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1
REGULATION OF HORMONAL CONTROL, CELL REPROGRAMING AND 1
PATTERNING DURING DE NOVO ROOT ORGANOGENESIS 2
Estefano Bustillo-Avendaño#, 1, Sergio Ibáñez#, 2, Oscar Sanz1, Jessica Aline Sousa Barros2, 3, 3
Inmaculada Gude1, Juan Perianez-Rodriguez1, José Luis Micol2, Juan Carlos Del Pozo1, Miguel 4
Angel Moreno-Risueno+$, 1, José Manuel Pérez-Pérez+$, 2 5
6 1Centro de Biotecnología y Genómica de Plantas (Universidad Politécnica de Madrid – Instituto 7
Nacional de Investigación y Tecnología Agraria y Alimentaria), Madrid, Spain 8 2Instituto de Bioingeniería, Universidad Miguel Hernández, 03202 Elche, Spain 9 3Current address: Departamento de Biologia Vegetal, Universidade Federal de Viçosa, 36570-10
900 Viçosa, Minas Gerais, Brazil 11
12
#: Co-first author 13
+: Co-senior author 14 $: Corresponding authors: M.A. Moreno-Risueno (e-mail: [email protected]) and 15
J.M. Pérez-Pérez (e-mail: [email protected]) 16
17
Short title: De novo root regeneration in Arabidopsis leaves 18
One-sentence summary: Distinctive developmental stages lead to de novo root organogenesis 19
in leaves guide genetically dissection of the primary developmental pathways 20
Keywords: regeneration; cell reprogramming; hormonal signaling; de novo organ formation; 21
adventitious rooting; root patterning 22
23
Word count breakdown: Abstract, 195; Introduction, 908; Results, 3749; Discussion, 2210; 24
Materials and methods, 1420; Acknowledgements, 34; References, 2844; Figure legends, 1042 25
Figures: 8 Supplemental figures: 826
Plant Physiology Preview. Published on December 12, 2017, as DOI:10.1104/pp.17.00980
Copyright 2017 by the American Society of Plant Biologists
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FOOTNOTES 27
List of author contributions: Conceptualization and Supervision: J.M.P.-P. and M.A.M.-R.; 28
Methodology, J.M.P.-P., M.A.M.-R., E.B.-A., and S.I.; Investigation, E.B.-A., S.I., O.S., J.A.S.B., 29
I.G., and J.P.; Formal Analysis: E.B.-A. and S.I.; Writing – Original Draft, J.M.P.-P., M.A.M.-R., 30
E.B.-A. and S.I.; Writing – Review & Editing, . J.M.P.-P., M.A.M.-R., J.L.M., and J.C.P.; Funding 31
Acquisition, J.M.P.-P. and M.A.M.-R.; Resources, . J.M.P.-P., M.A.M.-R., J.L.M., and J.C.P. 32
33
Funding information: This work was supported by grants from Ministerio de Economía y 34
Competitividad (MINECO) of Spain, ERDF and FP7 Funds of the European Commission, 35
BFU2013-41160-P, BFU2016-80315-P and PCIG11-GA-2012-322082 to M.A.M.-R., AGL2012-36
33610 and BIO2015-64255-R to J.M.P.-P. and BIO2014-52091-R to J.C.P. M.A.M.-R. was 37
supported by a Ramon y Cajal contract from MICINN. 38
39
Corresponding authors e-mail: M.A. Moreno-Risueno (e-mail: [email protected]) 40
and J.M. Pérez-Pérez (e-mail: [email protected]) 41
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ABSTRACT 42
Body regeneration through formation of new organs is a major question in developmental 43
biology. We investigated de novo root formation using whole leaves of Arabidopsis thaliana. Our 44
results show that local cytokinin biosynthesis and auxin biosynthesis in the leaf blade followed 45
by auxin long-distance transport to the petiole leads to proliferation of J0121-marked xylem-46
associated tissues and others through signaling of INDOLE-3-ACETIC ACID INDUCIBLE28 47
(IAA28), CRANE (IAA18), WOODEN LEG, and ARABIDOPSIS RESPONSE REGULATORS1 48
(ARR1), ARR10 and ARR12. Vasculature proliferation also involves the cell cycle regulator KIP-49
RELATED PROTEIN2 and ABERRANT LATERAL ROOT FORMATION4 resulting in a mass of 50
cells with rooting competence that resembles callus formation. Endogenous callus formation 51
precedes specification of postembryonic root founder cells, from which roots are initiated 52
through the activity of SHORT-ROOT (SHR), PLETHORA1 (PLT1) and PLT2. Primordia 53
initiation is blocked in shr plt1 plt2 mutant. Stem cell regulators SCHIZORIZA, JACKDAW, 54
BLUEJAY and SCARECROW also participate in root initiation and are required to pattern the 55
new organ, as mutants show disorganized and reduced number of layers and tissue initials 56
resulting in reduced rooting. Our work provides an organ regeneration model through de novo 57
root formation, stating key stages and the primary pathways involved. 58
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INTRODUCTION 59
Plants have striking regeneration capacities, and can produce new organs from postembryonic 60
tissues (Hartmann et al., 2010; Chen et al., 2014; Liu et al., 2014) as well as reconstitute 61
damaged organs upon wounding (Xu et al., 2006; Heyman et al., 2013; Perianez-Rodriguez et 62
al., 2014; Melnyk et al., 2015; Efroni et al., 2016). Intriguingly, root regeneration upon stem cell 63
damage recruits embryonic pathways (Hayashi et al., 2006; Efroni et al., 2016), while in 64
contrast, postembryonic formation of whole new organs, such as lateral roots, appears to use 65
specific postembryonic pathways (Lavenus et al., 2013). 66
Crosstalk between auxin and cytokinin signaling is required for many aspects of plant 67
development and regeneration (El-Showk et al., 2013) although how their synergistic interaction 68
is implemented at the molecular level has not been clarified (Skoog and Miller, 1957; Chandler 69
and Werr, 2015). Exogenous in vitro supplementation of these two hormones results in 70
continuous cell proliferation, to form a characteristic structure termed callus. Callus emerges as 71
a common regenerative mechanism for almost all plant organs through in vitro culture (Atta et 72
al., 2009; Sugimoto et al., 2010). There is increasing evidence that callus formation requires 73
hormone-mediated activation of a lateral and meristematic root development program in 74
pericycle-like cells defined by expression of the J0121 marker (Sugimoto et al., 2010). 75
Accordingly, many regulators of lateral root development, such as AUXIN RESPONSE 76
FACTOR7 (ARF7), ARF19, LATERAL ORGAN BOUNDARIES DOMAIN16 (LBD16), LBD17, 77
LBD18 and LBD29, are required for hormone-induced callus formation (reviewed in Ikeuchi et 78
al., 2013). 79
Many species can regenerate new organs from explants (e.g. roots from leaves) without 80
exogenous supplementation of hormones (Bellini et al., 2014). Making roots de novo requires 81
generating the different tissues and cell types of the new organ. All roots have the same tissues, 82
although the number of layers and cells types of these may vary (Kuroha et al., 2006; Lucas et 83
al., 2011). Tissues are continuously formed by asymmetric division of initial cells, which are 84
stem cells, followed by proliferative divisions of their daughter meristematic cells. Stem cell 85
activity is maintained by quiescent center (QC) (van den Berg et al., 1997; Drisch and Stahl, 86
2015) and auxin activity (Della Rovere et al., 2013). Auxin accumulation in the QC area triggers 87
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a dose-dependent and slow response that activates PLETHORA (PLT) factors. PLT proteins 88
form a gradient in the root meristem which is required to position the QC, maintain stem cell 89
activity and trigger proliferation of meristematic cells (Aida et al., 2004; Mahonen et al., 2014). 90
Position and activity of the QC also requires radial information delivered by the mobile factor 91
SHORT-ROOT and its downstream target SCARECROW (Sabatini et al., 2003; Levesque et al., 92
2006; Moubayidin et al., 2016). In addition, WUSCHEL-RELATED HOMEOBOX5 (WOX5) is 93
confined by auxin signaling into the QC and represses differentiation of the stem cell niche, 94
primarily from the QC (Sarkar et al., 2007; Forzani et al., 2014; Pi et al., 2015; Zhang et al., 95
2015). Tissue formation in the primary root meristem also requires lineage specific factors that 96
function as cell fate determinants and as tissue endogenous signaling factors to incorporate 97
positional information into patterning (Moreno-Risueno et al., 2015). However, little is known 98
about how tissues are formed de novo. 99
Recently, a hormone-free method to study de novo root organogenesis in excised leaf 100
blades has been described (Chen et al., 2014). YUCCA-mediated auxin biosynthesis was 101
shown to be ubiquitously enhanced in the leaf mesophyll and indirectly contribute to auxin 102
accumulation near the excision site to trigger localized auxin signalling in the vasculature (Liu et 103
al., 2014; Chen et al., 2016). Formation of new roots involves formation of competent cells 104
through auxin-induced expression of WOX11 transcription factor, which has been defined as a 105
first-step for cell fate transition during de novo organ regeneration (Liu et al., 2014). WOX11 and 106
its homolog WOX12 can in addition promote callus formation and upregulate the callus 107
formation factors LBD16 and LDB29 (Fan et al., 2012; Liu et al., 2014), suggesting that de novo 108
root formation might share similar regulatory mechanisms with callus formation. Subsequently in 109
leaf blade rooting, WOX11 and WOX12 activate WOX5 and WOX7 factors, which are 110
expressed in dividing cells forming root primordia, while WOX11/12 expression quickly 111
decreases in dividing cells (Liu et al., 2014; Hu and Xu, 2016). Activation and maintenance of 112
WOX5/7 expression also requires auxin signalling in an unknown pathway different from 113
WOX11/12. Mutants in these WOX factors reduce the number of roots regenerated per leaf 114
blades and affect rooting rate of leaf blades. As a considerably high percentage of leaf blades 115
still root in these mutants, additional regulation must exist. 116
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We have performed an extensive study to further understanding root regeneration from 117
aerial organs. Whole leaves of many species can regenerate entire functional plants in 118
hormone-free medium, and thus we used whole leaves with petioles of Arabidopsis thaliana 119
instead of excised leaf blades. We identified four developmental stages: 1) proliferation of some 120
xylem-associated tissues to form an endogenous callus; 2) specification of root founder cells 121
within the callus; 3) root primordia initiation from founder cells and patterning and 4) root 122
meristem activation and emergence. We have also characterized a number of factors regulating 123
these developmental stages. Some auxin and cytokinin signaling factors appear as critical for 124
endogenous callus initiation and formation while some stem cell regulators control initiation and 125
patterning of newly formed organs. These results define key stages and regulators required for 126
leaf rooting establishing a developmental framework for de novo organ formation in plants. 127
128
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129
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RESULTS 130
Vasculature-associated cell proliferation is required for de novo organ regeneration in 131
Arabidopsis 132
We found that excised whole leaves of Arabidopsis can root without hormone supplementation, 133
similarly to leaf blades as previously described (Chen et al., 2014) and at similar percentages 134
(Fig. S1A-B). Because some species can regenerate entire functional plants from whole leaves 135
without the aid of external hormones, we performed our studies using whole leaves. As de novo 136
formed roots emerged from the petiole base of whole leaves (Fig. 1A), petioles were 137
microscopically observed (Fig. 1B). All petioles showed the same morphological changes during 138
de novo organ regeneration. Although asynchrony was observed in the regeneration process, 139
by day 10 after excision (dae) most leaves (85-100%) had regenerated at least one root. At 2 140
dae cells adjacent to xylem started to proliferate, forming stratified layers from 3 dae onwards 141
(Fig. 1C-E) that pushed away xylem conducts and displaced the collenchyma. Vasculature 142
proliferation and subsequent formation of primordia caused the proximal petiole to thicken (Fig. 143
S1C). First primordia were visible at 4 dae, and located at external layers of proliferating 144
vasculature (Fig. 1F). At 5 dae root primordia showed a layered pattern (Fig. 1G). Eventually, 145
newly formed roots with well-organized meristems emerged through petiole tissues from 7 dae 146
onwards (Fig. 1H). 147
Pericycle-like cells (those expressing the J0121 reporter) have been associated to 148
regenerative and morphogenic processes as the source of reprogrammable cells (Sugimoto et 149
al., 2010; Chen et al., 2014). Sections of petioles at the time of excision revealed that the root-150
pericycle line J0661-GFP marks cells around xylem and procambium cells (Figure 1I-J), while 151
the J0121-GFP line (Fig. 1L-M) was restricted to a layer around xylem vessels, being excluded 152
from procambium. Number of cells marked with J0661 and J0121increased quickly during first 153
days of regeneration (Fig. 1K, N-P). We observed that all proliferating cells were marked with 154
J0661-GFP while some proliferating cells in the J0121 line did not have the GFP (Fig.1 K, O-P) 155
indicating that cell proliferation associated to the J0661 reporter. Although it cannot be ruled out 156
that J0661-GFP is activated in proliferating cells, it is possible that xylem and procambium 157
proliferate as part of the reprogramming process. In addition, we observed that primordia at 158
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early stages of development were marked with J0121-GFP (Fig. 1P), establishing an 159
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association between de novo primordia formation and J0121 identity, similarly to other 160
developmental or regenerative processes such as callus or lateral root formation (Dubrovsky el 161
al., 2006; Sugimoto et al., 2010). 162
We next studied mutants defective in cell cycle progression, at the G1/S transition, such 163
as the KIP-RELATED PROTEIN2 (KRP2) overexpressor, and at the G2/M transition, such as 164
cyclinB1;1 (cycb1;1) and cycb1;2 mutants and a dominant negative form of the CDKB1;1 kinase 165
(CDKB1;1 DN161). Percentage of petioles showing vasculature-associated proliferation was 166
reduced in Pro35S:KRP2 and CDKB1;1 DN161 lines (Fig. 1Q), while only size of proliferating 167
mass of cells was reduced in rest of lines. In addition, Pro35S:KRP2 blocked de novo organ 168
regeneration, while cycb1;1 and CDKB1;1 DN161 showed a significant reduction in the number 169
of petioles regenerating roots (Fig. 1R). We also chemically inactivated the G1/S transition by 170
incubating leaves with either 2.5 or 5 mM hydroxyurea (HU). We observed a significant 171
decrease in rate of petioles showing vascular-associated proliferation and subsequent new root 172
formation by the HU treatment (Fig. 1Q-R and Fig. S2A-B). HU treatment did not associate with 173
increased cell death around the vasculature near the leaf excision site upon trypan blue staining 174
(Fig. S2C). All together, these results indicate that cell division activation of vasculature cells is 175
the first and required stage for de novo organogenesis during rooting of leaves. 176
177
Cytokinin biosynthesis and response during de novo root regeneration 178
As we had found an association between de novo root regeneration and J0661 and J0121 179
identities, and callus originates from J0121-marked cells after hormonal induction (Sugimoto et 180
al., 2010), we hypothesized that vasculature proliferation required for leaf rooting could be a 181
type of callus. Cytokinin and auxin signalling are required for callus formation and regeneration, 182
and thus we tested if these two hormones were involved in the developmental pathway leading 183
to de novo organ regeneration from leaves. 184
First, we investigated cytokinin biosynthesis and signalling (Zürcher and Müller, 2016). 185
We found enriched expression of ProIPT3:GUS in petioles right after excision (Fig. 2A). 186
ProIPT5:GUS was highly expressed in vascular-associated cells in the petiole base at 2 dae (Fig. 187
2B), while ProLOG4:GUS, which was originally expressed in leaf vasculature at 0 dae, increased 188
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expression at the petiole base over time (Fig. 2C, arrowhead). Cytokinin signalling, as reported 189
by ProARR5:GUS (D'Agostino et al., 2000), was restricted to a subset of vascular-associated cells 190
near the petiole base at 2 dae, which associated with proliferation of vasculature, to decrease at 191
later time points (Fig. 2D) and it did not show expression during de novo primordia formation. 192
Consistent with ARR5 reporting primary cytokinin response during petiole vasculature 193
proliferation (D'Agostino et al., 2000), incubation of leaf explants with synthetic cytokinin 6-194
benzylaminopurine (6-BAP) increased ProARR5:GUS expression in the petiole vasculature and 195
basal region (Fig. 2E). 196
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Next, we studied the ability of several cytokinin signalling mutant combinations in 197
ARABIDOPSIS HISTIDINE KINASE4 (AHK4), ARABIDOPSIS HISTIDINE 198
PHOSPHOTRANSFER PROTEIN1 (AHP1) to AHP5 and ARR1, ARR10 and ARR12 genes in 199
regulating both vasculature proliferation and de novo root regeneration. We quantified petioles 200
regenerating as petioles showing vasculature proliferation/thickening, root primordia formation 201
or visible roots. Leaf petioles of wooden leg (wol, a dominant negative mutant in AHK4 202
receptor), and ahp1 ahp2 ahp3 and arr1 arr10 arr12 loss-of-function mutants displayed lower 203
regeneration percentage at 7 dae (Fig. 2F). Interestingly, wol, ahp1 ahp2 ahp3 and arr1 arr10 204
arr12 mutants were also defective in hormone-induced callus formation from different tissue 205
explants, such as leaves, cotyledons and roots (Fig. S3), indicating that specific cytokinin 206
signalling is required for both callus formation and vasculature proliferation in leaf petioles. 207
Despite cytokinin signalling was required for vasculature proliferation in petioles during rooting, 208
for those leaf petioles of cytokinin signalling mutants which regenerated, we detected higher 209
number of roots (which we categorized by frequencies in numbers of roots and designated as 210
rooting capacity) (Fig. 2F). Higher auxin-to-cytokinin ratios have been shown to induce 211
specification and growth of new root primordia (Müller and Sheen, 2008). Thus, we wondered if 212
we could alter new primordia initiation by altering hormone ratios. Cytokinin treatment increased 213
vascular proliferation on a concentration- and time-dependent manner (Fig. S2D-E). We 214
observed that regeneration deficiencies of most cytokinin signalling mutants could be 215
compensated by low levels of exogenous auxin (Fig. 2G) that also increased vasculature 216
proliferation at expenses of reducing rooting capacity in the ahp1 ahp2 ahp3 mutant (Fig. 2G). 217
All together, these results indicate a dual role for cytokinin first as a positive activator of 218
vasculature cell division, and second as a negative regulator of root primordia initiation. 219
220
Specific auxin signalling factors regulate de novo root regeneration 221
Next we investigated auxin signalling during rooting of leaves using the DR5 reporter line 222
(Ulmasov et al., 1997). ProDR5:GUS was expressed in vascular-associated cells at the proximal 223
region of the petiole, as early as 12 hours after excision (hae), to increase quickly to 1 dae, 224
remaining high during proliferative stages and decreasing over time coincident with deceleration 225
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of vasculature growth (Fig. 3A-B). We observed high localized expression of ProDR5:GUS in 226
clusters of cells at the time of primordia initiation and formation. Consistent with a regulatory role 227
of auxin in rooting, local IAA application significantly increased root formation, along with 228
expansion of ProDR5:GUS expression domain (Fig. 3A, C). In addition, the auxin-overproducing 229
mutant superroot2 (sur2) (Barlier et al., 2000) also showed increased number of de novo 230
formed roots at 7 dae, similarly to auxin treated petioles (Fig. 3C). 231
Auxin signalling is regulated by INDOLE-3-ACETIC ACID INDUCIBLE (IAA) co-factors 232
acting in combination with AUXIN RESPONSE FACTOR (ARF) transcriptional partners (Li et 233
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al., 2016). We reasoned that IAA factors regulating postembryonic (lateral) root formation (Goh 234
et al., 2012) could be involved in de novo organ formation. We assessed the mutants auxin 235
resistant2-1 (axr2-1), solitary-root-1 (slr-1), crane-2, iaa28-1, and the double mutant non-236
phototrophic hypocotyl4-1 (nhp4-1) arf19-1 (see Materials and Methods); which are, 237
respectively, gain-of-function mutants for the factors IAA7, IAA14, IAA18, IAA28 and a double 238
loss-of-function mutant for ARF7 and ARF19. crane-2 reduced de novo root regeneration, with 239
approximately 40% of petioles not showing any sign of vasculature proliferation or de novo root 240
formation (Fig. 3D). In addition, slr-1 and iaa28-1 showed reduced rooting capacity although all 241
petioles showed some vasculature proliferation (Fig. 3E). When we quantified vasculature 242
proliferation area of petioles regenerating we observed a reduction for iaa28-1 and crane-2 but 243
not for slr-1 (Fig. 3F). These results indicate that the auxin signalling module mediated by 244
IAA18 is required for de novo root regeneration at stages of vascular proliferation, that of IAA28 245
for vascular proliferation and root initiation, while IAA14 appears to be only required for de novo 246
root initiation. 247
We also investigated if these mutants were affected in hormone-induced callus formation. 248
We found that only crane-2 showed reduction in all explants assayed after hormonal incubation, 249
while axr2-1 intriguingly showed an increase for callus formed from root explants (Fig. S4A-C). 250
These results indicate that vasculature proliferation during rooting and hormone-induced callus 251
use the auxin signaling pathway mediated by IAA18. Our previous results also showed that 252
cytokinin signaling required for vasculature proliferation during de novo organogenesis was also 253
required for hormone-induced callus formation suggesting that vasculature proliferation is a type 254
of callus. We investigated ABERRANT LATERAL ROOT FORMATION4 (ALF4) during leaf 255
rooting, as alf4-1 mutants have been previously linked to callus formation (Sugimoto et al., 256
2010) and vascular connection during graft establishment (Melnyk et al., 2015). We observed 257
that during de novo root formation vasculature proliferation is reduced by 2.5 fold in alf4-1 258
mutants which is accompanied by 15 fold decrease in de novo formed root and primordia (Fig. 259
S5). Based on these results, we designated vasculature proliferation developmental stage as 260
endogenous callus formation. 261
262
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Auxin signalling factors are required for de novo organ regeneration in leaf blades 263
In contrast to endogenous callus formation observed in petioles of whole leaves during rooting, 264
limited vasculature proliferation was observed during rooting of leaf blades (Liu et al., 2014). We 265
wondered to what extent auxin and cytokinin signaling factors regulating proliferation at the 266
petiole base would be involved in rooting of leaf blades. When we assessed rooting capacity in 267
leaf blades of these mutants, we observed that crane-2 and iaa28-1 displayed a reduction in 268
rooting capacity while slr-1 presented moderate although non-significant reductions (Fig. S6A). 269
wol and arr1 arr10 arr12 mutants were similarly affected as slr-1, while no change was detected 270
for ahp1 ahp2 ahp3 and ahp2 ahp4 ahp5 mutants (Fig. S6B). As IAA18 and IAA28 are required 271
for endogenous callus formation during whole leaf rooting and are shared with leaf blade 272
rooting, it is possible that de novo root regeneration in leaf blades could also involve an 273
endogenous callus developmental program. 274
275
Local auxin accumulation at the petiole base is dependent on polar auxin transport 276
As localized auxin signalling was required for whole leaf rooting we wondered about the source 277
of auxin. YUCCA-mediated auxin biosynthesis was shown to be ubiquitously enhanced in the 278
mesophyll of leaf blades shortly after wounding (Chen et al., 2016). During rooting of whole 279
leaves, we detected ProYUC9:GUS enriched expression in leaf mesophyll cells at 12 hae, which 280
progressively decreased at later time points (Fig. S7A). ProYUC8:GUS expression was induced in 281
proliferating vascular-associated cells at the petioles base 2 dae (Fig. S7B). These results 282
indicate two possible sources of auxin during first stages of regeneration, one from the leaf 283
blade, and the other from the proliferating vasculature itself. To determine if there was 284
differential contribution of these two auxin sources, we removed the leaf blade and found a 285
significant decrease in regeneration and rooting capacity, which in most cases stopped at the 286
endogenous callus stage (Fig. 4A). Local auxin response (ProDR5:GUS expression) in petioles 287
without leaf blade was low or undetectable (Fig. 4B). We also locally inhibited polar auxin 288
transport through application of N-1-naphthylphthalamic acid (NPA) at the blade-petiole 289
junction, resulting in almost complete block of auxin response and subsequent regenerative 290
response (Fig. 4C-D). 291
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We next characterized PIN-FORMED (PIN) expression. ProPIN4:GUS was ubiquitously 292
expressed in the leaf vasculature at the time of excision; however at 1 dae it was only 293
expressed at the base of the petiole and in endogenous callus at 3 dae (Fig. 4E). 294
ProPIN3:PIN3:GFP expression in the petiole at the time of excision was polarized towards the 295
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base in epidermal cell membranes and both laterally- and basally-localized in some vascular-296
associated cell membranes (Fig. 4F, arrowheads). From 12 hae to 1 dae, we found enriched 297
expression of ProPIN3:PIN3:GFP in a subset of vascular-associated cells at the petiole base 298
region (Fig. 4G). Interestingly, PIN3-GFP protein in cells proximal to the excision was oriented 299
towards the apex (upper left direction in Fig. 4G, arrowheads) while its orientation changed to 300
the base in cells at the distal position from the excision. 301
We also investigated rooting in mutants of genes affected in auxin influx (AUX1) or auxin 302
efflux (PIN1, PIN2, PIN3 and PIN7), which have been described to have low auxin transport 303
rates (Petrasek et al., 2006). Consistent with our previous observations the regenerative 304
potential of leaves of pin1, pin2 pin3, pin2 pin3 pin7 and aux1 mutants was reduced (Fig. 4H). 305
GNOM loss-of-function mutants have altered polar auxin transport by interfering with PIN 306
internalization (Kleine-Vehn et al., 2009). When we studied the GNOM mutant fewer (fwr) 307
(Okumura et al., 2013), we observed significant differences in regeneration and rooting capacity 308
(Fig. 4H). fwr is a weak gnom allele but it is possible that several auxin transporters are 309
simultaneously affected, which could explain why there is also a reduction in rooting capacity 310
while no reduction was observed for single auxin transporter mutants. 311
312
Postembryonic root founder cells establish on endogenous callus prior primordia 313
formation 314
WOUND INDUCED DEDIFFERENTIATION1 (WIND1) is rapidly induced at the wound site to 315
promote callus formation through the ARR-dependent signaling pathway (Iwase et al. 2011). 316
From 1 to 4 dae, we found ProWIND1:GUS expression in vascular cells at the petiole near the 317
excision site, while expression was downregulated in new root primordia and no expression was 318
detected at the time of root emergence by 6 dae (Fig. 5A). Postembryonic development involves 319
specification of organ founder cells (Chandler, 2011). Thus, we hypothesized that root founder 320
cells (RFCs) could be specified within the endogenous callus to de novo form a root. It has been 321
proposed that RFCs during de novo root formation in leaf blades could be marked by 322
ProWOX11:GUS expression (Liu et al., 2014; Hu and Xu, 2016). We observed discrete 323
ProWOX11:GUS signals early detected, at 1 dae, in a few xylem-associated cells at the petiole 324
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base (Fig. 5B). From 1 dae onwards, during callus formation, high ProWOX11:GUS expression 325
was observed in many cells within this domain, but not simultaneously in all proliferating cells. 326
Later on, ProWOX11:GUS expression was observed near the central zone of the endogenous 327
callus but excluded from the dome-shape root primordia. WOX11 expression thus appears to 328
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19
associate with endogenous callus formation while not all marked cells resulted in primordia 329
initiation. We wondered whether cell division of vascular-associated cells was downstream of 330
the WOX11 signal. ProWOX11:GUS expression was increased in vasculature of petioles of 331
excised leaves incubated with the G1/S cell cycle inhibitor hydroxyurea, even when little 332
vasculature proliferation was observed (Fig. 5E-F). 333
ProSKPB2s:GUS expression has been shown to mark RFCs and their progeny during early 334
stages of lateral root formation (Manzano et al., 2012). We detected ProSKPB2s:GUS expression 335
at 3 dae restricted to few cells within the endogenous callus (Fig. 5C). From 4 to 5 dae, we 336
observed marker expression in developing root primordia while its expression disappeared from 337
functional primordia during the emerging process, remaining in some cells of the endogenous 338
callus (Fig. 5C). Interestingly, RFCs did not appear to be specified simultaneously, supporting 339
our initial observations about asynchrony in the regeneration process. In order to follow 340
primordia formation we used ProWOX5:GUS. WOX5 is expressed early during lateral root 341
primordia formation (Tian et al., 2014; Goh et al., 2016), and also following root primordia 342
initiation during de novo organogenesis (Liu et al., 2014; Hu and Xu, 2016). Expression of 343
ProWOX5:GUS was first detected at 4 dae in clusters of cells within the endogenous callus (Fig. 344
5D), similarly to ProSKPB2s:GUS after root primordia initiation. At 6 dae ProWOX5:GUS expression 345
was enriched at root meristem tip, coinciding with meristem activation and growth prior 346
emergence. Interestingly, we did not detect ProWOX5:GUS in the endogenous callus (Fig. 5D), 347
although WOX5 expression has been associated to hormone-induced callus (Sugimoto et al., 348
2010), suggesting specific regulation. 349
Cell division is required for primordia initiation and formation, and petioles of leaves 350
treated with the cell cycle inhibitor HU which started to form an endogenous callus remained 351
almost blocked at this stage (Fig. 5G). We wondered if general regulators of cell cycle 352
progression would be involved in de novo primordia formation. Our results indicate that rooting 353
capacity is compromised in cycb1;1, cycb1;2 and CDKB1;1 DN161 mutants. Accordingly, 354
overexpression of the cell cycle inhibitor KRP2 blocked progression to root initiation and 355
formation, along with reduction in callus formation by 60% previously shown in Fig. 1Q. Taken 356
together, our results indicate that WIND1 and WOX11 expression associates with vasculature 357
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proliferation leading to endogenous callus formation at the petiole base, while ProSKPB2s:GUS is 358
restricted to a few RFCs within the callus that quickly acquire WOX5 expression associated to 359
cell division, in turn mediated by CYCB1;1, CYCB1;2, CDKB1;1 and KRP2, to initiate de novo 360
root primordia regeneration (Fig. 5G). 361
362
SHORT-ROOT, PLETHORA1 and PLETHORA2 are required for de novo initiation of roots 363
We investigated whether factors specifying stem cell fate or their activity, such as PLETHORA 364
(PLT), JACKDAW (JKD), BLUEJAY (BLJ), SCARECROW (SCR), SHORT-ROOT (SHR), 365
SCHIZORIZA (SCZ) and WOX5 were required during de novo root regeneration. In addition to 366
single loss-of-function mutants, double plt1-4 plt2-2 and triple blj-1 jkd-4 scr-4, we generated 367
and tested the double mutant jkd-4 scz-1. We found significant differences in percentage of 368
leaves rooting and in rooting capacity for shr-2 and plt1-4 plt2-2 (Fig. 6A-B). In addition, scr-4, 369
blj-1 jkd-4 scr-4, scz-1 and jkd-4 scz-1 were impaired in rooting capacity. These phenotypes 370
suggested impairment in de novo root initiation or primordia formation. As in shr-2 and plt1-4 371
plt2-2 many leaves failed to regenerate any root, we generated shr-2 plt1-4 plt2-2. Notably, 372
leaves of shr-2 plt1-4 plt2-2 were not capable of rooting (Fig. 6A-C). 373
Petioles of shr-2 plt1-4 plt2-2 were observed through confocal microscopy at 6, 10 and 20 374
dae. We did not observed any primordia at 6 dae in shr-2 plt1-4 plt2-2, while most control 375
leaves had one or more primordia (Fig. 6D). Inspection at later days indicated that most leaves 376
(97%, n=90) of shr-2 plt1-4 plt2-2 did not form any primordia up to 20 dae. The few primordia 377
found (3%, n=90) remained blocked or developed aberrant shapes with presence of mature 378
xylem indicating premature differentiation. These primordia did not emerge through petiole 379
tissues. We observed endogenous callus formation, which maintained growth over time up to 20 380
dae in all observed petioles of shr-2 plt1-4 plt2-2. These results indicate that the combined 381
activity of SHR and PLT1 and PLT2 is required to de novo initiate root primordia. In addition, 382
these factors maintain proliferative activity in the forming primordia, as when these factors are 383
removed primordia differentiate. 384
385
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JACKDAW, BLUEJAY, SCARECROW AND SCHIZORIZA are required for de novo 386
formation of root primordia 387
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We observed formation of root primordia in petioles of jkd-4 scz-1 and blj-1 jkd-4 scr-4 through 388
confocal microscopy. We observed abnormal formative divisions in jkd-4 scz-1 primordia at 5 389
dae, which did not organize properly in layers or rows as compare to control roots (Fig. 6E). At 6 390
dae we observed reduced and disorganized number of cell rows in jkd-4 scz-1 and blj-1 jkd-4 391
scr-4 primordia. While endodermis, cortex and middle cortex could be identified in control roots 392
at this developmental stage based on position, corresponding rows in jkd-4 scz-1 and blj-1 jkd-4 393
scr-4 could not be identified (Fig. 6E). These results suggest that cell lineages or positional 394
identity could not be correctly established in these mutants during de novo root formation. 395
396
JACKDAW, BLUEJAY, SCARECROW, SHORT-ROOT AND SCHIZORIZA regulate 397
patterning of de novo formed roots and of lateral roots 398
Primordia formation occurred incorrectly in blj-1 jkd-4 scr-4 and jkd-4 scz-1, and therefore 399
patterning of these de novo formed roots could be compromised after emergence. Longitudinal- 400
and cross sections of de novo root meristems were examined through confocal microscopy after 401
emergence at 10 dae (encompassing roots at 1-3 days post emergence, dpe). de novo wild-402
type roots showed ground tissue with middle cortex formation, a layer located between 403
endodermis and cortex and associated to postembryonic development (Paquette and Benfey, 404
2005), while de novo roots of shr-2, scr-4, blj-1 jkd-4 scr-4 and jkd-4 scz-1 had a single layer of 405
ground tissue which we denoted as mutant (m) layer (Fig. 7A and Fig S8A-B). In addition, 406
number of cell rows making the ground tissue, which indicates number of tissue initials was 407
reduced in de novo emerged roots of blj-1 jkd-4 scr-4 and jkd-4 scz-1 as compared to wild-type 408
roots (Fig 7E and Fig. S8C). As a result, the stele region in these mutants was not delimited by 409
a closed ring of ground tissue as in wild-type roots as shown in cross sections (Fig 7B). In 410
addition, shr-2, scr-4, blj-1 jkd-4 scr-4 and jkd4 scz-1 mutants also had a smaller stele region 411
(Fig. 7B-C and Fig. S8D-E). 412
Lateral roots are organs formed postembryonically the same as de novo formed roots. 413
We decided to investigate if stem cell regulators regulating de novo root formation could also 414
regulate lateral root formation. We quantified lateral root capacity. In this assay the root tip is cut 415
to induce growth of lateral roots as described previously (Van Norman et al., 2014). We 416
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23
confirmed there were not unemerged primordia under the microscope. We observed reduced 417
lateral root capacity in shr-2 plt1-4 plt2-2, blj-1 jkd-4 scr-4 and jkd-4 scz-1 mutant combinations 418
indicating defects in founder cell specification or lateral root initiation (Fig. 7D), similarly to 419
defects observed in these mutants during de novo root formation. We also observed patterning 420
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24
defects in emerged lateral roots of these mutants, which showed a single mutant layer of 421
ground tissue and reduced number of cell rows as compared to wild type lateral roots (Fig. 7E). 422
In addition, we observed signs of premature differentiation in shr-2 plt1-4 plt2-2, with presence 423
of mature xylem in the meristem similarly to de novo formed roots. 424
425
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25
426
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26
DISCUSSION 427
Plants have the remarkable ability to regenerate a new entire individual, specific organs or their 428
tissues from explants or even few cells (Birnbaum and Sánchez Alvarado, 2008; Della Rovere 429
et al., 2016). Plant regeneration through de novo organogenesis can be achieved through 430
hormonal induction, directly or indirectly (Ikeuchi et al., 2013), but also in hormone free medium. 431
The molecular pathways involved (Ikeuchi et al., 2016; Kareem et al., 2016) and the relationship 432
between hormonal-induced and endogenous programmes are not well understood. Moreover 433
callus formation, which is a prerequisite for hormonal-induced regeneration, does not appear to 434
occur during endogenous organogenesis, although both processes share regulation (Sugimoto 435
et al., 2010; Liu et al., 2014; Perianez-Rodriguez et al., 2014; Ramirez-Parra et al., 2017). Using 436
a simple method to study de novo root organogenesis without hormone supplementation (Chen 437
et al., 2014; Liu et al., 2014) but applied to whole leaves (Fig. 8A) we found that formation of an 438
endogenous callus is a required step for de novo root organogenesis, and thus we established 439
a direct connection between both regenerative processes. Our research has also identified the 440
distinctive and required developmental stages leading to novo root organogenesis (Fig. 8B): 1) 441
vasculature proliferation and endogenous callus formation, 2) specification of root founder cells, 442
3) root primordia initiation and patterning, and 4) root meristem activation and emergence. 443
Furthermore, we have genetically dissected the primary developmental pathways involved in its 444
regulation and identified some of the key regulators involved (Fig. 8C). 445
446
Vasculature proliferation and endogenous callus formation 447
Vasculature division is the first morphological change we detect and by chemically or genetically 448
inhibiting vasculature proliferation we affected de novo root organogenesis. We also observed 449
that root primordia originated from vasculature. Pericycle-like cells, particularly those expressing 450
the J0121 marker, have been associated to regenerative and morphogenic processes as the 451
source of reprogrammable cells (Sugimoto et al., 2010; Chen et al., 2014). When we assessed 452
the J0661 and J0121 pericycle markers we observed that vasculature proliferation associated to 453
J0661 identity while some proliferating cells were devoid of J0121 expression. Intriguingly, 454
J0661 marks cells around xylem and procambium in petioles while J0121 only marks cells in 455
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27
contact to xylem. Proliferation competence, therefore, appears to involve different cell types. 456
Closer examination showed that all primordia expressed J0121 marker which indicates that 457
regeneration competence associates with J0121 identity and shows parallelisms with callus and 458
lateral root formation. Cell lineage tracing using clonal markers and live imaging could dissect 459
the exact source of reprogrammable cells during de novo root regeneration from whole leaves. 460
We also found that proliferating vascular cells express WOUND INDUCED 461
DEDIFFERENTIATION1 (WIND1), a positive regulator of wound-induced callus formation 462
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28
(Iwase et al., 2011), suggesting together with J0121 analysis that these proliferating tissues 463
could be a type of callus. Callus formation requires confluence of auxin and cytokinin responses 464
in the same set of cells (Gordon et al., 2007). In agreement with this idea, our results show 465
specific expression of auxin and cytokinin signalling reporters ProDR5:GUS and ProARR5:GUS, 466
respectively, in the vasculature. These results also indicate that specific regulation was required 467
to induce auxin and cytokinin signalling at the petiole base. In contrast to early notions that 468
cytokinins are produced only in roots, it is now recognised that they are synthesized throughout 469
the plant (Zürcher and Müller, 2016). Our results are consistent with 470
ISOPENTENYLTRANSFERASE3 (IPT3), IPT5 and LONELY GUY4 locally mediating cytokinin 471
biosynthesis at the petiole base to contribute to vascular proliferation during root regeneration. 472
Thus, cytokinin signalling mutants displayed reduced regeneration potential as well as defective 473
hormone-induced callus formation. Interestingly, the triple cytokinin signaling mutant ahp2 ahp4 474
ahp5 is affected in hormone-induced callus formation but not in vasculature proliferation during 475
rooting, indicating the existence of specific genetic differences between hormone-induced callus 476
formation and de novo root formation. 477
ProDR5:GUS expression in the proximal petiole vasculature could be indicative of auxin 478
accumulation. A study in leaf blades has shown that YUCCA1 (YUC1) and YUC4 appear to 479
mediate synthesis of auxin in mesophyll cells (Chen et al., 2016). If this occurred in our system, 480
auxin would need in addition to be transported to cells near the wound to induce de novo root 481
organogenesis. We observed that YUC9 expression was induced in the leaf blade mesophyll 482
but not in the petiole (Fig. 8A), suggesting a predominant function of the leaf blade mesophyll as 483
source of auxin for regeneration. We confirmed this by removing leaf blades which resulted in 484
inhibition of regeneration. YUC9 expression has been shown to respond to methyl-jasmonate 485
(MeJA) treatment in a COI1-dependent manner (Hentrich et al., 2013). As excision of whole 486
leaves or leaf blades involves wounding and therefore MeJA production, MeJA might activate 487
YUC9 expression to rapid increase auxin levels in leaf blades, likely in combination with YUC1 488
and YUC4 activity. We also found that YUC8 was specifically upregulated in the vascular region 489
of the petiole associated with proliferation, and thus, it is possible that YUC8 might have a 490
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29
specific role in maintaining auxin levels during vasculature proliferation or at later regenerative 491
steps. 492
Our results indicate that a long distance basipetal transport system concentrates auxin 493
generated in the leaf blade mesophyll towards defined vascular cells at the petiole base. We 494
showed that genetic and localized chemical inhibition of auxin transport significantly affected 495
regeneration. Despite of known redundancy among auxin transporters (Blilou et al., 2005; Péret 496
et al., 2012) we detected phenotypes in single mutants suggesting spatial 497
compartmentalization. Supporting this idea, PIN-FORMED3 (PIN3) was expressed in the petiole 498
vasculature while PIN4 was later restricted to the proliferating vascular region. In contrast, more 499
de-localized auxin transport is involved in rooting of leaf blades (Liu et al., 2014; Chen et al., 500
2016). As we observed predominant expression of ProDR5:GUS in the proximal petiole 501
vasculature, it is possible that auxin would need to be retained in this area. Our data suggest a 502
model in which subcellular PIN3 localization shifts from basal to apical membranes in vascular 503
cells near the wound to redirect auxin flow backwards and thus maintaining high auxin levels in 504
the proximal petiole vasculature. Interestingly, an auxin-dependent switch in PIN3 polarization 505
contributing to auxin-flow reversal is involved in the shoot gravitropic response (Rakusová et al., 506
2016), where basal-to-apical shift in PIN localization has been described to depend on 507
phosphorylation (Dai et al., 2012). It is thus tempting to speculate that auxin-dependent 508
phosphorylation of PIN3 would be involved in maintaining high auxin levels in the petiole base 509
vasculature during root regeneration. 510
The aberrant lateral root formation4 (alf4) mutant (DiDonato et al., 2004) has been linked 511
to hormone-induced callus formation (Sugimoto et al., 2010) but not to wound-induced callus 512
formation during graft stablishment (Melnyk et al., 2015). We observed reduced vasculature 513
proliferation along with great reduction in de novo root formation from whole leaves in alf4-1 514
mutants. As vasculature proliferation during de novo root formation associates to J0121- and 515
WIND1-marked cells, requires auxin and cytokinin signalling and involves ALF4, we propose it 516
is a type of callus, and therefore we refer to it as “endogenous callus” to differentiate it from 517
callus obtained by exogenous hormone supplementation. In our model, time-dependent auxin 518
accumulation in a subset of vascular cells activates proliferation, while cytokinins might regulate 519
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30
the expression of genes that are directly involved in callus formation (such as WIND1 or ALF4) 520
or that are downstream targets of the auxin signal involved in callus formation (LATERAL 521
ORGAN BOUNDARIES DOMAIN factors) (Schaller et al., 2015). Particularly, we found that 522
INDOLE-3-ACETIC ACID INDUCIBLE18 (IAA18) and IAA28 are both involved in endogenous 523
callus formation, although only mutations in IAA18 affect whole regeneration response, while 524
ARABIDOPSIS HISTIDINE PHOSPHOTRANSFER PROTEIN1 (AHP1) to AHP3 and 525
ARABIDOPSIS RESPONSE REGULATOR1 (ARR1), ARR10 and ARR12 were involved in 526
vasculature proliferation and regeneration response. 527
528
Specification of root founder cells 529
Hormone-induced callus is organized in layers showing root tissue identities that resemble a 530
root meristem and therefore a new organ could be theoretically initiated through a differentiation 531
process (Sugimoto et al., 20010). WUSCHEL-RELATED HOMEOBOX11 (WOX11) expression 532
has been associated to first cell-fate transition from regeneration-competent cells to root 533
founder cells during leaf blade rooting (Liu et al., 2014; Hu and Xu, 2016). WOX11 activates 534
WOX5 during root formation in leaf blades; however we did not find WOX5 expression in 535
endogenous callus during de novo regeneration from whole leaves, suggesting that WOX11 536
could be involved in an earlier step in the reprograming process. On the other hand, specific 537
expression associated to root founder cell specification (SKP2B) revealed the establishment of 538
a cell lineage capable of forming a new root. These results indicate that additional 539
reprogramming processes are required. 540
PLETHORA1 (PLT1) and PLT2 have been recently shown to be required to establish 541
pluripotency during de novo shoot regeneration (Kareem et al., 2015). Our results show that 542
during de novo root formation PLT1 and PLT2, in combination with SHORT-ROOT (SHR), could 543
also be involved in specification of root founder cells, which are pluripotent. In addition, 544
persistent expression of PLT1, PLT2 and SHR appears to be necessary during subsequent 545
formative stages to maintain primordia growth, as the very primordia found in shr-2 plt1-4 plt2-2 546
quickly differentiate. In contrast, in the de novo shoot regeneration system transient induction of 547
PLT2 has been shown to be sufficient to specify shoot progenitors, while subsequent 548
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31
expression of other regulators is required to accomplish de novo shoot formation from these 549
progenitors (Kareem et al., 2015). 550
551
Root primordia initiation and patterning 552
Multiple INDOLE-3-ACETIC ACID INDUCIBLE (IAA)-ARF modules cooperatively regulate 553
lateral root formation (Goh et al., 2012). We observed that factors regulating auxin signaling, 554
such as SOLITARY ROOT (IAA14) could be also involved in de novo root initiation, as we 555
detected decreased root capacity for slr-1, which could be indicative of fewer primordia 556
initiation. In addition, the IAA28 module, which is upstream of lateral root founder cell 557
specification (De Rybel et al., 2010), also regulates de novo root founder cell specification or 558
initiation, although further experiments could dissect more precisely at which stage IAA28 is 559
involved. Our results show that factors primarily involved in formation of lateral roots are also 560
affected in rooting of leaves, suggesting the existence of partially overlapping auxin signalling 561
modules during postembryonic root development. Conversely, cytokinin mutants (ahp1 ahp2 562
ahp3 and arr1 arr10 arr12) showed increased rooting capacity and thus, a repressor role in de 563
novo root initiation can be assigned for these factors, likely in an analogue manner as their role 564
during lateral root initiation (Lavenus et al., 2013; Chang et al., 2015). In agreement with this 565
model, we restored regeneration potential of cytokinin signalling mutants by a moderate 566
increase in auxin levels. 567
PLT1 and PLT2 expression is considered to be a slow read-out of auxin response and 568
prolonged auxin treatment results in PLT1 and PLT2 activation and the de novo specification of 569
WOX5-marked stem cells (Mahonen et al., 2014). We found severe impairment in de novo 570
primordia initiation in shr-2 plt1-4 plt2-2 and WOX5 expression during de novo primordia 571
formation requires auxin input through an unknown pathway (Hu and Xu, 2016). Therefore, it is 572
possible that PLT1 and PLT2 are required for specification of WOX5-marked cells downstream 573
of auxin during de novo organ initiation, which could occur in combination with activity of WOX5 574
transcription factor itself and/or SHORT-ROOT (SHR), in turn acting in an auxin-independent 575
pathway. Further experiments will be required to dissect the molecular pathway including PLTs, 576
SHR and WOX5. 577
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We have also identified specific factors involved in formation of de novo root primordia. 578
We have found that the stem cell regulators BLUEJAY (BLJ), JACKDAW (JKD), SCARECROW 579
(SCR), SHR and SCHIZORIZA (SCZ) regulate ground tissue patterning and vasculature 580
formation prior emergence at the step of dome-shape primordia. Subsequently, more developed 581
primordia are not properly organized in cell layers or rows and by the time of emergence, these 582
defects persist and aggravate. Our results indicate that ground tissue patterning appears to be 583
regulated in newly formed roots at two levels. First, we observed impairment in asymmetric 584
divisions specifying cortex, middle cortex and endodermis in mutants of SHR and SCR, 585
although still a few asymmetric divisions were observed. In agreement, shr mutants have been 586
shown to form endodermis in anchor roots (Lucas et al., 2011), which are a type of adventitious 587
roots. Furthermore, we observed that ground tissue asymmetric divisions were absent in mutant 588
combinations of scr-4 with bjl-1 and jkd-4 and in double mutants jkd-4 scz-1. Interestingly, when 589
we studied if these mutant combinations had defects in lateral roots, which are also organs 590
formed postembryonically, we also observed absence of ground tissue asymmetric divisions 591
suggesting a conserved developmental program for endodermis and cortex specification. 592
Secondly, we observed that SCR, JKD, BLJ and SCZ could function as ground tissue lineage 593
determinants during de novo root organogenesis. The combined action of BLJ, JKD and SCR is 594
required to maintain postembryonically the ground tissue lineage and lacking these three factors 595
results in missing ground tissue initials (hence fewer ground tissue cell rows are observed in 596
cross sections) (Moreno-Risueno et al., 2015). We observed that de novo formed roots in blj-1 597
jkd-4 scr-4 and jkd-4 scz-1 mutants were missing ground tissue cell rows shortly after 598
emergence, which indicates incorrect specification of ground tissue initials during primordia 599
formation or later on in the course of development.. It, thus, possible SCZ, SHR, PLT1 and 600
PLT2 function as lineage or cell fate determinants during postembryonic development, and 601
particularly during de novo organogenesis. 602
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33
MATERIALS AND METHODS 603
Growth conditions 604
Seeds were surfaced-sterilized in 10% (m/v) NaClO and rinsed with sterile water before being 605
transferred to 120×120×10 mm Petri dishes containing 65 mL of one-half-Murashige and Skoog 606
(MS) medium with 1% sucrose and 10 g/L Plant Agar (Duchefa). After two days of stratification 607
at 4°C in darkness, plates were transferred to Panasonic MLR-352-PE growth chamber at 22°C, 608
16/8 photoperiod or continuous light (50 µmol·m-2·s-1). Twelve days after germination, the first 609
pair of leaves was excised across the junction of the petiole with the stem and transferred to 610
Gamborg B5 medium with 2.5% sucrose, 10 g/L Difco Agar (BD) or 3 g/L Gelrite (Sigma) and 611
Gamborg B5 vitamin mixture (Duchefa). Leaves after excision were grown in darkness at 22°C, 612
routinely for 10 days, or for number of days indicated in corresponding experiment. 613
614
Hormonal and inhibition treatments 615
For exogenous hormone treatment, filter-sterilized indole-3-acetic acid (IAA), 6-616
benzylaminopurine (6-BAP) or thidiazuron (TDZ) stock solutions were added to warm growth 617
medium before pouring into plates to provide a final concentration of 1 μM IAA,, 5 μM 6-BAP or 618
4 μM TDZ respectively. N-1-naphthylphtalamic acid (NPA) was applied locally by preparing a 619
lanoline solution containing 1% w/w NPA. For cell cycle arrest, leaf explants were incubated 620
with growth medium supplemented with 0.5, 2.5 or 5 mM hydroxyurea (Sigma). 621
622
Plant material 623
Columbia-0 (Col-0), Landsberg erecta (Ler), and Wassilewskija-2 (Ws-2) accessions were used 624
as a genetic background as corresponding. The reporter lines ProIPT3:GUS, ProIPT5:GUS 625
(Miyawaki et al., 2004), ProLOG4:GUS s (Kuroha et al., 2009), obtained from RIKEN, and 626
ProARR5:GUS (D'Agostino et al., 2000) were used for tracing cytokinin biosynthesis and signaling 627
during rooting. ProYUC8:GUS, ProYUC9:GUS (Hentrich et al., 2013) and ProDR5:GUS (Ulmasov et 628
al., 1997) were used to investigate auxin biosynthesis and signaling; ProPIN3:PIN3:GFP (Xu et 629
al., 2006) and ProPIN4:GUS (Friml et al., 2004) for auxin transport. To trace the molecular 630
mechanisms during de novo root formation we used: ProWIND1:GUS (Iwase et al., 2011) 631
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34
ProWOX11:GUS (Liu et al., 2014), ProWOX5:GUS (Sarkar et al., 2007) and ProSKP2Bs:GUS version 632
(Manzano et al., 2012), which corresponds to a promoter deletion containing 0.5 Kb upstream 633
from the ATG. J0121 and J0661 lines (Laplaze et al., 2005) were used to locate pericycle-like 634
cells during rooting. The following mutant lines were used: wol-1 (Scheres et al., 1995), ahp1 635
ahp2 ahp3 (Hutchison et al., 2006), ahp2 ahp4 ahp5, arr1 arr10 arr12 (Mason et al., 2005), 636
aux1-22 (Bennett et al., 1996), axr2-1 (Timpte et al., 1994), slr-1 (Fukaki et al., 2005), crane-2 637
(Uehara et al., 2008), sur2-1 (Delarue et al., 1998) and iaa28-1 (Rogg et al., 2001) obtained 638
from NASC; pin1, pin2 pin3, pin2 pin3 pin7 (Blilou et al., 2005); fwr (Okumura et al., 2013); 639
cyclinb1;1 (cycb1;1) and cycb1;2 (Nowack et al., 2012) and CDKB1;1 DN161 and Pro35S:KRP2 640
(Boudolf et al., 2004; Verkest et al., 2005). We also used the following stem cell niche mutants, 641
blj-1 jkd-4 scr-4 (Moreno-Risueno et al., 2015), scr-4 (Fukaki et al., 1996), shr-2 ProSHR:SHR:GR 642
(Levesque et al., 2006), plt1-4 plt2-2 (Aida et al., 2004), jkd-4 (Welch et al., 2007), scz-1 643
(Mylona et al., 2002), wox5-1 (Sarkar et al., 2007), and jkd-4 scz-1 and shr-2 plt1-4 plt2-2 644
ProSHR:SHR:GR (generated in this study). 645
646
Genotyping 647
The triple mutant shr-2 plt1-4 plt2-2 ProSHR:SHR:GR was generated by crossing shr-2 648
ProSHR:SHR:GR and plt1-4 plt2-2. F2 seedlings were grown and pre-selected on one-half-MS 649
medium with 10 µM dexamethasone followed by DNA extraction and genotyping by PCR to 650
finally obtain homozygous lines. The oligonucleotides used for genotyping shr-2, while 651
discriminating ProSHR:SHR:GR, were: F 5´-CCAATACCATCCCGCCAC-3´ and R 5´-652
TGAACCGGTCATGCGGTTG-3´; for plt1-4: F 5´-AGACGGCCACGCCAAGAC-3´ and R 653
5´CTAGTATCACGACATTATTTGC-3´; for plt-2-2: F 5´-ACCTACAGTCGTCACTTGTGC-3´ and 654
R 5´-ACTCTTGTCTCGTCATGTTTTTC-3´. T-DNA insertions of PLT1 and PLT2 were amplified 655
using LB 5´-CATTTTATAATAACGCTGCGGACATCTAC-3´ and respective reverse primer. 656
Double mutant jkd-4 scz-1 was generated by crossing jkd-4 and scz-1. DNA extraction from F2 657
seedlings was used for genotyping by PCR. The oligonucleotides used for genotyping jkd-4 658
were: F 5´-GGATGAAAGCAATGCAAAACA-3´ and R 5´-AATGTCGGGATGATGAACTCC-3´. 659
The T-DNA insertion in the jkd-4 line was genotyped using respective forward primer and RB 5´-660
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35
TCAAACAGGATTTTCGCCTGCT-3´. For scz-1 genotyping, a SCZ fragment was amplified 661
using F 5’-CGAAGGTCAAGGCAAAGCTG-3’ and R 5´-GAGCAACAGGCTTGACATGG-662
3´primers, followed by digestion with NlaIII restriction enzyme. Upon digestion, SCZ fragment 663
from scz-1 renders a band of 900 bp, while the wild-type fragment is cut by the enzyme 664
resulting in two fragments of 530 and 360 bp. 665
666
Cell death, GUS staining and microscopy analysis 667
In leaf tissues, dying cells were visualized by lactophenol-trypan blue staining as described in 668
Pavet et al. (2005). For GUS staining, leaf explants were incubated at 37 °C for variable times 669
(4-24 h) in multi-well plates in the presence of the GUS staining solution as described in Perez-670
Perez et al. (2010) or Manzano et al. (2012). To bleach chlorophyll two different methods were 671
used indistinctly with similar results. First method: samples were dehydrated after GUS staining 672
using increasing ethanol concentrations (15, 50, 70, 96 and 100%) 15 minutes in each one, and 673
kept overnight in 100% ethanol. Next, samples were rehydrated following the same ethanol 674
concentration series to 15% ethanol and mounted in 15% glycerol. Second method: samples 675
were fixed in 96% ethanol for 48 h and washed with 0.1 M phosphate buffer (pH 6.8) before 676
being transferred them to clearing solution (80 g chloral hydrate and 30 mL distilled water). Leaf 677
explants were mounted on slides using a mixture of 80 g chloral hydrate, 20 mL distilled water 678
and 10 mL glycerol. After GUS staining, pictures were taken in a Leica DM 2000 with a DCF300 679
camera, or in a bright field Olympus AX70 microscope equipped with an Olympus PM-C35DX 680
microphotography system. The area of proliferating tissues (callus or vasculature) was manually 681
drawn from microscopic images using a Wacon Bamboo tablet and areas or diameters of their 682
best-fitting ellipses were measured with ImageJ (v1.50) software. 683
684
Microscopy analysis 685
Leaf petioles (for studying developmental stages during rooting) and mature embryos, were 686
stained using aniline blue staining method as described in (Bougourd et al., 2000). Processed 687
samples were observed by Leica TCS SP8 laser scanning confocal microscope (Heerbrugg, 688
Switzerland) with the settings described in Bougourd et al. (2000). J0121 fluorescent reporter 689
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36
line was processed prior imaging as indicated in Kurihara et al. (2015). Cross sections (10 µm 690
of thickness) of J0121 petioles were obtained using Vibratome 1000 Plus when indicated. Other 691
fluorescent reporter lines were incubated on methanol:acetone solution for 20 min at −20°C , 692
and immediately transferred to 0.1 M phosphate buffer (pH 6.8). Imaging was performed by 693
confocal microscopy using a Nikon Digital Eclipse C1 equipped with the EZ-C1 control software 694
(Nikon Instruments) or a Leica TCS SP8 laser scanning microscope. GFP was excited with an 695
Argon laser at 488 nm and emission was collected at 505-530 nm, while YFP was excited at 696
514 nm and emission collected at 535-560 nm. To exclude auto-fluorescence contamination, 697
sample emission was collected at 605-675 nm prior excitation with He-Ne laser at 543 nm, and 698
used as a reference for background. Root samples for radial and longitudinal analyses in 699
patterning studies were fluorescently stained with 10 mM propidium iodide (Sigma), and imaged 700
using standard settings on a Leica TCS SP8 confocal. 701
702
Quantification and statistical analysis 703
Regeneration percentage was scored as the number of leaf explants that showed swelling of 704
petioles, proliferation of vascular-associated cells and/or outgrowth of roots at the proximal 705
region of the petiole, at 7 days after excision (dae) or as indicated in corresponding 706
experiments. Rooting percentage was scored as the number of leaf explants showing roots at 707
10 dae or as indicated. Proliferation of vascular-associated cells and number of de novo formed 708
roots were scored on individual leaf samples and used to estimate rooting capacity categories 709
at indicated days in corresponding experiments. Ten dae de novo formed roots were used for 710
establishing differences in tissue patterning using number of cortical cells. Number of cortical 711
cells was quantified using confocal cross sections taken at the site of lateral root cap ending. To 712
determine lateral root capacity, root meristems of 4 days post-imbibition seedlings were 713
removed and number of lateral roots quantified 3 days later. Roots were observed at the 714
microscope to confirm there were not unemerged primordia. Data values referred to % rooting, 715
% regeneration, number of cortical cells, lateral root capacity and vasculature or callus area 716
were statistically analyzed by univariate General Linear Model (GLM) and ANOVA with DMS 717
Post-Hoc test, using IBM SPSS Statistics 21 software. For rooting capacity, Chi-square test was 718
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37
performed to assay if there were differences in distribution frequency between lines, analyzed 719
two by two, using STATGRAPHICS Centurion XVI.I software. Significant differences were 720
collected with 5% level of significance (p-value˂0.05). 721
722
ACKNOWLEDGEMENTS 723
We are especially indebted to M.A. Fernández-López for her expert technical assistance. We 724
thank Dr. M. Pernas for providing scz-1 mutant and advice about its genotyping, and two 725
anonymous reviewers for their useful suggestions. 726
727
Figure S1. Rooting of whole leaves and leaf blades of Arabidopsis wild-type 728
accessions. 729
Figure S2. Vascular proliferation and callus formation during rooting of whole leaves. 730
Figure S3. AHK4, AHP1, AHP2, AHP3, AHP4, AHP5 and ARR1, ARR10 and ARR12 731
cytokinin signaling factors regulate hormone-induced callus formation. 732
Figure S4. IAA7/AXR2 and IAA18 auxin signaling factors regulate hormone-induced 733
callus formation. 734
Figure S5. ALF4 is required for vasculature proliferation and de novo root formation. 735
Figure S6. IAA7/AXR2, CRANE/IAA18 and IAA28 signaling factors regulate de novo 736
root formation in leaf blades. 737
Figure S7. The YUC8 and YUC9 auxin biosynthesis genes are induced in excised 738
leaves. 739
Figure S8. SCARECROW and SHORT-ROOT regulate ground tissue patterning of de 740
novo formed roots. 741
742
743
FIGURE LEGENDS 744
Figure 1. Developmental stages during rooting of Arabidopsis leaves. (A) Whole leaf 745
rooting procedure. (B-H) Confocal microscopy pictures of petioles showing (C-E) proliferation of 746
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38
vasculature cells, (F, G) initiation and formation of primordia, and (H) de novo root emergence. 747
(I-K) J0661-GFP line in sections of petioles at (I, J) excision time or (K) 5 days after excision 748
(dae). (L-P) J0121-GFP line in sections of petioles at (L, M) excision time, (N, O) 2 dae and (P). 749
5 dae. (J, M, O) Transversal sections of petioles at the base. va, vasculature; co, collenchyma; 750
xy, xylem; pc, proliferating cells; RP, root primordium; AR, adventitious root; ph, phloem; pr, 751
procambium. Scale bar: 25 µm. (Q-R) Leaves of wild type and cell-cycle mutants, or upon 752
treatment with hydroxyurea (HU), were assessed for (Q) vasculature proliferation and (R) 753
rooting at 7dae. Asterisks: statistically significant (p-value <0.05) by General Linear Model 754
(GLM) and DMS Post-Hoc test 755
756
Figure 2. Cytokinin acts locally during de novo root regeneration. (A-C) Expression of (A) 757
ProIPT3:GUS, (B) ProIPT5:GUS, (C) ProLOG4:GUS and (D-E) ProARR5:GUS in petioles at indicated 758
days after excision (dae). (E) Leaves were incubated with 5 µM 6-benzylaminopurine (6-BAP). 759
(F) Mutants impaired in cytokinin signalling show defective regeneration (which accounts for 760
petioles with vasculature proliferation, primordia and roots; yellow barplots) and rooting capacity 761
(multi-coloured barplots show frequencies of vasculature proliferation and of roots or primordia, 762
indistinctly) at 7 dae. (G) Cytokinin signalling mutant leaf explants incubated with 1 µM indol-3-763
acetic acid (IAA) and measured for root regeneration responses as in (F). Scale bars: (A-C) 0.5 764
mm, (D-E) 0.25 mm. Asterisks: p-value<0.05 by General Linear Model (GLM) followed by DMS 765
Post-Hoc or Chi-square test. 766
767
Figure 3. Distinctive auxin signaling pathways regulate de novo root regeneration. (A-B) 768
Auxin response reported by (A) ProDR5:GUS expression in Arabidopsis leaf petioles and (B) 769
quantification of GUS stained area; dae, days after excision. (C) ProDR5:GUS expression and de 770
novo formed roots (AR) in leaves supplemented with 1 µM IAA (left) and in the sur2-1 auxin 771
overproduction mutant (right). (D) Regeneration percentage and (E) rooting capacity at 7 days 772
after excision of auxin signaling mutants (nph4-1 arf19-1, axr2-1, slr-1, crane-2, and iaa28-1) 773
and auxin overproducer sur2-1. (F) Vasculature proliferation area on Arabidopsis leaves 774
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39
cultivated for 14 days in hormone-free medium. Scale bars: 200 μm. Asterisks: p-value<0.05 by 775
General Linear Model (GLM) followed by DMS Post-Hoc or Chi-square test. 776
777
Figure 4. Polar auxin transport from the leaf blade to the petiole is required for rooting. 778
(A-B) Distal blade excision reduces (A) regeneration rate and rooting capacity at 7 days after 779
excision (dae and (B) ProDR5:GUS expression at 12 hours after excision (hae) and 3 dae. (C-D) 780
Local treatment at the blade-petiole junction with 1% N-1-naphthylphthalamic acid (NPA) 781
reduces (C) regeneration rate and rooting capacity at 7 dae, and (D) ProDR5:GUS expression. 782
(E-G) Expression of (E) ProPIN4:GUS and (F-G) ProPIN3:PIN3:GFP Arrowheads point to 783
membrane-localized PIN-GFP. Scale bars: (B, D, F, G) 0.2 and (E) 0.5 mm. (H) Regeneration 784
and rooting capacity in auxin transport mutants at 7 dae. Asterisks: p-value<0.05 by General 785
Linear Model (GLM) followed by DMS Post-Hoc or Chi-square test. 786
787
Figure 5. Root founder cells establish on endogenous callus prior de novo primordium 788
formation. (A-F) Expression of (A) ProWIND1:GUS, (B, E, F) ProWOX11:GUS, (C) ProSKP2Bs:GUS 789
and (D) ProWOX5:GUS in leaf petioles at indicated days after excision (dae). (E) Control 790
conditions or (F) upon 5 mM hydroxyurea (HU) treatment. GUS staining time in (E) was set for 791
no expression; xy, xylem; pc, proliferating cells; va, vasculature; RFC, root founder cell; RP, root 792
primordium; NR, de novo formed root. Scale bars: 200 µm. (G) Regeneration percentage and 793
rooting capacity at 7 dae of mutants impaired in cell cycle progression and hydroxyurea (HU)-794
treated Col-0 leaves. Asterisks: p-value<0.05 by General Linear Model (GLM) followed by DMS 795
Post-Hoc or Chi-square test. 796
797
Figure 6. SHORT-ROOT, PLETHORA1, PLETHORA2, JACKDAW, BLUEJAY, SCARECROW 798
and SCHIZORIZA are required for de novo root primordia initiation and formation. (A) 799
Rooting percentage and (B) rooting capacity at 10 days after excision (dae) for loss-of-function 800
mutants in stem cell regulators. Asterisks: p-value<0.05 by General Linear Model (GLM) 801
followed by DMS Post-Hoc or Chi-square test. (C) Triple shr-2 plt1-4 plt2-2 mutant does not 802
regenerate roots at 10 dae. (D) Confocal images of control and shr-2 plt1-4 plt2-2 during root 803
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40
regeneration. Malformed primordia (3%, n=90) in shr-2 plt1-4 plt2-2 do not emerge through 804
petiole tissues. (E) Confocal images of de novo root primordia of Wild type, jkd4-4 scz-1 and blj-805
1 jkd-4 scr-4 at 5 and 6 dae; co, collenchyma; AR, adventitious root; pc, proliferative cells; RP, 806
root primordium; xy, xylem; black dashed line corresponds to cells in the position of quiescent 807
center; white dashed line cells in the position of the ground tissue and in contact to quiescent 808
center; green asterisk: cortex ; turquoise asterisk: middle cortex; yellow asterisk: endodermis. 809
Scale bars: (C) 5 mm and (D-I) 50 µm. 810
811
Figure 7. JACKDAW, BLUEJAY, SCARECROW, SHORT-ROOT and SCHIZORIZA regulate 812
patterning of postembryonic roots. (A-B) Confocal (A) longitudinal and (B) transversal 813
sections of de novo formed roots at 10 days after excision (1-3 days post emergence). 814
Transversal sections were taken at end of lateral root cap. Dashed lines and asterisks: green, 815
cortex; turquoise, middle cortex; yellow, endodermis; brown, stele; white and m: mutant 816
undivided ground tissue; xy, xylem. (C) Number of cortical-like cell or undivided mutant cell rows 817
in control accessions, jkd-4 scz-1 and blj-1 jkd-4 scr-4. (D) Lateral root capacity in Col-0, shr-2 818
plt1-4 plt2-2, jkd-4 scz-1 and blj-1 jkd-4 scr-4 and at 4 days after seed imbibition. (E) Confocal 819
longitudinal sections of emerged lateral roots of Col-0, shr-2 plt1-4 plt2-2, jkd-4 scz-1 and blj-1 820
jkd-4 scr-4 at 7 days after seed imbibition. Scale bars: 50 μm. Asterisks: p-value<0.05 by 821
General Linear Model (GLM) and DMS Post-Hoc test. 822
823
Figure 8. Model of de novo root regeneration in Arabidopsis leaves. (A) Auxin is 824
transported to the petiole base and acts in combination with locally-produced cytokinin to induce 825
endogenous callus formation; LOG4, IPT3, IPT5: cytokinin biosynthesis; ARR5: cytokinin 826
response; YUCCA8/9: auxin synthesis; PIN1/2/3/7, AUX1: auxin transport; DR5: auxin 827
response. (B) Stages of de novo formation of roots based on morphological changes and 828
expression of markers. (C) Regulation of distinctive stages of de novo organogenesis based on 829
mutant phenotypes. Regulation or functions which could not be directly assigned are indicated 830
as hypothetical. 831
832
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41
833
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