Lateral root initiation and formation within the parental root ...

Annals of Botany 122: 873–888, 2018doi: 10.1093/aob/mcy052

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Lateral root initiation and formation within the parental root meristem of Cucurbita pepo: is auxin a key player?

Elena L. Ilina1, Alexey S. Kiryushkin1, Victoria A. Semenova1, Nikolay P. Demchenko1, Katharina Pawlowski3 and Kirill N. Demchenko1,2,*

1Laboratory of Cellular and Molecular Mechanisms of Plant Development, Komarov Botanical Institute, Russian Academy of Sciences, Professora Popova Str. 2, 197376, Saint-Petersburg, Russia, 2Laboratory of Molecular and Cellular Biology, All-

Russia Research Institute for Agricultural Microbiology, Podbelsky chaussee 3, 196608, Pushkin 8, Saint-Petersburg, Russia and 3Department of Ecology, Environment and Plant Sciences, Stockholm University, 106 91 Stockholm, Sweden

*For correspondence. E-mail [email protected]

Received: 31 January 2018 Returned for revision: 20 February 2018 Editorial decision: 19 March 2018 Accepted: 20 March 2018 Published electronically 19 April 2018

• Background and Aims In some plant families, including Cucurbitaceae, initiation and development of lateral roots (LRs) occur in the parental root apical meristem. The objective of this study was to identify the general mechanisms underlying LR initiation (LRI). Therefore, the first cellular events leading to LRI as well as the role of auxin in this process were studied in the Cucurbita pepo root apical meristem.• Methods Transgenic hairy roots harbouring the auxin-responsive promoter DR5 fused to different reporter genes were used for visualizing of cellular auxin response maxima (ARMs) via confocal laser scanning microscopy and 3-D imaging. The effects of exogenous auxin and auxin transport inhibitors on root branching were analysed.• Key Results The earliest LRI event involved a group of symmetric anticlinal divisions in pericycle cell files at a distance of 250–350 µm from the initial cells. The visualization of the ARMs enabled the precise detection of cells involved in determining the site of LR primordium formation. A local ARM appeared in sister cells of the pericycle and endodermis files before the first division. Cortical cells contributed to LR development after the anticlinal divisions in the pericycle via the formation of an ARM. Exogenous auxins did not increase the total number of LRs and did not affect the LRI index. Although exogenous auxin transport inhibitors acted in different ways, they all reduced the number of LRs formed.• Conclusions Literature data, as well as results obtained in this study, suggest that the formation of a local ARM before the first anticlinal formative divisions is the common mechanism underlying LRI in flowering plants. We propose that the mechanisms of the regulation of root branching are independent of the position of the LRI site relative to the parental root tip.

Key words: Auxin, Cucurbita pepo (squash), DR5, endodermis, lateral root, lateral root initiation, pericycle, root meristem, root primordia.

INTRODUCTION

Higher plants continually produce new lateral roots (LRs) throughout their lifetime, resulting in the generation of a root system. The main functions of roots are to supply the plant with water and mineral compounds and to anchor the plant in the soil. The analysis of how the different positions for LR initiation in the parental root are related is important for elu-cidating how Tracheophyta adapted to terrestrial life and for understanding the general evolution of organ development. For many years, Peter Barlow devoted his research to these issues (Barlow, 1984; Barlow and Adam, 1988; Newson et al., 1993; Zhang et al., 1999; Barlow et al., 2004).

The ancestral form of root branching is dichotomous, as shown by fossils of the ancestors of current ferns and Lycopodiophyta. In extant plants, the dichotomous branching occurs via a division of the apical meristem. Angiosperms exhibit monopodial root branching, i.e. the parental root forms the main axis from which

LRs emerge (Fig. 1), and LRs initiate in the pericycle, the outer layer(s) of the vascular cylinder (Esau, 1977). It is not clear how dichotomous and monopodial branching evolved from a com-mon ancestor (Dubrovsky and Laskowski, 2017). Since all ances-tral forms of root branching show endogenous LR initiation and development, it is assumed that LR initiation is triggered by com-pounds from the vascular cylinder (Goebel, 1928; Voronin, 1964).

The positions along the longitudinal axis of the parental root where founder cells are formed and the LR is initiated can dif-fer considerably among angiosperm groups. LR initiation and development can take place either in the apical meristem of the parental root or proximal to the elongation zone at a significant distance from the root tip (Table 1), the latter of which is typical for Arabidopsis thaliana (arabidopsis) and other Brassicaceae species, and several cereals.

The tissues involved in primordium formation can also differ. In arabidopsis, three pericycle cell files participate

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Ilina et al. – Auxin in lateral root initiation of Cucurbita pepo874

in primordium formation (Casimiro et al., 2003), while in Solanum lycopersicum (Ivanchenko et al., 2006) and in cere-als such as Triticum aestivum (Demchenko and Demchenko, 2001a), not only pericycle but also endodermal cell files are normally involved. Among Cucurbitaceae species, pericycle, endodermal and cortical cell files from several layers contrib-ute to primordium formation (Demchenko and Demchenko, 2001a).

During LR initiation in arabidopsis, lateral root primordia (LRPs) originate from a sub-set of pericycle founder cells, which undergo asymmetric anticlinal divisions (Casero et al., 1993, 1995; Dubrovsky et al., 2001; Casimiro et al., 2003). These divisions occur in three adjacent pericycle cell files

(Casimiro et al., 2003). In the basal part of the elongation zone, all pericycle cells are in proliferative arrest. Only such peri-cycle cells that had left the basal portion of the root apical meri-stem in the G1 phase of the cell cycle can subsequently resume the cycle, but only some will be involved in the initiation of LRPs (Demchenko and Demchenko, 2001b; Himanen et al., 2002; Vanneste et al., 2005; Alarcón et al., 2016). An interest-ing exception to this rule may be observed when the parental arabidopsis root is mechanically stimulated, resulting in the formation of a new LR outside of the acropetal root branching zone (Ditengou et al., 2008).

In some plant families, including Cucurbitaceae, LRP initi-ation and development occur directly in the apical meristem of the

A B C D E

Fig. 1. Lateral root primordia and the first cellular events in lateral root initiation in Cucurbita pepo DR5::GUS root tips. Visualization of auxin response maxima (ARMs) and GUS activity in parental roots; blue staining. (A) Overview of the parental root tip. (B) Longitudinal vibratome section of the root presented in (A). (C–E) Longitudinal section of a root embedded in plastic resin and counterstained with ruthenium red. (A) The ARMs are visible in the root cap, initial cells and in the developing lateral root primordia in the parental root apical meristem and elongation zone. (B) A close-up shows ARMs in the root tip, initial cells and in lateral root primordia (white arrows) in the parental root apical meristem. (C) Longitudinal root section and its close-ups (D, E) depict cell packets resulting from anticlinal divisions (black outline) as well as the first periclinal divisions in the pericycle (green outline). (D) Approximately 300 µm proximal to the initial cells, ARMs appear in pairs of pericycle sister cells and in the adjacent protoxylem. (E) Approximately 500 µm proximal to initial cells, anticlinal divisions have taken place in pairs of pericycle and endodermis sister cells (asterisks). The first periclinal divisions (green outline) in the central pericycle cells have been completed.

en, endodermis; pI, pericycle I; pII, pericycle II; px, protoxylem. Scale bars = 500 µm (A), 125 µm (B), 20 µm (C) and 10 µm (D, E)

Table 1. Lateral root initiation in parental root meristem

Class/Clade Order Family Species Reference

Angiosperms Eudicots Rosids Cucurbitales Cucurbitaceae Cucurbita pepo Gulyaev (1964)Cucurbita maxima Mallory et al. (1970)Cucumis sativus Dubrovsky, (1986b, 1987)

Superasterids Caryophyllales Polygonaceae Fagopyrum esculentum O’Dell and Foard (1969)Lamiids Solanales Convolvulaceae Ipomea purpurea Seago (1973)

Liliopsida (Monocots)

Commelinidae Commelinales Pontederiaceae Pontederia cordata Charlton (1975, 1983a, 1987)Eichhornia crassipes Mallory et al. (1970); Clowes (1985)

Alismatidae Alismatales Araceae Pistia stratiotes Clowes (1985); Charlton (1987)Butomaceae Butomus umbellatus Zhupanov and Brykov (2014)

Zhupanov and Brykov (2014)Alismataceae Sagittaria sagittifoliaPteridophyta Pteridopsida Polypodiales Pteridaceae Ceratopteris thalictrides

Ceratopteris richardiiMallory et al. (1970); Charlton (1983a)Hou et al. (2004); Hou and Blancaflor

(2009)Salviniales Marsileaceae Marsilea quadrifolia Charlton (1983a)

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parental root (Table 1). These species exhibit early and rapid mono-podial branching of the main root. Additionally, the initiation of the first LRP in Cucurbitaceae takes place already during embryo-genesis, and the first 3–5 groups of LRPs can be observed in the radicle (Dubrovsky, 1987; Dubrovsky and Laskowski, 2017).

The phytohormone auxin functions as a key regulator of LR development. Specifically, it contributes to the priming of the pericycle cells that will initiate LRs (Dubrovsky et al., 2008; Overvoorde et al., 2010; Parizot and Beeckman, 2012; Du and Scheres, 2018). There is considerable experimental evidence supporting the importance of auxin for the initiation and devel-opment of LRs (Vanneste and Friml, 2009; De Smet, 2012). Briefly, the addition of exogenous auxin to the growth medium led to the formation of new LRPs unrelated to acropetal branch-ing in nearly all examined plant species, including arabidopsis (Lloret and Casero, 2002). The auxin-overproducing arabi-dopsis mutant superroot produced an excessive amount of adventitious roots and LRs (Boerjan, 1995). Additionally, the overproduction of auxin in transgenic Petunia plants express-ing agrobacterial auxin biosynthesis genes enhanced the pro-duction of adventitious roots (Klee et al., 1987). Moreover, the application of the auxin transport inhibitor naphthylphthalamic acid (NPA) at the arabidopsis root–shoot junction decreased the number and density of LRs (Reed et al., 1998).

Analyses of auxin distribution in plants often involve the use of synthetic auxin-inducible promoters such as DIRECT REPEAT 5 (DR5; Ulmasov et al., 1997) as well as its variant DR5rev (Friml et al., 2003) and the novel version DR5v2 (Liao et al., 2015). The DR5–reporter gene system has been used to conduct studies on auxin distribution and transport in arabidopsis and many other species, including Nicotiana tab-acum (Chen et al., 2010), Cucurbita pepo (Ilina et al., 2012), rice (Zhou et al., 2014), the tree Populus tremula (Chen et al., 2013) and the moss Physcomitrella patens (Bierfreund et al., 2003). A fusion of DR5 and the GUS (β-glucuronidase) re-porter gene was also used to analyse the auxin response dur-ing LR development in arabidopsis (Casimiro et al., 2001; Benková et al., 2003). Importantly, the DR5 promoter activity pattern coincides with local maxima of auxin levels in arabi-dopsis cells and tissues (Benková et al., 2003). Using trans-genic arabidopsis plants with clonal sectors simultaneously expressing iaaM and the GUS reporter gene due to the action of a heat shock-inducible Cre recombinase gene, Dubrovsky et al. (2008) could demonstrate that the localized accumula-tion of GUS in response to auxin production precedes cell divisions during LRP initiation. Similar results were obtained in in vivo investigations of cellular events during LR initi-ation in a DR5::GFP-NLS arabidopsis line (De Rybel et al., 2010). Thus, auxin accumulation consistently precedes the first asymmetric division in the pericycle.

Additionally, immunolocalization has been applied to monitor auxin distribution (Benková et al., 2003; Schlicht et al., 2006). This method was used to analyse the distribu-tion of auxin in the root apical meristem of Cucurbita maxima where LR initiation occurs (Chiappetta et al., 2011). However, the method did not yield clear results in that it did not show any auxin maxima in the root apical meristem.

In spite of significant recent advances in the understand-ing of LR initiation in arabidopsis and cereals, several ques-tions remain unanswered. For example, it is unclear whether

a general physiological mechanism of LR initiation exists or whether different positions of LR initiation relative to the tip of the parental root indicate different mechanisms. LR initiation above the apical meristem of the parental root requires cells from the elongation zone to re-enter the cell cycle, and involves not only the pericycle, but also stelar parenchyma and endoder-mis (Demchenko and Demchenko, 2001a).

Most of the published studies on the genetics and physiology of LR initiation were carried out on plant species that initiate LRs above the elongation zone, mostly on arabidopsis. Thus, the genetic and physiological mechanisms mediating LR initi-ation in plant species where this process occurs in the root apical meristem remain poorly understood. Furthermore, determining whether both variants of LR initiation are closely related (i.e. both are derived from dichotomous branching) or if one of them evolved independently requires a more thorough understanding of the mechanism underlying LR initiation in meristems. In the present study, the role of auxin during the first steps of LR initi-ation and development was analysed in squash (Cucurbita pepo). The cellular response to auxin was examined in composite squash plants with transgenic roots expressing constructs comprising fluorescent reporters under control of the DR5 promoter. The first events leading to LRP initiation and formation were revealed for parental root meristem cells. Moreover, we examined the effects of exogenous auxin and auxin transport inhibitors on squash root branching patterns and on the initiation of LRPs.

MATERIALS AND METHODS

Plant materials and bacterial strains

Squash (Cucurbita pepo L. var. giromontina) cv. Beloplodniy and cucumber (Cucumis sativus L.) cv. Konkurent (Sortsemovosch, St. Petersburg, Russia) were used in this study. Squash seed-lings were transformed using Agrobacterium rhizogenes (strain R1000) as previously described (Ilina et al., 2012). Escherichia coli strain XL-1 Blue was used for molecular cloning.

Design of destination clones for plant transformations

The pKGW-RR-MGW-DR5 (DR5::GUS) construct con-taining the auxin-inducible synthetic promoter DR5, the gusA reporter gene and the pAtUBQ10::DsRED1 cassette as a screen-able marker was developed in a previous study (Ilina et al., 2012). To create the pKGW-RR-MGW-pCsSCR (CsSCR::GUS) construct, a 1684 bp fragment of the promoter of the C. sativus SCARECROW (SCR) gene (Wiśniewska et al., 2013) was ampli-fied by PCR using cucumber genomic DNA. The construct was cloned into pUC18-entry8 (Hornung et al., 2005). Additionally, pCsSCR was incorporated into the pKGW-RR-MGW destin-ation vector with an LR clonase reaction (Gateway LR Clonase II, Thermo Fisher Scientific, Waltham, MA, USA).

A series of entry vectors was generated to create the 242_pKGW-RR-MGW-DR5::NLS-GFP [DR5::eGFP-NLS; containing a nuclear localization signal (NLS)] and 236_pKGW-MGW-pAtUBQ10::H2B-Venus-DR5::tdTomato-H2B [DR5::tdTomato-H2B; containing the human histone H2B gene

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(Nam and Benezra, 2009)] constructs. A list of plasmids and vectors used is given in Table 2.

The destination binary vectors 242_pKGW-RR-MGW and 236_pKGW-MGW were kindly provided by Erik Limpens (Wageningen University, Wageningen, The Netherlands). The LR plus clonase reactions (Gateway LR Clonase II Plus, Thermo Fisher Scientific), which were prepared according to the manufacturer’s instructions, involved the entry and destin-ation vector combinations listed in Table 3. The 373_pENTRat-tR2attL3-T35S construct (Thermo Fisher Scientific) was used as the donor of the T35S terminator in all reactions. To create the intermediate construct 236_pKGW-MGW-UBQ10::H2B-Venus, the pAtUBQ10::H2B-Venus cassette was PCR ampli-fied using the 242_pKGW-RR-MGW-pAtUBQ10::H2B-Venus construct as the template and cloned into 236_pKGW-MGW at the KpnI restriction sites.

The genetic fusions in all constructs were verified by PCR amplification and sequencing of the products. Primer sequences are provided in Supplementary Data Table S1.

β-Glucuronidase activity assays

The GUS reporter was localized in transgenic hairy roots con-taining DR5::GUS or CsSCR::GUS by the histochemical stain-ing of the GUS reaction product as previously described (Ilina et al., 2012). Stained transgenic and wild-type root tips (3–4 mm long) were fixed using 3 % paraformaldehyde, 0.25 % glutaral-dehyde, 0.1 % Tween-20 and 0.1 % Triton X-100 in stabilizing buffer (15 mm 1,4-piperazinediethane sulphonic acid (PIPES), 1.5 mm MgSO4, 1.5 mm EGTA; pH 6.9), dehydrated in a graded ethanol series and embedded in Technovit 7100 hydrophilic resin (Heraeus Kulzer, Wehrheim, Germany) according to the manu-facturer’s instructions. Longitudinal sections (7 µm thick) were prepared using a Microm HM 360 rotary microtome equipped

with a tungsten carbide D-profile microtome knife (Thermo Fisher Scientific). The cell walls were counterstained for 30 min with 0.01 % ruthenium red (Sigma-Aldrich, St. Louis, MO, USA) in 0.1 m borate buffer, pH 9.2 (Gutierrez-Gonzalvez et al., 1984). The sections were mounted with Eukitt Mounting Medium (Sigma-Aldrich) under cover slips. Alternatively, GUS-stained and fixed root tips (7–10 mm long) were embedded in 1 % agarose, and sections (65 µm thick) were prepared with a Microm HM 650V vibrating-blade microtome (Thermo Fisher Scientific) equipped with a sapphire knife.

Fluorescent protein reporter assay

To localize the enhanced green fluorescent protein (eGFP)–NLS, Venus–H2B and tdTomato–H2B reporters, transgenic root tips (7–10 mm long) were gently infiltrated with a fixa-tive (McLean and Nakane, 1974) modified by Brian Lin (Tufts University, Boston, MA, USA) [1 % paraformaldehyde, 5 % dimethylsulphoxide (DMSO), 0.1 m l-lysine and 0.01 m sodium m-periodate in 0.05 m phosphate buffer pH 7.2]. The root tips were fixed for 1 h at room temperature, rinsed with 0.1 m phosphate buffer, and then sectioned with a vibrating-blade microtome as previously described. Nuclei and cell walls were counterstained for 30 min with 0.3 µg mL−1 4′,6-diamidino-2-phenylindole (DAPI) and 0.01 % Pontamine Fast Scarlet 4B (Sigma-Aldrich), respectively. Sections were mounted in con-secutive order on slides in a non-hardening antifade mountant CFMR2 (Citifluor, London, UK) under coverslips.

Detection of DNA-replicating cells

5-Ethynyl-2′-deoxyuridine (EdU), a thymidine analogue that is incorporated into DNA during the S-phase of the

Table 2. Construction of entry vectors

Name of insert Template Source of template Name of vector used Source of vector used

Resulting entry vector

DR5 promoter pJET1.2-DR5 Ulmasov et al. (1997); Ilina et al. (2012)

369_pENTRattL4attR1_ BSAI

Thermo Fisher Scientific

DR5-369_ pENTRattL4attR1_BSAI

Arabidopsis thaliana UBQ10 promoter

Addgene plasmid # 61168

Ivanov and Harrison (2014) 369_pENTRattL4attR1_ BSAI

Thermo Fisher Scientific

pAtUBQ10-369_ pENTRattL4attR1_BSAI

NLS-Egfp-gusA reporter fusion

pMK7S*NFM14G plasmid

Karimi et al. (2007) pUC18-entry8 Hornung et al. (2005)

NLS-Egfp-gusA–pUC18- entry8 vector

H2B-Venus Addgene plasmid # 20971

Nam and Benezra (2009) pUC18-entry8 Hornung et al. (2005)

H2B-Venus–pUC18-entry8

tdTomato-H2B Addgene plasmid # 58102

A gift from Michael Davidson

pUC18-entry8 Hornung et al. (2005)

tdTomato-H2B–pUC18- entry8

Table 3. Construction of binary vectors

Binary vector Destination vector Promoter in entry vector Reporter in entry vector

242_pKGW-RR-MGW-DR5::NLS-GFP 242 pKGW-RR-MGW DR5-369_pENTRattL4attR1_BSAI NLS-Egfp-gusA–pUC18-entry8242_pKGW-RR-MGW-pUBQ10::

H2B-Venus242_pKGW-RR-MGW pAtUBQ10-369_pENTRattL4attR1_

BSAIH2B-Venus–pUC18-entry8

236_pKGW-MGW-pAtUBQ10:: H2B-Venus-DR5::tdTomato-H2B

236_pKGW-MGW-UBQ10::H2B-Venus

DR5-369_pENTRattL4attR1_BSAI tdTomato-H2B–pUC18-entry8

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mitotic cycle, was used for cell proliferation assays. A pre-viously described method was used for the incorporation of EdU (30 min) and fixation of root tips (5–8 mm long) (Ilina et al., 2012). The fixed root tips were embedded in Steedman’s wax (Stumpe et al., 2006). Serial longitudinal sections (10 µm) were prepared with a Microm HM 360 rotary microtome (Thermo Fisher Scientific) equipped with single-use 35SEC-p blades. Sections were placed on slides and processed as described previously (Ilina et al., 2012). The incorporation of EdU was detected with the Alexa Fluor® 488 dye of the Click-iT® EdU Alexa Fluor® 488 Imaging Kit (Thermo Fisher Scientific) (Salic and Mitchison, 2008; Ilina et al., 2012). Nuclei were counterstained with 0.3 µg mL−1 propidium iodide in Tris-buffered saline (50 mm Tris–HCl, pH 7.3 and 150 mm NaCl).

Microscopy

Sections of wild-type and transgenic hairy roots stained for GUS activity were examined under an AxioImager.Z1 micro-scope (Carl Zeiss, Germany) using Plan-Apochromat objec-tives and bright-field microscopy. The same microscope was used for the localization by indirect fluorescence, with filter set 09 (EX 450–490 nm, EM LP 515 nm) applied for Alexa Fluor® 488 and propidium iodide. An MRc5 digital camera and ZEN2pro software (Carl Zeiss) were used for imaging. Optical sectioning with a structured illumination was performed with an AxioImager.Z1 microscope equipped with an ApoTome mod-ule and an AxioCam HR3 monochromatic camera (Carl Zeiss) using filter set 46 HE (EX BP 500/25 nm, EM BP 535/30 nm) for Alexa Fluor 488 and filter set 00 (EX 530–585 nm, EM LP 615 nm) for propidium iodide.

Composite plants were examined using a SteREO Lumar.V12 fluorescent stereomicroscope equipped with an MRc5 digital camera (Carl Zeiss). Filter set 43 HE (EX BP 550/25, EM BP 605/70) was used to observe DsRED1 or tdTomato fluorescence, while filter set 38 HE (EX BP 470/40, EM BP 525/50) was used to observe eGFP or Venus fluorescence. Total root systems and roots stained for GUS activity were examined with the same stereomicroscope using bright-field microscopy.

Examination of the localization of fluorescent proteins was performed using an LSM 780 confocal laser scanning micro-scope (Carl Zeiss) with the 488 nm laser line for eGFP or Venus, the 561 nm laser line for tdTomato or Pontamine Fast Scarlet 4B, and the 405 nm laser line for DAPI-stained nuclei. Three-dimensional reconstructions and animations as well as maximum intensity projections were obtained using ZEN2pro software (Carl Zeiss) on a Z800 graphic workstation (Hewlett Packard, Palo Alto, CA, USA). For optical sectioning, the pin-hole diameter setting was varied between 1 and 2 Airy units. For background elimination, the linear spectral unmixing tech-nique in the ZEN2pro software (Carl Zeiss) was used.

Application of exogenous auxins and auxin transport inhibitors

Naphthylacetic acid (NAA) or indole-3-butyric acid (IBA) were used as exogenous auxins at 0.1–100 nm. The auxin trans-port inhibitors used were NPA (0.1–1 µm), 2,3,5-triiodobenzoic

acid (TIBA; 1–100 µm) and 1-naphthoxyacetic acid (1-NOA; 25–100 µm). The concentrations of auxin transport inhibitors and auxins used in this study were selected empirically after testing published concentrations (Shen et al., 1988; Kamada et al., 2003; Laňková et al., 2010; Novickienė et al., 2010; Strader et al., 2011) to avoid concentrations that inhibited primary root growth.

Three-day-old seedlings with approx. 1 cm long roots were incubated in aerated 1/4 strength Hoagland’s medium supple-mented with the respective agents for 96 h at 25 °C under a 16 h light period. The root length of each seedling was meas-ured at the plating stage, after 48 and 96 h of incubation. Whole root systems were fixed and stained with Schiff’s reagent (Demchenko et al., 2004). The total numbers of LRs and LRPs were determined with n = 37–61 per experiment. The LR ini-tiation index (ILRI; Dubrovsky et al., 2009) was estimated in 1 cm root tips. Longitudinal root sections, 50 µm thick, were prepared using a vibrating-blade microtome, and mounted on slides under a coverslip. The number of cells in the three outer cortical layers (independently) between two of the most prox-imal LRPs was determined for the region located 2–6 mm from the root tip above the division zone of the cortex (Demchenko and Demchenko, 2001a). The ILRI was calculated as the number of LRPs corresponding to 100 cortical cells in one cell layer, the cortical cell file, with n = 12–33. Each experiment included at least 50 seedlings and was repeated at least three times inde-pendently. Data for individual experiments are given.

Statistical analyses

Data were analysed using Sigma Plot v.12.2 software (Systat Software, San Jose, CA, USA). Means were compared using one-way analysis of variance (ANOVA) on ranks (Kruskal–Wallis one-way ANOVA).

RESULTS

The first cellular events in lateral root initiation in Cucurbita pepo roots

Formative divisions occur within the parental root meri-stem. The localization of the first events during LR initiation in squash was performed using transgenic roots (DR5::GUS) of composite squash plants (Fig. 1). The distribution of the early stages of LR initiation can be shown based on the localization of GUS expression. The LRPs were initiated opposite the xylem poles (Fig. 2). The number of xylem poles in different squash roots varied from three to five. Primordia could be initiated in groups of four or five (Fig. 1A). Within a group, primordia were located nearly opposite each other.

Squash roots had two pericycle layers opposite xylem poles, namely the inner pericycle (pericycle I) and the outer pericycle (pericycle II) (Figs 1C–E and 2). These two layers began to form at a distance of 100–150 µm proximal to the initial cells.

The earliest LR initiation event detectable at the histological level was symmetric anticlinal divisions in two cells in two peri-cycle cell files (two cells from the inner pericycle and two from the outer pericycle) opposite a xylem pole (Fig. 1C, D). At a dis-tance of 250–350 µm from the initial cells, these divisions led to

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the formation of cell packets [stage I; see Malamy and Benfey (1997) and Benková et al. (2003)], representing two pairs of sister cells in each cell file. These divisions could be defined as forma-tive (Figs 1C, D and 3A; see Dubrovsky et al., 2001). Thus, the zone of LR initiation was determined as the root area 250–350 µm proximal to the initial cells. At a distance of approx. 500 µm prox-imal to the initial cells, the central cells of the packet underwent periclinal divisions (Figs 1C, E and 3B, C), resulting in the visually detectable formation of an LRP (stage II). Importantly, the forma-tion of founder cells occurred within three files of the inner peri-cycle cell layer and within 3–4 files of the outer pericycle cell layer (Fig. 2A), located opposite one protoxylem cell (Figs 1D and 2B).

Lateral root initiation occurs among proliferating cells of the parental root meristem. The distribution of DNA-synthesizing and dividing cells was analysed using longitudinal sections of the parental root (Fig. 3). Primordium formation stages 0–II occurred in groups of actively proliferating cells in different tissues. In pericycle cell files, DNA synthesis and cell division occurred throughout the LR initiation zone. Pericycle cells op-posite a xylem pole exited the mitotic cycle at a distance of 1000–1200 µm proximal to the initial cells of the respective cell files, i.e. after the formation of LRPs. Mitotic figures and DNA-synthesizing nuclei were detected in the pericycle and endo-dermis between the early stages of LRP formation in the parental root apical meristem as well as in stelar parenchyma cells of the vascular cylinder and in the cortical cell files (Fig. 3A, C). Thus, the early stages of founder cell specification and succes-sive formative divisions occurred within the area of actively proliferating cells of different tissues of the parental root. An analysis of the distribution of proliferating cells within peri-cycle cell files in the LR initiation zone showed that the founder cells did not leave the mitotic cycle prior to the acquisition of competence. Consequently, the formative divisions were not due to the founder cells re-entering the cell cycle. The average length of pericycle cells in the prophase was the same regard-less of whether the cells were involved in LR initiation (data not shown). Thus, the duration of the first mitotic cycles was unchanged in founder cells. If the duration of the mitotic cycle was significantly shorter in founder cells than in neighbouring

cells, the average length of the founder cells in prophase would have decreased significantly.

Cortical cell layers within the parental root meristem partici-pate in primordium formation. Adjacent endodermal layers became involved in LRP formation after the completion of anti-clinal divisions of founder cells in the pericycle at a distance of approx. 500 µm from the initials (stage I; Figs 1D and 2A). Periclinal divisions in the outer pericycle layer could be accom-panied by radial anticlinal divisions in the inner pericycle layer, resulting in an increase of the number of inner pericycle cell files (Fig. 2A). The subsequent involvement of inner cortical cell files occurred after the completion of the first periclinal divisions in the pericycle (stage II; Fig. 2A). The last mitosis in the cortical cells of the parental root meristem was observed at a distance of 491 ± 38 μm proximal to the initials (n = 126). After resumption of the cell cycle at a distance of approx. 700 µm proximal to the initials, cortical cells underwent several rounds of anticlinal (Fig. 2A) and in some cases periclinal divi-sions (Fig. 2B). In general, 3–4 layers of the inner cortex were involved in primordium formation and the three outer cortical cell layers did not participate in this process (Fig. 2B).

SHORTROOT (SHR) and SCR are the key genes respon-sible for endodermal specification in the root meristem (Di Laurenzio et al., 1996; Helariutta et al., 2000). SCR expression is well known as an endodermal fate marker (Heidstra et al., 2004). Transgenic hairy roots harbouring the promoter fusion construct SCR::GUS were used to investigate the involve-ment of endodermal cells in primordium formation in squash (Supplementary Data Fig. S1). The GUS expression pattern indicated that during primordium formation, endodermal cells did not lose SCR expression, i.e. endodermal identity. SCR expression was maintained even after the completion of the first anticlinal and periclinal divisions of the endodermal cells involved in LRP formation (Supplementary Data Fig. S1C).

A B

Fig. 2. Visualization of auxin response maxima in Cucurbita pepo DR5::GUS roots during the development of lateral root primordia. (A) Cross- and (B) longitu-dinal root sections stained for GUS activity and embedded in plastic resin were counterstained with ruthenium red. Formation of lateral root primordia at a distance of (A) 700 µm and (B) 1400 µm from the initial cells. (A) A primordium is formed opposite a xylem pole. Periclinal divisions within pericycle and endodermal layers have been completed. The number of pericycle cell files in the outer pericycle layer (pII) has doubled (black outline). The protoxylem is outlined in green. (B) Note that while the auxin response maximum is shifting towards the apical part of the primordium, the inner cortical cell layers become involved in LRP for-

mation. en, endodermis; pI, pericycle I; pII, pericycle II. Scale bars = 20 µm.

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Cellular responses to auxin related to lateral root initiation in Cucurbita pepo

To analyse the localization of cellular auxin response maxima (ARMs), transgenic squash roots harbouring the

auxin-responsive promoter DR5 fused to different reporter genes were obtained. The following reporter fusions were used: a GUS–eGFP fusion with cytosolic localization (Figs 1 and 2) as well as a GUS–eGFP–NLS fusion (Figs 4–7 and 9;

A B C

Fig. 3. Distribution of proliferating cells in Cucurbita pepo root tips. Formative divisions initiating a lateral root primordium are surrounded by DNA replicat-ing cells of the adjacent tissues. Longitudinal sections: green channel, EdU-labelled nuclei (DNA-synthesizing cells); red channel, nuclei counterstained with propidium iodide. (A) Formative anticlinal divisions at a distance of 300 µm from the initial cells in the outer pericycle layer initiating a lateral root primordium (white arrowhead). Pericycle cells have undergone the first periclinal divisions (black arrow) in another lateral root primordium (asterisk). Wide field fluorescence microscopy. (B, C) Periclinal divisions during primordium formation at a distance of 500 µm proximal to the initial cells. (B) and (C) show the same root portion. (B) A single section captured with differential interference contrast microscopy. (C) A maximum intensity projection of a z-stack (5 μm) of 19 optical sections created by structured illumination. A white arrow points at a metaphase cell, black arrows point at cells in the outer pericycle layer that have undergone periclinal

divisions, and a developed primordium is marked by an asterisk. Scale bars = 50 µm (A), 20 µm (B, C).

A B C D

Fig. 4. Visualization of auxin response maxima (ARMs) in a Cucurbita pepo DR5::eGFP-NLS root tip. Confocal laser scanning microscopy images of longitu-dinal vibratome sections. Green channel, ARMs (fluorescence of eGFP–NLS); magenta channel, DNA stained with DAPI; and red channel,cell walls counter-stained with Pontamine Fast Scarlet 4B. (A, B) Overview of the parental root tip shows the acropetal sequence of developing lateral root primordia (white arrows). ARMs are visible within the quiescent centre and the columella cells (asterisk), as well as in xylem cell files (black arrow) and lateral root primordia (white arrows). (С, D) Young lateral root primordia (white arrows) within the parental root meristem at a distance of 500–1000 µm from the initial cells are shown. Scale

bars = 100 µm (A, B), 50 µm (C, D).

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Supplementary Data Video S1) and a tdTomato–H2B fusion (Fig. 8) with nuclear localization. The most precise data on the localization of the auxin response were obtained using reporter cassettes with nuclear localization of fluorescent proteins. In particular, analysis of roots containing the tdTomato–H2B fusion permitted the most precise detection of the cellular auxin response during mitosis. This made it possible for the first time to identify the cells of the parental root meristem involved in the process called ‘priming’ (De Smet et al., 2007).

Distribution of auxin response maxima in the parental root meristem. Analyses of longitudinal (Fig. 4) and cross- (Fig. 5) sections of DR5::eGFP-NLS roots showed that in the apical part of the parental root meristem, the ARM was maintained in the quiescent centre and the initial cells as well as in the columella and the periphery of the root cap, the stelar paren-chyma of the vascular cylinder and the xylem cell files (Figs 4 and 5A). In cortical cell files, the ARM disappeared at a short distance from the initials (Figs 1B and 4A, B). Activity of DR5 persisted in the pericycle up to a distance of no more than 100–150 µm proximal to the initials of a cell file (Figs 4B, D and 6A). Stelar parenchyma cells lost the ARMs at a distance of 300–400 µm from their initials (Fig. 5A, B). At a distance of 400–500 µm from the initials, the ARMs remained present in the peripheral metaxylem and protoxylem cell files (Fig. 5C); in the peripheral metaxylem, the ARM was still visible even at a distance of 1000–1400 µm from the initials (Figs 4A, B and 5D). Furthermore, ARMs were clearly visible in LRPs develop-ing acropetally (Figs 1A, B, 4 and 5D).

Auxin response maxima occur before the formative divisions in pericycle and endodermis. Cell files in the stelar paren-chyma, pericycle, endodermis and cortex were analysed using longitudinal sections of parental roots to identify the earliest stages of LR initiation and examine the formation of the cor-responding ARMs. Cell files were traced from the initial cells up to the point where the first completed periclinal divisions associated with primordium formation were visible in the two pericycle layers (stage II). Three-dimensional reconstructions of a z-series of optical sections permitted the observation of the almost complete disappearance of the parental root meristem

ARM in pericycle cells approx. 200 µm proximal to the initials (Fig. 6). ARMs in pericycle and endodermal founder cells were detected again at a distance of 250–300 µm proximal to the initials in the LR initiation zone (stage 0, Fig. 6; Supplementary Data Video S1). Notably, ARMs formed simultaneously in two adjacent cells within the same cell file (Fig. 6). These adjacent pericycle and endodermis founder cells represented sister cells that underwent one or two rounds of anticlinal divisions, while maintaining their ARMs (Fig. 7). During LR initiation, ARMs were formed in cell pairs of one or two (or rarely three) cell files of the inner pericycle, three to four cell files of the outer pericycle and one cell file of the endodermis (Figs 5C and 6–8). Thus, the minimal cell group displaying an ARM consisted of 10–14 cells (Fig. 6; Supplementary Data Video S1). These cells subsequently underwent one or two formative anticlinal divi-sions, thereby initiating an LRP (Figs 7 and 8). No single inter-phase pericycle and/or endodermal cells with an ARM were observed in this zone. The first LRP initiation stage and the maintenance of an ARM always involved an even number of sister cells.

During developmental stages 0 (Fig. 6) and I (Figs 1C, D, 5D and 7A, B), the primary root ARM included the protoxylem and peripheral metaxylem as well as some stelar parenchyma cells adjacent to a new LRP initiation site. Three-dimensional recon-structions did not show any local LRP initiation-related ARMs in protoxylem cell files adjacent to the incipient LRP (Figs 5C, D and 6B; Supplementary Data Video S1).

Involvement of auxin in further primordium development. After completion of one or two formative anticlinal divisions (Figs 7 and 8), pericycle cells with an ARM underwent periclinal divi-sions (Figs 1E, 2A and 9A, B). Endodermal cells situated prox-imal and distal to the first formative division within their cell file and adjacent files of the endodermal layer became part of the LR-related ARM and underwent anticlinal divisions (Figs 1E, 6A, B, 8 and 9A, B) and then periclinal divisions (Figs 2C and 9C). The ARM expanded successively to include pairs of cells from the cortical cell files adjacent to the LRP (Figs 8A, B and 9A–C). Later on, these cortical cells that were part of the LR-related ARMs underwent divisions to form the

A B C D

Fig. 5. Formation of auxin response maxima (ARMs) in the apical part of the parental meristem of an individual Cucurbita pepo DR5::eGFP-NLS root. Confocal laser scanning microscopy images of vibratome cross-sections. A single optical section of a differential interference contrast microscopy image is combined with a maximum intensity projection of optical sections in the green channel, showing ARMs (fluorescence of eGFP–NLS). (A) ARMs in developing xylem cell files and xylem parenchyma at a distance of 65 µm proximal to the initial cells. (B) ARMs in peripheral metaxylem and protoxylem cells at a distance of 390 µm from the initial cells. (C) ARMs at the stage of the first anticlinal division in the pericycle. The auxin response is concentrated in peripheral metaxylem and protoxylem cells at a distance of 455 µm from the initial cells. (D) ARMs at the stage of the first periclinal division in the pericycle. ARMs are concentrated only in develop-ing primordia and in peripheral metaxylem cells at a distance of 780 µm from the initial cells. Z-stack thickness: 45 µm (A), 48 µm (B), 43 µm (C) and 37 µm (D). White arrowheads, protophloem sieve elements; black arrows, protoxylem cell file; black arrowheads, developing lateral root primordia. Scale bars = 50 µm.

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outer layers of the LRP. During the subsequent LRP develop-ment, the LR-related ARM gradually decreased in size until it no longer encompassed the peripheral cells, and shifted to the apical part of the LRP (Figs 2C and 9C).

Influence of exogenous auxins and auxin transport inhibitors on lateral root initiation in Cucurbita pepo

To analyse the role of auxin transport during LR initiation in the root apical meristem, 3-day-old wild-type squash seedlings were incubated for 96 h in solutions of auxins (NAA or IBA) or auxin transport inhibitors (NPA, TIBA or 1-NOA) at different concentrations.

The growth rate of the parental root did not change at 10 nm NAA, but decreased by almost a factor of two at 25 nm NAA (Fig. 10A; Supplementary Data Fig. S2). Moderate

A B C

Fig. 6. The first cellular events during lateral root initiation in the meristem of a Cucurbita pepo DR5::eGFP-NLS parental root with auxin response maxima (ARMs) in the pericycle and endodermis. Confocal laser scanning microscopy images of longitudinal vibratome sections. A 3-D reconstruction of z-stacks: 45 µm thick, 100 optical sections (A); 40 µm thick, 75 optical sections (B, C). Green channel: ARMs (fluorescence of eGFP–NLS). (A–C) A cell group initiating a lateral root primordium at a distance of 280 µm from the initial cells is depicted. (A) Overview and (B, С) close-ups at different angles to the longitudinal axis of the par-ental root show the cell files of the inner (I) and outer (II) pericycle and of the endodermis. Three groups of adjacent cell pairs in three inner pericycle I cell files,

two pairs of adjacent cells of outer pericycle II cell files and a pair of adjacent cells in the endodermis display ARMs. Scale bars = 20 µm.

A B C

Fig. 7. Anticlinal divisions in pairs of sister pericycle cells with auxin response maxima (ARMs) during lateral root initiation in the parental meristem of a Cucurbita pepo DR5::eGFP-NLS root. Confocal laser scanning microscopy images of longitudinal vibratome sections. (A, B) Single optical sections, (C) 3-D reconstruction of a z-stack: 48 µm thick, 90 optical sections. Green channel, ARMs (fluorescence of eGFP–NLS); magenta channel, DNA stained with DAPI. (A–C) Two cell groups initiating lateral root primordia. (A, B) Overview and (C) close-up of the upper cell group. Two sister cells in the outer pericycle (II) layer in the lower half of the figure show an ARM (arrowheads). In this cell file, the cells distal and proximal to the sister pair show a reduced auxin response. In the upper half of the figure, i.e. a little further from the root tip, the first anticlinal division can be observed in a cell below the pair of sister cells with the ARM (metaphase, white arrow). (C) 3-D overview of the cell group shown in the upper part of (A) and (B). Pairs of adjacent cells showing that ARMs can be observed in pericycle

II and endodermis cell files. Scale bars = 20 µm (A, B) and 10 µm (C).

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concentrations of IBA (0.1–10 nm) slightly decreased root length increment, while 100 nm IBA drastically reduced the root growth rate (Fig. 10A; Supplementary Data Fig. S2). Counting the total number of LRs, including all visible LRPs, under a stereo microscope (Fig. 10B; Supplementary Data Fig. S2) revealed that 10 nm NAA or 0.1–10 nm IBA, respectively, did not affect the total number of LRs and LRPs in comparison with control plants. Only upon treatment with 25–50 nm NAA or 100 nm IBA, respectively, was a slight decrease found in the total number of LRs and LRPs.

A similar situation was observed for roots treated with exogenous auxin transport inhibitors. Low concentrations of NPA (0.1 µm) or 1-NOA (25 µm) did not cause changes in root growth increment (Fig. 10A; Supplementary Data Fig. S2). However, an increase of NPA or 1-NOA concentrations, or TIBA concentrations of ≥1 µm, significantly inhibited root growth. The number of LRs and LRPs was not affected at 0.1 µm NPA or 1–10 µm TIBA, respectively, but decreased con-siderably at higher concentrations (Fig. 10B; Supplementary Data Fig. S2). Regarding 1-NOA, even a treatment with only 25 µm led to a decrease in the number of LRPs. In summary,

A B C D

Fig. 8. Participation of different pericycle cell files and layers with ARMs in the lateral root initiation of a Cucurbita pepo DR5::tdTomato-H2B parental root meri-stem. Confocal laser scanning microscopy images of radial longitudinal (A, B) and tangential longitudinal (C, D) sections of a parental root in the zone after the first anticlinal formative divisions in the pericycle. Maximum intensity projection of z-series of 16 (8.9 µm in depth) (A, B) and 3-D reconstructions of 12 (4.6 µm depth) (C) and 24 (10 µm depth) (D) optical sections. Green channel, ARMs (fluorescence of tdTomato–H2B); magenta channel, cell nuclei in different cell cycle stages (fluorescence of Venus–H2B). (A, B) ARMs in pairs of sister cells in the pericycle, endodermis and cortex layers initiating lateral root primordium. Two telophase cells in the outer pericycle layer show ARMs. (C, D) Basal part of an incipient primordium in the inner (C) and outer (D) layers of pericycle. (C) Two cells of the inner pericycle (I) in one cell file before the first formative division show an ARM. (D) Three cell files in the outer pericycle (II) with cells that have

completed formative divisions. Only four cells in each file show ARMs. Scale bars = 20 µm (A, B), 10 µm (C, D).

A B C

Fig. 9. Formation of lateral root primordia in Cucurbita pepo DR5::eGFP-NLS roots. First periclinal divisions in pericycle cell files and the formation of the primordium axis with a shift in the ARMs to the apical part of the primordium. Confocal laser scanning microscopy images of longitudinal vibratome sections. (A) Single optical section overlaid with differential interference contrast. 3-D reconstructions of a z-series of (B) 56 optical sections (16 µm depth) and (C) 90 optical sections (23 µm depth). Green channel, ARMs (fluorescence of eGFP–NLS); magenta channel, DNA stained with DAPI; yellow channel, autofluorescence of lig-nin. (A) Lateral root primordium at a distance of approx. 600 µm from the pericycle initials. The first periclinal divisions of the pericycle cells have commenced. A black arrow points at a cell in the metaphase, while white arrows point at daughter cells resulting from completed divisions. (B) 3-D reconstruction of a lateral root primordium at a distance of approx. 600 µm from the pericycle initials. The first periclinal division in the outer pericycle (II) layer has been completed (white arrow), and four cortical cells, including two central cells showing a high auxin response, are involved in lateral root initiation. (C) At a distance of approx. 2 mm from the pericycle initials, the ARM is undetectable in the outer parts of the primordium. In the parental root, protoxylem has differentiated. Scale bars = 20 µm.

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treatment with NPA led to a reduction of both root length incre-ment and the number of LRs and LRPs. Treatment with TIBA strongly suppressed root growth, but the number of LRs and

LRPs was reduced only at 100 µm TIBA. In contrast, treatment with 25 µm 1-NOA did not influence root growth but led to a decrease in the number of LRPs.

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Fig. 10. Primary root growth and lateral root initiation in Cucurbita pepo seedlings under the influence of exogenous auxins and auxin transport inhibitors. Exposure of seedlings to different concentrations of exogenous auxins (NAA, 1-naphthylacetic acid; IBA, indole-3-butyric acid) and auxin transport inhibitors (NPA, 1-N-naphthyphthalamic acid; TIBA, 2,3,5-triiodobenzoic acid; 1-NOA, 1-naphthoxyacetic acid) over a period of 96 h. (A) Primary root growth increment (mm) after treatments. (B) The total number of lateral roots, including all visible primordia, in 7-day-old seedlings. (C) Lateral root initiation index in the outer cor-tical layer. Asterisks indicate the presence of statistically significant differences between control and treatment groups following the one-way ANOVA (P < 0.05).

Values are presented as the mean ± s.e. n/a, data not available.

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For a more precise comparison of LR initiation under the influence of exogenous auxins or auxin transport inhibitors, we calculated the ILRI proposed by Dubrovsky et al. (2009). Data shown in Fig. 10C demonstrate that the ILRI did not change during treatment with exogenous NAA or IBA; neither was it affected by treatment with the auxin transport inhibitor 1-NOA. These results indicate that NAA, IBA and 1-NOA mainly affected root growth, but not LR initiation. However, the auxin transport inhibitors NPA and TIBA significantly increased the ILRI at concentrations of 1 and 1–10 µm, respectively, indicating that NPA and TIBA may directly affect LR initiation.

The LRPs formed in the presence of exogenous auxins, either NAA or IBA, did not differ significantly from the con-trol LRPs regarding shape and structure. However, the LRPs formed in the presence of auxin transport inhibitors (NPA, TIBA or 1-NOA) were substantially different from the LRPs of control plants. Specifically, they had a flattened shape because the number of pericycle and endodermal cells along the lon-gitudinal axis of the root that participated in LRP formation was strongly increased. In several cases, merged LRPs were observed (Supplementary Data Fig. S3A, C, D), especially in response to 100 µm TIBA (Fig. S3B, C). The number of cor-tical cells involved in LRP formation was also affected by auxin transport inhibitors. For example, NPA affected the involve-ment of cortical cell layers in LR formation, with six layers of the outer cortex not participating (Fig. S3A), while in the control group, only two to three layers of the outer cortex were not involved in this process (Fig. 2). Treatments with TIBA and 1-NOA had a weaker effect on the involvement of cortical lay-ers, but there were still three or four layers of the outer cortex that did not contribute to LRP formation (Fig. S3A, C, D).

DISCUSSION

Our results highlight the fundamental similarities in the control of LR initiation by the key regulator auxin in flowering plants with different positions of LR initiation.

In most flowering plants, the initiation of LRs starts with anticlinal divisions in pericycle cell files (McCully, 1975; Lloret et al., 1989; Casero et al., 1993, 1995, 1996). These divi-sions can be either equal or unequal (Lloret et al., 1989; Casero et al., 1996; Demchenko and Demchenko, 1996; Dubrovsky et al., 2001). It has been suggested that asymmetric divisions in two pericycle cells of the same file, preceded by co-ordinated nuclear migration accompanied by auxin accumulation, play a key role in LR initiation (Casimiro et al., 2003; De Rybel et al., 2010).

We have reported previously that transgenic hairy roots of C. pepo represent an excellent system for studying root devel-opment (Ilina et al., 2012), where root anatomy and the pattern of lateral root development do not differ between wild-type and transgenic hairy root systems (Supplementary Data Fig. S4).

Two pericycle layers and the endodermal layer simultan-eously take part in primordium initiation in C. pepo roots. The visualization of local ARMs allowed the precise detection of cells involved in the determination of the site of LRP forma-tion in C. pepo. The first event in the initiation of lateral roots was the formation of a local ARM in the cells of these lay-ers (Figs 1C–E and 6; Supplementary Data Video S1). The

initiation of an LR then started with equal anticlinal divisions in two pericycle founder cells in the same cell file (Figs 1D and 3A). This event occurred within the meristem of the par-ental root at a distance of 250–350 µm proximal to the initial cells. At least 12 cells were involved in primordium initiation, with four to six cells from the inner pericycle, six cells from the outer pericycle and two to four cells from the endodermis (Figs 4 and 6–8; Supplementary Data Video S1). There were no significant differences in pericycle cell lengths, regardless of whether the cells were involved in LR initiation (Figs 1C–D and 3A). This observation suggests that the duration of the first mitotic cycle during C. pepo LR initiation is not significantly shortened as was proposed earlier for some species (MacLeod and Thompson, 1979; Laskowski et al., 1995), including Cucumis sativus (Dubrovsky, 1986a). The number of cell files of the inner pericycle that participate in LR initiation seems to be three in C. pepo as in arabidopsis (Casimiro et al., 2003). In contrast to arabidopsis, not only the pericycle but also the endo-dermis and the cortex are involved in LRP formation in C. pepo (Fig. 9), leading to a more complex LRP structure.

Numerous studies have shown that in arabidopsis and other plants, auxin plays a crucial role in LRP initiation (De Smet et al., 2006; Overvoorde et al., 2010; Möller et al., 2017). In some species, treatment of root systems with exogenous aux-ins led to a dramatic increase in the number of LRs (Blakely et al., 1988; Laskowski et al., 1995). In plants with LR ini-tiation proximal to the elongation zone, application of inhibi-tors of auxin transport always led to decreases in LR number or even to the suppression of LR development (Lloret and Casero, 2002). However, treatment of C. pepo roots with exogenous auxins [the synthetic auxin NAA or the naturally occurring auxin IBA, a precursor of indole-3-acetic acid (IAA) synthe-sis related to LR initiation] did not stimulate LR initiation (Fig. 10). With increasing exogenous auxin concentrations, the total number of LRPs and LRs decreased along with the gen-eral suppression of primary root growth. Meanwhile, the ILRI proposed by Dubrovsky et al. (2009) remained unaltered. This can be explained by the high auxin content already present in the C. pepo LRP initiation zone. Consequently, the increase of auxin levels in the root meristem cells caused by treatment with exogenous auxin did not lead to the formation of new local ARMs required for the initiation of more LRPs. The observed decrease in the total number of LRPs in response to 25–50 nm NAA or 100 nm IBA was related to the general inhibition of cell proliferation in the parental root.

Therefore, in C. pepo, LRP initiation seems to be insensitive to exogenous auxins. The only comparable data on the insensi-tivity of LR initiation to exogenous auxins were obtained using the monocotyledonous aquatic plant Pontederia cordata (Charlton, 1983a) and the fern Ceratopteris richardii (Hou et al., 2004; Hou and Blancaflor, 2009). In both species, LR initiation also occurs in close proximity to the apical meristem of the parental root (Table 1). In C. richardii, exogenous appli-cation of either IAA or IBA did not affect the LR initiation pattern or development, although auxins had a strong inhibitory effect on the growth of the parental root (Hou et al., 2004). The authors proposed that different mechanisms control LR development in non-seed vascular plants compared with angio-sperms, considering the long independent evolutionary history of the two groups (Hou et al., 2004).

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The influence of three auxin transport inhibitors (NPA, TIBA and 1-NOA) on LR initiation in C. pepo roots was also tested. 1-NOA inhibits AUX1-mediated auxin influx (Laňková et al., 2010), while NPA and TIBA inhibit ABCB/PGP proteins and PIN-mediated auxin efflux (Geldner et al., 2001; Bailly et al., 2008). All three compounds were able to inhibit the growth of the parental root of C. pepo and caused a decrease of the total number of LRPs (Fig. 10). However, the ILRI remained unchanged after 1-NOA treatment, indicating that the decrease in the total number of LRPs was due to the general inhibition of proliferation in the parental root. In contrast, the application of NPA or TIBA increased the ILRI. Thus, the PIN-mediated auxin efflux plays a key role in the formation of a local ARM and the further development of LRPs in C. pepo. Treatments with auxin transport inhibitors also resulted in abnormal LRP development (Supplementary Data Fig. S3), with significant increases in the number of cells in a cell file involved in primordium forma-tion. This might imply that auxin transport helps mediate the determination of LRP borders. In a similar vein, NPA reduced the involvement of inner cortical cell layers in LRP formation (Supplementary Data Fig. S3).

Our results show that the resumption of the mitotic cycle from the G1 phase as observed consistently for LR forma-tion above the elongation zone (Demchenko and Demchenko, 2001b; Alarcón et al., 2016) did not occur during the initiation of LRPs in the meristem of the parental root of C. pepo. A pair of sister cells (designated as founder cells) were observed

following an anticlinal division in the pericycle cell file; a simi-lar observation was made for the endodermis. The progression of these cells through the mitotic cycle (G1–S–G2) would be accompanied by the formation of a local ARM, followed by the first anticlinal division, which represents the first division in the initiation of an LRP. The effects of NPA and TIBA vs. 1-NOA show that the local ARM is formed due to PIN-mediated auxin efflux from the central cylinder and thus does not seem to be cor-related with auxin flow from dying root cap cells, as suggested for arabidopsis (Möller et al., 2017). We propose that the local ARM in the parental root meristem of C. pepo is unaffected by exogenous application of auxin because the endogenous auxin concentrations are too high.

Recent studies have shown that the endodermis plays an essential role in the progression of pericycle cells to LRP ini-tiation (Marhavý et al., 2013;Vermeer et al., 2014). One of the most important unresolved questions regarding LR formation involves whether endodermal cells maintain their identity when involved in LRP formation or whether they become undiffer-entiated LRP cells. The SCR gene is a reliable endodermal marker (Di Laurenzio et al., 1996; Xiao et al., 2014). The SCR expression pattern in C. pepo roots shows that endodermal cells maintain their identity when involved in LRP formation (Supplementary Data Fig. S1). After emergence of the LRP from the parental root of C. pepo, derivatives of cortical cells form the temporary root cap, while all initial cells and, accord-ingly, all tissues of the LR originate from the pericycle of the parental root (Demchenko and Demchenko, 2001a).

In arabidopsis, PIN3-mediated reflux of pericycle-derived auxin between endodermis and pericycle and SHY2-mediated auxin signalling assist the progression of founder cells towards the next phase (Goh et al., 2012; Marhavý et al., 2013). Furthermore, the loss of volume of cortical cells is required for LR emergence (Vermeer et al., 2014). In C. pepo, an ARM is formed in the cortex adjacent to the developing LRP (Fig. 2B). This might imply a common mechanism of LR movement through the cortex of the parental root based on LAX3-dependent auxin influx into cortical and epidermal cells periph-eral to the LRP (Swarup et al., 2008).

All of the main functions of the root depend on the branching of the root axis. In the course of root system evolution in land plants, a transition from dichotomous branching to monopodial branching (the so-called herring-bone pattern) can be observed. Monopodial branching offers advantages in a dense medium. Goebel (1928) already proposed that in the transition from dichotomous to monopodial branching, the stimulus for the formation of LRPs was derived from the tissues of the central cylinder. Transition stages have been reported for lycophytes (Lycopodium) and ferns (Marsilea and Ceratopteris) (Voronin, 1964; Charlton, 1983a). In those plants, LRP initiation occurs in the pericycle just a few cells proximal to the initials, close to the tracheal elements (Voronin, 1964). This situation might represent the ancestral form of monopodial branching. Thus, in the course of evolution, the LRP initiation site shifted from the root apex to the root base, i.e. at least to the upper part of the meristem and in most flowering plants to an area above the elongation zone (Fig. 11B, C). This trend, combined with the endogenous initiation of LRPs in pericycle or endodermis, simplifies the movement of the root tip in dense soil during branching.

Common ancestor:monopodial branching

A B C

Elo

ngat

ion

zone

Mer

iste

m

Elo

ngat

ion

zone

Mer

iste

m

Elo

ngat

ion

zone

Mer

iste

m

In parentalroot meristem

Above theelongation zone

Fig. 11. Hypothetical scheme for the evolution of root branching in vascular plants (Tracheophyta). (A) Lateral root initiation and primordium formation in the apical part of the parental root meristem represent the evolutionary precur-sor of monopodial branching. (B, C) During the evolution of Tracheophyta, the lateral root initiation zone shifted upward, to (B) 250–350 µm proximal to ini-tials in Cucurbita pepo and (C) above the elongation zone of the parental root in most plant species. Zone of lateral root initiation, red; root cap, yellow; central

cylinder, blue. Modified after Goebel (1928) and Voronin (1964).

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In summary, there was one general tendency during the evo-lution of root systems, but in each angiosperm clade, different morphogenetic solutions were realized. Cucurbita pepo takes an intermediate position regarding the site of the determination of competence for LR initiation, which takes place in the apical part of the parental root meristem (Fig. 11B). LRs in Cucurbitaceae can develop rapidly, leading to the emergence of LRs directly above the zone of cell elongation in the parental root. This allows the development of a vast root system within a short time. In some hydrophytes, such as Pistia stratiotes (Araceae) or Eichhornia crassipes and Pontederia cordata (Pontederiaceae), LRP initiation occurs in the root apex which is suitable for an aquatic environ-ment (see Charlton, 1983b). In Cucurbitaceae and Polygonaceae, the proximal shift of the LRP initiation site from the initial cells of the parental root meristem was small, so in these families LRPs are still surrounded by proliferating cells in the meristem. However, in Poaceae, LRP initiation takes place above the elongation zone. Thus, cells in the basal part of the meristem must exit the mitotic cycle in the G1 phase and subsequently resume proliferation later to initiate an LRP (Demchenko and Demchenko, 2001b; Alarcón et al., 2016). In arabidopsis, LR initiation sites occupy an inter-mediate position. Analyses of arabidopsis and cereals have led to the assumption that some pericycle cells remain in the G2 phase in the basal part of the meristem and are more susceptible to LR ini-tiation than others, described as the concept of an ‘extended meri-stem’ (Beeckman et al., 2001; Beeckman and De Smet, 2014; Du and Scheres, 2018).

Auxin plays a critical role in plant development, being involved in the regulation of pattern formation, cell prolifer-ation and differentiation. The pathways of regulation of auxin signalling are common in all multicellular plants (Kato et al., 2018). Literature data as well as results obtained in this study suggest that the mechanism for the formation of a local max-imum of auxin for launching the programme of LRP formation is common to all Tracheophyta.

In summary, we propose that the different mechanisms underlying LRP initiation in various groups of land plants are due to a set of common principles enabling plants to adapt to diverse environmental conditions. The differences in these mechanisms influence the distance between the initial cells in the root apex and the LRP initiation site, and the involvement of the tissues supporting the pericycle (or in ferns, the endo-dermis) during LRP formation. What is still unclear is how the competence for LRP formation is established in individual cells. This question remains the most intriguing in the biology of root development. Future investigations aimed at clarifying this issue should involve model plants as well as diverse evolu-tionarily transitional species.

SUPPLEMENTARY DATA

Supplementary data are available online at htpps://academic.oup.com/aob and consist of the following. Figure S1: expres-sion pattern of pCsSCR::GUS in the root tip of Cucurbita pepo. Figure S2: 7-day-old seedlings of Cucurbita pepo exposed to exogenous auxins and auxin transport inhibitors. Figure S3: lateral root primordia in parental roots of 7-day-old Cucurbita pepo seedlings exposed to auxin transport inhibitors. Figure S4: structure of root tips from the wild type and two types of trans-genic hairy roots of Cucurbita pepo. Table S1: list of primers

used in this study. Video S1: animated 3-D model of an early stage of lateral root initiation in Cucurbita pepo DR5::eGFP-NLS that is presented in Fig. 6. The model shows the auxin response maximum in pairs of sister cells in two different layers of the pericycle and endodermis, respectively.

ACKNOWLEDGEMENTS

The authors gratefully acknowledge the handling editor Dr Nigel Chaffey and two anonymous reviewers for insightful comments on the manuscript. We thank Dr Nambirajan Govindarajan and the Edanz Group (www.edanzediting.com/ac) for editing drafts of this manuscript. We are grateful to Erik Limpens (Wageningen University, Wageningen, The Netherlands), Maria Harrison (Boyce Thompson Institute for Plant Research, New York, USA), Robert Benezra (Sloan-Kettering Memorial Cancer Center, Sloan-Kettering Institute, New York, USA) and Michael Davidson (Florida State University, USA) for providing plasmids. This study was performed using equipment of the Core Facility of Cell and Molecular Technologies in Plant Science at the Komarov Botanical Institute (Saint-Petersburg, Russia) and the Core Facility of Genomic Technologies, Proteomics and Cell Biology at the Research Institute for Agricultural Microbiology (Saint-Petersburg, Pushkin, Russia). This re-search was financially supported by the Russian Science Foundation (grant no. 16-16-00089). N.P.D., K.P. and K.N.D planned and designed the research. E.L.I., K.N.D., V.A.S. and A.S.K. performed experiments and analysed data. K.N.D. and E.L.I. carried out the design of all vectors and microscopy. K.N.D., E.L.I. and K.P. wrote the manuscript.

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