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Conserved regulatory mechanism controls thedevelopment of cells
with rooting functionsin land plantsThomas Ho Yuen Tam, Bruno
Catarino, and Liam Dolan1
Department of Plant Sciences, University of Oxford, Oxford OX1
3RB, United Kingdom
Edited by Philip N. Benfey, Duke University, Durham, NC, and
approved June 11, 2015 (received for review August 22, 2014)
Land plants develop filamentous cells—root hairs, rhizoids,
andcaulonemata—at the interface with the soil. Members of thegroup
XI basic helix–loop–helix (bHLH) transcription factors en-coded by
LOTUS JAPONICUS ROOTHAIRLESS1-LIKE (LRL) genespositively regulate
the development of root hairs in the angio-sperms Lotus japonicus,
Arabidopsis thaliana, and rice (Oryza sat-iva). Here we show that
auxin promotes rhizoid and caulonemadevelopment by positively
regulating the expression of PpLRL1and PpLRL2, the two LRL genes in
the Physcomitrella patens ge-nome. Although the group VIII bHLH
proteins, AtROOT HAIRDEFECTIVE6 and AtROOT HAIR DEFECTIVE
SIX-LIKE1, promote root-hair development by positively regulating
the expression of AtLRL3in A. thaliana, LRL genes promote rhizoid
development indepen-dently of PpROOT HAIR DEFECTIVE SIX-LIKE1 and
PpROOT HAIRDEFECITVE SIX-LIKE2 (PpRSL1 and PpRSL2) gene function
inP. patens. Together, these data demonstrate that both LRL andRSL
genes are components of an ancient auxin-regulated genenetwork that
controls the development of tip-growing cells withrooting functions
among most extant land plants. Although this net-work has diverged
in the moss and the angiosperm lineages, our datademonstrate that
the core network acted in the last common ancestorof themosses and
angiosperms that existed sometime before 420mil-lion years ago.
auxin | bHLH | evolution | rhizoids | root hairs
The evolution of rooting structures was a key
morphologicalinnovation that occurred when plants colonized the
relativelydry continental surfaces of the planet sometime before
470 millionyear ago. The rooting structures of the earliest
diverging groups ofland plants comprised systems of rhizoids.
Rhizoids are eitherunicellular filaments (in liverworts and
hornworts) or multicel-lular (in mosses) and elongate into the
growth substrate or airsurrounding the plant. Bryophyte (liverwort,
moss, and hornwort)rhizoids develop on gametophytes, the
multicellular haploid stagein the life cycle. The evolution of
vascular plants was accompaniedby 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 leastonce among the Lycophytes and at
least once in the Euphyllo-phyte clade (2). Roots generally grow
into the soil and expandthe plant–soil interface into different
soil horizons. For example,some root systems extend deep into the
soil (taproots), whereasothers proliferate in the nutrient-rich
horizons near the soil sur-face. However, unicellular, filamentous
protuberances called roothairs, which are morphologically similar
to rhizoids, develop onall roots with few exceptions (2–5). Despite
the different contextsin which they develop, root hairs and
rhizoids carry out similarrooting functions, including nutrient
uptake and anchorage (6, 7).Group VIII basic helix–loop–helix
(bHLH) transcription fac-
tors encoded by ROOT HAIR DEFECTIVE SIX-LIKE (RSL)genes
positively regulate root-hair development in the angio-sperm
Arabidopsis thaliana and both rhizoid and caulonema in
the moss Physcomitrella patens (8–12). To test whether the
con-servation of RSL function is unique or indicative of a more
gen-eral, conserved regulatory mechanism, we determined
whetherother components of the gene-regulatory network controlling
an-giosperm root-hair development are conserved among land
plants.Group XI bHLH transcription factors encoded by
LOTUSJAPONICUS ROOTHAIRLESS1-LIKE (LRL) genes are alsokey
regulators of root-hair development in angiosperms. LRLgenes encode
several proteins that regulate root-hair devel-opment in diverse
angiosperms, including ROOTHAIRLESS1(LjRHL1) in Lotus japonicus,
AtLRL1, AtLRL2, AtLRL3 inA. thaliana (13, 14), and ROOTHAIRLESS1
(OsRHL1) in Oryzasativa (15). Other LRL genes include OsPTF1,
involved inphosphate starvation tolerance in rice (16), and
AtUNE12, in-volved in female gametophyte development in
Arabidopsis(17). Here we define the function of LRL genes in the
mossP. patens and conclude that, together, LRL and RSL genes
formthe core of an ancient network that operated in the
commonancestor of the mosses and angiosperms that existed in
theSilurian Period 415–435 million years ago.
ResultsTwo LRL Genes Are Present in the Genome of the Moss P.
patens. TheLRL transcription factors belong to group XI of plant
bHLHtranscription factors (18) and share a conserved LRL domain
of36 amino acids between the bHLH domain and the C terminus(Fig.
1A). To identify LRL homologs in P. patens, we performedBLAST
searches in selected land plant genomes using theAtLRL3 (At5g58010)
sequence as a query. Phylogenetic anal-yses identified two LRL
genes in P. patens, designated PpLRL1
Significance
This work describes the discovery of an ancient genetic
mech-anism that was used to build rooting systems when plants
col-onized the relatively dry continental surfaces >470 million
yearsago. We demonstrate that a group of basic
helix–loop–helixtranscription factors—the LOTUS JAPONICUS
ROOTHAIRLESS1-LIKE proteins—is part of a conserved auxin-regulated
genenetwork that controls the development of tip-growing cells
withrooting functions among extant land plants. This result
suggeststhat this mechanism was active in the common ancestor of
mostland plants and facilitated the development of early land
plantfilamentous rooting systems, crucial for the successful
coloniza-tion of the land by plants.
Author contributions: T.H.Y.T. and L.D. designed research;
T.H.Y.T. and B.C. performedresearch; T.H.Y.T., B.C., and L.D.
analyzed data; and T.H.Y.T., B.C., and L.D. wrotethe paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Freely available online through the PNAS open access option.1To
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.
www.pnas.org/cgi/doi/10.1073/pnas.1416324112 PNAS | Published
online July 6, 2015 | E3959–E3968
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(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 proteinsincluding OsRHL1
(Os06g08500), and four Selaginella moellen-dorffii proteins into a
monophyletic group (Fig. 1B, Fig. S1, andDataset S1). The LRL
domains of PpLRL1 and PpLRL2 are97.2% identical to each other and
are 77.8% and 80.6% identicalto the LRL domain of the AtLRL3
protein, respectively. The twoPpLRL proteins have conserved bHLH
domains that are 100%identical to each other (Fig. 1A). The bHLH
domains of thePpLRL proteins are 92.4% identical to the bHLH domain
ofthe AtLRL3 protein. To infer the phylogenetic relationship of
thedifferent proteins containing the LRL domain, we
constructedphylogenetic trees using Bayesian and ML methods (Fig.
1C andFig. S2). These trees showed that LRL homologs are present in
all32 land plant species sampled (Fig. S3). The LRL proteins
com-prise three major groups, designated XIa, XIb, and XIc (Fig.
1Cand Fig. S2). Genes that control the development of root hairs
arepresent in group XIa (Fig. 1C, Fig. S2, and Dataset S2).
PpLRL1and PpLRL2 are not members of any of these three groups
(Fig.1C and Figs. S2 and S3). Together, these results indicate that
theLRL genes are a conserved group of transcription factors
whoseexistence predates the divergence of mosses and flowering
plants.
PpLRL Genes Have Different, but Overlapping, Expression
Patterns.We hypothesized that LRL genes controlled the development
ofrhizoids and caulonemata and that PpLRL1 and PpLRL2 wouldbe
expressed in these cell types. To define where LRL genes
areexpressed, we replaced the endogenous PpLRL1 and PpLRL2genes
with the coding sequence (CDS) of the UidA glucuroni-dase-encoding
gene (GUS) (Fig. S4). Consequently, the GUSexpression in these
lines identified regions of the protonema inwhich the PpLRL1 and
PpLRL2 promoters are active in re-spective loss-of-function Pplrl1
or Pplrl2 mutant background. InP. patens, the germination of spores
is followed by the development
of a uniseriate filamentous network (protonema) comprisingcells
with large chloroplasts and transverse cell walls (chlor-onema
cells). Subsequently, increasingly longer cells with obliquecell
walls (caulonema cells) develop. Caulonema cells are
mor-phologically and genetically similar to rhizoids (19).
BothPpLRL1 and PpLRL2 were expressed in the central region of
theprotonema, which is rich in chloronema and from which caulo-nema
develop (Fig. 2 A and B). No expression was detected inthe very
edges of the 30-d-old protonema, where caulonema isthe dominant
cell type. These observations suggest that PpLRL1and PpLRL2 are
active in cells that develop caulonema.PpLRL1 promoter was also
active in the rhizoid-forming re-
gion at the base of the gametophore in plants in which thePpLRL1
CDS was replaced by the GUS-reporter gene (Fig. 2 Cand D). However,
no GUS expression was detected in rhizoidsthemselves. This
expression pattern is consistent with a role forPpLRL1 in the
regulation of the earliest stages of rhizoid de-velopment. PpLRL1
promoter was also active in the gameto-phore apex and leaves,
suggesting that this gene likely controlsother aspects of
gametophore development. PpLRL2 promoteractivity was detected in
the epidermal cells of the gametophoreand leaves (Fig. 2 C and D).
Unlike PpLRL1, no GUS activitywas detectable in the buds or at the
base of the gametophore.Like PpLRL1, there was no detectable
expression of PpLRL2promoter in rhizoids.
PpLRL1 and PpLRL2 Are Positive Regulators of Rhizoid
Development.To determine whether PpLRL genes function in rhizoid
de-velopment, we generated mutants that lack LRL gene functionby
homologous recombination (Figs. S5 and S6). To confirm
thathomologous recombination had taken place, we characterizedthe
genomic organization of the transformants. PCR analyseswere
performed to select transformants in which the endogenousPpLRL gene
was replaced by the gene-targeting construct (Fig.S5) and thus
represent complete loss-of-function alleles. Fromthese, Southern
blot analyses were used to further select those
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 theclass 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 inFig. S1. (C) Unrooted
tree showing the relationships of different LRL proteins in land
plants using Bayesian statistics. The location of selected LRL
proteinsfrom 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 areshown in Fig. S2.
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transformants in which there were no additional transgene
insertionsin the genome (Fig. S5). Three independent lines for each
genewere maintained for phenotypic analyses. With the exception
ofPplrl2-1, all single-mutant lines and all three double-mutant
linesshowed a single band in the Southern blot analysis,
indicatingthat 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
rhizoidswere ∼35% and 18% shorter than WT, respectively (Fig.
3C).Pplrl1 Pplrl2 double mutants were completely rhizoidless, and
norhizoids developed on these plants (Fig. 3D). Together, thesedata
show that the activity of PpLRL1 and PpLRL2 together arerequired
for the initiation and development of rhizoids frombuds and
gametophores.To determine whether the bHLH and LRL domains of
the
LRL proteins—characteristic of group XI plant bHLHs—arerequired
for the control of rhizoid development in P. patens, wegenerated
additional mutants in which the region containing thebHLH–LRL
domain of each protein was deleted (Fig. S6). Theorganization of
the genomic DNA in the mutants confirmed thatthe bHLH–LRL domains
were deleted (Fig. S6). These mutantswere designated Pplrl1ΔbL and
Pplrl2ΔbL, respectively. Defectsin rhizoid and caulonema
development in these partial-deletionmutants were similar to the
defects in the complete gene-deletion mutants (Fig. 3E). To
independently verify that the bHLHand LRL domains are positive
regulators of rhizoid develop-ment, we overexpressed a fragment of
the PpLRL1 and PpLRL2CDS 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
CaMV35Spromoter (Fig. S6). Plants transformed with
35S:PpLRL1bL-1,which overexpressed the bHLH–LRL fragment of PpLRL1,
de-veloped longer and more intensively pigmented rhizoids thanWT
(Fig. 3E). In contrast, rhizoid pigmentation in
35S:PpLRL2bL-1–transformed plants, which overexpressed the bHLH–LRL
fragmentof PpLRL2, resembled WT (Fig. 3E and Fig. S6). This
difference inrhizoid phenotype in these lines suggests that PpLRL1
plays amore important role than PpLRL2 in rhizoid
development.Together, these data are consistent with the hypothesis
that thebHLH and LRL domains are responsible for PpLRL genefunction
in rhizoid development.
PpLRL Genes Positively Regulate Caulonema Development.
Thetransition from chloronema to caulonema, characteristic of
WTprotonema development, did not occur in Pplrl1 Pplrl2 double-
mutant plants (Fig. 4 A and B). Caulonemata did not develop,and,
instead, a dense mat of chloronemata constituted the
entireprotonema (Fig. 4A). The diameter of the Pplrl1 Pplrl2
double-mutant protonema was approximately half that of the WT,
whichcan be explained by the absence of caulonemata, because
thesecells have higher growth rates and are longer than
chloronemata(Fig. 4 B and C). To verify that the smaller diameter
of Pplrl1Pplrl2 mutants was due to the loss of caulonemata, we
comparedthe 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.
Thegrowth rates of protonema filaments in Pplrl1 Pplrl2
doublemutants were 8.48 ± 0.81 μm·h−1. That is, the growth rates
ofprotonema filaments of the Pplrl1 Pplrl2 double mutant were
notsignificantly different from those of WT chloronema. These
dataindicate that chloronemata, but not caulonemata, develop
inPplrl1 Pplrl2 double mutants. Pplrl1 and Pplrl2 single
mutantsdeveloped less severe phenotypes than the Pplrl1 Pplrl2
doublemutants. Protonema diameters of Pplrl1 and Pplrl2 mutants
were83% and 89% that of WT, respectively (Fig. 4 A and B).
To-gether, these data demonstrate that PpLRL1 and PpLRL2
pos-itively regulate the developmental transition from chloronema
tocaulonema during the development of moss protonemata.
Auxin Positively Regulates PpLRL Activity. Auxin positively
regu-lates the development of rhizoids and caulonemata in P.
patens(8, 9, 20–22). To test whether auxin promotes rhizoid and
cau-lonema development by positively regulating the expression
ofPpLRL genes, we determined the effect of auxin treatment onthe
activities of the PpLRL1 and PpLRL2 promoters. P. patensplants in
which the endogenous PpLRL CDS were replaced bythe GUS gene (Fig.
S4) were grown on Knops-GT medium(defined in Plant Material, Growth
Conditions, and PhenotypicAnalyses) for 30 d. No GUS expression was
detectable in theprotonema or buds, even when incubated in
glucuronidase re-action buffer for 24 h (Fig. 5A). Addition of
1-naphthaleneaceticacid (NAA) to the medium at a final
concentration of 100 nMresulted in detectable GUS expression in
both protonema andbuds of PpLRL1pro:GUS and PpLRL1pro:GUS plants
(Fig. 5A);this indicates that exogenous treatment with auxin
increases theactivity of the PpLRL1 and PpLRL2 promoters.
Together,these data suggest that auxin positively regulates
PpLRL1and PpLRL2 expression.If auxin positively regulates LRL
expression, and LRL genes in
turn promote caulonema development, we hypothesized that
Fig. 2. GUS staining pattern in plants transformed with
PpLRL1pro:GUS and PpLRL2pro:GUS and untransformed controls (WT). (A
and B) PpLRL1 and PpLRL2were expressed in the center of the
protonema, composed mainly of chloronema filaments. (C and D) Young
buds and gametophores isolated from 4-wk-oldprotonemata grown on
KNOPS-GT medium. PpLRL1, but not PpLRL2,was expressed in the early
buds. In the gametophores, PpLRL1was 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 thegametophores, but was expressed in the leaves.
[Scale bars: 2 mm (A), 1 mm (B and D), and 200 μm (C).]
Tam et al. PNAS | Published online July 6, 2015 | E3961
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Fig. 3. Rhizoid phenotypes of PpLRL loss- and gain-of-function
mutants. (A) Rhizoid phenotypes of complete PpLRL loss-of-function
mutants. (Scalebars: 2 mm.) (B) Quantification of rhizoid number.
(C ) Quantification of rhizoid length. Rhizoid length was
determined by measuring the length be-tween the base of the
gametophore and the tip of the rhizoid cluster. Both rhizoid number
and length were significantly reduced in the Pplrl1 andPplrl2
single mutants. Rhizoids are absent in the Pplrl1 Pplrl2 double
mutant. Asterisks represent P values from two-tailed unequal
variance (Welch’s) ttests 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 ) Phenotypesof
PpLRLΔbL partial-deletion mutants and 35S:PpLRLbL mutants. The
35S:PpLRL1bL-1 mutant showed an increased number of rhizoids and
increasedpigmentation.
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auxin-induced morphological changes in caulonema would re-quire
PpLRL1 and PpLRL2 activity. To test this hypothesis, wecompared the
morphology of Pplrl1 Pplrl2 double mutants grownin medium
containing 100 nM NAA with double mutants grownwithout added auxin.
Pplrl1 Pplrl2 protonema that developedin the presence of 100 nM NAA
were morphologically identicalto double mutants grown without auxin
(Fig. 5B). That is, Pplrl1Pplrl2 mutants were resistant to auxin.
Furthermore, auxin in-duced the development of rhizoids on buds of
WT, but rhizoidinduction did not occur on the buds of Pplrl1 Pplrl2
doublemutants (Fig. 5C). Together, these data demonstrate that
rhizoidand caulonema development in plants that lack both PpLRL1and
PpLRL2 function are resistant to auxin. This result is con-sistent
with the model in which auxin-regulated caulonema andrhizoid
development require PpLRL1 and PpLRL2 activity.The STYLISH1/SHI
transcription factors regulate auxin bio-
synthesis in both A. thaliana and P. patens (23, 24). To
testwhether auxin promotes the expression of PpLRL1 and PpLRL2in a
PpSHI-dependent fashion in P. patens, we quantified mRNAlevels in
the auxin biosynthesis mutants Ppshi1, PpOXSHI1-5,and Ppshi2. We
could not detect a difference in PpLRL1 orPpLRL2 mRNA levels
between WT and these mutants (Fig. S7
A–C). This result suggests that auxin-regulated expression
ofPpLRL1 and PpLRL2 is independent of the STYLISH/SHItranscription
factors.
PpLRL1 and PpLRL2 Are Required for
Low-Phosphate-InducedDevelopment of Caulonema. We hypothesized that
PpLRL geneswould be required for the adaptive response of protonema
tolow phosphate; growth of protonema in medium containinglow
phosphate promotes the transition from chloronema tocaulonema (25).
To test this hypothesis, we compared thephenotypes of Pplrl1 Pplrl2
double mutants with WT in mediawith different phosphate content.
Low phosphate promotes thechloronema-to-caulonema transition in WT
protonema, but theprotonema morphology of Pplrl1 Pplrl2 double
mutants wasidentical in replete and no phosphate (Fig. 6 A and B).
That is,the double mutant protonema was insensitive to changes
inexternal phosphate concentration, and caulonema did not de-velop.
These results suggest that the morphological changesthat take place
as part of the adaptive response to low phos-phate require PpLRL1
and PpLRL2 activity. However, wecould not detect a difference in
PpLRL1 and PpLRL2 mRNAlevels in different phosphate conditions
(Fig. S7D), suggesting
Fig. 4. Protonema phenotypes of PpLRL loss-of-function mutants.
(A) Caulonema phenotype of complete PpLRL loss-of-function mutants.
Caulonema de-velopment 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
Pplrl1Pplrl2). (C) Growth rate of caulonema and chloronema
filaments in the loss-of-function mutants. In all of the mutant
lines, the growth rate of chloronemafilaments was not significantly
different from WT, but the growth rate of caulonema was
significantly reduced. Asterisks represent the 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: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
forchloronema; n = 5 for Pplrl1 and Pplrl2 caulonema; and n = 3 for
WT caulonema).
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that the induction of caulonemal development in low-phos-phate
media does not increase the expression of LRL genes inP.
patens.
PpLRL and PpRSL Genes Act Independently in P. patens. The
groupVIII basic helix–loop–helix transcription factors, AtROOT
HAIRDEFECTIVE6 (AtRHD6) and AtROOT HAIR DEFECTIVE6-LIKE1 (AtRSL1),
promote the development of root hairsby positively regulating
AtLRL3 expression (13, 14). To de-termine whether there are
regulatory interactions betweenPpRSL and PpLRL genes, we compared
the expression of PpLRL1and PpLRL2 in Pprsl1 Pprsl2 double mutants
and WT, and theexpression of PpRSL1 and PpRSL2 in Pplrl1 Pplrl2
double mu-tants with WT. We found no evidence of transcriptional
regu-latory interactions between PpLRL and PpRSL genes in P.
patens(Fig. S7 E and F). Together, these data suggest that the LRL
andRSL genes are transcriptionally independent of each other inP.
patens.
DiscussionThe colonization of the land by streptophyte plants
sometimebefore 470 million years ago was a pivotal event in the
history ofthe Earth. Structures that anchor plants to their growth
substrateand provide access to mineral nutrients, water, and the
soil mi-croflora were key adaptations to life on the relatively dry
con-tinental surfaces. Recent phylogenetic analyses suggest that
therhizoidless algae of the Zygnematales or Colecochaetales (or
aclade consisting of Zygnematales and Coleochaetales) are sisterto
the land plants (26–29). If a clade consisting of Zygnematalesand
Coleochaetales is sister to land plants, then there are twoequally
parsimonious models for the evolution of rhizoids, eachinvolving
two changes: (i) rhizoids evolved independently in thecommon
ancestor of land plants and in the more distantly relatedalgal
lineages such as the Charales or (ii) rhizoids evolved in thecommon
ancestor of Charales, Zygnematales, Coleochaetales,and land plants,
but were lost in the common ancestor of theZygnematales and
Coleochaetales.
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
protonemagrown with 0.1 μM NAA. Auxin failed to induce rhizoid
development in the Pplrl1 Pplrl2 double mutants. (Scale bars: 500
μm.)
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We show here that LRL genes positively regulate the devel-opment
of the tip-growing rhizoids and caulonema in the mossP. patens. LRL
genes also positively regulate the developmentof root hairs in
angiosperms (13–15). Given that similar proteinscontrol the
development of filamentous rooting cells at theplant–soil interface
in mosses and angiosperms, we conclude thatLRL genes positively
regulated the development of tip-growingrooting cells in the common
ancestor of mosses and angiospermsthat 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
develop-ment. The class I RSL genes, AtRHD6 and AtRSL1,
promoteroot-hair development in A. thaliana; root hairs do not
developon Atrhd6 Atrsl1 double loss-of-function mutants (10).
AtRHD6and AtRSL1 positively regulate the expression of the class II
RSLgenes AtRSL2, AtRSL3, and AtRSL4 and the LRL gene AtLRL3;these
in turn modulate late stages of root-hair differentiation inA.
thaliana (12). AtRSL4 is a class II RSL gene that is necessaryand
sufficient for growth. Modulation of AtRSL4 expression byclass I
RSL genes, environment, and auxin determine root-haircell size.
AtRSL4 controls cell size by controlling the expressionof genes
that encode proteins involved in tip growth (12). Class IRSL genes
in P. patens—PpRSL1 and PpRSL2—positively reg-ulate rhizoid and
caulonema development (8–10). Class II RSLgenes—PpRSL3, PpRSL4,
PpRSL5, and PpRSL6—regulatecaulonema development, but have much
more subtle functionsthan class I RSL genes (11).
Our results demonstrate that the LRL and RSL genes representcore
components of a conserved gene-regulatory network that waspresent
in the common ancestor of mosses and angiosperms.However, although
we showed that the genetic components ofthis gene-regulatory
network have been conserved, the detailedpatterns of
transcriptional regulation within this network remainto be
resolved. For example, AtRHD6 and AtRSL1 (class I RSLgenes)
regulate the expression of AtLRL3 in A. thaliana (13, 14),but we
could not find evidence that PpRSL genes control PpLRLgene
expression in P. patens. If there is a genuine absence
oftranscriptional regulation between LRL and class I RSL genes inP.
patens, then two alternative evolutionary scenarios can
besuggested. Either (i) class I RSL genes acquired the ability
toregulate the expression of LRL genes in the lineage that gave
riseto angiosperm, or (ii) RSL-regulated LRL expression was
anancestral 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
auxin-signaling pathway that is conserved among land plants (8,
9,11, 20–23, 31). We showed here that auxin positively regulatesthe
expression of PpLRL1 and PpLRL2 and that the PpLRLgenes are
necessary for auxin-induced caulonema and rhizoiddevelopment.
Therefore, the observed GUS activity in thePpLRL1pro:GUS and
PpLRL2pro:GUS protonema may be dueto higher auxin concentration in
the center of the protonema.Previously, it was shown that auxin
promotes caulonema andrhizoid development by positively regulating
the expression ofthe class I PpRSL genes, PpRSL1 and PpRSL2 (8, 9).
We showedhere that auxin positively regulates PpLRL expression.
However,because we could not find evidence that class I PpRSL
genesregulate the expression of PpLRL genes, or that PpLRL
genesregulate the expression of PpRSL genes in P. patens, these
datasuggest that auxin positively regulates class I PpRSL and
PpLRLgenes independently.According to the classical P. patens
literature (22, 32), game-
tophores form only on caulonema. Our results suggest that
budformation can occur in chloronema. The morphology, chloro-plast
density, and growth rate of the Pplrl1 Pplrl2 protonemaresemble WT
chloronema, but buds develop from this mutant.Similarly, the
caulonema-less Pprsl1 Pprsl2 loss-of-function mu-tants (8, 33) also
develop buds from chloronema. Together, thesedata suggest that the
bud-formation process in P. patens is notlimited to caulonema, but
can develop from chloronema in certaingenetic backgrounds.LRL genes
are not general regulators of tip growth. There are
three types of tip-growing cells in P. patens: chloronema,
caulo-nema, and rhizoid. The caulonemata are cytologically similar
torhizoids (33–36). PpLRL genes positively control both rhizoidand
caulonema development, but not chloronema development;chloronema
development is indistinguishable from WT in Pplrl1Pplrl2 double
mutants. These results are consistent with theobservation that
caulonemata and rhizoids also have similar re-sponses to auxin and
low phosphate, and these responses aredifferent from the response
of chloronema to auxin (8, 9, 25).Together, these data suggest that
the PpLRL genes are specificregulators of the development of
tip-growing cells with a rootingfunction. Similarly, RSL genes are
specific regulators of caulo-nema and rhizoid development and do
not regulate chloronemadevelopment (8–10). Therefore, we concluded
that LRL andRSL genes specifically regulate rhizoid and caulonema
devel-opment and are not simply general regulators of tip growth.
Thisconclusion is supported by the finding that the tip-growth
de-fect in Atrsl1 Atrsl2 double mutants is restricted to root hairs
inA. thaliana; pollen tube development is identical to WT in
thesedouble mutants (10).LRL transcription factors are ancient and
were present in the
last common ancestor of mosses and angiosperms that was extantat
least 420 million years ago. Bayesian analysis indicates that
Fig. 6. PpLRL1 and PpLRL2 function are required for low
phosphateadaptive response. (A) Pplrl1 Pplrl2 double mutants are
insensitive to lowphosphate. Protonemata were grown in high (+P)
and low (−P) phosphatemedium for 2.5 wk. (Scale bars: 5 mm.) (B)
Low phosphate increased cau-lonema development significantly in WT
protonemata. *P = 0.034 [two-tailed unequal variance (Welch’s) t
tests)] but had no discernable effect onPplrl1 Pplrl2 mutants.
Error bars represent SEM (n = 20 for WT+P and Pplrl1Pplrl2 +P; n =
16 for WT-P; and n = 12 for Pplrl1 Pplrl2 −P).
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three monophyletic clades (XIa, XIb, and XIc) constitute
themajority of land plant LRL proteins. However, PpLRL proteinsare
not members of groups XIa, XIb, or XIc. Groups XIb andXIc each
contain a single gymnosperm LRL protein that is sisterto the other
angiosperm LRL proteins in each of these mono-phyletic groups,
suggesting that groups XIb and XIc originatedbefore the divergence
of gymnosperms and angiosperms (Figs.S2 and S3). The evolution of
group XIa LRL protein is morecomplex (Figs. S2 and S3), but the
presence of a group XIa LRLproteins in Amborella and other
angiosperm taxa suggests thatthis group was present in the common
ancestor of extant an-giosperms. Only group XIa LRL proteins are
known to regulateroot-hair development in angiosperms. These
results support ourconclusion that the role of LRL proteins in
regulating filamen-tous rooting cell development is ancient.
Further sampling ofLRL genes in well-annotated, high-quality
assemblies of mon-ilophyte (ferns and horsetails) and gymnosperm
genomes willallow the elucidation of the evolutionary history of
LRL genesduring land plant evolution.There is evidence that PpLRL1
plays a greater role than
PpLRL2 in rhizoid development. First, the PpLRL1 and
PpLRL2expression patterns are different. The strong expression
ofPpLRL1 in chloronema cells, young buds, and the base of
thegametophore is consistent with its role in controlling early
stagesof caulonema and rhizoid development. In contrast,
PpLRL2promoter activity was not detected in either young buds or
thebase of gametophores where rhizoids develop, suggesting
thatPpLRL2 regulates rhizoid development via a
nonautonomousmechanism. Second, Pplrl1 mutants have a stronger
rhizoid de-fect than Pplrl2 single mutants. Third, overexpression
of thePpLRL1bL partial transcript led to an increased number
ofrhizoids, whereas overexpression of the PpLRL2bL
partialtranscript had no discernable effect on rhizoid
development.Nevertheless, the activity of both proteins is required
for thedevelopment of rhizoids because only the double mutant
doesnot form rhizoids. Together these results suggest that,
althoughboth PpLRL1 and PpLRL2 function in rhizoid
development(because the double mutant does not form rhizoids),
PpLRL1plays a greater role than PpLRL2.
ConclusionLRL genes are part of an ancient gene-regulatory
network thatcontrols the development of tip-growing filamentous
cells withrooting functions—rhizoids, caulonema, and root hairs—at
theinterface between land plants and the soil. This
auxin-regulatednetwork was active early in land plant evolution and
was presentin the last common ancestor of the angiosperms and
mosses. Thisancient gene-regulatory network diverged since P.
patens andA. thaliana last shared a common ancestor (Fig. 7).
Materials and MethodsSequence Retrieval and Alignment. To
identify potential LRL proteins inP. patens, we performed BlastP
searches using AtLRL3 (At5g58010) aminoacid sequence as a query on
Phytozome (Version 9.1) (37) with a E-valuethreshold of 1e-6, and
we retrieved all of the resulting sequences. We usedonly the
primary transcripts where alternative transcripts exist. CDS
retrievedwere translated to amino acids and then aligned by using
Mafft with theL-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 fromfour 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 assemblybuild 2.5
(www.kazusa.or.jp/lotus/index.html), and the Amborella tricho-carpa
genome (41). Only sequences containing unambiguous LRL domainswere
included. The sequences were aligned with Mafft. The alignment
wastrimmed with TrimAl (Version 1.3) (39) as implemented in
Phylemon (Version
2.0) with automated parameters (40).The final amino acid
alignment, con-taining 164 sequences from 32 species and 121
columns, was used in sub-sequent analyses. This alignment was also
used to determine the similarity ofthe bHLH and LRL domains (as
shown in Fig. 1) of selected LRL proteins, byusing the Ident and
Sim program in the Sequence Manipulation Suite (42).
Phylogenetic Analyses. The best-fit model of amino acid
evolution for thetrimmed alignments were determined by Prottest
(Version 3) (43), andthe 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-Likeapproximate
likelihood-ratio test (aLRT) for branch support (45) and the bestof
nearest neighbor interchange and subtree pruning and regrafting
fortree improvement. Bayesian analyses were carried out with
MrBayes (Ver-sion 3.2) (46) for >15 million generations. One
tree was saved every 100generations, and the first 25% of trees
were discarded. A 50-majority-ruletree was generated from the
remaining trees. The default settings wereused for gap
treatment—i.e., as unknown character in PhyML and as missingdata in
MrBayes.
Plant Material, Growth Conditions, and Phenotypic Analyses. The
GransdenWT strain (47) of P. patens (Hedw.) Bruch & Schimp was
used in this study.Unless otherwise stated, all plants were grown
in a SANYO MLR-351growth chamber at 25 °C and a 16-h light/8-h dark
regime with a pho-tosynthesis photon flux density of ∼40 μmol
m−2·s−1. Plants were grownon 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/LFeSO4·7H2O, 0.055 mg/L CuSO4·5H2O, 0.055
mg/L ZnSO4·7H2O, 0.614 mg/LH3BO3, 0.389 mg/L MnCl2·4H2O, 0.055 mg/L
CoCl2·6H2O, 0.028 mg/L KI,0.025 mg/L Na2MoO4·2H2O, 25 mg/L KH2PO4
buffer (i.e., 1.84 mM atpH 6.5 with KOH), and 7 g/L agar (Formedium
AGA03). Then, 5 g/L glucoseand 0.5 g/L ammonium tartrate dibasic
(C4H6O6·2H3N) were added as sup-plements for routine subculture of
macerated protonema tissues (KNOPSmedium). Sterile cellophane discs
(80-mm 325P Cellulose discs; A.A. Pack-aging) were put on top of
the medium to facilitate phenotypic analysis andcollection of
tissues for subculture.
PEG-Mediated Transformation of P. patens. PEG-mediated
transformation wasperformed as described (10, 48). Antibiotic
selection was performed byadding 50 μg/mL G418 disulfate or 25
μg/mL hygromycin B to thegrowth medium.
Southern Blot Analysis. Genomic DNA was extracted from protonema
tissuesby using the cetyltrimethylammonium bromide (CTAB) method
and thendigested overnight with the appropriate restriction
endonuclease. A total of1 μg (for Pplrl2, Pplrl1 Pplrl2, and
Pplrl2pro:GUS) or 2 μg (for Pplrl1 and
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).
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PpLRL1pro:GUS) of digested genomic DNA, as quantified with a
NanoDropND1000 spectrometer (Thermo Scientific), was subjected to
agarose gelelectrophoresis before being transferred to a positively
charged nylonmembrane. Southern blots were carried out with the
digoxigenin (DIG)system for filter hybridization with DIG-labeled
probe generated by PCRaccording to the DIG Application Manual for
Filter Hybridization (Roche).DIG-labeled DNAMolecular Weight Marker
VII (Roche) was used throughoutthe analysis. Primers for probe
synthesis are given in Table S1.
Generation of P. patens GUS-Reporter Lines. The GUS-reporter
lines weregenerated by replacing the endogenous PpLRL1 or PpLRL2
gene with theuidA gene and a hygromycin-resistance cassette from
the plasmid pBHrev-GUS (pBHSNR-GUS) (9); this strategy for
constructing the reporter lines isillustrated in Fig. S4. To
generate the GUS reporter lines for PpLRL1, a 871-bpfragment
upstream and a 888-bp fragment downstream of the predictedPpLRL1
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 a880-bp fragment downstream of the predicted CDS
(Pp1s209_22V6) werecloned into the AscI–NcoI and BamHI–HindIII site
of the pBHrev–GUS plas-mid, respectively. These generated the
pBHrev-PpLRL1proGUS and pBHrev-PpLRL2proGUS 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 byPCR analyses
and then further selected with Southern blot analysis (Fig.
S4).Three independent lines, which show the same GUS-staining
pattern, wereused for each genotype.
Generation of P. patens Loss- and Gain-of-Function Mutants. The
plasmidspBHrev (pBHSNR) and pBNRF (10) were used to generate the
loss-of-functionconstructs in this study. To generate the Pplrl1
complete knockout mutants,an 812-bp DNA fragment upstream and a
819-bp fragment downstream ofthe PpLRL1 were cloned into the
SalI–BamHI and MluI–NcoI sites of thepBHrev plasmid, respectively
(Fig. S5). Similarly, to generate Pplrl2 knockoutmutants, a 697-bp
fragment upstream and a 655-bp fragment downstreamof the PpLRL2
locus were cloned into the SalI–BamHI and SpeI–AscI site ofthe
pBNRF plasmid, respectively (Fig. S5). These generated the
pBHrev-PpLRL1KO and pBNRF-PpLRL2KO constructs. The plasmids were
linearizedwith SalI + NcoI (for pBHreb-PpLRL1KO) and SalI + AscI
(for pBNRF-PpLRL2KO)before PEG-mediated transformation. The Pplrl1
Pplrl2 double mutants weregenerated by retransforming Pplrl1-3 with
the pBNRF-PpLRL2KO construct.Stable transformants were selected by
PCR analyses and then further selectedwith Southern blot analysis
(Fig. S5). Three independent lines, which showedthe same phenotype,
were used for each genotype. Two independent lines ofpartial
gene-deletion mutants (Pplrl1ΔbL and Pplrl2ΔbL), where only the
bHLHand LRL domains are deleted, were generated similarly (Fig. S6)
and werePCR-verified.
To generate the 35S:PpLRL1bL and 35S:PpLRL2bL mutants, partial
CDS forthe bHLH–LRL region of the PpLRL genes (Fig. S6) were first
amplified by RT-PCR and subcloned into the pCR8/GW/TOPO Vector
(Invitrogen), thentransferred to the p108GW35S vector through the
Gateway LR Clonase IIsystem (Invitrogen) for moss overexpression
(11). This construct includes the108 locus to facilitate insertion
into the neutral 108 locus in the P. patensgenome. The levels of
the PpLRL1 and PpLRL2 bHLH–LRL partial transcriptin 35S:PpLRLbL
lines were determined by quantitative reverse
transcrip-tion-polymerase chain reaction (qRT-PCR) (Fig. S6).
GUS Staining. Unless otherwise stated, protonema inocula were
grown onKNOPS medium with cellophane for 2.5 wk and then directly
incubated inGUS 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-
3-indolyl-β-D-glucuronic acid, and cyclohexyl ammonium salt
(X-GlcA; Mel-ford) dissolved in N,N-dimethylformamide at 37 °C for
24–48 h. Tissues werethen destained by incubation in 70–100%
(vol/vol) ethanol for at least 24 h.No staining was observed in WT
plants.
Time-Lapse Microscopy. Filaments at the edge of the protonema
growing oncellophane disk onminimal mediumwere photographed every 6
min under aLeica DFC310 FX camera mounted on a Leica M165 FC
stereomicroscope.Images were analyzed with ImageJ (49). The
increase in filament lengthevery 30 min was used to calculate the
growth rate.
qRT-PCR. Ten-day-old macerated protonema growing on minimal
mediumwas used for all qRT-PCRs. Approximately 100 mg of fresh
protonema thathad been gently squeezed to remove excess water was
frozen in liquid ni-trogen before RNA extraction with the RNeasy
Plant Mini Kit (Qiagen). Theamount of RNA was quantified with a
NanoDrop ND1000 spectrometer(Thermo Scientific) or a Qubit 2.0
Fluorometer (Invitrogen) before treatmentwith Turbo DNase (Ambion,
Life Technologies). First-strand cDNA synthesiswas performed with
the SuperScript III First-Strand Synthesis System for RT-PCR
(Invitrogen, Life Technologies) by using oligo(dT). The following
amountof DNase-treated RNA was used in the cDNA synthesis: 3.5 μg
for detectingPpLRL levels in Ppshi1, Ppshi2 and WT; 9.1 μg for
detecting PpLRL levels inPpOXSHI1-5 and WT; 1.3 μg for detecting
PpLRL levels in WT treated withhigh and low phosphate; 2.7 μg for
detecting PpRSL levels in Pplrl1 Pplrl2double mutants and WT; and
>1 μg for detecting PpLRL levels in Pprsl1Pprsl2 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 useddirectly in
a 20-μL qRT-PCR. qRT-PCR was performed with the SYBR GreenPCR
Master Mix (Applied Biosystems) and the Applied Biosystems 7300
Real-Time PCR system. Three biological replicates, each with three
technicalreplicates, were used for each plant sample. The cycling
conditions were asfollows: 50 °C for 2 min; 95 °C for 10 min; then
40 cycles of 95 °C for 15 s and60 °C for 1 min. Dissociation curve
analyses were performed at the end ofthe cycles to ensure the
formation of only a single amplicon for each rep-licate. LinRegPCR
(50) was used to estimate the baseline, threshold, andprimer
efficiencies. The starting concentration of the target transcript
in asample, expressed in arbitrary fluorescence units (N0), was
computed fromthe following formula: N0 = threshold/[(mean
efficiency of each primerpair)̂ Cq] (50). PpGAPDH (11) was used as
the internal reference for all ex-periments, except for the
determination of bHLH–LRL partial transcripts inthe 35S:PpLRL1bL
and 35S:PpLRL2bL lines, where PpAdePRT was used as theinternal
references (51). Primers used are given in Table S1.
Auxin and Low-Phosphate Treatment. For auxin treatments, plants
weregrown on KNOPS-GT supplemented with 0, 0.1, or 1 μM NAA
dissolved indimethyl sulfoxide. High-phosphate treatment was
carried out by growingplants in KNOPS-GT medium supplemented with 1
mM Mes. The low-phos-phate medium (−P) was prepared by replacing
the KH2PO4 buffer in theKNOPS-GT medium with 0.918 mM K2SO4.
ACKNOWLEDGMENTS. We thank Dr. Katarina Lundberg from Prof.
EvaSundberg’s group (Swedish University of Agricultural Sciences)
for providingthe Ppshi1, Ppshi2, and the PpOXSHI1 lines (23); Dr.
Krzysztof Szczyglowski(Agriculture and Agri-Food Canada) for
providing the Atlrl mutant lines forobservation; Nuno Pires for
technical assistance and scientific input at thebeginning of the
project; and Helen Prescott and Lida Chen for laboratorysupport.
T.T.H.Y. was supported by a Clarendon Fund Scholarship andthe
Overseas Research Students Awards Scheme; B.C. and L.D. were
sup-ported by the PLANTORIGINS Marie Curie Network and the EVO500
ERC-Advanced Grant.
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