VIIIc (1) encoded by the class I RSL genes, and group VIIIc (2) encoded by the class II RSL genes, are labeled in red and green, respectively. The full ML and Bayesian trees are shown in Fig. S1. (C) Unrooted tree showing the relationships of different LRL proteins in land plants using Bayesian statistics. The location of selected LRL proteins from the moss P. patens (Pp), the monocot O. sativa (Os), and the eudicot A. thaliana (At) are indicated in the diagram. The full ML and Bayesian trees are shown in Fig. S2.

(Pp1s173_83V6.1) and PpLRL2 (Pp1s209_22V6.2), respectively (Fig. 1B). Maximum- likelihood (ML) analysis resolved PpLRL1, PpLRL2, AtLRL1 (At2g24260), AtLRL2 (At4g30980), AtLRL3, AtLRL4 (At1g03040), AtUNE12 (At4g02590), five rice proteins including OsRHL1 (Os06g08500), and four Selaginella moellendorffii proteins into a monophyletic group (Fig. 1B, Fig. S1, and Dataset S1). The LRL domains of PpLRL1 and PpLRL2 are 97.2% identical to each other and are 77.8% and 80.6% identical to the LRL domain of the AtLRL3 protein, respectively. The two PpLRL proteins have conserved bHLH domains that are 100% identical to each other (Fig. 1A). The bHLH domains of the PpLRL proteins are 92.4% identical to the bHLH domain of the AtLRL3 protein. To infer the phylogenetic relationship of the different proteins containing the LRL domain, we constructed phylogenetic trees using Bayesian and ML methods (Fig. 1C and Fig. S2). These trees showed that LRL homologs are present in all 32 land plant species sampled (Fig. S3). The LRL proteins comprise three major groups, designated XIa, XIb, and XIc (Fig. 1C and Fig. S2). Genes that control the development of root hairs are present in group XIa (Fig. 1C, Fig. S2, and Dataset S2). PpLRL1 and PpLRL2 are not members of any of these three groups (Fig. 1C and Figs. S2 and S3). Together, these results indicate that the LRL genes are a conserved group of transcription factors whose existence predates the divergence of mosses and flowering plants. PpLRL Genes Have Different, but Overlapping, Expression Patterns.

We hypothesized that LRL genes controlled the development of rhizoids and caulonemata and that PpLRL1 and PpLRL2 would be expressed in these cell types. To define where LRL genes are expressed, we replaced the endogenous PpLRL1 and PpLRL2 genes with the coding sequence (CDS) of the UidA glucuronidase-encoding gene (GUS) (Fig. S4). Consequently, the GUS expression in these lines identified regions of the protonema in which the PpLRL1 and PpLRL2 promoters are active in respective loss-of-function Pplrl1 or Pplrl2 mutant background. In P. patens, the germination of spores is followed by the development 2 of 10 | www.pnas.org/cgi/doi/10.1073/pnas.1416324112

of a uniseriate filamentous network (protonema) comprising cells with large chloroplasts and transverse cell walls (chloronema cells). Subsequently, increasingly longer cells with oblique cell walls (caulonema cells) develop. Caulonema cells are morphologically and genetically similar to rhizoids (19). Both PpLRL1 and PpLRL2 were expressed in the central region of the protonema, which is rich in chloronema and from which caulonema develop (Fig. 2 A and B). No expression was detected in the very edges of the 30-d-old protonema, where caulonema is the dominant cell type. These observations suggest that PpLRL1 and PpLRL2 are active in cells that develop caulonema. PpLRL1 promoter was also active in the rhizoid-forming region at the base of the gametophore in plants in which the PpLRL1 CDS was replaced by the GUS-reporter gene (Fig. 2 C and D). However, no GUS expression was detected in rhizoids themselves. This expression pattern is consistent with a role for PpLRL1 in the regulation of the earliest stages of rhizoid development. PpLRL1 promoter was also active in the gametophore apex and leaves, suggesting that this gene likely controls other aspects of gametophore development. PpLRL2 promoter activity was detected in the epidermal cells of the gametophore and leaves (Fig. 2 C and D). Unlike PpLRL1, no GUS activity was detectable in the buds or at the base of the gametophore. Like PpLRL1, there was no detectable expression of PpLRL2 promoter in rhizoids. PpLRL1 and PpLRL2 Are Positive Regulators of Rhizoid Development.

To determine whether PpLRL genes function in rhizoid development, we generated mutants that lack LRL gene function by homologous recombination (Figs. S5 and S6). To confirm that homologous recombination had taken place, we characterized the genomic organization of the transformants. PCR analyses were performed to select transformants in which the endogenous PpLRL gene was replaced by the gene-targeting construct (Fig. S5) and thus represent complete loss-of-function alleles. From these, Southern blot analyses were used to further select those Tam et al.

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transformants in which there were no additional transgene insertions in the genome (Fig. S5). Three independent lines for each gene were maintained for phenotypic analyses. With the exception of Pplrl2-1, all single-mutant lines and all three double-mutant lines showed a single band in the Southern blot analysis, indicating that there was a single insertion into the genome (Fig. S5). Pplrl1 and Pplrl2 single-mutant plants developed ∼40% fewer rhizoids than wild-type (WT) (Fig. 3 A and B), and these rhizoids were ∼35% and 18% shorter than WT, respectively (Fig. 3C). Pplrl1 Pplrl2 double mutants were completely rhizoidless, and no rhizoids developed on these plants (Fig. 3D). Together, these data show that the activity of PpLRL1 and PpLRL2 together are required for the initiation and development of rhizoids from buds and gametophores. To determine whether the bHLH and LRL domains of the LRL proteins—characteristic of group XI plant bHLHs—are required for the control of rhizoid development in P. patens, we generated additional mutants in which the region containing the bHLH–LRL domain of each protein was deleted (Fig. S6). The organization of the genomic DNA in the mutants confirmed that the bHLH–LRL domains were deleted (Fig. S6). These mutants were designated Pplrl1ΔbL and Pplrl2ΔbL, respectively. Defects in rhizoid and caulonema development in these partial-deletion mutants were similar to the defects in the complete genedeletion mutants (Fig. 3E). To independently verify that the bHLH and LRL domains are positive regulators of rhizoid development, we overexpressed a fragment of the PpLRL1 and PpLRL2 CDS containing the essential bHLH and LRL domains (Fig. S6). Plants were transformed with constructs in which the bHLH– LRL domain was placed under the control of the CaMV35S promoter (Fig. S6). Plants transformed with 35S:PpLRL1bL-1, which overexpressed the bHLH–LRL fragment of PpLRL1, developed longer and more intensively pigmented rhizoids than WT (Fig. 3E). In contrast, rhizoid pigmentation in 35S:PpLRL2bL1–transformed plants, which overexpressed the bHLH–LRL fragment of PpLRL2, resembled WT (Fig. 3E and Fig. S6). This difference in rhizoid phenotype in these lines suggests that PpLRL1 plays a more important role than PpLRL2 in rhizoid development. Together, these data are consistent with the hypothesis that the bHLH and LRL domains are responsible for PpLRL gene function in rhizoid development. PpLRL Genes Positively Regulate Caulonema Development. The transition from chloronema to caulonema, characteristic of WT protonema development, did not occur in Pplrl1 Pplrl2 doubleTam et al.

mutant plants (Fig. 4 A and B). Caulonemata did not develop, and, instead, a dense mat of chloronemata constituted the entire protonema (Fig. 4A). The diameter of the Pplrl1 Pplrl2 doublemutant protonema was approximately half that of the WT, which can be explained by the absence of caulonemata, because these cells have higher growth rates and are longer than chloronemata (Fig. 4 B and C). To verify that the smaller diameter of Pplrl1 Pplrl2 mutants was due to the loss of caulonemata, we compared the growth rate of Pplrl1 Pplrl2 double mutants with WT (Fig. 4C). In WT plants, chloronema growth rates were 7.94 ± 0.45 μm·h−1, and caulonemal growth rates were 16.86 ± 0.23 μm·h−1. The growth rates of protonema filaments in Pplrl1 Pplrl2 double mutants were 8.48 ± 0.81 μm·h−1. That is, the growth rates of protonema filaments of the Pplrl1 Pplrl2 double mutant were not significantly different from those of WT chloronema. These data indicate that chloronemata, but not caulonemata, develop in Pplrl1 Pplrl2 double mutants. Pplrl1 and Pplrl2 single mutants developed less severe phenotypes than the Pplrl1 Pplrl2 double mutants. Protonema diameters of Pplrl1 and Pplrl2 mutants were 83% and 89% that of WT, respectively (Fig. 4 A and B). Together, these data demonstrate that PpLRL1 and PpLRL2 positively regulate the developmental transition from chloronema to caulonema during the development of moss protonemata. Auxin Positively Regulates PpLRL Activity. Auxin positively regulates the development of rhizoids and caulonemata in P. patens (8, 9, 20–22). To test whether auxin promotes rhizoid and caulonema development by positively regulating the expression of PpLRL genes, we determined the effect of auxin treatment on the activities of the PpLRL1 and PpLRL2 promoters. P. patens plants in which the endogenous PpLRL CDS were replaced by the GUS gene (Fig. S4) were grown on Knops-GT medium (defined in Plant Material, Growth Conditions, and Phenotypic Analyses) for 30 d. No GUS expression was detectable in the protonema or buds, even when incubated in glucuronidase reaction buffer for 24 h (Fig. 5A). Addition of 1-naphthaleneacetic acid (NAA) to the medium at a final concentration of 100 nM resulted in detectable GUS expression in both protonema and buds of PpLRL1pro:GUS and PpLRL1pro:GUS plants (Fig. 5A); this indicates that exogenous treatment with auxin increases the activity of the PpLRL1 and PpLRL2 promoters. Together, these data suggest that auxin positively regulates PpLRL1 and PpLRL2 expression. If auxin positively regulates LRL expression, and LRL genes in turn promote caulonema development, we hypothesized that PNAS Early Edition | 3 of 10

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Fig. 2. GUS staining pattern in plants transformed with PpLRL1pro:GUS and PpLRL2pro:GUS and untransformed controls (WT). (A and B) PpLRL1 and PpLRL2 were expressed in the center of the protonema, composed mainly of chloronema filaments. (C and D) Young buds and gametophores isolated from 4-wk-old protonemata grown on KNOPS-GT medium. PpLRL1, but not PpLRL2, was expressed in the early buds. In the gametophores, PpLRL1 was expressed in the base, where basal rhizoids develop. It was also expressed in the stem and at the apex. PpLRL2, in contrast, was not expressed in the early buds or the stem of the gametophores, but was expressed in the leaves. [Scale bars: 2 mm (A), 1 mm (B and D), and 200 μm (C).]

Fig. 3. Rhizoid phenotypes of PpLRL loss- and gain-of-function mutants. (A) Rhizoid phenotypes of complete PpLRL loss-of-function mutants. (Scale bars: 2 mm.) (B) Quantification of rhizoid number. (C ) Quantification of rhizoid length. Rhizoid length was determined by measuring the length between the base of the gametophore and the tip of the rhizoid cluster. Both rhizoid number and length were significantly reduced in the Pplrl1 and Pplrl2 single mutants. Rhizoids are absent in the Pplrl1 Pplrl2 double mutant. Asterisks represent P values from two-tailed unequal variance (Welch’s) t tests comparing individual mutant lines with WT. **P < 0.01; ***P < 0.000005. Exact P values are as follows. For rhizoid number, Pplrl1 and WT: P = 2.6 × 10−6; Pplrl2 and WT: P = 3.1 × 10−6; and Pplrl1 Pplrl2 and WT: P = 1.5 × 10−8. For rhizoid length, Pplrl1 and WT: P = 1.4 × 10−6; and Pplrl2 and WT: P = 1.6 × 10−3. NA, not applicable (the Pplrl1 Pplrl2 mutant has no measurable rhizoids). Error bars represent SEM (n = 9 for WT, n = 15 for Pplrl1, n = 12 for Pplrl2, and n = 11 for Pplrl1 Pplrl2). (D) No rhizoids developed at the base of the Pplrl1 Pplrl2 double-mutant gametophore. (Scale bars: 500 μm.) (E) Phenotypes of PpLRLΔbL partial-deletion mutants and 35S:PpLRLbL mutants. The 35S:PpLRL1bL-1 mutant showed an increased number of rhizoids and increased pigmentation.

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Fig. 4. Protonema phenotypes of PpLRL loss-of-function mutants. (A) Caulonema phenotype of complete PpLRL loss-of-function mutants. Caulonema development was defective in the Pplrl1 and Pplrl2 single mutants, and no caulonemata developed in the Pplrl1 Pplrl2 double mutants. [Scale bars: 2 mm (column 1) and 500 μm (column 2).] (B) Quantification of protonema development in terms of green (chloronema-rich) and nongreen (caulonema-rich) contributions to diameter. Asterisks represent P values from two-tailed unequal variance (Welch’s) t tests comparing individual mutant lines with WT. *P < 0.05; **P < 0.01. The exact P values are as follows. For chloronema, Pplrl1 and WT: P = 0.052; Pplrl2 and WT: P = 0.10; and Pplrl1 Pplrl2 and WT: P = 7.3 × 10−6. For caulonema, Pplrl1 and WT: P = 0.0011; and Pplrl2 and WT: P = 0.074. Error bars represent SEM (n = 8 for WT and Pplrl1; n = 7 for Pplrl2; and n = 6 for Pplrl1 Pplrl2). (C) Growth rate of caulonema and chloronema filaments in the loss-of-function mutants. In all of the mutant lines, the growth rate of chloronema filaments was not significantly different from WT, but the growth rate of caulonema was significantly reduced. Asterisks represent the P values from twotailed unequal variance (Welch’s) t tests comparing individual mutant lines with WT. *P < 0.05; **P < 0.01. The exact P values are as follows: for chloronema: P > 0.25 for all mutant lines compared with WT; for caulonema, Pplrl1 and WT: P = 0.017; and Pplrl2 and WT: P = 0.0024. Error bars represent SEM (n = 3 for chloronema; n = 5 for Pplrl1 and Pplrl2 caulonema; and n = 3 for WT caulonema).

auxin-induced morphological changes in caulonema would require PpLRL1 and PpLRL2 activity. To test this hypothesis, we compared the morphology of Pplrl1 Pplrl2 double mutants grown in medium containing 100 nM NAA with double mutants grown without added auxin. Pplrl1 Pplrl2 protonema that developed in the presence of 100 nM NAA were morphologically identical to double mutants grown without auxin (Fig. 5B). That is, Pplrl1 Pplrl2 mutants were resistant to auxin. Furthermore, auxin induced the development of rhizoids on buds of WT, but rhizoid induction did not occur on the buds of Pplrl1 Pplrl2 double mutants (Fig. 5C). Together, these data demonstrate that rhizoid and caulonema development in plants that lack both PpLRL1 and PpLRL2 function are resistant to auxin. This result is consistent with the model in which auxin-regulated caulonema and rhizoid development require PpLRL1 and PpLRL2 activity. The STYLISH1/SHI transcription factors regulate auxin biosynthesis in both A. thaliana and P. patens (23, 24). To test whether auxin promotes the expression of PpLRL1 and PpLRL2 in a PpSHI-dependent fashion in P. patens, we quantified mRNA levels in the auxin biosynthesis mutants Ppshi1, PpOXSHI1-5, and Ppshi2. We could not detect a difference in PpLRL1 or PpLRL2 mRNA levels between WT and these mutants (Fig. S7 Tam et al.

A–C). This result suggests that auxin-regulated expression of PpLRL1 and PpLRL2 is independent of the STYLISH/SHI transcription factors. PpLRL1 and PpLRL2 Are Required for Low-Phosphate-Induced Development of Caulonema. We hypothesized that PpLRL genes

would be required for the adaptive response of protonema to low phosphate; growth of protonema in medium containing low phosphate promotes the transition from chloronema to caulonema (25). To test this hypothesis, we compared the phenotypes of Pplrl1 Pplrl2 double mutants with WT in media with different phosphate content. Low phosphate promotes the chloronema-to-caulonema transition in WT protonema, but the protonema morphology of Pplrl1 Pplrl2 double mutants was identical in replete and no phosphate (Fig. 6 A and B). That is, the double mutant protonema was insensitive to changes in external phosphate concentration, and caulonema did not develop. These results suggest that the morphological changes that take place as part of the adaptive response to low phosphate require PpLRL1 and PpLRL2 activity. However, we could not detect a difference in PpLRL1 and PpLRL2 mRNA levels in different phosphate conditions (Fig. S7D), suggesting PNAS Early Edition | 5 of 10

Fig. 5. Auxin-regulated caulonema and rhizoid development require PpLRL1 and PpLRL2 function. (A) Four-week-old protonema grown with 0 μM (−NAA) or 0.1 μM (+NAA) NAA. Auxin treatment induced the activity of the PpLRL1 and PpLRL2 promoters. [Scale bars: 5 mm (columns 1 and 4) or 500 μm (columns 2, 3, 5, and 6).] (B) The Pplrl1 Pplrl2 mutants are partially insensitive to auxin. (Scale bars: 5 mm.) (C) Close-up of a gametophore from 2-wk-old protonema grown with 0.1 μM NAA. Auxin failed to induce rhizoid development in the Pplrl1 Pplrl2 double mutants. (Scale bars: 500 μm.)

that the induction of caulonemal development in low-phosphate media does not increase the expression of LRL genes in P. patens. PpLRL and PpRSL Genes Act Independently in P. patens. The group VIII basic helix–loop–helix transcription factors, AtROOT HAIR DEFECTIVE6 (AtRHD6) and AtROOT HAIR DEFECTIVE6LIKE1 (AtRSL1), promote the development of root hairs by positively regulating AtLRL3 expression (13, 14). To determine whether there are regulatory interactions between PpRSL and PpLRL genes, we compared the expression of PpLRL1 and PpLRL2 in Pprsl1 Pprsl2 double mutants and WT, and the expression of PpRSL1 and PpRSL2 in Pplrl1 Pplrl2 double mutants with WT. We found no evidence of transcriptional regulatory interactions between PpLRL and PpRSL genes in P. patens (Fig. S7 E and F). Together, these data suggest that the LRL and RSL genes are transcriptionally independent of each other in P. patens. 6 of 10 | www.pnas.org/cgi/doi/10.1073/pnas.1416324112

Discussion The colonization of the land by streptophyte plants sometime before 470 million years ago was a pivotal event in the history of the Earth. Structures that anchor plants to their growth substrate and provide access to mineral nutrients, water, and the soil microflora were key adaptations to life on the relatively dry continental surfaces. Recent phylogenetic analyses suggest that the rhizoidless algae of the Zygnematales or Colecochaetales (or a clade consisting of Zygnematales and Coleochaetales) are sister to the land plants (26–29). If a clade consisting of Zygnematales and Coleochaetales is sister to land plants, then there are two equally parsimonious models for the evolution of rhizoids, each involving two changes: (i) rhizoids evolved independently in the common ancestor of land plants and in the more distantly related algal lineages such as the Charales or (ii) rhizoids evolved in the common ancestor of Charales, Zygnematales, Coleochaetales, and land plants, but were lost in the common ancestor of the Zygnematales and Coleochaetales. Tam et al.

Fig. 6. PpLRL1 and PpLRL2 function are required for low phosphate adaptive response. (A) Pplrl1 Pplrl2 double mutants are insensitive to low phosphate. Protonemata were grown in high (+P) and low (−P) phosphate medium for 2.5 wk. (Scale bars: 5 mm.) (B) Low phosphate increased caulonema development significantly in WT protonemata. *P = 0.034 [twotailed unequal variance (Welch’s) t tests)] but had no discernable effect on Pplrl1 Pplrl2 mutants. Error bars represent SEM (n = 20 for WT+P and Pplrl1 Pplrl2 +P; n = 16 for WT-P; and n = 12 for Pplrl1 Pplrl2 −P).

We show here that LRL genes positively regulate the development of the tip-growing rhizoids and caulonema in the moss P. patens. LRL genes also positively regulate the development of root hairs in angiosperms (13–15). Given that similar proteins control the development of filamentous rooting cells at the plant–soil interface in mosses and angiosperms, we conclude that LRL genes positively regulated the development of tip-growing rooting cells in the common ancestor of mosses and angiosperms that existed sometime before 420 y ago (30). Basic helix–loop–helix proteins encoded by RSL genes also form a network that controls filamentous rooting cell development. The class I RSL genes, AtRHD6 and AtRSL1, promote root-hair development in A. thaliana; root hairs do not develop on Atrhd6 Atrsl1 double loss-of-function mutants (10). AtRHD6 and AtRSL1 positively regulate the expression of the class II RSL genes AtRSL2, AtRSL3, and AtRSL4 and the LRL gene AtLRL3; these in turn modulate late stages of root-hair differentiation in A. thaliana (12). AtRSL4 is a class II RSL gene that is necessary and sufficient for growth. Modulation of AtRSL4 expression by class I RSL genes, environment, and auxin determine root-hair cell size. AtRSL4 controls cell size by controlling the expression of genes that encode proteins involved in tip growth (12). Class I RSL genes in P. patens—PpRSL1 and PpRSL2—positively regulate rhizoid and caulonema development (8–10). Class II RSL genes—PpRSL3, PpRSL4, PpRSL5, and PpRSL6—regulate caulonema development, but have much more subtle functions than class I RSL genes (11). Tam et al.

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Our results demonstrate that the LRL and RSL genes represent core components of a conserved gene-regulatory network that was present in the common ancestor of mosses and angiosperms. However, although we showed that the genetic components of this gene-regulatory network have been conserved, the detailed patterns of transcriptional regulation within this network remain to be resolved. For example, AtRHD6 and AtRSL1 (class I RSL genes) regulate the expression of AtLRL3 in A. thaliana (13, 14), but we could not find evidence that PpRSL genes control PpLRL gene expression in P. patens. If there is a genuine absence of transcriptional regulation between LRL and class I RSL genes in P. patens, then two alternative evolutionary scenarios can be suggested. Either (i) class I RSL genes acquired the ability to regulate the expression of LRL genes in the lineage that gave rise to angiosperm, or (ii) RSL-regulated LRL expression was an ancestral trait that was lost in the moss lineage. The mechanism of auxin-regulated caulonema and rhizoid development in P. patens involves key components of the auxinsignaling pathway that is conserved among land plants (8, 9, 11, 20–23, 31). We showed here that auxin positively regulates the expression of PpLRL1 and PpLRL2 and that the PpLRL genes are necessary for auxin-induced caulonema and rhizoid development. Therefore, the observed GUS activity in the PpLRL1pro:GUS and PpLRL2pro:GUS protonema may be due to higher auxin concentration in the center of the protonema. Previously, it was shown that auxin promotes caulonema and rhizoid development by positively regulating the expression of the class I PpRSL genes, PpRSL1 and PpRSL2 (8, 9). We showed here that auxin positively regulates PpLRL expression. However, because we could not find evidence that class I PpRSL genes regulate the expression of PpLRL genes, or that PpLRL genes regulate the expression of PpRSL genes in P. patens, these data suggest that auxin positively regulates class I PpRSL and PpLRL genes independently. According to the classical P. patens literature (22, 32), gametophores form only on caulonema. Our results suggest that bud formation can occur in chloronema. The morphology, chloroplast density, and growth rate of the Pplrl1 Pplrl2 protonema resemble WT chloronema, but buds develop from this mutant. Similarly, the caulonema-less Pprsl1 Pprsl2 loss-of-function mutants (8, 33) also develop buds from chloronema. Together, these data suggest that the bud-formation process in P. patens is not limited to caulonema, but can develop from chloronema in certain genetic backgrounds. LRL genes are not general regulators of tip growth. There are three types of tip-growing cells in P. patens: chloronema, caulonema, and rhizoid. The caulonemata are cytologically similar to rhizoids (33–36). PpLRL genes positively control both rhizoid and caulonema development, but not chloronema development; chloronema development is indistinguishable from WT in Pplrl1 Pplrl2 double mutants. These results are consistent with the observation that caulonemata and rhizoids also have similar responses to auxin and low phosphate, and these responses are different from the response of chloronema to auxin (8, 9, 25). Together, these data suggest that the PpLRL genes are specific regulators of the development of tip-growing cells with a rooting function. Similarly, RSL genes are specific regulators of caulonema and rhizoid development and do not regulate chloronema development (8–10). Therefore, we concluded that LRL and RSL genes specifically regulate rhizoid and caulonema development and are not simply general regulators of tip growth. This conclusion is supported by the finding that the tip-growth defect in Atrsl1 Atrsl2 double mutants is restricted to root hairs in A. thaliana; pollen tube development is identical to WT in these double mutants (10). LRL transcription factors are ancient and were present in the last common ancestor of mosses and angiosperms that was extant at least 420 million years ago. Bayesian analysis indicates that

three monophyletic clades (XIa, XIb, and XIc) constitute the majority of land plant LRL proteins. However, PpLRL proteins are not members of groups XIa, XIb, or XIc. Groups XIb and XIc each contain a single gymnosperm LRL protein that is sister to the other angiosperm LRL proteins in each of these monophyletic groups, suggesting that groups XIb and XIc originated before the divergence of gymnosperms and angiosperms (Figs. S2 and S3). The evolution of group XIa LRL protein is more complex (Figs. S2 and S3), but the presence of a group XIa LRL proteins in Amborella and other angiosperm taxa suggests that this group was present in the common ancestor of extant angiosperms. Only group XIa LRL proteins are known to regulate root-hair development in angiosperms. These results support our conclusion that the role of LRL proteins in regulating filamentous rooting cell development is ancient. Further sampling of LRL genes in well-annotated, high-quality assemblies of monilophyte (ferns and horsetails) and gymnosperm genomes will allow the elucidation of the evolutionary history of LRL genes during land plant evolution. There is evidence that PpLRL1 plays a greater role than PpLRL2 in rhizoid development. First, the PpLRL1 and PpLRL2 expression patterns are different. The strong expression of PpLRL1 in chloronema cells, young buds, and the base of the gametophore is consistent with its role in controlling early stages of caulonema and rhizoid development. In contrast, PpLRL2 promoter activity was not detected in either young buds or the base of gametophores where rhizoids develop, suggesting that PpLRL2 regulates rhizoid development via a nonautonomous mechanism. Second, Pplrl1 mutants have a stronger rhizoid defect than Pplrl2 single mutants. Third, overexpression of the PpLRL1bL partial transcript led to an increased number of rhizoids, whereas overexpression of the PpLRL2bL partial transcript had no discernable effect on rhizoid development. Nevertheless, the activity of both proteins is required for the development of rhizoids because only the double mutant does not form rhizoids. Together these results suggest that, although both PpLRL1 and PpLRL2 function in rhizoid development (because the double mutant does not form rhizoids), PpLRL1 plays a greater role than PpLRL2. Conclusion LRL genes are part of an ancient gene-regulatory network that controls the development of tip-growing filamentous cells with rooting functions—rhizoids, caulonema, and root hairs—at the interface between land plants and the soil. This auxin-regulated network was active early in land plant evolution and was present in the last common ancestor of the angiosperms and mosses. This ancient gene-regulatory network diverged since P. patens and A. thaliana last shared a common ancestor (Fig. 7). Materials and Methods Sequence Retrieval and Alignment. To identify potential LRL proteins in P. patens, we performed BlastP searches using AtLRL3 (At5g58010) amino acid sequence as a query on Phytozome (Version 9.1) (37) with a E-value threshold of 1e-6, and we retrieved all of the resulting sequences. We used only the primary transcripts where alternative transcripts exist. CDS retrieved were translated to amino acids and then aligned by using Mafft with the L-INS-I option (38). The alignment was trimmed with TrimAl (Version 1.3) (39) as implemented in Phylemon (Version 2.0) with automated parameters (40). The resulting alignment consisted of 139 sequences and 54 columns from four species. To extend our analysis and resolve the relationship among LRL proteins, sequences were retrieved from BLAST searches on genomes on Phytozome (Version 9.1) (37), the Pinus taeda v1 transcriptome on the Dendrome project (dendrome.ucdavis.edu/resources/blast/), the L. japonicus genome assembly build 2.5 (www.kazusa.or.jp/lotus/index.html), and the Amborella trichocarpa genome (41). Only sequences containing unambiguous LRL domains were included. The sequences were aligned with Mafft. The alignment was trimmed with TrimAl (Version 1.3) (39) as implemented in Phylemon (Version

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Fig. 7. Models indicating the transcriptional regulation between auxin, LRL, and class I RSL genes in rhizoid and caulonema development in P. patens (A) and root hair development in A. thaliana (B).

2.0) with automated parameters (40).The final amino acid alignment, containing 164 sequences from 32 species and 121 columns, was used in subsequent analyses. This alignment was also used to determine the similarity of the bHLH and LRL domains (as shown in Fig. 1) of selected LRL proteins, by using the Ident and Sim program in the Sequence Manipulation Suite (42). Phylogenetic Analyses. The best-fit model of amino acid evolution for the trimmed alignments were determined by Prottest (Version 3) (43), and the JTT+Γ (for Dataset S1) or the JTT+Γ+I (for Dataset S2) model was chosen. The ML trees were calculated with PhyML (Version 3.0) (44), with the SH-Like approximate likelihood-ratio test (aLRT) for branch support (45) and the best of nearest neighbor interchange and subtree pruning and regrafting for tree improvement. Bayesian analyses were carried out with MrBayes (Version 3.2) (46) for >15 million generations. One tree was saved every 100 generations, and the first 25% of trees were discarded. A 50-majority-rule tree was generated from the remaining trees. The default settings were used for gap treatment—i.e., as unknown character in PhyML and as missing data in MrBayes. Plant Material, Growth Conditions, and Phenotypic Analyses. The Gransden WT strain (47) of P. patens (Hedw.) Bruch & Schimp was used in this study. Unless otherwise stated, all plants were grown in a SANYO MLR-351 growth chamber at 25 °C and a 16-h light/8-h dark regime with a photosynthesis photon flux density of ∼40 μmol m −2 ·s−1. Plants were grown on a modified KNOPS medium (referred to as “minimal medium” or “KNOPS-GT”) (47): 0.8 g/L CaN2O6·4H2O, 0.25 g/L MgSO4·7H2O, 0.0125 g/L FeSO4·7H2O, 0.055 mg/L CuSO4·5H2O, 0.055 mg/L ZnSO4·7H2O, 0.614 mg/L H3BO 3, 0.389 mg/L MnCl 2·4H2 O, 0.055 mg/L CoCl 2·6H2O, 0.028 mg/L KI, 0.025 mg/L Na 2MoO4·2H2O, 25 mg/L KH2 PO 4 buffer (i.e., 1.84 mM at pH 6.5 with KOH), and 7 g/L agar (Formedium AGA03). Then, 5 g/L glucose and 0.5 g/L ammonium tartrate dibasic (C4H6O6·2H3N) were added as supplements for routine subculture of macerated protonema tissues (KNOPS medium). Sterile cellophane discs (80-mm 325P Cellulose discs; A.A. Packaging) were put on top of the medium to facilitate phenotypic analysis and collection of tissues for subculture. PEG-Mediated Transformation of P. patens. PEG-mediated transformation was performed as described (10, 48). Antibiotic selection was performed by adding 50 μg/mL G418 disulfate or 25 μg/mL hygromycin B to the growth medium. Southern Blot Analysis. Genomic DNA was extracted from protonema tissues by using the cetyltrimethylammonium bromide (CTAB) method and then digested overnight with the appropriate restriction endonuclease. A total of 1 μg (for Pplrl2, Pplrl1 Pplrl2, and Pplrl2pro:GUS) or 2 μg (for Pplrl1 and

Tam et al.

Generation of P. patens GUS-Reporter Lines. The GUS-reporter lines were generated by replacing the endogenous PpLRL1 or PpLRL2 gene with the uidA gene and a hygromycin-resistance cassette from the plasmid pBHrevGUS (pBHSNR-GUS) (9); this strategy for constructing the reporter lines is illustrated in Fig. S4. To generate the GUS reporter lines for PpLRL1, a 871-bp fragment upstream and a 888-bp fragment downstream of the predicted PpLRL1 CDS (Pp1s173_83V6) were cloned into the AscI–NcoI and BamHI site, in the correct orientation, of the pBHrev-GUS plasmid, respectively. Similarly, for the GUS-reporter lines for PpLRL2, a 903-bp fragment upstream and a 880-bp fragment downstream of the predicted CDS (Pp1s209_22V6) were cloned into the AscI–NcoI and BamHI–HindIII site of the pBHrev–GUS plasmid, respectively. These generated the pBHrev-PpLRL1proGUS and pBHrevPpLRL2proGUS constructs (Fig. S4). The plasmids were linearized with AscI (for pBHrev-PpLRL1proGUS) and AscI + HindIII (for pBHrev-PpLRL2proGUS) before PEG-mediated transformation. Stable transformants were selected by PCR analyses and then further selected with Southern blot analysis (Fig. S4). Three independent lines, which show the same GUS-staining pattern, were used for each genotype. Generation of P. patens Loss- and Gain-of-Function Mutants. The plasmids pBHrev (pBHSNR) and pBNRF (10) were used to generate the loss-of-function constructs in this study. To generate the Pplrl1 complete knockout mutants, an 812-bp DNA fragment upstream and a 819-bp fragment downstream of the PpLRL1 were cloned into the SalI–BamHI and MluI–NcoI sites of the pBHrev plasmid, respectively (Fig. S5). Similarly, to generate Pplrl2 knockout mutants, a 697-bp fragment upstream and a 655-bp fragment downstream of the PpLRL2 locus were cloned into the SalI–BamHI and SpeI–AscI site of the pBNRF plasmid, respectively (Fig. S5). These generated the pBHrevPpLRL1KO and pBNRF-PpLRL2KO constructs. The plasmids were linearized with SalI + NcoI (for pBHreb-PpLRL1KO) and SalI + AscI (for pBNRF-PpLRL2KO) before PEG-mediated transformation. The Pplrl1 Pplrl2 double mutants were generated by retransforming Pplrl1-3 with the pBNRF-PpLRL2KO construct. Stable transformants were selected by PCR analyses and then further selected with Southern blot analysis (Fig. S5). Three independent lines, which showed the same phenotype, were used for each genotype. Two independent lines of partial gene-deletion mutants (Pplrl1ΔbL and Pplrl2ΔbL), where only the bHLH and LRL domains are deleted, were generated similarly (Fig. S6) and were PCR-verified. To generate the 35S:PpLRL1bL and 35S:PpLRL2bL mutants, partial CDS for the bHLH–LRL region of the PpLRL genes (Fig. S6) were first amplified by RTPCR and subcloned into the pCR8/GW/TOPO Vector (Invitrogen), then transferred to the p108GW35S vector through the Gateway LR Clonase II system (Invitrogen) for moss overexpression (11). This construct includes the 108 locus to facilitate insertion into the neutral 108 locus in the P. patens genome. The levels of the PpLRL1 and PpLRL2 bHLH–LRL partial transcript in 35S:PpLRLbL lines were determined by quantitative reverse transcription-polymerase chain reaction (qRT-PCR) (Fig. S6).

Time-Lapse Microscopy. Filaments at the edge of the protonema growing on cellophane disk on minimal medium were photographed every 6 min under a Leica DFC310 FX camera mounted on a Leica M165 FC stereomicroscope. Images were analyzed with ImageJ (49). The increase in filament length every 30 min was used to calculate the growth rate. qRT-PCR. Ten-day-old macerated protonema growing on minimal medium was used for all qRT-PCRs. Approximately 100 mg of fresh protonema that had been gently squeezed to remove excess water was frozen in liquid nitrogen before RNA extraction with the RNeasy Plant Mini Kit (Qiagen). The amount of RNA was quantified with a NanoDrop ND1000 spectrometer (Thermo Scientific) or a Qubit 2.0 Fluorometer (Invitrogen) before treatment with Turbo DNase (Ambion, Life Technologies). First-strand cDNA synthesis was performed with the SuperScript III First-Strand Synthesis System for RTPCR (Invitrogen, Life Technologies) by using oligo(dT). The following amount of DNase-treated RNA was used in the cDNA synthesis: 3.5 μg for detecting PpLRL levels in Ppshi1, Ppshi2 and WT; 9.1 μg for detecting PpLRL levels in PpOXSHI1-5 and WT; 1.3 μg for detecting PpLRL levels in WT treated with high and low phosphate; 2.7 μg for detecting PpRSL levels in Pplrl1 Pplrl2 double mutants and WT; and >1 μg for detecting PpLRL levels in Pprsl1 Pprsl2 double mutants and WT. The resulting cDNA reaction mixture (∼20 μL) was diluted 6.5-fold, and 5 μL of the diluted cDNA mixture was used directly in a 20-μL qRT-PCR. qRT-PCR was performed with the SYBR Green PCR Master Mix (Applied Biosystems) and the Applied Biosystems 7300 RealTime PCR system. Three biological replicates, each with three technical replicates, were used for each plant sample. The cycling conditions were as follows: 50 °C for 2 min; 95 °C for 10 min; then 40 cycles of 95 °C for 15 s and 60 °C for 1 min. Dissociation curve analyses were performed at the end of the cycles to ensure the formation of only a single amplicon for each replicate. LinRegPCR (50) was used to estimate the baseline, threshold, and primer efficiencies. The starting concentration of the target transcript in a sample, expressed in arbitrary fluorescence units (N0), was computed from the following formula: N0 = threshold/[(mean efficiency of each primer pair)^ Cq] (50). PpGAPDH (11) was used as the internal reference for all experiments, except for the determination of bHLH–LRL partial transcripts in the 35S:PpLRL1bL and 35S:PpLRL2bL lines, where PpAdePRT was used as the internal references (51). Primers used are given in Table S1. Auxin and Low-Phosphate Treatment. For auxin treatments, plants were grown on KNOPS-GT supplemented with 0, 0.1, or 1 μM NAA dissolved in dimethyl sulfoxide. High-phosphate treatment was carried out by growing plants in KNOPS-GT medium supplemented with 1 mM Mes. The low-phosphate medium (−P) was prepared by replacing the KH2PO4 buffer in the KNOPS-GT medium with 0.918 mM K2SO4.

GUS Staining. Unless otherwise stated, protonema inocula were grown on KNOPS medium with cellophane for 2.5 wk and then directly incubated in GUS staining solution: 100 mM NaH2PO4 (pH 7.0 with NaOH), 0.5 mM K3Fe (CN)6, 0.5 mM K4Fe(CN)6·3H2O, 0.05% Triton X-100, 1 mM 5-bromo-4-chloro-

ACKNOWLEDGMENTS. We thank Dr. Katarina Lundberg from Prof. Eva Sundberg’s group (Swedish University of Agricultural Sciences) for providing the Ppshi1, Ppshi2, and the PpOXSHI1 lines (23); Dr. Krzysztof Szczyglowski (Agriculture and Agri-Food Canada) for providing the Atlrl mutant lines for observation; Nuno Pires for technical assistance and scientific input at the beginning of the project; and Helen Prescott and Lida Chen for laboratory support. T.T.H.Y. was supported by a Clarendon Fund Scholarship and the Overseas Research Students Awards Scheme; B.C. and L.D. were supported by the PLANTORIGINS Marie Curie Network and the EVO500 ERCAdvanced Grant.

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7. Gahoonia T, Nielsen NE (2003) Phosphorus (P) uptake and growth of a root hairless barley mutant (bald root barley, brb) and wild type in low- and high-P soils. Plant Cell Environ 26:1759–1766. 8. Jang G, Dolan L (2011) Auxin promotes the transition from chloronema to caulonema in moss protonema by positively regulating PpRSL1and PpRSL2 in Physcomitrella patens. New Phytol 192(2):319–327. 9. Jang G, Yi K, Pires ND, Menand B, Dolan L (2011) RSL genes are sufficient for rhizoid system development in early diverging land plants. Development 138(11): 2273–2281. 10. Menand B, et al. (2007) An ancient mechanism controls the development of cells with a rooting function in land plants. Science 316(5830):1477–1480. 11. Pires ND, et al. (2013) Recruitment and remodeling of an ancient gene regulatory network during land plant evolution. Proc Natl Acad Sci USA 110(23):9571–9576.

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PNAS PLUS

3-indolyl-β-D-glucuronic acid, and cyclohexyl ammonium salt (X-GlcA; Melford) dissolved in N,N-dimethylformamide at 37 °C for 24–48 h. Tissues were then destained by incubation in 70–100% (vol/vol) ethanol for at least 24 h. No staining was observed in WT plants.

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PLANT BIOLOGY

PpLRL1pro:GUS) of digested genomic DNA, as quantified with a NanoDrop ND1000 spectrometer (Thermo Scientific), was subjected to agarose gel electrophoresis before being transferred to a positively charged nylon membrane. Southern blots were carried out with the digoxigenin (DIG) system for filter hybridization with DIG-labeled probe generated by PCR according to the DIG Application Manual for Filter Hybridization (Roche). DIG-labeled DNA Molecular Weight Marker VII (Roche) was used throughout the analysis. Primers for probe synthesis are given in Table S1.

12. Yi K, Menand B, Bell E, Dolan L (2010) A basic helix-loop-helix transcription factor controls cell growth and size in root hairs. Nat Genet 42(3):264–267. 13. Karas B, et al. (2009) Conservation of lotus and Arabidopsis basic helix-loop-helix proteins reveals new players in root hair development. Plant Physiol 151(3): 1175–1185. 14. Bruex A, et al. (2012) A gene regulatory network for root epidermis cell differentiation in Arabidopsis. PLoS Genet 8(1):e1002446. 15. Ding W, et al. (2009) A transcription factor with a bHLH domain regulates root hair development in rice. Cell Res 19(11):1309–1311. 16. Yi K, et al. (2005) OsPTF1, a novel transcription factor involved in tolerance to phosphate starvation in rice. Plant Physiol 138(4):2087–2096. 17. Pagnussat GC, et al. (2005) Genetic and molecular identification of genes required for female gametophyte development and function in Arabidopsis. Development 132(3): 603–614. 18. Pires N, Dolan L (2010) Origin and diversification of basic-helix-loop-helix proteins in plants. Mol Biol Evol 27(4):862–874. 19. Duckett JG, Schmid AM, Ligrone R (1998) Protonema morphogenesis. Bryology for the Twenty-First Century, eds Bates JW, Ashton NW, Duckett JG (Maney, Leeds, U.K.). 20. Lavy M, Prigge MJ, Tigyi K, Estelle M (2012) The cyclophilin DIAGEOTROPICA has a conserved role in auxin signaling. Development 139(6):1115–1124. 21. Prigge MJ, Lavy M, Ashton NW, Estelle M (2010) Physcomitrella patens auxin-resistant mutants affect conserved elements of an auxin-signaling pathway. Curr Biol 20(21): 1907–1912. 22. Ashton NW, Grimsley NH, Cove DJ (1979) Analysis of gametophytic development in the moss, Physcomitrella patens, using auxin and cytokinin resistant mutants. Planta 144(5):427–435. 23. Eklund DM, et al. (2010) Homologues of the Arabidopsis thaliana SHI/STY/LRP1 genes control auxin biosynthesis and affect growth and development in the moss Physcomitrella patens. Development 137(8):1275–1284. 24. Eklund DM, et al. (2010) The Arabidopsis thaliana STYLISH1 protein acts as a transcriptional activator regulating auxin biosynthesis. Plant Cell 22(2):349–363. 25. Wang Y, Secco D, Poirier Y (2008) Characterization of the PHO1 gene family and the responses to phosphate deficiency of Physcomitrella patens. Plant Physiol 146(2): 646–656. 26. Wickett NJ, et al. (2014) Phylotranscriptomic analysis of the origin and early diversification of land plants. Proc Natl Acad Sci USA 111(45):E4859–E4868. 27. Finet C, Timme RE, Delwiche CF, Marlétaz F (2010) Multigene phylogeny of the green lineage reveals the origin and diversification of land plants. Curr Biol 20(24): 2217–2222, and erratum (2012) 22(15):1456–1457. 28. Timme RE, Bachvaroff TR, Delwiche CF (2012) Broad phylogenomic sampling and the sister lineage of land plants. PLoS ONE 7(1):e29696. 29. Wodniok S, et al. (2011) Origin of land plants: Do conjugating green algae hold the key? BMC Evol Biol 11:104. 30. Clarke JT, Warnock RCM, Donoghue PCJ (2011) Establishing a time-scale for plant evolution. New Phytol 192(1):266–301. 31. Sakakibara K, et al. (2003) Involvement of auxin and a homeodomain-leucine zipper I gene in rhizoid development of the moss Physcomitrella patens. Development 130(20):4835–4846.

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32. Cove D, Bezanilla M, Harries P, Quatrano R (2006) Mosses as model systems for the study of metabolism and development. Annu Rev Plant Biol 57:497–520. 33. Menand B, Calder G, Dolan L (2007) Both chloronemal and caulonemal cells expand by tip growth in the moss Physcomitrella patens. J Exp Bot 58(7):1843–1849. 34. Vidali L, Bezanilla M (2012) Physcomitrella patens: A model for tip cell growth and differentiation. Curr Opin Plant Biol 15(6):625–631. 35. Rounds CM, Bezanilla M (2013) Growth mechanisms in tip-growing plant cells. Annu Rev Plant Biol 64(1):243–265. 36. Pressel S, Ligrone R, Duckett JG (2008) Cellular differentiation in moss protonemata: A morphological and experimental study. Ann Bot (Lond) 102(2):227–245. 37. Goodstein DM, et al. (2012) Phytozome: A comparative platform for green plant genomics. Nucleic Acids Res 40(Database issue, D1):D1178–D1186. 38. Katoh K, Standley DM (2013) MAFFT multiple sequence alignment software version 7: Improvements in performance and usability. Mol Biol Evol 30(4):772–780. 39. Capella-Gutiérrez S, Silla-Martínez JM, Gabaldón T (2009) trimAl: A tool for automated alignment trimming in large-scale phylogenetic analyses. Bioinformatics 25(15):1972–1973. 40. Sánchez R, et al. (2011) Phylemon 2.0: A suite of web-tools for molecular evolution, phylogenetics, phylogenomics and hypotheses testing. Nucleic Acids Res 39(Web Server issue, suppl 2):W470–W474. 41. Amborella Genome Project (2013) The Amborella genome and the evolution of flowering plants. Science 342(6165):1241089. 42. Stothard P (2000) The sequence manipulation suite: JavaScript programs for analyzing and formatting protein and DNA sequences. Biotechniques 28(6):1102–1104, 1104. 43. Darriba D, Taboada GL, Doallo R, Posada D (2011) ProtTest 3: Fast selection of best-fit models of protein evolution. Bioinformatics 27(8):1164–1165. 44. Guindon S, et al. (2010) New algorithms and methods to estimate maximum-likelihood phylogenies: assessing the performance of PhyML 3.0. Syst Biol 59(3):307–321. 45. Anisimova M, Gascuel O (2006) Approximate likelihood-ratio test for branches: A fast, accurate, and powerful alternative. Syst Biol 55(4):539–552. 46. Ronquist F, et al. (2012) MrBayes 3.2: Efficient Bayesian phylogenetic inference and model choice across a large model space. Syst Biol 61(3):539–542. 47. Ashton NW, Cove DJ (1977) The isolation and preliminary characterisation of auxotrophic and analogue resistant mutants of the moss, Physcomitrella patens. Mol Gen Genet 154(1):87–95. 48. Schaefer DG, Zrÿd JP (1997) Efficient gene targeting in the moss Physcomitrella patens. Plant J 11(6):1195–1206. 49. Schneider CA, Rasband WS, Eliceiri KW (2012) NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9(7):671–675. 50. Ruijter JM, et al. (2009) Amplification efficiency: Linking baseline and bias in the analysis of quantitative PCR data. Nucleic Acids Res 37(6):e45. 51. Le Bail A, Scholz S, Kost B (2013) Evaluation of reference genes for RT qPCR analyses of structure-specific and hormone regulated gene expression in Physcomitrella patens gametophytes. PLoS ONE 8(8):e70998.

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Conserved regulatory mechanism controls the development of cells with rooting functions in land plants.

Land plants develop filamentous cells-root hairs, rhizoids, and caulonemata-at the interface with the soil. Members of the group XI basic helix-loop-h...
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