Journal of Experimental Botany Advance Access published April 23, 2015 Journal of Experimental Botany doi:10.1093/jxb/erv174

Review Paper

Interplay of HD-Zip II and III transcription factors in auxinregulated plant development L. Turchi1, S. Baima2, G. Morelli2,* and I. Ruberti1,* 1  2 

Institute of Molecular Biology and Pathology, National Research Council, Piazzale Aldo Moro 5, 00185 Rome, Italy Food and Nutrition Research Centre, Agricultural Research Council, Via Ardeatina 546, 00178 Rome, Italy

*  To whom correspondence should be addressed. E-mail: [email protected] or [email protected]

Abstract The homeodomain-leucine zipper (HD-Zip) class of transcription factors is unique to plants. HD-Zip proteins bind to DNA exclusively as dimers recognizing dyad symmetric sequences and act as positive or negative regulators of gene expression. On the basis of sequence homology in the HD-Zip DNA-binding domain, HD-Zip proteins have been grouped into four families (HD-Zip I–IV). Each HD-Zip family can be further divided into subfamilies containing paralogous genes that have arisen through genome duplication. Remarkably, all the members of the HD-Zip IIγ and -δ clades are regulated by light quality changes that induce in the majority of the angiosperms the shade-avoidance response, a process regulated at multiple levels by auxin. Intriguingly, it has recently emerged that, apart from their function in shade avoidance, the HD-Zip IIγ and -δ transcription factors control several auxin-regulated developmental processes, including apical embryo patterning, lateral organ polarity, and gynoecium development, in a white-light environment. This review presents recent advances in our understanding of HD-Zip II protein function in plant development, with particular emphasis on the impact of loss-of-function HD-Zip II mutations on auxin distribution and response. The review also describes evidence demonstrating that HD-Zip IIγ and -δ genes are directly and positively regulated by HD-Zip III transcription factors, primary determinants of apical shoot development, known to control the expression of several auxin biosynthesis, transport, and response genes. Finally, the interplay between HD-Zip II and III transcription factors in embryo apical patterning and organ polarity is discussed. Key words:   Auxin, embryo apical patterning, gynoecium development, HD-Zip II proteins, HD-Zip III proteins, lateral organ polarity, shade-avoidance response.

Introduction The basic body plan is laid down during embryogenesis in both animals and plants. In Arabidopsis, formation of the apical–basal axis is followed by establishment of the radial axis and subsequently of bilateral symmetry; during the transition from the globular to the heart stage, the shoot apical meristem (SAM) and the root apical meristem are specified in the apical and basal domain of the embryo, respectively. However, in contrast to animal development, plant development is continuous, and new organs are formed throughout the life cycle through the activity of the stem-cell niches located in the

apical meristems. Furthermore, as sessile organisms, plants have evolved the capacity to accurately perceive and respond to external cues, adapting their growth to the environment. Auxin plays a prominent role in plant development, and there is plenty of evidence that auxin biosynthesis, transport, and response are all critical for pattern formation (Möller and Weijers, 2009; De Smet et al., 2010; Zhao et al., 2011). Interestingly, experimental and modelling work has recently lead to the proposal that local auxin sources (Ljung, 2013) in conjunction with polar auxin transport (Zažímalová et al.,

Abbreviations: bHLH, basic helix–loop–helix; EAR, ERF-associated amphiphilic repression; FR, far red; HD-Zip, homeodomain-leucine zipper; PAS, Per-Arnt-Sim; R, red; SAM, shoot apical meristem; START, steroidogenic acute regulatory protein-related lipid transfer domain; ZPR, LITTLE ZIPPER. © The Author 2015. Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved. For permissions, please email: [email protected]

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Received 26 January 2015; Revised 9 March 2015; Accepted 17 March 2015

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Structural characteristics and target sequences of members of the HD-Zip II and III families The HD-Zip class of proteins is unique to plants, and is characterized by the presence of a homeodomain closely linked to a leucine zipper motif (Ruberti et al., 1991). Several HD-Zip genes were identified in Arabidopsis at the beginning of the 1990s and, on the basis of homology in the HD-Zip domain, grouped into four families, HD-Zip I–IV (Sessa et al., 1994). Completion of the Arabidopsis genome sequence revealed that it codes for 48 HD-Zip proteins all belonging to the four families recognized previously (Sessa et al., 1994; Morelli et al., 1998; Baima et al., 2001; Henriksson et al., 2005; Nakamura et  al., 2006; Ciarbelli et  al., 2008; Brandt et  al., 2014). The HD-Zip genes have subsequently also been identified in charophyte algae, but seem to be absent from the genomes of chlorophyte algae (Floyd et al., 2006; Prigge and Clark, 2006; Mukherjee et  al., 2009). Each of the four protein families can be distinguished by the elevated conservation within the HD-Zip domain, the presence of additional conserved motifs, and specific intron and exon positions (Henriksson et al., 2005; Ariel et  al., 2007; Agalou et  al., 2008; Ciarbelli et  al., 2008; Harris et al., 2011). Phylogeny reconstruction has revealed that each family can be divided into subfamilies containing paralogous genes that have arisen through genome duplication, as they are associated with duplicated regions of chromosomes in the Arabidopsis and rice genomes (Henriksson et al., 2005; Agalou et al., 2008; Ciarbelli et al., 2008; Harris et al., 2011). The homeodomain, a 60 aa region, consists of three α-helices and a flexible N-terminal arm. The HD makes base contacts in both grooves, with α-helix-3 in the major groove of the DNA and the N-terminal arm in the adjacent minor groove. Most of the HD proteins recognize the core sequence TAAT (Gehring et al., 1994). The leucine zipper is a special case of a coiled-coil motif with a series of leucine residues spaced by exactly 7 aa. The start of each heptad repeat is designated by the letter a, such that the hydrophobic residues occur in positions a and d, with leucine residues usually at position d. This distribution of residues permits the formation of an amphipathic α-helix, which can pair by interaction of the aligned hydrophobic residues (Landschulz et al., 1988). The exact spatial register between the HD and the leucine zipper motif in the HD-Zip proteins is analogous to that observed between the DNA binding and the dimerization domains in the basic leucine zipper proteins, and at the time of their identification it was suggested that the HD-Zip proteins might use the dimerization domain to juxtapose a pair of DNA contacting surfaces, each of which fits into half of a dyad symmetric recognition sequence (Ruberti et al., 1991). Experimental work then demonstrated that the HD-Zip domain, but not HD alone, binds to DNA (Sessa et al., 1993). Furthermore, it has also been shown that a correct spatial relationship between HD and the leucine zipper motif is crucial for DNA binding (Sessa et al., 1993). Finally, binding-site selection experiments and subsequent chromatin immunoprecipitation sequencing experiments have determined that HD-Zip proteins recognize dyad symmetric sequences (Sessa

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2010; Grones and Friml, 2015) constitute the mechanism to initiate and orient the growth axes during embryonic and probably post-embryonic plant development (Robert et  al., 2013; Wabnik et al., 2013). Changes in auxin concentration are transduced by auxin signalling systems to mediate cellular responses such as division, elongation, and differentiation (Ljung, 2013; Grones and Friml, 2015). The auxin response is very complex; the developmental stage, the cell type, and the environmental context all contribute to determine a specific output. Auxin responsiveness can be specified at multiple levels including specificity of interactions among the auxin response components and specificity in interaction with regulatory factors controlling transcriptional repression (Lokerse and Weijers, 2009; Kieffer et  al., 2010; Pierre-Jerome et  al., 2013; Boer et al., 2014). In the past decade, molecular genetic studies have uncovered master regulatory genes involved in plant development, such, for example, as homeodomain-leucine zipper (HD-Zip) III and PLETHORA (PLT) transcription factors, which act as determinants of shoot and root development, respectively (Emery et  al., 2003; Aida et  al., 2004; Blilou et  al., 2005; Prigge et  al., 2005; Long et  al., 2006; Galinha et  al., 2007; Grigg et  al., 2009; Smith and Long, 2010). Consistent with the crucial role of auxin in organ patterning, it has been shown that PLT1 and PLT2 gene expression is stimulated by auxin and is dependent on auxin response transcription factors (ARFs) (Aida et  al., 2004). However, PLT proteins are required for the expression of multiple PIN-FORMED (PIN) proteins generating a positive-feedback loop, which reinforces the root auxin gradient and thus their own expression (Galinha et al., 2007). More recently, evidence has been provided that REVOLUTA (REV), a member of the HD-Zip III protein family, directly and positively regulates the expression of several auxin biosynthesis, transport, and response genes (Brandt et al., 2012; Huang et al., 2014). Interestingly, in the last few years, new patterning roles for PLT genes have emerged, indicating that PLT3, PLT5, and PLT7 act as regulators of phyllotaxis via control of local auxin biosynthesis in the central region of the SAM (Prasad et al., 2011; Pinon et  al., 2013). Together, these data indicate complex relationships between auxin and master regulators of plant development. This review describes recent advances that highlight the crucial role of HD-Zip IIγ and -δ transcription factors, mostly known for their function in the shade-avoidance response, in several developmental processes in a white-light environment (Bou-Torrent et  al., 2012; Reymond et  al., 2012; Ruberti et al., 2012; Zúñiga-Mayo et al., 2012; Turchi et al., 2013; Zhang et al., 2014). The review also reports on the finding that HD-Zip IIγ and -δ genes are directly and positively regulated by REV (Brandt et  al., 2012; Turchi et  al., 2013), which together with PHABULOSA (PHB) and PHAVOLUTA (PHV) control embryo apical patterning and specify adaxial identity of lateral organs (Emery et  al., 2003; Prigge et  al., 2005; Smith and Long, 2010). Finally, the review discusses the interplay between HD-Zip II and III transcription factors in auxin-regulated plant development.

HD-Zip II and HD-Zip III proteins in plant development  |  Page 3 of 11

Function of HD-Zip II transcription factors in auxin-regulated developmental processes HD-Zip IIγ and -δ proteins act as positive regulators of the shade-avoidance response The HD-Zip II protein family includes ARABIDOPSIS THALIANA HOMEOBOX 2 [ATHB2 (Ruberti et al., 1991); also known as HOMEOBOX ARABIDOPSIS THALIANA 4 (HAT4) (Schena and Davis, 1992)], the first gene shown to be rapidly induced by changes in the red (R) to far-red

(FR) light ratio that promotes the shade-avoidance response in the majority of the angiosperms (Carabelli et  al., 1996). Multiple phytochromes are involved in the regulation of ATHB2 by low R:FR (Carabelli et al., 1996; Franklin et al., 2003), and there is evidence that ATHB2 is a direct target of the basic helix–loop–helix (bHLH) transcription factor PHYTOCHROME INTERACTING FACTOR 5 (PIF5) (Hornitschek et  al., 2012). Loss-of-function athb2 mutants display reduced hypocotyl elongation in low R:FR compared with controls, whereas the phenotypes of plants overexpressing ATHB2 in high R:FR are reminiscent of those displayed by wild type grown in low R:FR, thus indicating that ATHB2 functions as a positive regulator of the shade-avoidance response (Steindler et  al., 1999; Carabelli et  al., 2013). The expression of the β-glucuronidase (GUS) fused to ATHB2 (ATHB2::ATHB2:GUS) is rapidly and strongly induced by low R:FR light in all the cell layers of the elongating region of the hypocotyl, suggesting that ATHB2 acts in this organ, at least in part, to regulate the shade-avoidance response (Carabelli et  al., 2013). Links between ATHB2 and auxin, which has a crucial role in the shade-avoidance response, have been established (Ruzza et al., 2014). Both of the other two HD-Zip II genes belonging together with ATHB2 to clade γ [HAT1, also known as JAIBA (JAB) (Zúñiga-Mayo et  al., 2012) and HAT2] and the HD-Zip IIδ genes (HAT3 and ATHB4) are induced by light quality changes that promote the shade-avoidance response. The kinetics of low R:FR induction and reversibility by high R:FR have suggested that HAT1, HAT3, and ATHB4, as for ATHB2, are regulated by the phytochrome(s), whereas HAT2, originally identified as an auxin-inducible gene (Sawa et al., 2002), is likely to be induced by light quality as a consequence of the changes in the auxin signalling pathway provoked by a shaded environment (Ciarbelli et  al., 2008). Furthermore, overexpression of HAT1, HAT2, HAT3, and ATHB4, as for ATHB2, results in phenotypes resembling those of wild type in low R:FR, further underlining the redundancy of the HD-Zip IIγ and -δ proteins in the regulation of shade avoidance (Ruberti et al., 2012). Interestingly, it has recently become apparent that, apart from their function in the shade-avoidance response, the HD-Zip IIγ and -δ transcription factors control several developmental processes in a white-light environment (BouTorrent et  al., 2012; Reymond et  al., 2012; Zúñiga-Mayo et al., 2012; Turchi et al., 2013; Zhang et al., 2014).

HD-Zip IIγ and -δ proteins act downstream of the bHLH transcription factor SPATULA in gynoecium development Fig. 1.  HD-Zip IIγ and -δ transcription factors contain an EAR repression domain. (A) The LxLxL type of EAR motif. The amino acids most frequently occurring in positions 2 and 4 of the LxLxL motif are shown. The height of the letter representing an amino acid in positions 2 and 4 reflects the difference in the frequency of its occurrence in the protein set analysed (Kagale et al., 2010). The amino acids are coloured according to their chemical properties: red (hydrophobic residues), green (polar residues), blue (residues with positive charge), violet (residues with negative charge). (B) Aa 1–15 of the ATHB2, HAT1, HAT2, HAT3, and ATHB4 proteins. The amino acids of the EAR motif are highlighted with the colour code used in (A).

HAT1, ATHB2, HAT3, and ATHB4 have recently been identified as genes positively regulated by SPATULA (SPT), a bHLH protein related to PIF transcription factors but lacking an active phytochrome-binding domain required for PIF negative regulation by the phytochrome in high R:FR (Reymond et al., 2012). SPT plays a crucial role in promoting growth of tissues derived from the carpel margins (Alvarez and Smyth, 1999). Loss of SPT function in fact results in alterations in the

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et  al., 1993, 1998; Tron et  al., 2001; Brandt et  al., 2012). Interestingly, HD-Zip II and III binding sites share the same core sequence [AAT(G/C)ATT] (Sessa et  al., 1993, 1998), thus suggesting that members of the two protein families may regulate common target genes (Turchi et al., 2013). The HD-Zip II protein family consists of 10 members and can be subgrouped into four clades (α–δ; Ciarbelli et  al., 2008). Interestingly, all the HD-Zip IIγ and -δ proteins contain an LxLxL type of ERF-associated amphiphilic repression (EAR) motif (Ciarbelli et al., 2008; Kagale et al., 2010) (Fig. 1), and evidence exists that at least some of them function as negative regulators (Steindler et  al., 1999; Ohgishi et  al., 2001; Sawa et  al., 2002; Turchi et  al., 2013). The promoter regions of the HD-Zip II genes are significantly enriched for HD-Zip cis elements (Ciarbelli et al., 2008), and it has been observed that inducible proteins containing the DNA-binding domain of HD-Zip II proteins and the transactivation domain of VP16 directly induce all the HD-Zip IIγ and -δ genes in vivo, thus suggesting the existence of an intricate negative-feedback network within the HD-Zip II protein family (Ohgishi et al., 2001; Sawa et al., 2002; Ciarbelli et al., 2008). By contrast, there is evidence that the HD-Zip III proteins act as positive regulators of gene expression (Wenkel et al., 2007; Kim et al., 2008; Brandt et al., 2012; Turchi et al., 2013; Baima et al., 2014).

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HD-Zip IIγ and -δ proteins control apical embryo development and organ polarity HD-Zip IIγ and -δ genes have also recently been identified as targets of REV, a member of the HD-Zip III protein family with a crucial role in several developmental processes (see below) (Brandt et al., 2012; Turchi et al., 2013). Consistently, it has been shown that progressive loss of HAT3, ATHB4, and ATHB2 activity causes developmental defects from embryogenesis onwards in a white-light environment (Brandt et al., 2012; Turchi et al., 2013). Cotyledon development and number are altered in hat3 athb4 and, to an even greater extent, in hat3 athb4 athb2 mutant embryos (Turchi et  al., 2013) (Fig.  2), and these defects correlate with changes in auxin distribution and response (Turchi et al., 2013). The formation of auxin maxima at the sites of cotyledon initiation depends mainly on changes in auxin flow, largely driven by PIN1, occurring at the transition stage of embryo development. PIN1 is expressed throughout the apical embryo during the globular stage, but by the transition and heart stages, PIN1 expression is higher at the sites of cotyledon initiation (Benková et  al., 2003). Mutations that alter PIN1 expression or polarity during the transition stage of embryo development lead to defects in bilateral symmetry (Izhaki and Bowman, 2007; Michniewicz et  al., 2007; Ploense et  al., 2009). It has been shown that hat3 athb4 mutant embryos with fused/single expanded cotyledon(s) display either two very close auxin maxima or

Fig. 2.  The mutants hat3 athb4 and hat3 athb4 athb2 display defects in embryo development. DIC images of mature embryos derived from hat3-3 athb4-1/+ (A, B) and hat3-3/+ athb4-1 athb2-3 (C, D) plants. (A, C) Wild-type-like embryo from hat3-3 athb4-1/+ (A) and hat3-3/+ athb4-1 athb2-3 (C) plants. (B, D) Altered embryos with two cotyledons displaying different degrees of expansion from hat3-3 athb4-1/+ plants (B) and altered embryos with one or two radialized cotyledons from hat33/+ athb4-1 athb2-3 plants (D). Embryo phenotypes emerged with a 1:3 ratio, as expected for a single segregating recessive mutation. Altered embryos from hat3-3 athb4-1/+ plants display varying degrees of defects in cotyledon development. Representative embryos of the phenotypic classes observed most frequently are shown in (B). Embryos with fused/ single cotyledon(s) were also detected at a low frequency (Turchi et al., 2013). Bars, 50 μm.

a single auxin maximum, suggesting that loss of HAT3 and ATHB4 activity may cause defects in PIN1 expression pattern at the transition stage (Turchi et al., 2013). In agreement, it was observed that a significant number of hat3 athb4 athb2 embryos failed to establish proper PIN1 polarity at the transition stage to direct auxin flow outward from the central apical region to the incipient cotyledons (Fig. 3). A connection between HD-Zip II transcription factors and auxin is further indicated by the finding that loss-of-function mutations in HAT3 and ATHB4 strongly enhance leaf organ fusion in the presence of 1-naphthylphthalamic acid, an inhibitor of polar auxin transport (Turchi et  al., 2013). PIN1 has a key role in organ separation at the SAM, and pin1 mutants display leaf fusion defects (Reinhardt et  al., 2003). Failure in leaf separation is also evident in plants lacking MONOPTEROS (MP)/ARF5 in the presence of 1-naphthylphthalamic acid (Donner et al., 2009). Evidence for a role of the HAT3, ATHB4, and ATHB2 proteins in adaxial identity specification of lateral organs has also been provided (Bou-Torrent et  al., 2012; Turchi et  al., 2013). Extensive work in recent years has identified a transcriptional regulatory network containing several leaf abaxial- and adaxial-promoting genes (Bowman and Floyd, 2008; Husbands et  al., 2009; Barton, 2010; Braybrook and Kuhlemeier, 2010; Efroni et  al., 2010). However, recently the existence of a transient low-auxin

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septum and transmitting tract, a decrease in stigmatic tissue development, and defects in carpel fusion in the upper part of the organ (Alvarez and Smyth, 2002). In agreement with a function of HD-Zip IIγ and -δ genes as SPT targets in the development of carpel margin tissues, it has been shown that simultaneous loss of HAT3 and ATHB4 results in incomplete apical fusion in the gynoecium and reduced development of the septum (Reymond et  al., 2012; Carabelli et  al., 2013). Defects in gynoecium development have also been observed in jab mutants (Zúñiga-Mayo et al., 2012). Interestingly, a recent report determined that SPT, together with INDEHISCENT (IND), regulates the transition of the apical style region from a bilaterally symmetric stage to a radially symmetric structure during gynoecium development by modifying the direction of auxin transport (Moubayidin and Østergaard, 2014). It was shown previously that the SPT and IND transcription factors directly repress the expression of PINOID (PID), which encodes a protein kinase A/protein kinase G/protein kinase C 3 (AGC3)-type protein kinase that regulates PIN protein localization (Sorefan et al., 2009; Girin et  al., 2011). Moubayidin and Østergaard (2014) have now provided evidence that repression of PID-mediated polar localization of PIN1 by SPT and IND proteins is required for the symmetry transition of the apical style region during gynoecium development. In this context, it will be of interest to determine how the HAT3 and ATHB4 HD-Zip II transcription factors act as targets of SPT in promoting the bilateral-to-radial symmetry switch during gynoecium development.

HD-Zip II and HD-Zip III proteins in plant development  |  Page 5 of 11 mutations strongly enhanced the hat3 athb4 phenotype. Almost all hat3 athb4 athb2 mutants display one or two radialized cotyledons and lack an active SAM, resembling phenotypes associated with loss-of-function mutations in multiple HD-Zip III genes. In agreement, bilateral symmetry and SAM defects are enhanced when hat3 athb4 is combined with mutations in PHB, PHV, or REV (Turchi et al., 2013).

Function of HD-Zip III transcription factors in auxin-regulated developmental processes HD-Zip III proteins act as positive regulators of apical development

zone in the adaxial domain of young leaf primordia has been shown to contribute to adaxial specification (Qi et al., 2014). It is well known that the distribution of auxin in the SAM depends largely on PIN1, and PIN1-mediated auxin transport in the SAM towards an incipient primordium triggers leaf organogenesis (Reinhardt et al., 2003; Heisler et  al., 2005). However, PIN1 polarity then reverses and directs back towards the meristem centre from young leaf primordia (Heisler et  al., 2005; Bayer et  al., 2009; Wang Q et al., 2014; Wang Y et al., 2014). Qi et al. (2014) found that the auxin flux reversal is tightly associated with leaf initiation and is required for the formation of the transient adaxial low-auxin domain, which may confer robustness to leaf polarity specification. HAT3 and ATHB4 expression is restricted to the adaxial domain of young leaf primordia (Turchi et al., 2013), and it will be intriguing to see if these HD-Zip II transcription factors contribute to the dynamic changes in PIN1 polarity occurring at the early stages of leaf primordia development. Interestingly, it has been shown that the synergistic interaction of HAT3 and ATHB4 with ATHB2 involves crossregulation between the HD-Zip IIγ and -δ subfamilies, and evidence has been provided that HAT3 directly and negatively regulates ATHB2 (Turchi et al., 2013). Consistently, ATHB2 became induced in the HAT3/ATHB4 expression domain in the apical part of the embryo and in the SAM of hat3 athb4 loss-of-function mutants, partially compensating for HAT3/ATHB4 function. By contrast, athb2 loss-of-function

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Fig. 3.  Loss of HAT3, ATHB4, and ATHB2 activity affects the expression pattern of PIN1 early during embryo development. (A, C, E, G, I, K) Fluorescent images of PIN1::PIN1:GFP (A, E, I) and hat3-3 athb4-1 athb2-3 PIN1::PIN1:GFP (C, G, K) embryos. Globular (A, C), transition (E, G), and heart (I, K) stage embryos are shown. Bars, 10 μm. (B,D,F,H,J,L) Schematic representations of putative PIN1-dependent auxin fluxes (arrows) in wild-type (B, F, J) (Benková et al., 2003; Izhaki and Bowman, 2007) and hat3-3 athb4-1 athb2-3 (D, H, L) embryos at the globular (B, D), transition (F, H), and heart (J, L) stages. In wild-type embryos at the transition stage, auxin flow reverses from the central apical region of the embryo outwards towards the incipient cotyledon primordia (E, F) (Benková et al., 2003). The reversal in auxin flow outward from the region where the shoot apical meristem will form does not occur in hat3-3 athb4-1 athb2-3 embryos at the transition stage (G, H).

The HD-Zip III protein family is the smallest Arabidopsis HD-Zip family consisting of five members: ATHB8, CORONA (CNA) [also known as ATHB15 or INCURVATA4 (ICU4)], PHB, PHV, and REV. Loss-offunction rev mutants fail to produce axillary meristems and functional floral meristems; gain- and loss-of-function mutations in REV also lead to alteration of vascular patterning within the stem (Talbert et al., 1995; Otsuga et al., 2001). By contrast, loss-of-function mutations in PHB, PHV, CNA, or ATHB8 have subtle mutant phenotypes. However, when higher-order mutants are generated, it becomes obvious that the HD-Zip III proteins act as regulators of embryonic apical fate (Smith and Long, 2010) and are crucial for establishing adaxial/abaxial polarity in the embryo and in the lateral organs during post-embryonic development (Emery et al., 2003; Prigge et al., 2005). For example, the Arabidopsis triple mutant rev phb phv develops a single radial cotyledon with vascular bundles showing amphicribral symmetry and lacks the SAM, indicating that HD-Zip III genes are required to properly pattern the apical region of the globular embryo (Emery et  al., 2003; Prigge et al., 2005). Consistently, HD-Zip III expression is restricted to the apical central region by the globular stage and to the SAM, the adaxial region of the cotyledons, and the vasculature during the heart stage (McConnell et  al., 2001; Otsuga et al., 2001; Emery et al., 2003; Prigge et al., 2005). During the post-embryonic growth phase, HD-Zip III proteins are also required for the maintenance of an active SAM, polarization of newly forming leaf primordia, and the initiation of lateral meristems (McConnell et  al., 2001; Emery et al., 2003; Prigge et al., 2005). In addition, the HD-Zip III genes are necessary for xylem formation and specification, at least during primary growth (Zhong and Ye, 1999; Baima et al., 2001; Ohashi-Ito and Fukuda, 2003; Ohashi-Ito et  al., 2005; Carlsbecker et  al., 2010; Ilegems et al., 2010). Interestingly, the function of the HD-Zip III proteins is not always redundant because multiple loss-of-function mutant plants display synergistic but also antagonistic phenotypes in Arabidopsis. For example, CNA and ATHB8 act oppositely to REV in regulating the formation of lateral shoots and floral meristems (Prigge et al., 2005).

Page 6 of 11 | Turchi et al. Multiple mechanisms control the activity of HD-Zip III transcription factors

Fig. 4.  Schematic representation of the feedback loop between auxin and HD-Zip III and the interplay between HD-Zip II and III proteins. The positive-feedback loop between auxin and REV (through the regulation of LAX2, LAX3, YUC5, and TAA1) and the negative-feedback loop operating within the HD-Zip II family are indicated.

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The important role that HD-Zip III transcription factors play in several aspects of plant growth and development is exerted by a fine-tuned regulation of their expression and function operating at multiple levels. The expression pattern of the HD-Zip III genes essentially coincides with the pattern of auxin distribution (Heisler et al., 2005; Floyd et al., 2006; Floyd and Bowman, 2006) (Fig.  4); ATHB8 expression is regulated by auxin (Baima et al., 1995, 2014) through MP (Donner et  al., 2009), whereas the expression of other HD-Zip III members is associated with the differentiation of vascular tissues as a consequence of changes in the auxin signalling pathway (Ohashi-Ito et al., 2005; Baima et al., 2014). In addition, Arabidopsis HD-Zip III genes are post-transcriptionally regulated by the microRNAs miR165/166, which repress these genes mainly through mRNA cleavage (Emery et  al., 2003; Tang et  al., 2003), resulting in a spatially dynamic regulation of HD-Zip III expression. In Arabidopsis, as well as in other species, nucleotide changes in the coding region result in dominant mutations, due to a loss of miR165- or miR166-mediated regulation (McConnell and Barton, 1998; McConnell et  al., 2001; Emery et  al., 2003; Floyd and Bowman, 2004; McHalea and Koningb, 2004; Juarez et al., 2004; Zhong and Ye, 2004). Failure to restrict HD-Zip III expression in the apical domain via a microRNA-dependent pathway prevents proper establishment of the embryonic root pole (Grigg et al., 2009). Semi-dominant gain-of-function mutations in the miR-binding site of PHB, REV, and CNA restore the topless-1 (tpl-1) double-root phenotype, possibly by excluding PLT1 and PLT2, master regulators of root development whose expression depends partially on MP activity (Aida et al., 2004; Blilou et al., 2005; Galinha et  al., 2007), from the embryo apical domain (Long et  al., 2006; Smith and Long, 2010). Furthermore, mis-expression of miR165- and miR166-resistant variants of REV, PHB, or CNA in the basal cells of the embryo produces a transformation of the root pole into a second shoot pole, indicating that HD-Zip III proteins act as master regulators of embryonic apical fate (Smith and Long, 2010). These dominant alleles all lead to the development of adaxialized leaves with excessive accumulation of the corresponding HD-Zip III transcripts throughout the leaf after emergence of leaf primordia (McConnell et al., 2001; Emery et al., 2003; McHalea and Koningb, 2004; Juarez et al., 2004; Zhong and Ye, 2004). It is noteworthy that the gene encoding ARGONAUTE 10

(AGO10), an AGO protein able to sequester miR165/166 in the central region of the shoot tip (Liu et al., 2009; Zhu et  al., 2011), is positively regulated by REV (Brandt et  al., 2013), thus suggesting another level of regulation that could reinforce the expression of HD-Zip III genes in the adaxial region of lateral organs. The HD-Zip III activity is negatively regulated by LITTLE ZIPPER (ZPR) proteins that prevent HD-Zip III dimerization, an obligate requirement for HD-Zip binding to DNA (Sessa et  al., 1993, 1998; Wenkel et  al., 2007; Kim et  al., 2008). ZPR genes were first identified as direct targets of REV (Wenkel et al., 2007) and, very recently, were shown to evolutionarily derive, through degenerative mutations, from the HD-Zip III transcription factors that they regulate (Floyd et  al., 2014). Double ZPR loss-of-function mutants exhibit an altered SAM activity with abnormal stem-cell maintenance, disruption of normal phyllotaxis, and production of extra cotyledons and leaves, as well as ectopic axillary meristems and sterile flowers (Kim et al., 2008). Together, these data suggest that HD-Zip III and ZPR proteins establish a negative-feedback loop that is required for normal SAM maintenance and function in Arabidopsis (Kim et al., 2008; Brandt et al., 2013). The structure of HD-Zip III proteins suggests other mechanisms of post-translational regulation. In addition to the homeodomain-leucine zipper domain, HD-Zip III proteins contain other domains including a steroidogenic acute regulatory protein-related lipid transfer domain (START), followed by a conserved region of unknown function referred to as the start adjacent domain (Ponting and Aravind, 1999; Schrick et al., 2004). START domains in animals bind cholesterol, phospholipids, and carotenoids (Radauer et  al., 2008), and some are known to be involved in shuttling these compounds between different subcellular compartments (Stocco, 2001). Interestingly, whereas a mouse START domain can functionally replace in vivo the Arabidopsis START domain of GLABRA2 (a member of the HD-Zip IV family), the Arabidopsis REV-START does not (Schrick et  al., 2014). Furthermore, the carboxyl-terminal region of the HD-Zip III proteins, named MEKHLA, possesses sequence similarity to the PAS (Per-Arnt-Sim) domain (Mukherjee and Bürglin, 2006). PAS domains are sensors able to perceive light, oxygen, or redox potentials (Möglich et al., 2009). There is evidence that the MEKHLA domain of REV acts as a negative regulatory domain that might prevent dimerization of this transcription factor (Magnani and Barton, 2011). Interestingly, the APETALA2-like (AP2like) transcription factors DORNROSCHEN (DRN) and DORNROSCHEN-like (DRNL) have been reported to interact with HD-Zip III proteins through the MEKHLA domain (Chandler et  al., 2007). The functional relevance of this interaction is provided by a missense mutation in the MEKHLA domain of CNA, responsible for a complex shoot regeneration phenotype (Duclercq et al., 2011). Intriguingly, DRN is a direct target of MP, and different lines of evidence indicate that DRN and DRNL might control auxin transport in cotyledon development (Chandler et al., 2007; Cole et al., 2009).

HD-Zip II and HD-Zip III proteins in plant development  |  Page 7 of 11

Towards an understanding of the function of HD-Zip II proteins

Fig. 5.  Schematic representation of the possible interactions between HD-Zip II and III proteins in the regulation of apical development and adaxial polarity specification of lateral organs. (A) HD-Zip II proteins, positively regulated by HD-Zip III proteins, act as main direct effectors of the processes. (B) HD-Zip II proteins co-operate together with HD-Zip III factors in the regulation of fundamental aspects of the processes. (C) HD-Zip II proteins negatively regulate the expression of genes that either restrict HD-Zip III expression [i.e. miR165/166; KANADI (KAN)] or reduce HD-Zip III activity (ZPR).

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Auxin not only plays a critical role in pattern specification but is also a trigger of developmental changes induced by environmental cues (Vanneste and Friml, 2009). For example, under shade conditions, the levels of auxin in the Arabidopsis shoot increase rapidly (Tao et  al., 2008; Hornitschek et  al., 2012; Li et al., 2012), and it has been shown that PIF proteins directly and positively regulate the auxin biosynthetic genes YUCCA5 (YUC5), YUC8, and YUC9 (Hornitschek et  al., 2012; Li et  al., 2012). Polar auxin transport is also actively regulated during shade avoidance (Ruzza et al., 2014). Interestingly, there is evidence that REV binds directly to the promoters of the auxin biosynthetic genes TRYPTOPHAN AMINOTRANSFERASE OF ARABIDOPSIS 1 (TAA1) and YUC5, suggesting that part of its role in several developmental processes as well as in the shade-avoidance response is exerted by regulating auxin biosynthesis (Brandt et al., 2012; Huang et al., 2014) (Fig. 4). Recent work has also shown that genes involved in auxin transport, including the influx carriers LIKE AUXIN RESISTANT 2 (LAX2) and LAX3, and response are also directly regulated by REV (Baima et  al., 2014; Huang et al., 2014) (Fig. 4). The positive regulation of auxin influx carriers might contribute to reinforcing the efficiency of polar auxin transport, as predicted by theoretical models (Kramer, 2004) and recently observed experimentally (Robert et al., 2015). It is not presently known if other members of the HD-Zip III family directly regulate auxin homeostasis. However, at least ATHB8 seems to have a role in the process. In leaf primordia, ATHB8 is required to stabilize PIN1 expression against auxin transport perturbation in the procambial cells, resulting in the formation of xylem precursor cells and promoting their differentiation into conducting cells (Baima et al., 2001; Donner et al., 2009). In xylem precursor cells, ATHB8 activates ACAULIS5 (ACL5), a gene encoding a polyamine biosynthetic enzyme, whose product, thermospermine, attenuates xylem differentiation through a negative-feedback loop involving other HD-Zip III genes (Milhinhos et al., 2013; Baima et al., 2014). Intriguingly, this negative regulatory circuit affects other main components of the auxin machinery including TAA1, YUC2, MP, TARGET OF MONOPTEROS 5 (TMO5), TMO5-LIKE 1 (T5L1), PIN6, LAX2, and LAX3. This mechanism is likely to modulate auxin biosynthesis through the action of REV because the rev-5 mutant, whose hypocotyl is significantly shorter than that of the wild type, suppresses the acl5 long-hypocotyl phenotype (Baima et al., 2014). Although ATHB8 and ACL5 are both expressed in the embryo provascular cells (Baima et  al., 1995; Clay and Nelson, 2005), it is not known if the negative-feedback loop is active during embryogenesis. Among the gene targets of REV are HAT3, ATHB4, ATHB2, and HAT2, and there is evidence that HAT3 is regulated by PHB and PHV (Brandt et  al., 2012; Turchi et  al., 2013) (Fig. 4). Consistently, HAT3 and ATHB4 exhibit overlapping expression patterns with PHB, PHV, and REV in the adaxial domain of cotyledons and young leaf primordia, in the vasculature, and in the SAM. By contrast, ATHB2

expression is restricted to procambial cells at the early stages of embryo development and leaf formation; however, ATHB2 becomes induced in the HAT3/ATHB4 expression domains of hat3 athb4 loss-of-function mutants, thus compensating at least in part for HAT3/ATHB4 function (Turchi et al., 2013). The expression pattern of the HD-Zip IIγ and -δ genes in a white-light environment is likely to depend on their responsiveness to the different HD-Zip III proteins, whose levels are tightly regulated through positive and negative feedbacks. The negative-feedback loop within the HD-Zip II protein family further increases the complexity of the HD-Zip III/ HD-Zip II regulatory network (Fig. 4). The molecular interactions between HD-Zip II and III proteins together with the finding that the phenotype of multiple loss-of-function HD-Zip II mutants resembles that of rev phb phv imply that they function in the same pathways. It is, however, still not clear whether HD-Zip IIγ and –δ genes, being positively regulated by HD-Zip III proteins, act as main direct effectors of embryo apical development and adaxial polarity specification of lateral organs or cooperate together with HD-Zip III factors in these processes (Fig. 5). Among other possibilities, the HD-Zip II proteins may negatively regulate molecules that restrict HD-Zip III expression (i.e. miR165/166) or HD-Zip III activity (ZPR genes) (Fig. 5). However, this seems unlikely at least in the vasculature where the expression of gain-of-function REV

Page 8 of 11 | Turchi et al.

Conclusions In the last decade, remarkable progress has been made in dissecting the molecular mechanisms underlying auxin biosynthesis, transport, and response, and in understanding of how they converge to specify different developmental outputs. Our knowledge of master regulators of plant development has increased tremendously, and their connections with the auxin machineries have started to emerge. However, many questions remain to be addressed about how these signals are integrated. The recent findings described in this review highlight a crucial role for HD-Zip II transcription factors in several auxinmediated developmental processes and provide evidence that HD-Zip II genes are directly and positively regulated by HD-Zip III proteins, master regulators of shoot development. However, there are many gaps in our understanding of the HD-Zip III/HD-Zip II network and its interaction with the auxin machineries. The linking of HD-Zip II and III proteins to their targets in a context-specific manner can not only help to untangle this regulatory network but can also contribute to unravel the complex interplay between auxin and master transcriptional regulators.

Acknowledgements We thank all our collaborators who made the work on HD-Zip II and III transcription factors a rewarding experience. Our apologies to the many researchers whose work or original publications could not be cited here because of space constraints. The authors’ work is funded by the Italian Ministry of Education, University and Research, PRIN Program; the Italian Ministry of Agricultural, Food and Forestry Policies, NUTRIGEA Program; and the Italian Ministry of Economy and Finance, Project FaReBio di Qualità.

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dominant (REVd) or ATHB8d under the CNA promoter or ectopic expression of ATHB8 results in a slightly increased production of xylem cells (Baima et al., 2001; Ilegems et al., 2010), whereas overexpression of ATHB2 (ATHB2-OE) causes a reduction in the number of xylem cells compared with controls (Steindler et  al., 1999). ATHB2-OE also results in increased elongation of the hypocotyl mimicking the effect of low R:FR light; radial cotyledon expansion is strongly reduced in ATHB2-OE plants, whereas cotyledon thickness is increased as a result of polarized growth in the cotyledon-thickness direction (Steindler et al., 1999). Seedlings overexpressing other HD-Zip II proteins display similar phenotypes including, as observed in ATHB2-OE, a reduced expansion of the leaf blade (Sawa et  al., 2002; Ciarbelli et al., 2008; Sorin et al., 2009; Köllmer et al., 2011; Ruberti et al., 2012; Turchi et al., 2013). Intriguingly, at least in the first pair of true leaves, this is associated with a reduced cell proliferation that might be responsible for the elongated leaves observed in these transgenic plants (Sawa et al., 2002; Wyrzykowska et  al., 2002; Ciarbelli et  al., 2008; Köllmer et  al., 2011; Turchi et  al., 2013). A  similar phenotype has also been observed in shaded plants in which the reduction of cell proliferation is clearly associated with the auxin gradient from the tip to the base of the leaf (Carabelli et al., 2007), and in multiple cytokinin receptor mutants (Nishimura et al., 2004; Riefler et al., 2006). Interestingly, ATHB2 and HAT3 are regulated by cytokinin (Köllmer et  al., 2011; V.  Forte, G.  Sessa, G.  Morelli, and I.  Ruberti, unpublished data). It is therefore tempting to speculate that, at least in the postembryonic development, some HD-Zip II proteins may be part of a feedback regulatory circuit(s) of cytokinin signalling. By contrast, the alteration of auxin maxima and of cell division planes in hat3 athb4 athb2 mutant embryos clearly indicates a connection between auxin-mediated patterning and HD-Zip II function (Turchi et al., 2013; L. Turchi and I. Ruberti, unpublished data). Auxin and cytokinin have been known for a long time to control several fundamental developmental processes, and recent studies have begun to shed light on the molecular mechanisms underlying auxin/cytokinin action and interaction. The cross-talk between these hormones is exerted at various levels including biosynthesis, degradation, transport, and signalling (Pernisová et al., 2011; Vanstraelen and Benková, 2012; El-Showk et  al., 2013). Interestingly, for instance, an antagonistic interaction between auxin and cytokinin in the cells deriving from the asymmetric division of the hypophysis controls the establishment of the Arabidopsis embryonic root stem-cell niche (Müller and Sheen, 2008). Very recently, seminal studies have started to provide insights into auxindependent control of cell division through direct regulation of cytokinin biosynthesis. In particular, it has been shown that TMO5 and LONESOME HIGHWAY (LHW), by promoting cytokinin biosynthesis within the xylem precursor cells, regulate periclinal divisions in adjacent cells of the vasculature (De Rybel et al., 2013, 2014; Ohashi-Ito et al., 2014). It will be of great interest in the future to determine whether HD-Zip II proteins play a part in the auxin/cytokinin crosstalk during embryo development.

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