Temporal regulation of vegetative phase change in plants

Summary

During their vegetative growth plants reiteratively produces leaves, buds and internodes at the apical end of the shoot. The identity of these organs changes as the shoot develops. Some traits change gradually, but others change in a coordinated fashion, allowing shoot development to be divided into discrete juvenile and adult phases. The transition between these phases is called vegetative phase change. Historically, vegetative phase change has been studied because it is thought to be associated with an increase in reproductive competence. However, this is not true for all species; indeed, heterochronic variation in the timing of vegetative phase change and flowering has made important contributions to plant evolution. In this review, we describe the molecular mechanism of vegetative phase change, how the timing of this process is controlled by endogenous and environmental factors, and its ecological and evolutionary significance.

Introduction

A plant shoot develops by producing leaves, buds, and internodes at regular intervals at the shoot apex. Each of these organs is morphologically or physiologically distinct from the organs produced earlier in development^1,2, which raises the question of how this reiterative process is modified over time to generate morphological and physiological diversity. This question has been studied extensively in both vertebrates and short germ band insects, which also develop by producing organs/segments sequentially at one end of their body axis. In these animals, segmental identity is thought to be specified by the interaction between a segmentation clock and a morphogen gradient.³ Here we discuss the temporal regulation of organ identity during the vegetative phase of shoot development.

Some traits change gradually as a shoot develops, whereas others change abruptly at predictable times in shoot development and, thus, at specific positions on the shoot. This latter phenomenon was first recognized by Hildebrand⁴ and Goebel⁵, who used it to divide vegetative development into juvenile and adult phases; the transition between these phases is called “vegetative phase change”. As the shoot develops it also acquires the ability produce structures involved in sexual reproduction (e.g., flowers or cones). Whether it actually produces these structures depends on a complex interaction between many internal and environmental factors.⁶ Because most plants produce reproductive structures after the transition to the adult vegetative phase, vegetative phase change and the transition from a reproductively-incompetent to a competent phase of shoot development have frequently been considered part of a single program of shoot maturation. As a result, the terms “juvenile” and “adult” are most often used to refer to, respectively, shoots bearing only leaves (vegetative shoots) and shoots that produce reproductive structures such as flowers or cones (reproductive shoots).⁷ However, the length of time between the onset of the adult vegetative phase and the production of reproductive structures varies tremendously within^8,9 and between species. Many trees flower years after the transition to the adult vegetative phase, whereas others—specifically, some species of Acacia¹⁰, Eucalyptus¹¹, and many woody species native to New Zealand¹²–remain permanently juvenile and flower in this condition. These observations suggest that vegetative phase change and the acquisition of reproductive competence are separate, interacting processes, rather than part of a single program of shoot maturation. Unfortunately, very little is known about the regulation of reproductive competence, compared to the regulation of floral induction, which has been extensively investigated.¹³ This review will therefore focus primarily on the mechanism of vegetative phase change. We will first describe some of the key features of vegetative phase change and then discuss the role of MIR156 and SPL genes in this process and how these genes are regulated.

The phenomenon of vegetative phase change

For many years, vegetative phase change was studied primarily in the vine, Hedera helix (English ivy), and in Acacia, Eucalyptus, and woody species native to New Zealand because these species undergo particularly dramatic changes in leaf and shoot morphology during development.^2,14,15 More recent studies have been conducted primarily with short-lived herbaceous plants such as Arabidopsis, maize, and rice because of the advantages these species offer for molecular genetic analysis. Despite the significant difference in the length of their life cycles, many aspects of vegetative phase change are quite similar in these two groups of plants.

In woody plants, phase-specific traits have traditionally been distinguished from other traits that change during shoot development by their stability.¹⁶ For example, lateral buds produced during the juvenile phase continue to produce juvenile traits even after the primary shoot has transitioned to the adult phase. Similarly, adult shoots of woody plants rarely revert to producing juvenile traits, even when grafted to a juvenile root stock. This is in contrast to other age-related traits in woody plants (e.g. growth rate), which can be reversed by grafting a shoot tip to a root stock, or by rooting the shoot tip.¹⁷ With the discovery of genes that regulate vegetative phase change it is now possible to distinguish phase-specific from age-related traits by whether a trait is affected by variation in the expression of these genes in mutant or transgenic plants. As will be described in more detail later in this review, this approach has revealed the tremendous diversity of phase-specific traits, and has facilitated research on the function of these traits in plant biology.

Phase-specific traits vary between species but commonly include the shape and anatomy of the leaf blade, the relative lengths of the leaf sheath/petiole and leaf blade, branching patterns, the capacity for adventitious root production, the presence of epicuticular wax, anthocyanin, trichomes, thorns and other specialized cells and organs, the rate of photosynthesis, and the degree or type of disease and insect resistance.^2,5,14,18 Most species of Acacia, for example, initially produce compound leaves, but then switch during vegetative phase change to producing a simple, vertically oriented leaf known as a phyllode (Figure 1A). In contrast, the ant-acacia, Vachellia collinsii, only produces compound leaves.¹⁹ However, in this species, leaves produced early in shoot development are subtended by small stipules and either lack or have a very small extrafloral nectary, whereas the leaves produced later in development have large thorns, one or more large extrafloral nectaries, and produce Beltian bodies (nutrient-rich structures that serve as food for the ants that live in association with these trees) at the tips of their leaflets (Figure 1B). Juvenile and adult leaves in maize are similar in shape but differ in many epidermal traits (Figure 1C). Specifically, juvenile maize leaves are covered with epicuticular wax, and have bulbous epidermal cells with a thin cuticle and weakly interdigitated cell walls that stain purple with toluidine blue. Adult leaves lack epicuticular wax but possess trichomes and bulliform cells; their epidermal cells are relatively flat and have a thick cuticle and tightly interdigitated cell walls that stain aqua with toluidine blue.²⁰ Juvenile maize leaves also have a lower rate of photosynthesis than adult leaves^21,22 and are more susceptible to common rust and European corn borer.²³ In Arabidopsis, juvenile leaves lack trichomes on their abaxial (lower) surface and are rounder and less serrated^24–26 than adult leaves, and have smaller palisade cells²⁷, a less complex vascular system^28,29 and more complex transfer cells³⁰ than these leaves (Figure 1D, 2A). They also have a lower rate of photosynthesis²¹ and are more susceptible to Pseudomonas syringae³¹ and insect larvae³² than adult leaves.

Figure 1: Species-specific patterns of vegetative phase change.

A) Acacia saligna is typical of over 1,000 species of Acacia in Australia. These species initially produce compound leaves, and then produce leaves with a vertically expanded base and a compound tip before finally producing a simple, vertically oriented leaf known as a phyllode.

B) Vachellia collinsii initially produces compound leaves with a small extra floral nectary (EFN) on the petiole and small stipules. At the 5–10^th node, it begins to produce leaves with a large EFN, large thorn-like stipules, and Beltian Bodies (BB) on the tips of leaflets.¹⁹ The juvenile leaves of this plant have abscised, but their small stipules are still visible.

C) Juvenile and adult leaves in maize are distinguished by a variety of epidermal traits including the presence or absence of epidermal hairs and bulliform cells, the staining of the cell wall with Toluidine blue, cuticle thickness, and the shape of the lateral and outer cell walls of pavement cells.

D) Rosette leaves of the Columbia accession of Arabidopsis grown under long day conditions. As described in the text, the morphology of these leaves and the distribution of abaxial trichomes (shown schematically as white marks) allows shoot development to be divided into the 4 phases shown here.

Figure 2: miR156 is expressed in a similar pattern in a short-lived annual and in a perennial tree.

A) The relative level of miR156 in successive rosette leaves of Arabidopsis plants (Columbia) grown in short days to delay flowering. Modified with permission from Fig. 2 in He et al.³⁸

B) The relative level of miR156 in successive leaves of seed-grown plants of the aspen Populus tremula x alba. A cross section of the petiole is shown below the leaf blade to illustrate its change in shape (round to eliptical) and vascular pattern. Modified with permission from Fig. 1 in Lawrence et al.⁶⁰

C) Arabidopsis plant transformed with a genomic SPL9-GUS construct. There is no visible GUS activity in the cotyledons (C) and in leaves 1&2, consistent with the very high level of miR156 in these organs. GUS activity increases in successive leaves starting with leaf 3. Modified with permission from Fig. 2 in Xu et al.⁵¹

Although it is common to divide vegetative development into a juvenile and an adult phase, it is more accurate to divide this process into 4 phases—a brief “seedling” phase, a juvenile phase that is similar to the seedling phase but is distinct from this phase and longer in duration, a transition phase in which leaves are divided into a distal juvenile domain and a proximal adult domain, and an adult phase (Figure 1D).^29,33–35 In maize, for example, the first two leaves express all of the epidermal traits characteristic of juvenile leaves but are morphologically and anatomically distinct from juvenile leaves²⁰ and also have a different pattern of gene expression.³⁶ Transition leaves possess juvenile epidermal traits, such as epicuticular wax, at the leaf tip and along the margins of the leaf, and adult epidermal traits at the base of the leaf; adult leaves possess these traits throughout the length of the leaf blade.²⁰ Similarly, the first two leaves in Arabidopsis have all of the traits of juvenile leaves but are smaller, rounder, and less responsive to GA than juvenile leaves²⁵ and have a characteristic pattern of gene expression.^37,38 Transition leaves in Arabidopsis have abaxial trichomes at the base of the leaf, but not at the tip, and adult leaves have abaxial trichomes along their entire abaxial surface (Figure 1D). As we will discuss later in greater detail, the apical-basal distribution of juvenile and adult traits in transition leaves indicates that leaf identity is determined relatively late in leaf development.³⁹

The molecular mechanism of vegetative phase change

Regulation of vegetative phase change by the miR156/SPL module

The first insight into genetic mechanism of vegetative phase change came from 3 spontaneous dominant mutations in maize--Teopod1, Teopod2 and Corngrass/Teopod3—whose pleiotropic phenotype was found to reflect the prolonged expression of traits normally restricted to the first nodes of the shoot⁴⁰. This result revealed that maize possesses juvenile and adult vegetative phases, and that the transition between these phases is under genetic control. It opened the way to subsequent genetic analyses of vegetative phase change in both maize^20,41–48 and Arabidopsis^25,49–51 and to the discovery that the microRNA miR156 is the master regulator of this transition.^52–57

In Arabidopsis³⁸, maize^36,54, rice^58,59, Populus tremula x alba⁶⁰ and Vachellia collinsi¹⁹, miR156 is expressed at very high levels in the first 1–5 leaves produced after germination, and at a much lower and slowly declining level in subsequent leaves. In Arabidopsis, for example, the level of miR156 drops 70% from leaf 1 to leaf 3, and then another 10–20% from leaf 3 to leaf 15 (Figure 2A).^38,61 A similar pattern is observed in the tree, Populus tremula x alba, where a major change in miR156 expression occurs between leaves 1 and 5 (Figure 2B).⁶⁰ This pattern is reflected in the expression of the targets of miR156, which accumulate at very low levels in the first few leaves of the shoot, and at increasingly higher levels in later leaves. In Arabidopsis, for example, reporter constructs for all 10 of the genes directly repressed by miR156 are not detectable in cotyledons and leaves 1 and 2, and only begin to be visible starting with leaf 3 or later (Figure 2C).³⁷ This expression pattern explains the observation that leaves 1 and 2 are much smaller and qualitatively different from other juvenile leaves, and that leaves produced later in shoot growth display gradual variation in traits such as leaf shape, complexity of leaf veination, the number of leaf serrations, and the density of abaxial and adaxial trichomes.^25,26,28,29 It also explains why these first two leaves are developmentally more robust than later leaves.^25,62

Exactly how this expression pattern is achieved is still unclear. The abundance of the primary transcripts of MIR156A and MIR156C in Arabidopsis demonstrate that these genes are transcribed more strongly in juvenile leaves than in adult leaves, and is consistent with the amount of mature miR156 in these leaves.^38,63,64 On the other hand, experiments in maize^41,44, Arabidopsis⁶⁵ and potato⁶⁶, indicate that miR156 is diffusible between organs, raising the possibility that at least some of the miR156 in more apical leaves could have come from juvenile leaves with higher levels of miR156. Although there is good evidence that the timing of vegetative phase change is regulated by a change in the transcription of miR156 genes^64,67,68, factors that affect the diffusion of miR156 could also be important regulators of this process.

miR156, and the closely related miRNAs, miR157 and miR529, act by repressing the expression of an ancient plant-specific family of transcription factors, known as SQUAMOSA PROMOTER BINDING PROTEINS (SBP/SPL).^69–71 As their name indicates, SBP family members were first identified by their ability to bind to the promoter of the snapdragon floral identity gene, SQUAMOSA, the orthologue of APETALA1 (AP1) in Arabidopsis.⁷² SBP-LIKE (SPL) genes were subsequently identified in Arabidopsis^73,74, the alga Chlamydomonas reinhardtii⁷⁵, in the moss Physcomitrium patens^76,77, and in all other surveyed land plants.^78–80 In Arabidopsis, miR156/157 regulate the expression of 10 SPL genes^37,70,79 through a combination of transcript cleavage^81,82 and translational repression^83,84, with translational repression being the dominant mechanism.³⁸ A recent genetic analysis suggests the cellular site of translational repression by miR156 may differ from that of other miRNAs.⁸⁵

Some of the effects of the miR156/SPL module are mediated by miR172. In Arabidopsis, miR172 has a complex expression pattern that depends both on inputs from the miR156/SPL module as well as genes involved in leaf expansion and floral induction.^61,86,87 The AP2-like transcription factors targeted by miR172 are important repressors of flowering time, and the function of the miR172/AP2-like module has been studied primarily from this perspective.^61,86–90 These transcription factors also promote juvenile epidermal traits and repress adult epidermal traits in both Arabidopsis^53,61,91,92 and maize.^45,47,48

Vegetative phase change and flowering

It is often assumed that the acquisition of reproductive competence is dependent on vegetative phase change, although there is very little evidence for this in trees⁹³, where flowering occurs long after vegetatively phase change, and some species remain permanently juvenile and flower in this condition.^10,11 Analyses of natural variation in vegetative phase change and flowering in both woody⁹ and herbaceous species^8,94 has revealed that these processes are largely independent under floral inductive conditions, indicating that evolution can manipulate this relationship to adapt species to specific ecological conditions.

It was initially proposed that the developmental increase in reproductive competence in Arabidopsis is regulated by miR156/SPL through its effect on miR172 (the “age-dependent pathway”) because miR172 is downstream of miR156/SPL and has a major effect on flowering time.⁵³ However, subsequent studies showed that under environmental conditions conducive to flowering, variation in the level of miR156 has almost no effect on the sensitivity of plants to a floral inductive stimulus⁶¹ and is not correlated with variation in flowering time in different natural accessions.⁸ These results indicate that the age-dependent increase in reproductive competence in Arabidopsis is not regulated by the vegetative phase change pathway in plants that are sensitive to photoperiodic induction.

The effect of the miR156/SPL module on flowering can be difficult to determine in some species because SPL genes have a significant effect on the rate of leaf initiation and on the expression of genes (e.g. AP1, LFY) involved in the floral meristem identity transition (i.e. the transformation of vegetative buds into flowers or flower-bearing shoots during the development of the inflorescence). Consequently, leaf number—which is commonly used as a proxy for flowering time—is a poor predictor of flowering time in plants with elevated or reduced levels of miR156 or SPL proteins. The Tp2 mutant of maize, for example, has an elevated level of miR156 due to the over-expression of Zma-miR156h.^56,57 This mutant has an increased number of leaves and a small unbranched tassel with only a few flowers, but does not exhibit a major delay in its sensitivity to a floral inductive photoperiodic stimulus, and terminates shoot development at same time as wild type plants⁴³; the “leafy” phenotype of this mutant is attributable to a delay in the floral meristem identity transition, not to a delay in the acquisition of reproductive competence. Similarly, in wheat⁹⁵ and snapdragon⁹⁶, over-expression of miR156 or suppression of SPL gene expression does not significantly delay the timing of the transition from a vegetative to an inflorescence growth pattern, although these genotypes produce an increased number of leaves because of an accelerated rate of leaf initiation and/or a delay in the floral meristem identity transition.

In contrast, over-expression of miR156 delays floral induction in tobacco⁹⁷, alfalfa⁹⁸ and switchgrass⁹⁹, and silencing of PhSBP1 and PhSBP2 delays floral induction in petunia.¹⁰⁰ In switchgrass and alfalfa, floral induction was only delayed significantly in transgenic lines that had extremely high levels of miR156. Lines with lower levels of miR156 had aberrant vegetative morphology but flowered normally. This result suggests that in these species miR156 may inhibit flowering during the seedling stage of development, when miR156 is present at very high levels, but have relatively little effect on flowering during the juvenile phase. To determine if reproductive competence (i.e., the ability to respond to floral inductive stimuli) is regulated by the same mechanism as vegetative phase change, it is important to know if varying the level of miR156 within its normal range has an effect on flowering time, and this is not usually examined. It is clear that the miR156/SPL module has the potential to regulate reproductive competence in some species and in some conditions, but it is unlikely to be a major contributor to variation in flowering time in all plants.

Genetic analysis of the function of SBP/SPL genes

The phenotype of plants over-expressing miR156—that is, plants with reduced expression of miR156-regulated SBP/SPL genes--has been characterized in many taxa, including Arabidopsis^{32,52,53,101–104}, maize^23,40,54, rice^59,105,106, wheat⁹⁵, tobacco^97,107, alfalfa⁹⁸, Populus^55,60, switchgrass^108,109 and tomato.^110,111 In addition to the prolonged expression of juvenile morphological traits and juvenile patterns of photosynthesis, disease resistance, and insect resistance, this phenotype typically includes an increase in the rate of leaf initiation, an increase in shoot branching and lateral root production, and a variety of floral and fruit defects, including the transformation of floral buds into leafy shoots. The appearance of these phase-specific traits may not be completely coordinated, and some traits are more sensitive to environmental conditions or experimental manipulation than others^8,19,112. This result would be explained if the many SPL genes regulated by miR156 were functionally distinct and had different sensitivities to miR156, were regulated differently at a transcriptional level, or interacted with unique sets of RNAs or proteins. In this case, developmental or environmentally-induced variation in the level of miR156 could affect some SPL genes more than others, resulting in a lack of coordination between different phase-specific traits. As described below, research in Arabidopsis and a few other species has shown that SPL genes are indeed functionally differentiated and have different transcription patterns, but whether they are differentially sensitive to miR156 remains to be determined.

The function of individual SPL genes has most often been inferred from the phenotype of plants expressing a miR156-resistant version of the gene under the regulation of the endogenous or a constitutive promoter. However, this approach can produce misleading information because it causes genes to be expressed at unusually high levels at inappropriate times and in inappropriate tissues. Methods that reduce or eliminate SPL gene expression (e.g. over-expression of miR156 or loss-of-function mutations in SPL genes) provide more reliable information about gene function, so our discussion will be limited to studies that have employed this approach. The phenotypes of loss-of-function mutations and the expression patterns of miR156-insensitive reporters for the 10 miR156-targeted SPL genes in Arabidopsis reveal that these genes can be divided into several distinct groups.³⁷ SPL9 and SPL13 are strongly transcribed and have similar and quite significant effects on most aspects of vegetative phase change, demonstrating that they are the major regulators of vegetative phase change and have shared functions in this process. SPL15 is transcribed at a lower level than SPL9 and SPL13 and has a smaller effect on vegetative phase change than these genes, but has larger effect on floral induction, particularly under short days.^37,113,114 SPL2, SPL10 and SPL11 are expressed at a relatively low level and have a relatively small effect on morphological aspects of vegetative phase change^37,53,115, but are exclusively responsible for mediating adult resistance to Pseudomonas syringae.³¹ The function of SPL6 is unclear because mutations in this gene have no obvious morphological phenotype and although they have been reported to affect disease resistance¹¹⁶, this result has not been confirmed.³¹ SPL3, and its paralogues SPL4 and SPL5, were initially thought to be important for floral induction because over-expression of SPL3 causes early flowering.⁷³ However, this phenotype was later shown to be an artefact of over-expression because plants mutant for all three of these genes flower at the same time as wild type plants.³⁷ The inflorescence phenotype of this triple mutant³⁷, and an analysis of the direct targets of SPL3¹¹⁷, demonstrate that SPL3/4/5 promote the floral meristem identity transition, a process that occurs after floral induction and is regulated by AP1 and LFY, which are direct targets of SPL3,. SPL3 is strongly transcribed during the adult vegetative phase, but whether it has a function in vegetative development is unknown because spl3 mutants have no obvious morphological phenotype.³⁷

Three functionally redundant SBP/SPL genes in maize—tasselsheath4 (tsh4), unbranched2 (ub2) and unbranched3 (ub3)—control many aspects of lateral organ development. Loss-of-function mutations in tsh4 cause the development of bracts in the inflorescence¹¹⁸, while plants triply mutant for tsh4, ub2 and ub3 have numerous tillers, an unbranched leafy inflorescence, and flower at the same time as wild type but produce more leaves than normal because they have an elevated rate of leaf initiation.¹¹⁹ This phenotype is very similar to that of Corngrass, Teopod1, and Teopod2⁴⁰, all of which cause the overexpression of miR156^54,56,57. Whether tsh4, ub2 and ub3 regulate leaf traits associated with vegetative phase change is unknown because these traits were not examined in these studies. The rice gene, OsSPL14—also known as Wealthy Farmer’s Panicle (WFP)¹²⁰, and Ideal Plant Architecture (IPA)¹²¹—is closely related to tsh4, ub2 and ub3, and has a similar function. Like these maize genes, loss-of-function mutations in OsSPL14 dramatically increase tillering and produce a leafy inflorescence.^122,123 Another SPL gene involved in the development of the maize inflorescence is Teosinte glume architecture1 (Tga1). Tga1 regulates the development of the glumes surrounding the maize kernel and, along with its upstream regulator Teosinte branched1¹²⁴, was an important target of selection during maize domestication.¹²⁵

The snapdragon genes, AmSBP1, the tomato gene, SlCNR, and the petunia genes, PhCNR, PhSBP1 and PhSBP2, are in the same clade as SPL3, SPL4 and SPL5 in Arabidopsis.^79,100 Viral-induced gene silencing of AmSBP1 produces a leafy inflorescence, a phenotype that is similar—albeit much more extreme—to the phenotype of loss-of-function mutations in SPL3, SPL4 and SPL5.⁹⁶ In petunia, viral-induced silencing of PhSB1 delays inflorescence initiation, flower production, and leaf initiation, whereas silencing of PhSBP2 had no effect on the timing of inflorescence initiation, but delays flower production.¹⁰⁰ In contrast, silencing of SlCNR delays fruit ripening, but has no obvious effects on flowering time or inflorescence development.^126,127 These results suggest that members of this clade primarily regulate floral meristem identity and fruit development, although they may regulate floral initiation in some species.

How do SBP/SPL genes promote adult vegetative traits?

In many species, a change in leaf shape is the most obvious indicator of vegetative phase change. This change can be quite dramatic, as in Acacia (Figure 1A), but most often involves an increase in leaf serration or leaf lobing, or a change in the angle of the leaf base or the shape or length of the petiole or leaf sheath. In Arabidopsis, for example, the first few leaves of the rosette have long, thin petioles, and have a round leaf blade with few serrations. Leaves produced later in shoot development become increasingly serrated and more elongated, their petioles become thicker and shorter, and the boundary between the leaf blade and petiole becomes less distinct (Figure 1D, 2A).²⁶ The increase in leaf serration is regulated, in part, by post-translational interactions between SPL proteins and two families of transcription factors that regulate leaf serration, CUC and TCP¹²⁸. CUC proteins promote serration, but are prevented from doing so in juvenile leaves by their interaction with TCP proteins. This interaction is disrupted in adult leaves by SPL9, which binds to TCP4, thus preventing it from interacting with CUC proteins (Figure 3).

Figure 3: The mechanisms by which phase-specific traits are regulated by miR156/miR157.

Active genes/functions are represented by black lines and black text, and inactive genes and functions are represented by gray lines and text. The result of these genetic interactions is provided at the bottom of the figure.

The change in the length and shape of the leaf blade and petiole is regulated by BOP1 and BOP2 (Figure 3).¹²⁹ In juvenile leaves, these transcription factors promote petiole development by repressing the expansion of the leaf blade. In adult leaves their transcription is repressed by SPL9 and SPL13, which leads to the extension of the leaf blade onto the petiole and produces the acute leaf base and relatively short petiole characteristic of adult leaves. The basal expansion of adult leaf blades is facilitated by enhanced anisotropic cell growth, which is associated with increased SPL activity.¹³⁰ This mechanism also operates in rice. In rice, juvenile leaves have a long leaf sheath and very short leaf blade, whereas adult leaves have a long leaf blade and a relatively short leaf sheath. This difference is regulated by OsBOP genes, which promote the development of the leaf sheath and repress the development of leaf blade.¹³¹ As in Arabidopsis, these genes are expressed at high levels in juvenile leaves, but are repressed in adult leaves by SPL transcription factors, allowing for the development of the leaf blade.

The distribution of trichomes on the leaf blade is regulated by SPL genes in part through their effect on miR172 expression. SPL9 and SPL15 promote the transcription of MIR172B^53,113,132, and thus contribute to the relatively high level of miR172 in adult leaves compared to juvenile leaves.⁶¹ This expression pattern is expected to produce higher levels of AP2-like transcription factors in juvenile leaves than in adult leaves. One of these transcription factors is TOE1. TOE1 interacts with the transcription factor, KANADI1--which is expressed in abaxial leaf cells—to repress the transcription of the trichome regulator, GLABRA1, on this surface of the leaf.^91–92 As TOE1 is expected to be present at higher levels in juvenile leaves than in adult leaves, this would explain the absence of abaxial trichomes on juvenile leaves (Figure 3). However, SPL regulation of miR172 cannot be the only explanation for the effect of SPL genes on abaxial trichome production because the trichome phenotype of plants with elevated SPL levels is only partially suppressed by mutations that nearly eliminate miR172 expression.⁶¹ This result suggests that SPL genes also regulate this phenotype independently of miR172.

In maize⁴⁰ and rice¹⁰⁶, elongated branches known as tillers are only produced by juvenile nodes. It is unknown if the capacity for branch outgrowth is also a juvenile trait in species that do not branch excessively, but this may be the case because constitutive over-expression of miR156 produces excessive branching in every species in which this trait has been engineered. This result suggests that SPL proteins repress branch outgrowth during the adult phase, when miR156 levels are low. The mechanism by which they do this has been elucidated in both rice^123,133 and wheat.⁹⁵ In both species, SPL proteins repress branching by promoting the transcription of orthologues of the maize branching repressor TEOSINTE BRANCHED1. The activity of these SPL proteins is also regulated by the hormone strigolactone through its effect on the repressor, D53.^95,133 In the absence of strigolactone, SPL proteins are inactivated by their association with D53, which enables branch outgrowth. D53 is degraded in the presence of strigolactone, allowing SPL proteins to promote the transcription of TB1 and thus repress branch outgrowth.

Epigenetic regulation of miR156 expression

In 1962, Brink made the prescient suggestion that vegetative phase change is regulated by “self-perpetuating accessory materials” he termed “parachromatin”.¹⁶ He also suggested that these materials were responsible for stable developmental states in many other organisms, including animals. This paper was published before the discovery of transcription factors and other forms of gene regulation in eukaryotes, and so was largely ignored. It is therefore remarkable that he was essentially correct. Based largely on work in Arabidopsis, it is now clear that epigenetic factors play a central role in coordinating the temporal decline in miR156 expression, and therefore the timing of vegetative phase change (Figure 4). Core to this epigenetic timing mechanism is the balance between active (e.g. H3K27ac) and repressive (e.g. H3K27me3) histone marks at MIR156 loci. Levels of H3K27ac are determined by the activity of histone acetyl, and histone deacetyl, transferases.¹³⁴ On the other hand, H3K27me3 levels are mediated by the activity of the transcriptional repressor complexes POLYCOMB REPRESSIVE COMPLEX 2 (PRC2) and PRC1. PRC2 contains a methyltransferase and directly deposits H3K27me3, whereas PRC1 represses gene expression via H2A ubiquitination and also promotes PRC2 activity at a subset of its targets.¹³⁵

Figure 4: Model for the epigenetic regulation of MIR156 transcription in Arabidopsis.

This model combines data from studies of MIR156A and MIR156C, which are regulated by slightly different epigenetic mechanisms that have not been completely elucidated. MIR156 transcription is regulated by nucleosome remodeling and histone modifications. During the juvenile phase, the chromatin remodeler, Brahma (BRM), repositions the +2 and −1 nucleosome to promote transcription, while PKL prevents repositioning of the +1 nucleosome. It is unclear whether BRM remains at MIR156 genes during the adult phase, but in any case, PKL appears to have a more significant effect at this stage. PKL represses transcription both by inhibiting nucleosome repositioning and through its association with histone deacetylases, most importantly, HDA9. The loss of the active mark, H3K27ac, is associated with an increase in the repressive mark, H3K27me3, which is deposited by PRC2. Binding of PRC2 to MIR156 is promoted by PRC1 but can also occur independently of PRC1. In addition to promoting the activity of PRC2, PRC1 may represses MIR156 transcription via H2A ubiquitination. The transcription factor, VAL1, is present at MIR156 throughout development and binds to the BMI1A component of PRC1, facilitating its association with MIR156. However, VAL1 is not responsible for the temporal regulation of PRC1 or PRC2 activity.

During vegetative development, H3K27me3 marks are gradually deposited in place of H3K27ac, leading to the transcriptional repression of MIR156A/C and the transition to adult development.⁶⁴ Binding of PRC2 to MIR156A/C increases during the period when the transcription of these genes is down-regulated, and loss-of-function mutations in the PRC2 methyltransferases SWINGER (SWN) delay vegetative phase change, elevate miR156 accumulation and reduce H3K27me3 levels at MIR156A/C – with subtly different effects on MIR156A and MIR156C.⁶⁴ PRC1 is important for PRC2 activity at MIR156A/C as loss of the PRC1 recruitment factors VIVIPAROUS1 ABI3-LIKE1 (VAL1) and VAL2, and of PRC1 components themselves, increases the overall level of miR156 and reduces H3K27me3 deposition (predominantly at MIR156C).^136,137 In addition to its role in recruiting PRC2, the H2A ubiquitination function of PRC1 also appears to be important in the repression of MIR156A/C.^68,136 Loss of the PRC accessory protein LIKE-HETEROCHROMATIN 1 (LHP1) in rice leads to elevated transcription of MIR156 loci concomitant with reduced H3K27me3¹³⁸, suggesting PRC-mediated silencing of MIR156 is conserved across flowering plants.

H3K27ac removal is largely determined by HISTONE DEACETYLASE 9 (HDA9).^68,139 The rate of both H3K27ac removal by HDA9, and H3K27me3 deposition by PRC2, is promoted by the CHD3 chromatin remodeler PICKLE (PKL). Genetic analyses have shown that pkl enhances the hda9 and swn delayed vegetative phase change phenotypes, with a far stronger effect of the respective double mutant on MIR156A/C expression than either single mutant.^64,68 PKL therefore plays a critical role in coordinating the temporal exchange of histone modifications during vegetative phase change, at least in part through its effects on nucleosome occupancy.⁶⁴ In animals, the PKL homolog Mi2β/CHD4 forms part of the nucleosome remodeling and deacetylation (NuRD) complex, which facilitates PRC2-recruitment via precursory H3K27ac removal.¹⁴⁰ Whether or not an equivalent NuRD complex functions in plants has been somewhat controversial.¹⁴¹ However, the recent discovery that PKL physically interacts with HDA6 and HDA9⁶⁸, and functions in both H3K27 deacetylation and methylation⁶⁴, strongly suggests that plants possess a NuRD-like complex.

The repressive effects of PKL and PRC2 are antagonized by chromatin remodeling complexes that promote MIR156A/C expression. The SWI2/SNF2 chromatin remodeling ATPase BRAHMA (BRM) reduces nucleosome occupancy and H3K27me3 levels at MIR156A, and loss of function brm mutants exhibit early phase change.¹⁴² Likewise, the SWR1 chromatin remodeling complex promotes MIR156A/C expression through maintenance of the H2A.Z histone variant and deposition of the positive H3K4me3 histone mark.^143,144

Despite recent insights from epigenetic studies, how the timing of the deposition of H3K27me3 at MIR156A/C is controlled remains to be determined. Based on the maintenance of a MIR156C reporter construct in quiescent cells relative to actively dividing ones, and chemical inhibition of the cell cycle, it has recently been proposed that the deposition of H3K27me3 is triggered by cell division.¹³⁹ Although this model would provide a parsimonious explanation for how miR156 activity decreases with age, it conflicts with the observed rates of phase change in mutants with slower^25,65 and faster⁸⁵ rates of cell division. Furthermore, a cell division-dependent decline in miR156 expression within cells of the SAM is not consistent with the almost permanently quiescent nature of some SAM cells¹⁴⁵, nor is cell division-dependent repression of miR156 expression consistent with the re-initiation of miR156 expression during leaf development¹⁰⁵, reproductive development⁶⁷ or culture-induced rejuvenation.³⁹

How other genetic factors found to regulate miR156 expression might interact with a putative cell division model is unclear. For example, the transcription factors AGL15/18, MYB33, ABI3/5, FUS3 and NF-YB8/NF-YA8^146–151 all promote miR156 expression whereas PLT2, PNY/PNF, the CDK8 mediator complex and the barley histone acetyltransferase MND8 all repress miR156 expression.^152–156 Which MIR156 loci these factors regulate, and whether they do so in the context of vegetative phase change, is not always apparent. An intriguing additional layer of genetic regulation at MIR156A/C was revealed by the discovery that micropeptides encoded by open reading frames upstream of the miRNA hairpin initiate self-activating loops that promote expression of the primary miRNA transcripts.^157–159

The likely complexity of the phase change clock is highlighted by the remarkable robustness of the temporal expression of MIR156A/C despite extensive attempts to genetically disrupt it. In almost all cases, loss of function mutations in regulators of MIR156A/C lead to changes in the overall level of miR156, rather than a change in the rate of its decline.^{64,68,137,138,142,144} Consistent with its role in both H3K27 deacetylation and trimethylation, genetic combinations featuring pkl appear to have the strongest reduction in the rate of decline, as pkl swn and pkl hda9 mutant plants maintain a relatively consistent level of MIR156C expression.^64,68 Importantly, MIR156A expression—while increased overall—still declines in a temporal pattern in these plants. Despite their recent duplication and sequence similarity, MIR156A and MIR156C are regulated by only partially overlapping mechanisms. This divergence likely reinforces the rigid timing mechanism of phase change and may provide an excellent model for the study of molecular evolution.

Ultimately, the phase change clock needs to be reset between generations. Although the nature of this clock is still unknown, a cis-regulatory region that is conserved between MIR156A/C and is required for their embryonic re-activation has recently been identified and may provide some clues.⁶⁷ Genetic and biochemical assays suggest the embryogenesis regulator LEAFY COTYLEDON 2 (LEC2) binds this region and is critical for resetting MIR156A/C expression. How LEC2 and the closely related VAL1/2, which bind the same DNA motif and have opposing effects on the timing of vegetative phase change¹³⁷, coordinate activation vs repression of MIR156A/C remains to be determined.

How is the timing of vegetative phase change regulated?

Although there is good evidence that vegetative phase change is the result of the transcriptional repression of miR156 and miR157 genes by epigenetic factors, this does not answer the question of how the timing of this process is regulated. The endogenous and exogenous factors responsible for variation in leaf morphology during shoot development (heteroblasty) were of considerable interest in the first half of the 20^th century¹, but these studies were never framed in terms of the more general process of vegetative phase change because it was assumed that these factors acted specifically on leaf morphogenesis, not on the developmental phase of the shoot as a whole. More recent studies have shown that at least some of these factors (e.g. defoliation, sugar, low light intensity) affect the expression of miR156/miR157 or the transcription of SPL genes, indicating that the literature on heteroblasty is a good source of inspiration for research on the mechanism of vegetative phase change. Linking these factors to changes in the epigenetic landscape of miR156 and miR157 genes remains a challenge, however.

Role of the shoot apical meristem

Whether the shoot apical meristem (SAM) controls the timing of vegetative phase change autonomously, or simply responds to external cues, is a classic question in plant biology.^160,161 It was traditionally thought that the juvenile-to-adult transition is regulated within the SAM¹⁶², but more recent work has demonstrated that pre-existing leaves promote both the initiation and the maintenance of the adult phase. For example, below a threshold number of leaf primordia, maize apex explants will rejuvenate during culture, suggesting that a leaf-derived signal is actively required to maintain apices in the adult phase.^39,163 In apices cultured with 6 leaf primordia, the existing primordia underwent rejuvenation while the SAM remained in the adult phase³⁹, demonstrating that leaf identity in maize does not require a change in the state of the SAM. Leaf ablation experiments in Arabidopsis have demonstrated that the transition to the adult phase is promoted by a leaf-derived signal that represses the expression of miR156.¹⁶⁴ Other experiments showed that this signal is diffusible and suggested that it might be glucose or another carbohydrate.^63,165 Additionally, cytokinin regulates the timing of abaxial trichome production in leaves, not in the SAM.¹⁶⁶ On the other hand, the observation that mutant plants with defective SAMs produce adult leaves precociously^25,65,167 demonstrates that the SAM is not an entirely neutral player. Importantly, the phase change phenotypes of these mutants, and plants with reduced miR156 activity, can be partly rescued by expression of miR156 specifically within the SAM.⁶⁵ Together, these results suggest that vegetative phase change is regulated by a complex interaction between leaves and the SAM, with the SAM being important for the specification of juvenile identity very early in development, whereas later shoot identity is regulated primarily by leaves.

Regulation of vegetative phase change by sugar

As noted earlier in this review, ablation studies indicate that leaves produce a stable diffusible factor that represses miR156¹⁶⁴, and subsequent studies have suggested this factor might be sugar. One attractive idea is that the “phase change clock” is the increase in the amount of sugar produced by the shoot as its photosynthetic capacity increases as a result of an increase leaf number or leaf expansion. Given the increased energetic cost of producing adult leaves²², carbohydrate-based signaling represents an attractive way to synchronize vegetative phase change with nutritional state, and the striking effect of low light conditions on the timing of vegetative phase change (described below) supports this idea.¹⁶⁸

Glucose, fructose and sucrose all suppress MIR156A/C expression, in part through the glucose sensor HEXOKINASE1.^63,165 The sucrose level in a plant influences the timing of vegetative phase change via the signaling metabolite trehalose-6-phosphate, which is synthesized by the enzyme TREHALOSE PHOSPHATE SYNTHASE1 (TPS1). Loss of TPS1 function leads to delayed phase change and elevated MIR156A/C expression.^169,170 The sugar sensor SUCROSE NON-FERMENTING1 RELATED-KINASE (SnRK1) likely functions downstream of TPS1, as inhibition of SnRK1 activity suppresses the increased miR156 accumulation of tps1.¹⁷¹ Intriguingly, the master integrator of nutrient status TARGET OF RAPAMYCIN (TOR) kinase has recently been shown to regulate PRC2 activity through phosphorylation of the core PRC2 component FIE¹⁷², suggesting a potential mechanism for how plant energy status coordinates the timing of phase change.

Hormonal regulation of vegetative phase change

Plant hormones were found to regulate the timing of vegetative phase change long before the molecular mechanism was elucidated.¹⁷³ Gibberellic acid (GA) accelerates or delays phase change in woody species and accelerates vegetative phase change in both Arabidopsis and maize.^25,46 The miR156 pathway operates at least partly downstream of GA as constitutive expression of miR156 and PKL are both epistatic to the effects of GA signaling.^174,175 However, SPL9 and SPL15 have been shown to physically interact with GA-sensitive DELLA proteins^113,174,176, suggesting that the regulation of vegetative phase change by GA is multi-layered. More recently, jasmonic acid (JA) has been shown to delay phase change by promoting the expression of miR156 in rice and maize.^36,177 SPL proteins also interact with JA signaling factors³².

Interactions between plant hormone networks and the miR156/SPL pathway appear to occur more frequently at the protein level as SPL proteins physically interact with brassinosteroid^178,179, strigolactone^123,180, ABA¹⁸¹ and cytokinin¹⁸² signaling components. SPL proteins and cytokinin signaling also function complementarily to promote miR172 expression.¹⁶⁶ The observation that hormones act downstream of miR156/miR157, as well as the absence of evidence linking developmental changes in the level of these hormones with vegetative phase change, suggest that plant hormones mainly affect the manifestation of vegetative phase change, rather than its timing.

Regulation of vegetative phase change by environmental conditions

The control of developmental timing is intrinsically robust but also sensitive to environmental conditions. In many species the timing of vegetative phase change is delayed by low light intensity.^1,19,168,183 The discovery that juvenile leaves are photosynthetically more efficient under low light intensity than adult leaves^21,22 provides a physiological explanation for this delay, and its molecular basis is suggested by the observation that low light intensity increases the abundance of miR156 in Arabidopsis and Vachellia collinsii.^19,168 In the case of Arabidopsis, the delay is independent of phytochrome and cryptochrome signaling and is associated with a reduction in H3K27me3 at MIR156C, although miR156-independent regulation of SPL gene expression is also a contributing factor.¹⁶⁸ The effects of low light are at least in part mediated through nutritional status, as exogenous sucrose partly suppresses the low light phenotype. Shade-adapted species photosynthesize efficiently under low light intensity throughout their life and have many morphological traits (the “shade tolerance syndrome”) typical of juvenile shoots.¹⁸⁴ It may be that the phenotype of these plants reflects the extended expression of the juvenile phase resulting from either elevated levels of miR156 or the reduced transcription (e.g. via mutation) of one or more SPL genes. Levels of miR156 are also regulated by soil nutrient conditions. Consistent with the idea that vegetative phase change is delayed until conditions are favorable, phosphate and nitrogen deficiency induce miR156 accumulation.^143,185

Although phytochrome signaling does not play a role in the response to light intensity, it does regulate the miR156/SPL module in response to light quality. Under far-red enriched light, Arabidopsis undergoes a “shade-avoidance syndrome”, which is distinct from the shade tolerance syndrome induced by overall low light levels¹⁶⁸. As part of the shade-avoidance syndrome, petioles lengthen, branching is inhibited and plants flower earlier. Constitutive expression of miR156 suppresses the shade-avoidance syndrome, which is induced by the PHYTOCHROME INTERACTING FACTOR (PIF) family of transcription factors. As PIF family members repress miR156 expression, it has been suggested that PIFs promote the shade-avoidance response via miR156.¹⁸⁶ However, the MIR156 loci that PIF proteins regulate (i.e. MIR156B/D/E/F/H) have limited roles in Arabidopsis development^38,67, making the significance of the interaction between these proteins and MIR156 loci unclear. PIFs¹⁸⁷, and the phytochrome signaling components FHY3 and FAR1^180,188, directly interact with SPL proteins, suggesting multi-layered regulation of the miR156/SPL network by light quality. This regulatory complexity is highlighted by the finding that UV-B wavelengths elevate miR156 levels via inhibition of H3K27me3 deposition.¹⁸⁹

Vegetative phase change is also sensitive to temperature and drought. Arabidopsis grown at 16°C exhibits delayed phase change and increased levels of miR156 relative to plants grown at 23°C^190,191. Interestingly, the elevated level of miR156 in 16°C plants is regulated post-transcriptionally as pri-MIR156A transcript levels are lower in these plants than in plants grown at 23°C.¹⁹² Extremely high temperatures also increase miR156 levels, and this may lead to increased tolerance to heat stress.¹⁹³ Similarly, miR156 expression is induced by drought while over-expression of miR156 promotes drought tolerance.¹⁹⁴ Downstream of miR156, SPLs limit plant responses to increased temperature by reducing sensitivity to auxin.^191,195 As temperature and shade are both mediated by PIF signaling, SPL-dependent auxin sensitivity may also contribute to the potential role of miR156 in the shade-avoidance response.

We have focused here on the regulation of miR156. It is important to emphasize that the timing of vegetative phase change can also be affected by factors that affect the transcription^196–198 or activity^{128,178,199–202} of the SPL proteins regulated by miR156. The relative importance for vegetative phase change of these mechanisms vs. post-transcriptional regulation of SPL transcripts by miR156 remains to be determined.

The evolution and ecological significance of vegetative phase change

The ancestral functions of the miR156/SPL module

SBP/SPL genes can be traced back to the algae whereas miR156—one of the most ancient and highly conserved miRNAs in plants—emerged when plants first colonized land.^76,203 As the ancestral land plants looked very different from the flowering plants in which miR156/SPL function has mainly been studied, identifying the ancestral function of miR156 and its role in the establishment of land plants is a key area of research. Studies of two experimentally tractable bryophyte models, the moss, Physcometrium (Physcomitrella) patens, and the liverwort, Marchantia polymorpha—whose lineage diverged from the rest of the plant kingdom approximately 500 million years ago^204,205—have started to yield answers.

Unlike flowering plants, these organisms spend most of their life cycle as haploid gametophytes, and the functions of the miR156/SPL module have been investigated during this phase of the life cycle. Early moss development consists of a 2D filamentous growth phase, during which gametophores – leafy shoots derived from a single apical meristem cell – subsequently initiate. The growth of gametophores terminates following the production of reproductive organs which, after fertilization, nurture the diploid sporophyte. Gametophores produce morphologically different leaves along their axis, which have been interpreted as juvenile and adult forms²⁰⁶. In Tortula, the timing of the transition between these two leaf types differs between species, and some species initiate reproductive development having only produced juvenile leaves²⁰⁶. In Physcomitrium, miR156 levels increase slightly during the filamentous-to-gametophore transition, and then decrease dramatically as the gametophore develops; consistent with this pattern, SPL/SBP gene expression decreases and then increases dramatically during gametophore maturation²⁰⁷. Transgenic lines with reduced levels of miR156 produce gametophores much less frequently than wild type, whereas Ppsbp3 mutants produce more gametophores than normal²⁰⁸, indicating that PpSBP3 represses meristem development. Whether this module also plays a role in the juvenile-to-adult transition during gametophore development is unknown because the phenotype of the gametophores in these lines was not characterized.

In contrast to mosses, gametophytes of M. polymorpha grow as a branching thallus from which umbrella-like gametangia arise. M. polymorpha has 4 SPL/SBP genes, one of which, MpSPL2, is orthologous to the miR156-regulated genes in higher plants. MpSPL2 is regulated by miR529c, a miRNA closely related to miR156.²⁰⁹. Loss of miR529c or over-expression of MpSPL2 causes the production of gametangia in the absence of an inductive far red light stimulus; loss of MpSPL2 has minor effects on gametangia morphology and the timing of gametangia formation, but does not affect gamete production or function.²¹⁰ These results suggest that MpSPL2 promotes, but is not required for the transition to reproductive development. A second SPL/SBP gene, MpSPL1, is regulated by the liverwort-specific miRNA, Mpo-MR13, and affects thallus branching by promoting meristem dormancy.²¹¹

The functions of the miR156/SPL module in the gametophyte of bryophytes closely resemble their functions in the sporophytes of angiosperms. For example, the role of PpSBP3 in suppressing the development of apical initials that give rise to the gametophore is similar to the function of SPL genes in Arabidopsis, which suppress the growth of the shoot apical meristem.⁶⁵ Although the function of the miR156/SPL module during gametophore development is unknown, the complementary temporal expression patterns of these genes is equivalent to their expression during shoot development in flowering plants, so it would not be surprising if they regulated the juvenile-to-adult transition during this phase of moss development. The role of MpSPL2 in gametangia development in Marchantia is similar to the roles of SPL genes in the regulation of flowering time and floral meristem identity. Although MpSPL1 is not in the same clade as miR156-regulated SPL family members, it has an analogous function in branch development as these genes. It is clear, therefore, that a role for miR156 in the regulation of critical developmental processes, including developmental identity and meristematic growth, is widely conserved across land plants. Furthermore, miR156 likely regulated phenomena equivalent to vegetative phase change in the ancestral land plant. Whether or not miR156 regulated development in both the gametophyte and sporophyte phases of early land plants, or if its sporophytic role in angiosperms was co-opted from ancestral gametophytic genetic networks²¹², is a topic for future research.

The evolution of vegetative phase change in higher plants

Plants face different challenges at different times in their development and must evolve temporally specific morphological and physiological adaptations to these conditions. Much of the biomass on earth consists of perennial species—specifically trees and shrubs--that spend most of their life in the vegetative phase of development. These species must contend with a wide range of changing environmental conditions during their lifespan, and phase-specific traits likely evolved in response to these conditions. The striking conservation of juvenile leaf morphology in Acacia compared to the enormous diversity in adult leaf morphology in this genus¹⁰, as well as genetic analysis in Arabidopsis⁶², indicate that the juvenile phase is ecologically and evolutionarily more robust than the adult phase. We suspect that this is because there is a conserved set of phase-specific traits that contributes to the survival of juvenile plants and is therefore under strong selection. Identifying these traits, as well as the traits shared by adult shoots, is an important topic for future research.

Variation in the timing of vegetative phase change can be an important mechanism for the evolution of adaptive traits because the timing of vegetative phase change can vary relative to the timing of floral initiation. This means that variation in the timing of different phase-specific traits, or of vegetative phase change as a whole, can enable plants to adapt to specific environmental conditions without altering the entire life history of a plant. For example, neotenous species of Acacia are found specifically in the southeast and southwest corners of Australia, which have a cooler and wetter climates than the rest of the subcontinent, where most phyllodinous species are found¹⁰. Similarly, precocious ecotypes of Arabidopsis are most often found in regions with significant seasonal variation and range in temperature⁸. In the Azores, neotenous accessions of Cardamine hirsuta are found predominantly in regions with warm, wet summers⁹⁴, which is similar to the climate in which neotenous species of Acacia are found. Interestingly, these neotenous accessions flower earlier than accessions with a relatively short juvenile phase. The vegetative phenotype of these neotenous accessions of C. hirsuta is attributable to a loss-of-function mutation in SPL9.⁹⁴ In Eucalyptus globulus, natural variation in the timing of vegetative phase change was correlated with variation in the abundance of miR156 and mapped to miR156.5.²¹³ These and other results discussed in this review suggest that the miR156/SPL module has been the target of natural selection many times during evolution, and could be a major contributor to morphological and physiological diversity within the plant kingdom.

Concluding Remarks

The timing of vegetative phase change can be modulated by a change in the rate of decline in miR156/miR157 during shoot growth or by an overall increase or decrease in the abundance of these miRNAs. Factors that affect the rate of decline represent components of a developmental clock, the identity of which is still unknown. As discussed here, this clock could be the increase in the photosynthetic capacity of the shoot, given that conditions that reduce this capacity (defoliation, low light intensity, photosynthetic mutations) delay this transition. It is also possible that there are two clocks—one that measures the changing physiology of the shoot, and a second that is independent of this input. Whatever the case, these clocks are expected to affect the abundance of H3K25me3, given that this modification is required for the down-regulation of miR156 genes.⁶⁴ An obvious possibility is that the deposition of this mark is controlled by factors that target chromatin modifying complexes to these genes at a particular time in shoot development. However, this leaves open the question of how the temporal expression of these targeting factors is regulated. We favor the hypothesis that the decline in the transcription of miR156/miR157 genes is the result of a difference in the constitutive rate at which H3K27ac and H3K27me3 or other activating and repressive chromatin modifications are added to these genes. This rate might be mostly dependent on intrinsic differences in the activity of the complexes that deposit these marks, and may have an amplification mechanism (e.g. H3K27me3 promotes the addition of more H3K27me3). Testing this hypothesis will require sophisticated measurements of these marks in the SAM and leaf primordia, combined with mathematical modelling based on these parameters. With the increasing interest in vegetative phase change, answers to these and other questions about this important process may not be long in coming.