The Plant Vascular System: Evolution, Development and FunctionsF

Journal of Integrative Plant Biology 2013, 55 (4): 294–388 

Invited Expert Review 

The Plant Vascular System: Evolution, Development 

and Functions F 

William J. Lucas1∗, Andrew Groover2 , Raffael Lichtenberger3 , Kaori Furuta3 , Shri-Ram Yadav3 , 

Ykä Helariutta3 , Xin-Qiang He4 , Hiroo Fukuda5 , Julie Kang6 , Siobhan M. Brady1 , 

John W. Patrick7 , John Sperry8 , Akiko Yoshida1 , Ana-Flor López-Millán9 , Michael A. Grusak10 and Pradeep Kachroo11 1Department of Plant Biology, College of Biological Sciences, University of California, Davis, CA 95616, USA 

2US Forest Service, Pacific SW Res Stn, Institute for Forest Genetics, Davis, CA 95618, USA 

3Institute of Biotechnology/Department of Biology and Environmental Sciences, University of Helsinki, FIN-00014, Finland 

4State Key Laboratory of Protein and Plant Gene Research, College of Life Sciences, Peking University, Beijing 100871, China 

5Department of Biological Sciences, Graduate School of Science, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, 

Japan 

6Biology Department, University of Northern Iowa, Cedar Falls, IA, USA 

7School of Environmental and Life Sciences, The University of Newcastle, Callaghan, NSW 2308, Australia 

8Biology Department, University of Utah, 257S 1400E Salt Lake City, UT 84112, USA 

9Plant Nutrition Department, Aula Dei Experimental Station (CSIC), P.O. Box 13034, E-50080, Zaragoza, Spain 

10USDA-ARS Children’s Nutrition Research Center, Department of Pediatrics, Baylor College of Medicine, 1100 Bates Street, Houston, TX 

77030, USA 

11Department of Plant Pathology, College of Agriculture, University of Kentucky, Lexington, KY 40546, USA 

All authors contributed equally to the preparation of this manuscript. 

∗ 

Corresponding author 

Tel: +1 530 752 1093; Fax: +1 530 752 5410; E-mail: wjlucas@ucdavis.edu 

F Articles can be viewed online without a subscription. 

Available online on 5 March 2013 at www.jipb.net and www.wileyonlinelibrary.com/journal/jipb 

doi: 10.1111/jipb.12041 

Contents 

I. Introduction 295 

II. Evolution of the Plant Vascular System 295 

III. Phloem Development & Differentiation 300 

IV. Molecular Mechanisms Underlying Xylem Cell Differentiation 307 

V. Spatial & Temporal Regulation of Vascular Patterning 311 

VI. Secondary Vascular Development 318 

VII. Physical and Physiological Constraints on Phloem Transport Function 321 

VIII. Physical & Physiological Constraints on Xylem Function 328 

IX. Long-distance Signaling Through the Phloem 339 

X. Root-to-shoot Signaling 347 

XI. Vascular Transport of Microelement Minerals 351 

XII. Systemic Signaling: Pathogen Resistance 356 

XIII. Future Perspectives 361 

XIV. Acknowledgements 362 

XV. References 362 

C○ 2013 Institute of Botany, Chinese Academy of Sciences

William J. Lucas 

(Corresponding author) 

Abstract 

Insights into Plant Vascular Biology 295 

The emergence of the tracheophyte-based vascular system of land plants 

had major impacts on the evolution of terrestrial biology, in general, 

through its role in facilitating the development of plants with increased 

stature, photosynthetic output, and ability to colonize a greatly expanded 

range of environmental habitats. Recently, considerable progress has 

been made in terms of our understanding of the developmental and 

physiological programs involved in the formation and function of the 

plant vascular system. In this review, we first examine the evolutionary 

events that gave rise to the tracheophytes, followed by analysis of the 

genetic and hormonal networks that cooperate to orchestrate vascular 

development in the gymnosperms and angiosperms. The two essential 

functions performed by the vascular system, namely the delivery of resources (water, essential mineral 

nutrients, sugars and amino acids) to the various plant organs and provision of mechanical support are 

next discussed. Here, we focus on critical questions relating to structural and physiological properties 

controlling the delivery of material through the xylem and phloem. Recent discoveries into the role of 

the vascular system as an effective long-distance communication system are next assessed in terms 

of the coordination of developmental, physiological and defense-related processes, at the whole-plant 

level. A concerted effort has been made to integrate all these new findings into a comprehensive picture 

of the state-of-the-art in the area of plant vascular biology. Finally, areas important for future research 

are highlighted in terms of their likely contribution both to basic knowledge and applications to primary 

industry. 

Keywords: Evolution; vascular development; phloem; xylem; nutrient delivery; long-distance communication; systemic signaling. 

Lucas WJ, Groover A, Lichtenberger R, Furuta K, Yadav SR, Helariutta Y, He XQ, Fukuda H, Kang J, Brady SM, Patrick JW, Sperry J, 

Yoshida A, López-Millán AF, Grusak MA, Kachroo P (2013) The plant vascular system: Evolution, development and functions. J. Integr. Plant 

Biol. 55(4), 294–388. 

Introduction 

The plant vascular system carries out two essential functions, 

namely the delivery of resources (water, essential mineral 

nutrients, sugars and amino acids) to the various plant organs, 

and provision of mechanical support. In addition, the vascular 

system serves as an effective long-distance communication 

system, with the phloem and xylem serving to input information 

relating to abiotic and biotic conditions above and below 

ground, respectively. This combination of resource supply and 

delivery of information, including hormones, peptide hormones, 

proteins and RNA, allows the vascular system to engage in the 

coordination of developmental and physiological processes at 

the whole-plant level. 

Over the past decade, considerable progress has been 

made in terms of our understanding of the developmental and 

physiological programs involved in the formation and function of 

the plant vascular system. In this review, we have made every 

effort to integrate these new findings into a comprehensive 

picture of the state-of-the-art in this important facet of plant 

biology. We also highlight potential areas important for future 

research in terms of their likely contribution both to basic 

knowledge and applications to primary industry. 

Evolution of the Plant Vascular System 

Why the need for a vasculature system? 

For plants as photosynthetic autotrophs, the evolutionary step 

from uni- to multi-cellularity conferred an important selective 

advantage in terms of division of labor; i.e., functional specialization 

of tissues/organs to more effectively extract, and 

compete for, essential resources in aquatic and terrestrial 

environments. Successful colonization of terrestrial environments, 

by plants, depended upon positioning of organs in both 

aerial and soil environments to meet their autotrophic requirements. 

For example, for photosynthetic efficiency, sufficient 

levels of light only co-occur with a supply of CO2 in aerial

296 Journal of Integrative Plant Biology Vol. 55 No. 4 2013 

environments, whereas water and mineral requirements are 

primarily acquired from soil environments. Thus, aerial and soil 

organs of early land plants were nutritionally interdependent 

and, consequently, there was intense selection pressure for the 

evolution of an inter-organ transport system to allow access to 

the complete spectrum of essential resources for cell growth 

and maintenance. 

An important feature of early multicellular plants was the 

acquisition of plasmodesmata (PD), whose cytoplasmic channels 

established a symplasmic continuum throughout the body 

of the plant (Lucas et al. 1993). This symplasm allowed for 

the exchange of nutrients between the different plant organs. 

However, this symplasmic route, in which intracellular cytoplasmic 

streaming is arranged in series with intercellular diffusion 

through PD, is effective only over rather short distances. For 

example, a PD-mediated sucrose flux of 2 × 10 −4 mol m −2 s −1 

into heterotrophic cells would satisfy their metabolic demand. 

Maximum reported permeability coefficients for sucrose diffusion 

through PD are on the order of 6 × 10 −6 ms −1 (Fisher 

and Wang 1995). Using this value, diffusion theory predicts that 

a significant sucrose concentration drop would be required, 

across each adjoining cell wall interface, to sustain this flux 

of sucrose from the autotrophic (photosynthetic) cells into the 

heterotrophic (water and mineral nutrient acquiring) cells. Thus, 

the path length would be limited to only a few cells, arranged in 

series, and the size of the organism would be limited to a few 

millimeters. 

In order for multicellular autotrophs to overcome these 

diffusion-imposed size constraints, a strong selection pressure 

existed to evolve an axially-arranged tissue system, located 

throughout the plant body, with a greatly increased conductivity 

for intercellular transport. The solution to this problem began 

over 470 Mya and, in combination with prevailing global climate 

change, including dramatic changes in atmospheric CO2 levels, 

gave rise to the development of the cuticle and stomata, 

important adaptations that both reduced tissue dehydration 

and increased the capacity for exchange of CO2, thereby 

enhancing the rates of photosynthesis (Franks and Brodribb 

2005; Ruszala et al. 2011; but see Duckett et al. 2009). 

Following acquisition of these two traits, early land plants 

evolved cells specialized for long-distance transport of food 

and water (Ligrone et al. 2000, 2012; Raven 2003; van Bel 

2003; Pittermann 2010). Irrespective of plant group, these cells 

became arranged end-to-end in longitudinal files having a simplified 

cytoplasm and modified end walls designed to increase 

their intra- and intercellular conductivities, respectively. 

In land plants, the degree of cellular modifications of transport 

cells increases from the bryophytes (pretracheophytes—also 

termed non-vascular plants—the liverworts, mosses and hornworts), 

to the early tracheophytes, the vascular cryptogams (lycophytes 

and pterophytes), on through to seed plants (Ligrone 

et al. 2000, 2012; Raven 2003; van Bel 2003). These cell 

specializations neatly scale with maximal sizes attained by 

each group of land plants. Interestingly, impacts of enhancing 

conductivities of cells transporting sugars converges with a 

greater influence imposed by evolving water conducting cells 

to sustain hydration of aerial photosynthetic tissues. 

Evolutionary origins and diversification of food 

and water transport systems 

Studies based on fossil records and extant (living) bryophytes 

have established that developmental programs evolved to form 

specialized water and nutrient conducting tissues. Based on 

the fossil record, early pretracheophyte land plants appeared 

to have developed simple water-conducting conduits having 

smooth walls with small pores, likely derived from the presence 

of PD. Similar structures are present, for example, in some of 

the mosses, the most ancient being termed water-conducting 

cells (WCCs) and the more advanced being the hydroids of the 

peristomate mosses (Mishler and Churchill 1984; Kenrick and 

Crane 1997; Ligrone et al. 2012). 

Hydroids often form a central strand in the gametophyte 

stem/sporophyte seta in the mosses (Figure 1A, B). During 

their development, these hydroid cells undergo various structural 

modifications to the cell wall and are dead at maturity 

(Figure 1C, D). Although in some cases the hydroid wall may 

become thickened, these are considered to be primary in 

nature and lack lignin. However, recent studies have indicated 

that bryophyte cell walls contain lignin-related compounds, 

but these do not impart mechanical strengthening properties 

(Ligrone et al. 2012). Although this absence of mechanical 

strength served as an impediment to an increase in body size, 

it allowed for hydroid collapse during tissue desiccation, and 

rapid rehydration following a resupply of water (Figure 1E, F), a 

feature that likely minimized cavitation of these WCCs (Ligrone 

et al. 2012) (see also later section). This trait may also have 

allowed peristomate mosses to expand into dryer habitats. 

The evolution of hydroids could have involved modification of 

existing WCCs. However, based on the distribution of WCCs 

in the early land plants (Figure 2), it seems equally probable 

that they arose through an independent developmental 

pathway after the loss of perforate WCCs (Ligrone et al. 

2012). 

The fossil record contains less information on the evolution 

of specialized food-conducting cells (FCCs), due in large 

part to their less robust characteristics that limited effective 

preservation. However, insights can be gained from studies 

on extant bryophyte species. As with WCCs, early FCCs were 

represented by files of aligned elongated cells in which the 

cytoplasmic contents underwent a series of positional and 

structural modifications (Figure 3). Here, we will use moss as an 

example; in some species (members of the order Polytrichales),

Figure 1. Water-conducting cells of early non-vascular (pretracheophyte) 

land plants. 

(A) Light micrograph of a leafy stem from the moss Plagiomnium 

undulatum illustrating the prominent central strand of hydroid cells 

(h) surrounded by parenchyma cells (p). 

(B–D) Transmission electron micrographs illustrating hydroids having 

unevenly thickened walls in the leaf shoot of the moss Polytrichum 

juniperinum (B), a differentiating hydroid (C) and mature 

hydroids (D) in the leafy shoot of Polytrichum formosum. 

(E, F) Transverse sections of moss hydroids in the hydrated (E) 

and dehydrated condition (F); note that in the presence of water, 

dehydrated hydroids become rehydrated and functional. Images 

(A–D) reproduced from Ligrone et al. (2000), with permission of 

The Royal Society London; (E, F) reproduced from Ligrone et al. 

(2012), with permission of Oxford University Press. Scale bars: 

50 µm in(A), 5µm in(B–D) and 5 µm in(E), common to (F). 

the FCCs gave rise to a group of more specialized cells, termed 

leptoids and associated specialized parenchyma cells. During 

development, leptoids undergo a series of cytological changes, 

including cytoplasmic polarization and microtubule-associated 


alignment of plastids, mitochondria and the endoplasmic reticulum 

(ER), in a longitudinal pattern. At maturity, the FCCs of 

the bryophytes generally lack a large central vacuole and, in 

some species, there is partial degradation of the nucleus. In 

addition, the end walls of cells within these files of aligned FCCs 

develop a high density of PD (Figure 4), presumably to optimize 

symplasmic continuity for cell-to-cell diffusion of photosynthate 

(Ligrone et al. 2000, 2012; Raven 2003). Finally, FCCs/leptoids 

often develop in close proximity to WCCs/hydroids; in 

some species, the WCCs/hydroids are centrally located 

in the tissue/stem being ensheathed by FCCs/leptoids 

(Figure 3E). 

As with hydroids, the evolutionary events leading to the 

development of FCCs, and leptoids in particular, appear to 

have been driven by the necessity to withstand periods in 

which the early land plants underwent desiccation. Insight sinto 

the presence and importance of such traits were offered by 

recent physiological and anatomical studies performed on the 

desiccation-resistant moss Polytrichum formosum. Here, the 

unique resilience of the lepoids to an imposed dehydration was 

shown to be associated with the unique role of microtubules 

in control over the special cytological features of these FCCs 

(Pressel et al. 2006). The properties of cavitation-resistant 

hydroids and desiccation-tolerant leptoids would likely have 

had an important impact on these mosses in terms of the ability 

to penetrate into diverse ecological niches. 

Emergence of xylem with lignified tracheids 

and vessels 

As indicated in Figure 2, xylem tissues may well have evolved 

independently from WCCs/hydroids. Although hydroids have a 

number of similar features to the early tracheary elements, 

including functioning after death, there are many important 

differences. Perhaps the most critical was the acquisition 

of a developmental program for the deposition of patterned 

secondary cell wall material. Of equal importance was the 

development of lignin and its deposition within the secondary 

wall of tracheary cells. Collectively, these evolutionary events 

imparted biomechanical support and compressive strength, 

with an ability to withstand tracheid collapse when the water column 

was placed under tension (see later section). Acquisition of 

biomechanical strength afforded the opportunity for an increase 

in plant height, with the benefit of enhanced competition for 

sunlight. 

A further defining feature of the early vascular plants was that 

their tracheary (water conducting) elements had pits of varying 

architecture that spanned the secondary wall. In contrast to 

the WCCs of the bryophytes, the formation of these pits is not 

dependent upon the dissolution of PD (Barnett 1982; Lachaud 

and Maurousset 1996), and in the early tracheary element, the 

tracheid (Edwards et al. 1992), the primary cell wall remains


Figure 2. Cladogram illustrating the distribution of water-conducting cells (WCCs) in early land plants. 

Note that hydroids in the peristomate mosses lack perforate cell walls. Reproduced from Ligrone et al. (2012), with permission of Oxford 

University Press. 

imperforate. However, in the advanced form, the vessel element 

or vessel member, the primary wall is removed in discrete 

regions between adjacent members, thereby giving rise to a 

perforation plate. This evolutionary adaptation allows water to 

flow through many mature vessel members that collectively 

form a vessel, unimpeded by the primary cell wall; i.e., the 

perforation plate reduces the overall resistance to water flow 

through vessels. 

Evolutionary relationship between FCCs and early 

tracheophyte sieve elements 

The cytological features of FCCs are widespread in the 

bryophytes and many are also present in the phloem sieve 

elements of the lycophytes, pterophytes and gymnosperms 

(Esau et al. 1953) (Table 1). It is also noteworthy that the ER is 

present in PD located in the adjoining transverse walls between 

FCCs, leptoids and the sieve elements of ferns (Evert et al. 

1989) and conifers (Schulz 1992). Furthermore, both leptoids 

and early sieve elements, termed sieve cells, have supporting 

parenchyma cells. These features, held in common between 

the more advanced FCCs and the phloem sieve elements of 

the early tracheophytes, raise the possibility of a developmental 

program having components shared between these nutrient 

delivery systems of the plant kingdom. 

Evolution of molecular mechanisms regulating 

vascular development 

Significant progress has been made in elucidating the molecular 

mechanisms regulating vascular development. In most 

cases, a modest number of angiosperm model species have 

been the focus of molecular-genetic and genomic analysis 

of vascular development. At present, individual genes 

regulating specific aspects of vascular development have 

been characterized in detail. In addition, models of how 

vascular tissues are initiated, patterned, balance proliferation 

and differentiation, and acquire polarity have been 

developed. 

Vascular development is currently being modeled at new 

levels of complexity in Arabidopsis and Populus, using computational 

and network biology approaches that make use 

of extensive genomic gene expression and gene regulation 

datasets. While incomplete, new models representing important 

phylogentic positions in land plant evolution are also being 

developed, and will provide important insights into the origins 

and diversification of mechanisms regulating vascular development. 

Importantly, many of the key gene families that regulate 

vascular development predate tracheophytes. Thus, one major 

challenge for understanding the evolution of vascular development 

will be to determine the evolutionary processes by which

Figure 3. Cytological details of moss food-conducting cells. 

(A) Cytoplasmic polarity in a leafy stem of Plagiomnium undulatum; 

most of the organelles are in the top end of the lower cell. 

(B) Sphorophyte seta of Mnium hornum showing the longitudinal 

alignment of elongated plastids (p) and the highly elongated nucleus 

(n). 

(C, D) Longitudinal arrays of microtubules associated with tubules 

and vesicles in a leafy stem of Plagiomnium undulatum (C) and 

Polytrichum juniperinum (D). 

(E) Transverse section of leptoids and adjacent hydroids (h) in a 

stem of Polytrichum commune. 

Scale bars: 4 µm in(A), 2µm in(B, E), 0.5µm in(C, D). 

Reproduced from Ligrone et al. (2000), with permission of The Royal 

Society London. 

regulatory genes and modules were duplicated, modified, or 

directly co-opted to function in vascular development (Pires and 

Dolan 2012). Even more challenging will be determining the 

evolutionary steps underlying the many biochemical processes 

required for the production of vascular tissues and lignified 

secondary cell walls. 


Figure 4. Abundant plasmodesmata in the trumpet-shaped end 

walls between food-conducting cells in the moss Sphagnum 

cuspiatum. 

Scale bar: 10 µm. Reproduced from Ligrone et al. (2000), with 

permission of The Royal Society London. 

Table 1. Comparison of cytological features present in moss 

food-conducting cells and sieve cells in ferns and conifers 

Similaritiesa Differences (in sieve cells) 

Absence of vacuoles No cytoplasmic polarization 

Nacreous wallsb Apparent lack of polyribosomes 

Nuclear degenerationb Presence of endoplasmic 

reticulum (ER) within 

plasmodesmata (PD) 

Callose associated with PDc a Modified after Ligrone et al. (2000). 

b Restricted to the Polytrichales in mosses. 

c Restricted to the Polytrichales in mosses, absent in some lower 

tracheopytes. 

Auxin is an evolutionarily ancient regulator of vascular 

development 

In the following sections we present some examples of the 

genes and mechanisms regulating specific aspects of vascular 

development. This is not a complete review of the literature, 

but rather we aim to highlight some of the molecular-genetic 

models of vascular development. We begin with the enigmatic 

plant hormone auxin, which has been known to play 

fundamental roles in vascular development for decades, but 

only recently have insights been gleaned at the moleculargenetic 

level as to how it exerts its many influences on vascular 

development. To understand the myriad of ways that auxin 

influences plant development, it is necessary to understand 

its synthesis, conjugation, transport, perception, and effects 

on gene expression. Fundamental insights into all of these 

processes have been gained, and have been summarized


in recent reviews. Importantly, auxin can apparently be synthesized 

by all plants (Johri 2008; Lau et al. 2009) in which 

it plays various roles in promoting growth, and has thus 

been recruited to participate in vascular development in the 

tracheopytes. Here, we consider one specific role for auxin 

during vascular development: how auxin transport proteins act 

during the establishment and propagation of interconnected 

vascular strands. 

Vascular strands consist of interconnected files of cells. It 

is critical that vascular strands be properly spaced and patterned 

within tissues, and that functional cell types (e.g. water 

conducting tracheary elements) coordinate differentiation to 

produce interconnected conduits for transport. Auxin has long 

been known to play fundamental roles in both the induction of 

vascular strands and in the differentiation of vascular cell types 

(Sachs 1991). How auxin is transported through tissues has 

been recognized as a primary factor in determining the location 

and propagation (or canalization) of vascular strands. Major 

insights into how auxin transport is regulated have been gained 

through the identification and characterization of PIN-FORMED 

(PIN) proteins, which include plasma membrane spanning 

auxin efflux carriers. PIN proteins act through asymmetric 

subcellular localization to direct routes of auxin flow through 

cells and tissues (Petráˇsek et al. 2006, Petráˇsek and Friml 

2009). 

Importantly, PIN proteins are found in all land plants (Krecek 

et al. 2009) including the pretracheophytes, and polar auxin 

transport (PAT) already existed in the Charophyta (Boot et al. 

2012), suggesting that ancestral functions of PIN proteins were 

not in vascular development, per se. The fossil record also 

provides insights into the role of auxin transport in the evolution 

of vascular tissues. Routes of auxin transport can be indirectly 

inferred from anatomical changes in extant angiosperms and 

gymnosperms, in which circular patterns of tracheary element 

develop in secondary xylem above branches, which impede 

auxin transport. Amazingly, fossils of wood from 375-millionyear-old 

Archaeopteris (a progymnosperm) also show this 

pattern (Rothwell and Lev-Yadun 2005). 

A general expectation is that the auxin-related mechanisms 

regulating vascular differentiation are shared (i.e. are homologous) 

among vascular plants, but this remains to be verified 

through functional studies in all the major vascular plant lineages. 

Perhaps more intriguing will be the characterization of 

ancestral auxin-related mechanisms in non-vascular plants and 

the determination of the evolutionary steps through which they 

were co-opted and modified during vascular plant evolution. 

CLASS III HD-ZIP transcription factors 

Gene transcription is a major mechanism for regulating 

vascular development. One increasingly well characterized 

transcriptional module is defined by the Class III homeodomain- 

leucine zipper (HD-ZIP) transcription factors. The genes encoding 

these transcription factors are evolutionarily ancient, and 

are found in all land plants (Floyd et al. 2006). Interestingly, 

transcript levels of Class III HD ZIPs are negatively regulated by 

miRNAs which are also highly conserved (Floyd and Bowman 

2004). The Class III HD-ZIP gene family expanded and diversified 

in land plant lineages, acquiring new expression patterns 

and functions along the way. Indeed, the functions of Class 

III HD-ZIPs in regulating vascular tissues are undoubtedly 

derived, since Class III HD-ZIPs predate the appearance of 

vasculature in land plant evolution. 

In Arabidopsis, the Class III HD-ZIP gene family is comprised 

of five genes, REVOLUTA (REV), PHABULOSA (PHB), 

PHAVOLUTA (PHV), ATHB8, and CORONA/ATHB15. Phylogenetic 

and functional relationships among them support the 

conclusion that ATHB8 and ATHB15, and REV, PHAB, and 

PHAV represent subclades (Prigge et al. 2005, 2006; Floyd 

et al. 2006). Functional relationships among the Class III HD- 

ZIPs are complex, and different family members have been 

implicated in shoot apical meristem formation, lateral organ 

initiation, embryo patterning, and leaf polarity (Floyd et al. 

2006). However, all five Arabidopsis Class III HD-ZIPs have 

been implicated in some aspect of vascular development. The 

role for the Class III HD-ZIPs in regulating vascular polarity will 

be discussed in depth later in the review. 

The interplay of transcription and hormones is just one of 

the many areas ripe for exploration in terms of the evolution 

and development of vascular biology. Powerful new tools are 

available or quickly being developed that will result in dramatic 

changes to both the scope and level of complexity that can 

be addressed in future studies. For example, new sequencing 

technologies now allow for comprehensive cataloguing of gene 

expression in vascular tissues from virtually any species. These 

and related genomic technologies are also being used to 

provide massive datasets for new computational approaches, 

including network biology, which can model the complex interactions 

of genes that together regulate fundamental features 

of vascular development. Importantly, these new technologies 

must be integrated within the framework of paleobotany, plant 

anatomy, and plant physiology to provide meaningful models of 

the evolutionary steps that occurred, at the molecular-genetic 

level, to provide the diversity of vascular biology that we see in 

extant plants. 

Phloem Development & Differentiation 

Recently, several novel regulatory mechanisms that control the 

specification of vascular patterning and differentiation have 

been uncovered. Through the use of novel genomics and 

molecular techniques in several model plant systems, such as 

Arabidopsis, Populus and Zinnia, new insights have become

available regarding the regulation of vascular development. 

The importance of signaling in the control of vascular morphogenesis 

has become increasingly apparent. The primary agents 

involved include well-known phytohormones such as auxin, 

cytokinin and brassinosteroids, as well as other small regulatory 

molecules. This level of understanding also involves the 

transporters and receptors of these factors. There is increasing 

evidence for the notion that xylem and phloem development 

are highly coordinated. While many of the factors which will be 

described in this section of the review act cell-autonomously, 

the emerging importance of mobile regulatory factors will also 

be highlighted. 

Embryonic provascular and shoot vascular 

development 

During embryogenesis, the progenitor cells that will eventually 

become the vascular tissues are first established as undifferentiated 

procambial tissues. Surrounded by the epidermal and 

ground tissue layers, this procambial tissue forms the innermost 

domain of the plant embryo. By the late globular embryonic 

stage, the four procambium cells have undergone periclinal 

divisions to generate the future pericycle and the vascular 

primordium. The complete root promeristem with all initials and 

derived cell types is contained already in the early torpedo 

stage embryo (Scheres et al. 1995). Asymmetric cell divisions 

within the vascular primordium go on to establish the number 

of vascular initials present in the seedling root meristem. 

In the aerial parts of a seedling, the protovascular elements 

are first specified and start to differentiate in the cotyledons and, 

only subsequently, in the axis (Bauby et al. 2007). Protophloem 

differentiation can first be observed in the midvein by the 

typical cell elongation, followed by extension to the distal 

loops and cotyledonary node. Before vasculature differentiation, 

continuous procambial strands can already be observed 

(Figure 5). The main agent in the establishment of this pattern 

is the phytohormone auxin. The accumulation of auxin, through 

polar transport mechanisms, as shown by the synthetic auxin 

reporter DR5 and the auxin-induced pre-procambial marker 

AtHB8, precedes the formation of vascular strands in leaves 

(Scarpella et al. 2004). Facilitating this highly localized auxin 

accumulation is the auxin transporter PIN1, which channels 

auxin to the provascular regions. PIN1 is already expressed 

before procambium formation. 

After the initial differentiation, the vasculature develops in 

bundles (Esau 1969). These bundles have a very distinct 

radial pattern of abaxial phloem and adaxial xylem divided by 

an active procambium (Figure 6). The radial patterning of the 

vascular bundles is already established during embryogenesis 

by the main factors of radial patterning, KANADI (KAN) and the 

Class III HD-ZIPs, PHB, PHV, REV and CORONA/ATHB15 

(McConnell et al. 2001; Emery et al. 2003). Auxin also plays 


a role in this determination of the radial vascular patterning 

(Izakhi and Bowman 2007). PIN1 localization is affected in kan 

mutants, showing the integration of the auxin transport pathway 

and KAN signaling. The triple mutant phb phv rev has radialized 

as well as abaxialized leaves and vascular bundles. In contrast, 

gain-of-function Class III HD-ZIP mutants with faulty microRNA 

(miRNA) regulation and kan1 kan2 kan3 have radialized and 

adaxialized leaves and bundles. In addition, the Class III HD- 

ZIP genes also appear to regulate vascular tissue proliferation. 

As will be described in more detail later, the phloem translocation 

stream, or phloem sap, contains not only photosynthate 

but also a wide array of macromolecules, such as mRNA, 

small RNAs and proteins. Both in animals and plants, small 

RNAs have already been identified as important regulatory 

factors controlling cell fate. A bidirectional cell-to-cell communication 

network involving the mobile transcription factor 

SHORTROOT (SHR) and microRNA165/166 species specifies 

the radial position of two types of xylem vessels in Arabidopsis 

roots (Carlsbecker et al. 2010; Miyashima et al. 2011). Since 

microRNA165/166 is a factor restricting PHB activity, it also 

regulates phloem development. 

Recent studies have shown that CALLOSE SYNTHASE 

3 (CALS3), a membrane-bound enzyme which synthesizes 

callose, a β-1,3-glucan (Verma and Hong 2001; Colombani 

et al. 2004), appears to be involved in regulating the cell-to-cell 

movement of microRNA165/166 (Vatén et al. 2011). CALS3 is 

expressed both in phloem and meristematic tissues. Gain-offunction 

mutations in CALS3 result in increased accumulation 

of PD callose, a decrease in the PD aperture, multiple defects 

in root development, and reduced intercellular trafficking of 

various molecules. Using an inducible expression system for a 

modified version of CALS3 (CALS3m), Vatén et al. (2011) were 

able to show that increased callose deposition inhibited SHR 

and microRNA165 movement between the stele and the endodermis. 

This interesting result suggested that regulated callose 

biosynthesis, at the PD level, may be essential for control over 

cell-to-cell communication and cell fate determination. 

Root vascular development 

In the root, protophloem initially differentiates from an independent 

differentiation locus in the upper hypocotyl; only later 

does protophloem differentiation from the root apical meristem 

begin. As the root grows, the cellular pattern is established and 

maintained by the self-renewal of pluripotent root meristem 

cells. Different cell identities are initiated from the stem cells 

around the quiescent center (QC): the provascular initials of 

the stele, the cortex/endodermal initials, the epidermal/lateral 

root initials, and the columella initials. In the Arabidopsis root, 

a central vascular cylinder (consisting of xylem, phloem and 

procambium) is surrounded by radially symmetric layers of 

pericycle, endodermis, cortex and epidermal cells. In this root


Figure 5. Schematic of the primary phloem organization in Arabidopsis shoot and root. 

(A) Longitudinal section through the shoot apex, shoot and root. 

(B) Cross section of an early developing leaf showing the preprocambial bundles which precede the vascular bundles. 

(C) Cross section of a leaf showing established vascular bundles where primary phloem and xylem differentiate asymmetrically from a 

separating layer of procambium. 

(D) Cross section of the stem showing primary vascular bundles. 

(E) Cross section of the root showing the primary vascular patterning with two phloem poles consisting of sieve elements and companion 

cells flanking the xylem axis. 

(F) Cross section of the root tip showing two poles of protophloem sieve elements flanking the xylem axis. 

system, the vascular tissue is comprised of a central axis of 

water-conductive xylem tissue that is flanked by two poles of 

photoassimilate-conductive phloem tissue. 

In the root apical meristem, the phloem cell lineages arise 

from two domains of initials through asymmetric cell divisions 

(Mähönen et al. 2000). Periclinal divisions establish companion 

cells (CCs) and tangential divisions establish sieve elements 

(SEs). This asymmetry allows these initials to give rise to 

multiple cell lineages with different fates; in addition to the 

phloem lineage, they also precede undifferented procambial 

cell lineages. As opposed to the invariant pattern of cell 

lineages in the endodermis and outer layers, the number 

and exact pattern of these procambial divisions vary between 

individual seedlings. 

Mähönen et al. (2000) have described these initial asymmetric 

cell divisions in great detail through sequential 

cross-sections of the region immediately above the quiescent 

center, allowing the precise determination of the first true 

phloem domains. At first, although cell divisions and early xylem 

specification can be observed (though not yet the complete


Figure 6. Vascular patterning is regulated by KANADI and Class III HD-ZIP genes and the phytohormones auxin and cytokinin. 

(A) In shoot vascular bundles, the default radial pattern has phloem located abaxially and xylem adaxially. 

(B) In the phb phv rev mutant, phloem surrounds xylem. 

(C) Conversely, in Class III HD-ZIP gain-of-function mutants and kan1 kan2 kan3, xylem surrounds phloem. 

(D) Class III HD-ZIP genes are regulated by miR165/166 and interact with auxin and brassinosteroids. 

(E) In the root, auxin is restricted to the xylem axis by the presence of cytokinin. 

(F) In mutants with defective cytokinin signaling such as wol, auxin is abundant throughout the stele, leading to ubiquitous protoxylem 

differentiation and loss of phloem identity. 

xylem axis), phloem identity is not observable directly above 

the QC. The first newly formed cell walls associated with 

phloem development are only visible 27 µm above the QC. 

At a distance of 69 µm, the first protophloem SEs are clearly 

present. 

Hormonal balance determines the development 

of vascular poles in the root 

Cytokinin, an essential phytohormone for development in the 

root, is required for vascular patterning and the differentiation 

of all cell types except the protoxylem. Recently, it has been 

shown that the root vascular pattern is defined by a mutually 

inhibitory interaction between cytokinin and auxin (Bishopp 

et al. 2011a, 2011b). If cytokinin signaling is disturbed, as 

in the WOODEN LEG mutant, wol or the triple cytokinin 

receptor mutant ahk2 ahk3 ahk4 (ARABIDOPSIS HISTIDINE 

KINASE), or if cytokinin levels are reduced, as is found in 

transgenic plants overexpressing CYTOKININ OXIDASE, the 

effect is always an increased number of protoxylem cell files 

and the loss of other cell types in the root vasculature. The 

domain of cytokinin activity is restricted by the action of 

the cytokinin signaling inhibitor, ARABIDOPSIS HISTIDINE 

PHOSPHOTRANSFER PROTEIN 6 (AHP6). Only through this 

mechanism does protoxylem differentiation occur in a spatially 

specific manner, allowing for the proper development of the 

phloem cell types.


Phloem differentiation 

Sieve elements comprise the main conductive tissue of the 

phloem. Like the CCs, they originate from phloem precursor 

cells in the procambium. However, very early in primary phloem 

development, they undergo dramatic changes in their morphology. 

As the SEs mature, they experience extensive degradation 

of their organelles. The nucleus, vacuoles, rough endoplasmic 

reticulum (ER) and Golgi are degraded in a process which 

has not yet been characterized at the molecular level. This 

reduction in cellular contents establishes an effective transport 

route through the sieve tubes. However, the SEs still remain 

living, as they retain a plasma membrane and a reduced 

number of other organelles, such as smooth ER, plastids and 

mitochondria. The residual ER is localized near the PD which 

interconnect the SEs to their neighboring CCs. 

The cell walls of the SEs also undergo drastic changes in 

structure. The first observable process is an increase in callose 

that is deposited in platelet form around the PD of the SEs, 

replacing the already present cellulose. The cell walls which 

form the interface to adjoining SEs contain a high density 

of these callose-ensheathed PD. As these SEs mature, both 

these callose deposits and the middle lamella in these regions 

of the cell wall are removed, thereby forming a sieve plate 

with enlarged pores (Lucas et al. 1993). The lateral cell walls 

of SEs also develop specialized areas of PD-derived pores, 

which are called lateral sieve areas. Recent studies have 

shown that CALS3 and CALS7 are involved in depositing PDcallose 

during this developmental process (Vatén et al. 2011; 

Xie et al. 2011). The newly formed sieve plates, in combination 

with the lateral sieve areas, enable each individual SE to 

become a component of an integrated sieve tube system that 

can facilitate effective fluid transport by bulk flow. It is also 

noteworthy that these pores increase considerably in size as 

tissues age, thus increasing the transport potential of the more 

mature vasculature (Truernit et al. 2008). 

The survival and differentiation of SEs depends on a close 

association with their neighboring CCs, a specialized type of 

parenchyma cell. The cytoplasm of the CC is unusually dense, 

due in part to an increased number of plastids, mitochondria 

and free ribosomes (Cronshaw 1981). The CCs are connected 

to their adjacent SEs by numerous branched PD. Through 

these connections, the enucleate SEs are supplied with energy, 

assimilates and macromolecular compounds, such as proteins 

and RNA (Raven 1991; Lough and Lucas 2006). The size 

exclusion limit of these PD connections usually lies between 

10 and 40 kDa (Kempers and van Bel 1997), giving credence 

to the concept of protein transport from CCs to SEs. 

The morphological and physiological uniqueness of the 

phloem cell types described above is also a result of specific 

gene expression patterns, as shown by recent transcriptome 

studies (Lee et al. 2006; Brady et al. 2007, 2011). These 

transcriptional programs are exquisitely controlled in space 

and time. To understand how these unique cell identities 

are acquired, a deeper understanding of these programs is 

absolutely essential. Microarray analyses of a high-resolution 

set of developmental time points, and a comprehensive set of 

cell types within the root, has resulted in the most detailed root 

expression map to date. 

Numerous distinct expression patterns have been identified 

through these analyses, several of which are specific to SEs 

or to SEs and CCs together. More than a thousand genes 

have been identified as having phloem-specific expression, 

highlighting the phloem as a highly specialized tissue within 

the stele. The data from these microarray studies has been 

made publicly available in the AREX LITE: The Arabidopsis 

Gene Expression Database (arexdb.org), providing an invaluable 

resource for future studies on phloem function. 

Bauby et al. (2007) identified several phloem markers, which 

they named Phloem Differentiation 1–5 (PD1-PD5) by screening 

the Versailles collection of gene trap mutants for plant lines 

expressing the uidA reporter gene in immature vascular tissues. 

PD1-4 were restricted to protophloem cells, as determined by 

their cell shape. However, PD5 was expressed in both protophloem 

and metaphloem cells. Using these markers, these 

authors could track the onset of phloem development directly 

after embryogenesis. PD4 is expressed in the tips of leaf 

primordia some 3 days after germination (dag). Spreading from 

there, by 7 dag, PD4 has already traced out the entire future 

leaf vasculature. The first expression of PD1 and PD3 was 

detected 3 dag in the proximal protophloem of leaf primordia. 

These results establish that PD1 and PD3 are expressed 

during the differentiation of protophloem. PD4 and PD5 gene 

reporter-based expression was also detected in the location 

of the midveins and higher order veins before procambium 

differentiation, thereby defining the pre-patterning of the future 

veins. 

Regulation of phloem differentiation 

Currently, only two factors are known which specify phloem 

identity: ALTERED PHLOEM DEVELOPMENT (APL) and OC- 

TOPUS (OPS). APL is a MYB coiled-coil transcription factor 

essential for the proper differentiation of both SEs and CCs 

(Bonke et al. 2003). Additionally, APL contributes to the spatial 

limiting of xylem differentiation, is expressed both in SEs and 

CCs, and has been shown to be nuclear localized. Loss of 

APL function has an extraordinary effect on the phloem, as 

can be observed in the loss-of-function mutant apl1 (Figure 7). 

This mutant is seedling-lethal and results in short-rooted plants. 

Neither CCs nor SEs can be detected in cross-sections of 

apl1 plants. Furthermore, phloem-specific reporters, such as 

the CC-specific sucrose transporter, SUC2, or the proto SE 

reporter, J0701, cease to be expressed entirely in these mutant

Figure 7. Model of OCTOPUS (OPS) and ALTERED PHLOEM DEVELOPMENT (APL) action. 


(A) OPS is located at the apical plasma membrane in the procambium and phloem lineage. OPS interprets vascular signals for phloem 

differentiation, such as APL. 

(B) In the ops mutant, phloem differentiation is delayed. Procambial cell number is increased and gaps of undifferentiated cells are visible in 

the protophloem strand. 

(C) In the apl mutant, the initiation of phloem differentiation is largely unperturbed, however proper protophloem fails to emerge. In their 

place, protophloem/protoxylem hybrid cells appear. 

plants. This dramatic effect is restricted to the phloem poles; 

other cell types appear similar to wild-type (WT) plants. 

As mentioned above, asymmetrical cell divisions establish 

the phloem poles in wild-type plants; periclinal divisions establish 

CCs and tangential divisions establish SEs. In the apl 

mutant, these cell divisions are often delayed, but they still 

take place, so APL does not appear to be required for these 

asymmetric cell divisions. However, since the subsequent 

differentiation of SEs cannot be observed, it can be concluded 

that APL is responsible for phloem differentiation, rather than 

the establishment of the phloem cell lineage. 

It has also been proposed that APL acts as an inhibitor to 

xylem differentiation. In the apl mutant, ectopic xylem strands 

are seen in the place of the phloem poles. When APL is 

expressed ectopically in vascular bundles, xylem formation is 

inhibited. Recently, novel imaging techniques were employed 

to analyze the apl mutant in more detail (Truernit et al. 2008). 

With this increased resolution, it was discovered that protophloem 

differentiation proceeds normally in this mutant until 

2 dag. At this time, cells in the protophloem position display the 

normal characteristic shape and cell wall thickening. However, 

after this period, the previously described acquisition of xylem 

characteristics was observed, although the cells in place of 

the SEs still formed sieve plates. This finding suggests that 

these cells can be classified as hybrids between phloem and 

xylem (Truernit et al. 2008). Thus, APL is absolutely required 

for the later stages of phloem development, although it is 

not the earliest factor acting in this process. The identity of 

such a factor, or factors, has yet to be determined. Additional 

support for the presence of such factor(s) was provided by Lee 

et al. (2006), who pointed out that there are numerous phloem 

markers with earlier SE expression than APL.


In addition to APL, OPS has also been identified as a gene 

related to phloem cell differentiation; this gene is required 

for phloem continuity during phloem development (Truernit 

et al. 2012). Interestingly, OPS was first reported based on 

its vascular expression pattern (Nagawa et al. 2006). Further 

detailed analysis revealed that its expression is initially in the 

provascular cells at the heart stage of embryo development, 

and it subsequently becomes restricted to the phloem lineage 

cells following phloem cell-type specification (Bauby et al. 2007; 

Truernit et al. 2012). Unlike APL, OPS expression can be 

observed in the phloem and procambium initials near the QC. 

Vascular patterning of the cotyledons, in the mature embryo 

of the ops mutant, is delayed, and the number of completed 

vascular loops in the developing cotyledon is reduced (Truenit 

et al. 2012). In contrast, the progression of vascular patterning 

is accelerated by OPS overexpression in the cotyledons of 

mature embryos, and the number of completed vascular loops 

in the developing cotyledon is increased. This suggests that 

OPS is involved in promoting the progression of vascular 

patterning. Furthermore, Truernit et al. (2012) found that, in ops 

hypocotyls and roots, the phloem SE cell file was interrupted 

by undifferentiated SEs, which failed to undergo an increase 

in cell wall thickness, callose deposition, or nuclear degradation; 

these cells failed to acquire SE-specific PD1 marker 

expression (Truernit et al. 2012). These cellular differentiation 

defects caused inefficient phloem transport in the root. These 

phenotypes indicated that OPS was required for continuous 

phloem development. 

OPS encodes a membrane-associated protein (Benschop 

et al. 2007) specific to higher plants (Nagawa et al. 2006). The 

function of OPS is currently unknown; no functional domain 

has been identified in this protein. However, a functionally 

complementing OPS-GFP protein was located at the apical 

end of the SEs (Truernit et al. 2012). Inductive cell-to-cell communication 

from differentiated to undifferentiated neighboring 

cells is known to occur during xylem differentiation; XYLO- 

GEN is a polar-localized proteoglycan-like factor required to 

direct continuous xylem differentiation in Zinnia elegans L. and 

A. thaliana (Motose et al. 2004). It is thought that, in a similar 

manner, OPS contributes to longitudinal signaling, thereby 

inducing SE differentiation in undifferentiated SE precursor 

cells. Further study of OPS function and identification of factors 

relating to OPS function will advance our understanding of how 

phloem continuity is organized and how the phloem develops. 

The Arabidopsis LATERAL ROOT DEVELOPMENT 3 

(LRD3) is another important gene which has been reported 

to regulate early phloem development and to control transport 

function of phloem (Ingram et al. 2011). The LRD3 gene 

encodes a LIM-domain protein which is specifically expressed 

in the CCs. The normal function of LRD3 is to maintain a 

balance between primary and lateral root growth and phloemmediated 

resource allocation within the root system. The lrd3 

loss of function mutant has decreased primary and increased 

lateral root growth and density, without having a significant 

effect on sucrose uptake. Additionally, aniline blue staining 

of the lrd3 primary root shows an overall reduction in the 

callose level in root meristems, developing SEs, and the PD 

that connect CCs to SEs, suggesting a non-cell-autonomous 

role for LRD3 in early phloem development. 

A detailed analysis of long-distance transport using different 

transport assays, including 14 C-sucrose, the fluorescent tracer 

dye carboxyfluorescein diacetate (CFDA) and CC-driven GFP 

(AtSUC2::GFP), demonstrated that phloem translocation to 

the primary root tip is severely limited in young lrd3 plants, 

whereas phloem loading and export from the shoot appear to 

be normal. Notably, these phloem defects were subsequently 

rescued, spontaneously, in older plants, along with a subsequent 

increase in phloem delivery to and growth of the primary 

root. Importantly, continuous exogenous auxin treatment could 

rescue the early phloem developmental defects and transport 

function in the primary roots of lrd3. This finding suggested 

either that auxin functions downstream of LRD3, or that it may 

have an independent key role in early phloem development. 

Interestingly, this study of the effects of lrd3 on root system 

architecture and the pattern of phloem translocation in the root 

system suggests that there might be some tightly regulated 

mechanism(s) which selectively supports a biased phloemmediated 

resource allocation in the lateral roots when the 

primary root is compromised. 

Phloem: A conduit for delivery of photosynthate and 

information molecules 

Phloem is an important organ, vital for more than just the wellestablished 

function of photoassimilate transport from the photosynthetic 

organs to the sink tissues. New transport functions 

continue to be discovered, such as the phloem-based transport 

of phytohormones, small RNAs, mRNAs and proteins. As will 

be discussed later, this transport of macromolecules appears 

to play a role in facilitating the coordinated developmental 

programs in meristematic regions located at various locations 

around the body of the plant. 

In addition to its specialized transport functions, phloem also 

stands out by the highly distinctive morphology of its cell types. 

In the angiosperms, the interaction between the enucleate SEs 

and the CCs needs to be highly integrated in order to maintain 

the operation of the sieve tube system (van Bel 2003). This 

process likely involves the cell-to-cell trafficking of a wide range 

of molecules via the PD that interconnect the SE-CC complex. 

Future studies on the nature of this molecular exchange will be 

greatly assisted by the use of the modified CALS3m system 

(Vatén et al. 2011) which can serve as an effective tool to 

modulate transport between specific cell types.

Molecular Mechanisms Underlying Xylem 

Cell Differentiation 

In the shoot apical meristem, stem cells differentiate into various 

cell types that comprise the shoot, while still proliferating 

in order to maintain themselves (Weigel and Jürgens 2002). 

Similarly, in the vascular meristem, procambial and cambial 

cells differentiate into specific vascular cells, such as tracheary 

elements, xylem fiber cells, xylem parenchyma cells, SEs, CCs, 

phloem parenchyma and phloem fiber cells, while again maintaining 

activity to proliferate (Figure 8). Therefore, procambial 

and cambial cells are considered as vascular stem cells (Hirakawa 

et al. 2010, 2011; Miyashima et al. 2012). Recent studies 

have revealed that local communication between vascular 

stem cells and differentiated vascular cells directs the wellorganized 

formation of vascular tissues (Lehesranta et al. 2010; 

Hirakawa et al. 2011). During this vascular formation, plant 

hormones, including auxin, cytokinin and brassinosteroids, act 

as signaling molecules that mediate in this process of cell-cell 

communication (Fukuda 2004). In addition, recently, a tracheary 

element differentiation inhibitory factor (TDIF), a small 

peptide, was found to function as a signaling molecule both inhibiting 

xylem cell differentiation from procambial cells and promoting 

procambial cell proliferation (Ito et al. 2006; Hirakawa 

et al. 2008, 2010). TDIF belongs to the CLAVATA3/EMBRYO 

SURROUNDING REGION-related (CLE) family, some of 

whose members are central players in cell-cell communication 

within meristems (Diévart and Clark 2004; Matsubayashi and 

Sakagami 2006; Fiers et al. 2007; Fukuda et al. 2007; Jun et al. 

2008; Butenko et al. 2009; Betsuyaku et al. 2011). 

Further insight into the differentiation of procambial cells 

into xylem cells has been gained from recent comprehensive 

gene expression and function analyses (Kubo et al. 2005; 

Zhong et al. 2006; Yoshida et al. 2009; Ohashi-Ito et al. 2010; 

Yamaguchi et al. 2011). In particular, the discovery of master 

genes that induce differentiation of various xylem cells greatly 

enhanced our understanding of xylem formation (Kubo et al. 

2005; Zhong et al. 2006; Mitsuda et al. 2007). Further analysis 

of these master genes revealed transcriptional networks that 

control xylem cell differentiation, which involves specialized 

secondary wall formation. Tracheary element differentiation 

also involves programmed cell death (PCD) (Fukuda 2000). In 

this section of the review, we will evaluate advances in our understanding 

of xylem cell differentiation from procambial cells, 

with a focus on cell-cell signaling, the underlying transcriptional 

network and the onset of PCD. 

Intercellular signaling pathways regulating xylem 

differentiation 

The TDIF-TDR signaling pathway regulates vascular stem cell 

maintenance. TDIF is a CLE-family peptide composed of twelve 

Figure 8. Xylem cells. 


(A) Poplar vascular tissue in which tracheary elements (TE) and 

xylem fiber (XF) cells are formed. 

(B) VND6-induced tracheary elements. 

(C) SND1-induced xylem fiber cells. 

Scale bars: 25 µm in(B) and (C). 

amino acids with hydroxylation on two proline residues (Ito 

et al. 2006). In the Arabidopsis genome, this TDIF sequence 

is encoded by two genes, CLE41 and CLE44 (Hirakawa 

et al. 2008). The TDIF RECEPTOR/PHLOEM INTERCALATED 

WITH XYLEM (TDR/PXY) is a receptor for TDIF, which belongs 

to the Class XI LEUCINE-RICH REPEAT RECEPTOR-LIKE 

KINASE (LRR-RLK) family (Hirakawa et al. 2008). 

Genetic and physiological analyses have revealed that the 

TDIF-TDR signaling pathway is crucial for vascular stem cell 

maintenance, by inhibiting xylem differentiation from procambial 

cells and promoting procambial cell proliferation (Hirakawa 

et al. 2008) (Figure 9). TDR is expressed preferentially in the 

procambium and cambium (Fisher and Turner 2007; Hirakawa 

et al. 2008), whereas CLE41 and CLE44 are expressed 

specifically in the phloem and more widely in its neighbors, 

respectively. Defects in TDR or CLE41 cause the exhaustion 

of procambial cells located between the phloem and xylem, 

resulting in formation of xylem vessels adjacent to phloem 

cells in the hypocotyl (Fisher and Turner 2007; Hirakawa et al. 

2008, 2010). Ectopic expression of CLE41, under either a


Figure 9. Regulation of procambial cell fates by the tracheary 

element differentiation inhibitory factor (TDIF) –TDIF receptor 

(TDR) signaling pathway. 

TDIF is produced in phloem cells, secreted from phloem cells, and 

perceived by TDR in procambial cells. TDR signaling is diverged 

into two pathways: one promotes self-renewal via WOX4, and the 

other inhibits tracheary element (TE) differentiation from procambial 

cells, probably indirectly via the suppression of VND6/VND7. 

xylem-specific or 35S promoter, disrupts the normal pattern 

of vascular tissues, indicating the importance of the synthesis 

site for this signaling peptide (Etchells and Turner 2010). Thus, 

phloem-synthesizing TDIF regulates procambial cell fate in a 

non-cell-autonomous fashion. 

The TDIF peptide signal activates expression of WOX4, 

a member of the WUSCHEL-related HOMEOBOX (WOX) 

gene family, in procambial and cambial cells (Hirakawa et al. 

2010; Ji et al. 2010; Suer et al. 2011). Interestingly, WOX4 

is required for TDIF-dependent enhancement of procambial 

cell proliferation, but not for the TDIF-dependent suppression 

of xylem differentiation (Hirakawa et al. 2010). Ethylene/ERF 

signaling is reported to be another pathway to regulate procambial/cambial 

cell division and may function in parallel to the 

CLE41-TDR/PXY pathway, and, under normal circumstances, 

TDR/PXY signaling acts to repress the ethylene/ERF pathway 

(Etchells et al. 2012). Hence, at least two intracellular signaling 

pathways that diverge after TDIF recognition by TDR 

may regulate, independently, the behavior of vascular stem 

cells. Lastly, TDIF, which is produced mainly by CLE42, has 

also been shown to play a role in axillary bud formation in 

Arabidopsis, indicating that it is a multifunctional peptide signal 

in plants (Yaginuma et al. 2011). 

CLE peptides inhibit protoxylem vessel formation through 

activating cytokinin signaling. Cytokinin is a key regulator of 

xylem development (Mähönen et al. 2000, 2006; Mok and Mok 

2001; Matsumoto-Kitano et al. 2008; Bishopp et al. 2011b). 

Recent studies have revealed crosstalk between CLE peptide 

and cytokinin signaling, which regulates xylem differentiation 

(Kondo et al. 2011). In roots, TDIF does not significantly 

affect vascular development (Kondo et al. 2011). In contrast, 

treatment with some CLE peptides, including CLE9/CLE10, 

inhibits formation of protoxylem but not of metaxylem vessels in 

Arabidopsis roots. CLE9 and CLE10, which encode the same 

CLE peptide, are preferentially expressed in vascular cells of 

roots (Kondo et al. 2011). Microarray analysis revealed that the 

CLE9/CLE10 peptide specifically reduces expression of type-A 

ARABIDOPSIS RESPONSE REGULATERs (ARRs) which are 

known as negative regulators of cytokinin signaling (Kiba et al. 

2003; To et al. 2004, 2007). 

The ARR5 and ARR6 are particular CLE9/CLE10 targets 

and, consistent with this finding, in the root of arr5arr6 double 

mutant plants, protoxylem vessel formation is often inhibited 

(Kondo et al. 2011). Conversely, arr10arr12, a double mutant 

for two type-B ARRs, which function positively in cytokinin 

signaling, displayed ectopic protoxylem vessel formation. Furthermore, 

arr10arr12 was resistant to the CLE9/CLE10 peptide 

in terms of protoxylem vessel formation. Interestingly, 

other combinations of type-B ARR mutants, such as arr1arr12 

and arr1arr10, showed much weaker resistance against the 

CLE9/CLE10 peptide compared with arr10arr12. This result 

implies that ARR10 and ARR12 act as major Type-B ARRs. 

Thus, the CLE9/CLE10 peptide activates cytokinin signaling 

through the repression of ARR5 and ARR6, resulting in the 

inhibition of protoxylem vessel formation. Genetic analysis 

suggests that the CLV2 membrane receptor and its partner 

CRN/SOL2 kinase (Miwa et al. 2008; Müller et al. 2008) 

may act in protoxylem vessel formation, downstream of the 

CLE9/CLE10 peptide signaling (Kondo et al. 2011). 

For cell-to-cell communication, plant cells send signaling 

molecules via the symplasmic pathway. A GRAS-family transcription 

factor, SHR, is a signal that moves cell to cell selectively 

through PD. SHR proteins are known to move from the 

stele into the endodermis to induce another GRAS-family transcription 

factor, SCARECROW (SCR), and then, together with 

SCR, they up-regulate expression of target genes, including the 

miR165/166 genes (Levesque et al. 2006; Cui et al. 2007; Gallagher 

and Benfy 2009). The mature miR165/166 moves back 

from the endodermis into the pericyle and protoxylem vessel 

poles in the stele, most likely through PD. Here, miR165/166 

degrades the transcripts of PHB and its family of Class III HD- 

ZIP genes (Carlsbecker et al. 2010). These transcripts within

xylem precursors specify central metaxylem vessels, at high 

levels, and peripheral protoxylem vessels, at low levels. This 

reciprocal signaling between the inner vascular tissues and the 

surrounding cell layers allows the domain of response to be 

confined to one of two tissue compartments (Scheres 2010). 

Genome-wide analysis revealed the presence of direct targets 

of SHR not only in the endodermis but also in the xylem and 

pericycle, suggesting a complex function of this transcription 

factor in vascular development (Cui et al. 2012). 

Thermospermine, a structural isomer of spermine (Ohsima 

1979), has been shown to act as a suppressor of xylem development. 

ACAULIS 5 (ACL5) encodes a thermospermine synthase 

(Kakehi et al. 2008) that is expressed specifically in early 

developing vessel elements (Muñiz et al. 2008). ACL5 lossof-function 

mutants cause excessive differentiation of xylem 

cells (Hanzawa et al. 1997; Clay and Nelson 2005; Muñiz et al. 

2008). Exogenously applied themospermine suppresses xylem 

vessel differentiation in both Arabidopsis plants and a Zinnia 

xylogenic culture (Kakehi et al. 2010). Genetic analysis of acl5 

identified a suppressor of the acl5 phenotype, sac51, whose 

causal gene encodes a basic helix-loop-helix (bHLH) transcription 

factor (Imai et al. 2006). Thermospermine is considered 

to regulate translational activity of SAC51 mRNA, resulting 

in the suppression of xylem development (Imai et al. 2008). 

A recent chemical biology approach also indicated that the 

SAC51-mediated thermospermine signaling pathway can limit 

auxin mediated promotion of xylem differentiation (Yoshimoto 

et al. 2012). Thus, the possibility exists that ACL5 may control 

xylem specification through the prevention of premature cell 

death (Muñiz et al. 2008; Vera-Sirera et al. 2010). 

Transcriptional regulation of xylem cell differentiation 

The Class III HD-ZIP genes have been shown to regulate 

xylem differentiation. In the phb phv rev cna athb8 mutant 

background, procambial cells fail to differentiate into xylem 

cells, but proliferate actively to produce many procambium 

cells. However, every quadruple loss-of-function mutant of the 

five Class III HD-ZIP genes exhibits ectopic xylem formation 

in the roots (Carlsbecker et al. 2010). In contrast, a gain-offunction 

mutant of PHB induces ectopic metaxylem vessel 

formation (Carlsbecker et al. 2010), and overproduction of 

ATHB8 promotes xylem differentiation (Baima et al. 1995). 

These findings indicate that the Class III HD-ZIP members function 

positively in xylem specification. However, the regulation 

of xylem differentiation by these genes is more complicated. 

In roots, miR165/166, which degrades Class III HD-ZIP transcripts, 

promotes protoxylem vessel differentiation. Therefore, 

it is proposed that high transcript levels of these genes inhibit 

protoxylem vessel formation, but promote metaxylem vessel 

formation. Because exogenously applied brassinosteroids promote 

the expression of Class III HD-ZIP genes (Ohashi-Ito and 


Fukuda 2003) and xylem cell differentiation (Yamamoto et al. 

1997), brassinosteroids may promote xylem differentiation, at 

least partly, through activation of these genes. These Class III 

HD-ZIP and KANADI transcription factors were also reported to 

regulate cambium cell differentiation, in which KANADI might 

act by inhibiting auxin transport and Class III HD-ZIPs by 

promoting xylem differentiation (Ilegems et al. 2010; Robischon 

et al. 2011). 

Members of a subgroup of NAM/ATAF/CUC (NAC) domain 

proteins, namely the VASCULAR-RELATED NAC-DOMAINs 

(VNDs) and NAC SECONDARY WALL THICKENING PRO- 

MOTING FACTORs/SECONDARY WALL-ASSOCIATED NAC 

DOMAIN PROTEINs (NSTs/SNDs), function as master transcription 

factors that can induce xylem cell differentiation by 

their ectopic expression (Demura and Fukuda 2007; Zhong and 

Ye 2007). VND6 and VND7 initiate metaxylem and protoxylem 

vessel differentiation, respectively (Kubo et al. 2005). Similarly, 

SND1/NST3 and NST1 induce xylem fiber differentiation 

(Mitsuda et al. 2005, 2007; Zhong et al. 2006). However, a 

single loss-of-function mutant of each gene shows no morphological 

defects, suggesting that other family members may 

have redundant functions to induce xylem differentiation, although 

each does not induce xylem cell differentiation when 

overexpressed (Kubo et al. 2005). 

The activity of these master transcription factors appears to 

be regulated by the following three mechanisms. (1) Expressions 

of VND7 and two genes for AS2/LBD domain-containing 

proteins, ASL20/LBD18 and ASL19/LBD30, form a positive 

feedback loop to amplify their expression (Soyano et al. 2008). 

This rapid amplification of the master transcription factor may 

drive xylem cell differentiation promptly and irreversibly. (2) 

VND7 activity is also regulated at the protein level by its 

proteasome-mediated degradation (Yamaguchi et al. 2008). (3) 

A NAC domain transcription repressor, VND-INTERACTING2 

(VNI2), represses VND activity by protein-protein interaction 

(Yamaguchi et al. 2010). VNI2 has an unstable property because 

of the PEST proteolysis target motif in its C-terminal 

region, which may allow VND7 to exert its function promptly 

when it is required. 

Xylem cells form characteristic secondary walls. These morphological 

events are regulated by master regulators such as 

SND1/NST3, VND6 and VND7. Microarray experiments revealed 

that VND6, VND7 and SND1 induce a hierarchical gene 

expression network (Zhong et al. 2008; Ohashi-Ito et al. 2010; 

Yamaguchi et al. 2010). These master transcription factors 

induce, probably directly, the expression of genes for other 

transcription factors, such as MYB46, MYB83, and MYB103. 

MYB46 and MYB83 regulate redundant biosynthetic pathways 

for all three major secondary wall components, namely cellulose, 

lignin, and xylan (Zhong et al. 2007; McCarthy et al. 

2009). Here, two NACs and 10 MYBs appear to act downstream 

of SND1 (Zhong et al. 2008; Zhou et al. 2009). Of them,


MYB58, MYB63 and MYB85, which might be target genes of 

MYB46 and/or MYB83, specifically upregulate genes related 

to the lignin biosynthetic pathway (Zhou et al. 2009). Some of 

these key transcription factors are also induced by VND6 and 

VND7, suggesting that this hierarchal structure is also true in 

the case of VND6 and VND7 (Ohashi et al. 2010; Yamaguchi 

et al. 2010). However, each master regulator also induces the 

expression of distinct genes, including transcription factors. 

Thus, SND1/NST3, VND6 and VND7, as master regulators, 

switch at the top of the hierarchy to upregulate transcription 

factors such as MYBs, which, in turn, functioning as the second 

and third regulators, upregulate expression of genes encoding 

enzymes catalyzing secondary wall thickening during specific 

stages of xylem cell differentiation. 

Interestingly, VND6 and VND7 can directly upregulate the 

expression of genes for enzymes such as XCP1 and CESA4, 

which are ranked lowest, as well as genes for transcription 

factors, such as MYB46, which are ranked higher in the 

gene expression hierarchy (Ohashi-Ito et al. 2010; Yamaguchi 

et al. 2011). Similarly, genes for enzymes such as 4CL1 are 

direct targets of SND1 (Zhong et al. 2008; McCarthy et al. 

2009). These findings indicate a sophisticated transcriptional 

regulatory network, by the master regulator, over a hierarchy. 

Tracheary elements and xylem fibers possess different characteristics, 

such as cell wall structure and PCD. In accordance 

with these characters, VND6, but not SND1, induces the 

expression of genes related to rapid PCD, such as XCP1 

and XCPs, while SND1, but VND6 preferentially, upregulates 

genes for lignin monomer synthesis, such as PAL1, 4CL3, and 

CCoAOMT (Ohashi-Ito et al. 2010). 

It is well-established that an 11-bp cis-element named 

the tracheary-element-regulating cis-element (TERE), which 

is found in upstream sequences of many genes expressed 

in xylem vessel cells, is responsible for xylem vessel cellspecific 

expression (Pyo et al. 2007). VND6 binds the TERE 

sequence and activates the TERE-containing promoter, in 

planta, but not a mutated promoter having substitutions in the 

TERE sequence (Ohashi-Ito et al. 2010). VND7 also binds 

TERE (Yamaguchi et al. 2011). These results demonstrate 

that TERE is one of the target sequences contained within 

the VND6 promoter. In contrast, SND1 specifically binds to a 

19-bp sequence named SECONDARY WALL NAC BINDING 

ELEMENT (SNBE), to activate its target genes (Zhong et al. 

2010). 

Cellular events underlying xylem cell formation 

Xylem cell differentiation involves temporal and spatial regulation 

of secondary cell wall deposition. A number of xylem 

cell types exist, such as those with annular and spiral patterns 

in protoxylem vessels, reticulate and pitted patterns in 

metaxylem vessels, and a smeared pattern in xylem fibers. 

The cortical microtubules regulate the spatial pattern of the 

secondary cell wall by orientating cellulose deposition. By 

using cultures expressing GFP-tubulin it was discovered that 

cortical microtubules became gradually bundled, which, in turn, 

was followed by secondary wall deposition (Oda et al. 2005, 

2006, 2010). It is important to find microtubule associated 

proteins (MAPs) involved in secondary wall formation and to 

know their function. In this context, some important secondary 

wall-related MAPs that regulate cortical microtubule orientation 

have been discovered. For example, AtMAP70 family proteins 

appear to be involved in the formation of the secondary wall 

boundary (Pesquet et al. 2010). The plant-specific microtubule 

binding protein MIDD1/RIP3 promoted microtubule depolymerization 

in the future secondary wall pit area, resulting in a 

secondary wall-depletion domain (Oda et al. 2010). Further 

analysis revealed that ROPGEF4 and ROPGAP3 mediate local 

activation of the plant Rho GTPase ROP11, and this activated 

ROP11 then recruits MIDD1 to induce local disassembly of 

cortical microtubules (Oda and Fukuda 2012b). Interestingly, 

and conversely, cortical microtubules eliminate active ROP11 

from the plasma membrane through MIDD1. Such a mutual 

inhibitory interaction between active domains of ROP and 

cortical microtubules gives rise to distinct patterns of secondary 

cell walls. These findings shed new insights into the microtubule 

organizing mechanism regulating secondary wall patterning 

(Oda and Fukuda 2012a). 

PCD is a genetically regulated cell suicide process involved 

in many aspects of plant growth, such as seed germination, 

vascular differentiation, aerechyma tissue formation, reproductive 

organ development and leaf senescence (Kuriyama and 

Fukuda 2002). During xylem development, rapid and slow PCD 

occurs in tracheary elements and xylem fiber cells, respectively, 

in order to facilitate the removal of cellular content for the 

formation of dead cells with secondary walls (Bollhoner et al. 

2012). PCD during tracheary element differentiation has long 

been recognized as an example of developmental PCD in 

plants (Fukuda 2004; Turner et al. 2007). This process includes 

cell death signal induction, accumulation of autolytic enzymes 

in the vacuole, vacuole swelling and collapse, and degradation 

of cell contents followed by mature tracheary element formation 

(Fukuda 2000). 

It has been suggested that the signals for xylem cell death are 

produced early during xylem differentiation, and that cell death 

is prevented through the action of inhibitors and the storage 

of hydrolytic enzymes in the vacuole (Bollhoner et al. 2012). 

According to the morphological process, the death of xylem 

tracheary elements is defined as a vacuolar type of cell death 

(Kuriyama and Fukuda 2002; Van Doorn et al. 2011a). Vacuolar 

membrane breakdown is the crucial event in tracheary element 

PCD, and bursting of the central vacuole triggers autolytic 

hydrolysis of the cell contents, thereby leading to cell death 

(Bollhoner et al. 2012).

Microarray analyses of gene expression have revealed a 

simultaneous expression of many genes involved in both secondary 

wall formation and PCD (Demura et al. 2002; Milioni 

et al. 2002; Kubo et al. 2005; Pesquet et al. 2005; Ohashi 

et al. 2010). As mentioned above, recent results have demonstrated 

that a transcriptional regulatory system, composed of 

transcription factors such as VND6 and a TERE-cis sequence, 

regulates the simultaneous expression of genes related to 

both secondary wall formation and PCD in tracheary elements 

(Ohashi-Ito et al. 2010). These findings indicate that tracheary 

element-differentiation-inducing master genes initiate at least 

a part of PCD directly by activating PCD-related genes through 

binding the TERE sequence in their promoters. This suggests 

that, in contrast to apoptosis in animals, in which a common 

intracellular signaling system induces PCD, in plants, various 

developmental processes involving PCD may be regulated independently 

involving their own specific developmental steps. 

Nitric oxide (NO) and polyamine have been suggested as 

signals involved in the cell death induction in xylem development. 

NO production is largely confined to xylem cells; removal 

of NO from the cultured Zinnia cells, with its scavenger PTIO, 

results in dramatic reductions in both PCD and in the formation 

of tracheary elements (Gabaldon et al. 2005). Thus, NO might 

well be an important factor mediating PCD during tracheary 

element differentiation. 

Execution of PCD in developing tracheary elements involves 

expression and vacuole-accumulation of several hydrolytic 

enzymes, such as the cysteine proteases XCP1 and XCP2 

(Zhao et al. 2000; Funk et al. 2002; Avci et al. 2008), the Zn 2+ - 

dependent nuclease ZEN1 (Ito and Fukuda 2002) and RNases 

(Lehmann et al. 2001). Ca 2+ -dependent DNases were also 

detected in secondary xylem cells, and their activity dynamics 

were closely correlated with secondary xylem development 

(Chen et al. 2012a). ATMC9 encodes a caspase-like protein, 

which does not function as a caspase but as an arginine/lysinespecific 

cysteine protease (Vercammen et al. 2004). ATMC9 

was specifically expressed in differentiating vessels but not 

in fully-differentiated vessels (Ohashi-Ito et al. 2010). This 

result suggested the involvement of ATMC9 in PCD. However, 

application of caspase inhibitors significantly delays the time of 

tracheary element formation and inhibits DNA breakdown and 

appearance of TUNEL-positive nuclei in Zinnia xylogenic cell 

culture (Twumasi et al. 2010). 

Recently, the protease responsible for developing xylemrelated 

caspase-3-like activity was purified and identified to 

be 20S proteasome (Han et al. 2012). The fact that treatment 

with a caspase-3 inhibitor Ac-DEVD-CHO causes a defect in 

veins in Arabidopsis cotyledons, and the proteasome inhibitor 

clasto-lactacystin β-lactone delays tracheary element PCD in 

VND6-induced Arabidopsis xylogenic culture, strongly suggest 

that the proteasome is involved in PCD during this process of 

differentiation (Han et al. 2012). Consistent with this notion, the 


26S proteasome inhibitors lactacystin and MG132 also delay 

or block the differentiation of suspension-cultured tracheary 

elements (Woffenden et al. 1998; Endo et al. 2001). Autophagy 

has also been suggested to be involved in tracheary element 

PCD (Weir et al. 2005). A recent finding that a small GTP 

binding protein RabG3b plays a positive role in PCD during tracheary 

element differentiation by activating autophagy (Kwon 

et al. 2010) provided support for this notion. 

The PCD of xylem fibers is less well characterized compared 

to that of xylem tracheary elements, likely due to the fact that 

this process proceeds slowly in these cell types. Microarray 

analyses revealed that a large number of genes encoding 

previously-uncharacterized transcription factors, as well as 

genes involved in ethylene, sphingolipids, light signaling and 

autophagy-related factors, are expressed preferentially during 

xylem fiber development (Courtois-Moreau et al. 2009). Further 

comparison of genes related to PCD between xylem fibers 

and tracheary elements, in a model species like poplar, may 

help in advancing our understanding of PCD as it occurs in 

plants. 

Control over master transcription factors and crosstalk 

between signaling pathways 

It is now well established that xylem cell differentiation is 

regulated by various factors, both at the cell-autonomous and 

non-cell-autonomous level. Auxin, cytokinin, brassinosteroids 

and CLE peptides act, cooperatively, at different stages of 

xylem cell differentiation. Importantly, an as-yet-unidentified 

intracellular signaling system initiates the expression of genes 

for master transcription factors such as VND6, VND7 and 

SND1/NST1, each of which in turn induces distinctive xylem 

cell-specific gene expression. Further advances in our understanding 

of the events underlying xylem differentiation will 

be gained by studies on the related intracellular signaling 

pathways and the nature of the crosstalk that occurs between 

these specific signaling pathways. 

Spatial & Temporal Regulation 

of Vascular Patterning 

Vascular organization in leaves 

The leaf vascular system is a network of interconnecting 

veins, or vascular strands, consisting of two main conducting 

tissue types: the xylem and the phloem. While the specialized 

conducting elements are composed of tracheary or vessel 

elements in the xylem and SEs in the phloem, the vascular 

system also contains non-conducting supporting cells, such as 

parenchyma, sclerenchyma and fibers. Thus, the development 

of cell types within the radial arrangement of the vascular 

bundle must be precisely spatially coordinated along with the


temporal longitudinal vein pattern in order to efficiently carry 

out their function as the long-distance transport system of the 

plant (Dengler and Kang 2001). 

The spatial organization of the leaf vascular system is both 

species- and organ-specific. Despite the diverse vein patterns 

found within leaves, the one commonality that is present during 

the ontogeny of the vascular system is the organization of the 

vascular bundles into a hierarchical system. Veins are organized 

into distinct size classes, based on their width at the most 

proximal point of attachment to the parent vein (Nelson and 

Dengler 1997). Primary and secondary veins are considered 

to be major veins, not only due to their width, but because 

they are typically embedded in rib parenchyma, whereas higher 

order, or minor, veins such as tertiary and quaternary veins are 

embedded in mesophyll (Esau 1965a). The highest order veins, 

the freely ending veinlets, are the smallest in diameter and end 

blindly in surrounding mesophyll (Figure 10A). 

The presence of this hierarchical system in leaves reflects the 

function of the veins such that larger diameter veins function 

in bulk transport of water and metabolites, whereas smaller diameter 

veins function in phloem loading (Haritatos et al. 2000). 

In both the juvenile and adult phase leaves of Arabidopsis, 

the vein pattern is characterized by the major secondary veins 

that loop in opposite pairs in a series of conspicuous arches 

along the length of the leaf (Hickey 1973). This looping pattern, 

termed brochidodromous, is present in both juvenile and adult 

phase leaves. However, the hierarchical pattern is well defined 

in the adult leaves; there is a higher vein density and vein order 

(up to the 6 th order) when compared with the juvenile leaves 

(Kang and Dengler 2004). Despite this increasing vascular 

complexity, the overall vein pattern within a given species is 

highly conserved and reproducible, yet the vasculature itself 

is highly amenable to changes and re-modification during leaf 

development (Kang et al. 2007). 

Longitudinal vein pattern—procambium 

As indicated above, the procambium is a primary meristematic 

tissue that develops de novo from ground meristem cells to 

form differentiated xylem and phloem. In a temporal sense, the 

longitudinal vein pattern in Arabidopsis develops basipetally. 

However, the individual differentiating strands of the preprocambium, 

procambium, and xylem develop in various directions 

(basipetally, acropetally or perpendicular to/or from the 

leaf midvein) depending on the stage of vascular development, 

as well as the local auxin levels (Figure 10B). Based strictly 

on its anatomical appearance, procambium is first identifiable 

by its cytoplasmically dense narrow cell shape and continuous 

cell files that seemingly appear either simultaneously or 

progressively along the length of the vascular strand (Esau 

1965b; Nelson and Dengler 1997). 

Figure 10. Longitudinal and radial vein patterning in leaves. 

(A) Venation pattern in the lamina of a mature Arabidopsis leaf. 

Vein size hierarchy is based on diameter of the veins at their 

most proximal insertion point. Vein size classes are color coded 

as follows: Orange, mid (primary) vein; purple, secondary/marginal 

veins; blue, tertiary veins; red, quaternary/freely ending veinlets. 

(B) Development of vein pattern in young leaves, as indicated by 

AtHB-8 (Kang and Dengler 2004; Scarpella et al. 2004). Establishment 

of the overall vein pattern in Arabidopsis is basipetal (black 

arrow). Secondary pre-procambium of the first pair of loops develop 

out from the midvein (dotted pink arrows, arrow indicates direction 

of pre-procambial strand progression). Pre-procambium of the 

second pair of secondary vein loops progresses either basipetally or 

acropetally. Third and higher secondary vein loop pairs progress out 

from the midvein towards the leaf margin and reconnect with other 

extending strands (dotted black arrows). Procambium differentiates 

simultaneously along the procambial strand (blue solid lines). Xylem 

differentiation occurs approximately 4 d later and can develop 

either continuously, or as discontinuous islands, along the vascular 

strand (purple lines, arrow indicates direction of xylem strand 

progression). 

(C) Differentiation of procambial cells. Pre-procambium is isodiametric 

in cell shape and is anatomically indistinguishable 

from ground meristem cells (maroon cell). Cell divisions of 

the pre-procambium are parallel to the direction of growth 

(light blue cells) of the vascular strand, resulting in elongated 

shaped cells characteristic of the procambium (dark blue 

cell). 

(D) Radial vein pattern in leaves. (Left to right): Procambial cells 

(as indicated by AtHB-8) are present within the vascular bundle. 

In a typical angiosperm leaf, xylem cells are dorsal to phloem 

cells (collateral vein pattern). In severely radialized leaf mutants, 

vein cell arrangement also becomes radialized. In adaxialized 

mutants such as phabulosa (phb), phavoluta (phv), and revolute 

(rev), xylem cells surround phloem cells (amphivasal), whereas in 

abaxialized mutants, such as those in the KANADI gene family, 

phloem cells surround the xylem cells (amphicribral) (Eshed et al. 

2001; McConnell et al. 2001; Emery et al. 2003).

The elongated procambium cells develop through distinct 

cell division patterns in which they divide parallel to the vascular 

strand (Kang et al. 2007) (Figure 10C). Although the 

anatomical distinction of procambium is clearly evident by its 

elongated shape, the precursor cells, pre-procambium, are 

isodiametric and are anatomically indistinguishable from surrounding 

ground meristem cells. Due to the difficulty of clearly 

identifying pre-procambium and ground meristem through 

anatomy alone, the use of molecular markers such as Arabidopsis 

thaliana HOMEOBOX 8 (AtHB-8), MONOPTEROS 

(MP), and PINFORMED 1 (PIN1) to identify procambium and 

pre-procambium has facilitated the visualization of these early 

stages of vascular development (Mattsson et al. 2003; Kang 

and Dengler 2004; Scarpella et al. 2004; Wenzel et al. 2007). 

Auxin has been shown to regulate many aspects of plant 

development and play a critical role during vascular patterning, 

specifically vascular cell differentiation and vascular strand 

formation (Aloni 1987; Aloni et al. 2003; Berleth et al. 2000; 

Mattsson et al. 2003). Early classical experiments showed 

that auxin is capable of inducing new strands in response to 

wounding by promoting the transdifferentiation of parenchyma 

cells into continuous cell files towards the basal parts of the 

plant (Sachs 1981). These early observations led to the “auxin 

canalization hypothesis” which suggested that auxin is transported 

directionally through a cell, progressively narrowing into 

discrete canals and operating through self-reinforcing positive 

feedback (Sachs 1981). 

In recent years, it has been shown that auxin is synthesized 

predominantly in leaf primordia and transported unidirectionally 

from apical to basal regions of the plant. Polar auxin transport 

(PAT) is accomplished by translocating auxin in a targeted 

manner through the cell, via auxin influx and efflux carriers 

(Lomax and Hicks 1992). Several gene families are known 

to affect vascular strand formation by modulating auxin levels 

during leaf development. The Arabidopsis family of efflux 

carriers, PIN1, regulates the polarity and elevated auxin levels 

from the shoot apical meristem into developing leaf primordia 

(Reinhardt et al. 2000; Benkova et al. 2003). The subcellular 

epidermal localization and convergence of auxin flow to the 

tip of the leaf primordia and subsequent basal transport of 

auxin directs the location of the future midvein and the sites of 

vascular strand formation (Reinhardt et al. 2003; Petráˇsek et al. 

2006). 

In young leaf primordia, the lateral marginal convergence 

points of PIN1 are required for vascular strand positioning and 

arrangement (Sieburth 1999; Wenzel et al. 2007). Specifically, 

the initial broad expression domain of auxin in the developing 

leaf converges and tapers to narrow cell files (presumptive vascular 

strands) that are dependent on auxin transport (Scarpella 

et al. 2006; Sawchuk et al. 2008). The large family of auxin 

response factors, which include transcription factors such as 

MP/AUXIN RESPONSE FACTOR 5 (ARF5), plays a key role 


in vascular strand formation (Wenzel et al. 2007). It is now 

well documented that mp loss-of-function mutants have reduced 

vasculature, discontinuous veins, and also affect embryo 

polarity and root meristem patterning (Hardtke and Berleth 

1998; Hardtke et al. 2004; Wenzel et al. 2007; Schuetz et al. 

2008). 

MP regulates vascular formation by inducing PIN1 expression, 

and recently, MP has been shown to directly target AtHB- 

8 through an activator that binds to the TGTCTG element 

in the AtHB-8 promoter to induce pre-procambial expression 

(Donner et al. 2009). Expression of AtHB-8 is simultaneously 

present along with the expression of SHR to demarcate presumptive 

vascular cells (Wenzel et al. 2007; Donner et al. 

2009; Gardiner et al. 2011). Although AtHB-8 expression is 

specifically localized to and remains in pre-procambial and 

procambial cells, the expression domain of SHR is localized 

beyond the vascular strand, suggesting an alternative function 

beyond vascular development in leaves (Gardiner et al. 2011). 

The Class III HD-ZIP family of transcription factors, which 

includes AtHB-8, act as known regulators of both longitudinal 

and radial vascular patterning. The AtHB-8 gene is one of the 

earliest expressed in pre-procambial and procambial strands 

to set up vascular patterning (Baima et al. 1995; Kang and 

Dengler 2004; Scarpella et al. 2004). In Arabidopsis leaves, 

longitudinal vein pattern is initiated early in development 

through the acquisition of pre-procambial cells along a presumptive 

procambial strand (Kang and Dengler 2004; Scarpella 

et al. 2004; Sawchuck et al. 2007). Here, AtHB-8 is expressed 

in pre-procambial cells that are genetically identifiable from 

surrounding ground meristem cells. The distinct spatial organization 

of the secondary loops, characteristic of Arabidopsis 

vein patterning, develop uniquely based on the position of the 

secondary loop. 

The pre-procambium of the first secondary loop pair develops 

progressively away from the point of origin, the central midvein, 

to form a continuous loop (Figure 10B) (Kang and Dengler 

2004; Scarpella et al. 2004; Sawchuck et al. 2007). Later 

formed secondary pre-procambial strands (i.e. second or third 

secondary loop pairs) also develop away from the point of 

origin; however, the direction of the extending strand can 

develop either acropetally or basipetally (Kang and Dengler 

2004). Procambial strands develop simultaneously along the 

entire length of the vascular strand (Sawchuck et al. 2007). This 

simultaneous occurrence of the procambial strand is commonly 

seen in the first secondary loop pair. Importantly, procambial 

strands of later formed secondary loop pairs can differentiate 

towards the leaf margin and reconnect with other strands 

(Figure 10B). 

Approximately four d after procambium differentiation, xylem 

begins to develop both continuously from a point of origin 

or as discontinuous islands that connect either acropetally 

or basipetally with other strands (Figure 10B). To date, many


vascular pattern mutants have been identified (Scarpella and 

Meijer 2004; Scarpella and Helariutta 2010), and invariably, 

these mutants have disrupted and/or discontinuous vascular 

strands, suggesting that proper formation or continuity of vascular 

strands occurs early during the pre-procambial stages of 

development (Scarpella et al. 2010). 

Radial vein pattern—polarity and cell proliferation 

The spatial and temporal coordination of both longitudinal and 

radial vein pattern is essential for proper functioning of the vascular 

system. As leaves arise from the shoot apical meristem, 

the incipient leaf primordia are initially radialized, but internal 

tissues quickly become polarized acquiring adaxial (dorsal) 

and abaxial (ventral) cellular identities. The juxtaposition of 

adaxial and abaxial characteristics allows the leaf to grow 

out into a flattened lateral organ. In a typical eudicot leaf, the 

vascular tissues are arranged in a collateral pattern with xylem 

adaxial to the phloem. This differentiation must occur from 

a uniform procambium cell population (Figure 10D). Much of 

what we currently know concerning vascular polarity is derived 

from work conducted on leaf polarity mutants, as alterations in 

leaf polarity often result in vascular bundle defects (Scarpella 

and Meijer 2004; Husbands et al. 2009). In a completely 

radialized polarity mutant, vascular tissue is also radialized in 

that either xylem tissue surrounds a central cylinder of phloem 

(amphivasal) or phloem tissue surrounds a central cylinder of 

xylem (amphicribral) (Figure 10D). 

The establishment of adaxial-abaxial polarity is temporally 

regulated in the shoot apical meristem. Early experiments 

showed that meristem-derived signals may act to promote 

adaxial cell fate, as leaf primordia that were altered surgically 

became abaxialized (Sussex 1954). However, it is unlikely that 

meristem-derived factors alone are sufficient in establishing 

organ (and vascular) polarity, and that patterning of adaxialabaxial 

cell fate requires a number of genetic inputs. Transcription 

factors, such as those in the Class III HD-ZIP family, were 

identified as adaxial determinants based on radialized mutant 

phenotypes in Arabidopsis. Of these, the gain-of-function mutants 

in PHB, PHV, and REV display radialized leaves with 

amphivasal vascular bundles (McConnell et al. 2001; Emery 

et al. 2003). Their abaxial counterparts, such as the KAN genes 

(MYB-like GARP transcription factors) are expressed in abaxial 

tissues, promoting abaxial identity. Gain-of-function mutations 

in these genes can result in amphicribral (phloem cells outside 

of a ring of xylem cells) vascular bundles (Kerstetter et al. 2001; 

Emery et al. 2003), or in the most extreme case, result in the 

complete elimination of vascular tissue within the radialized 

organ (McConnell and Barton 1998; Sawa et al. 1999). 

The intimate connection between the development of the 

procambium and the surrounding ground meristem (mesophyll) 

during tissue histogenesis has yet to be deciphered. The 

genetic mechanisms coupling vascular cell proliferation with 

organ formation during tissue histogenesis are also largely 

unknown. However, it is known that cell proliferation is a 

critical developmental process that is required during tissue 

histogenesis. Attaining proper cell numbers within the radial 

vascular bundle is essential in order for vein size hierarchy 

to be properly established during vascular development. The 

organization of this vein hierarchy is controlled, at least in part, 

by cell cycle regulators such as CyclinB1;1 (Kang and Dengler 

2002). Expression of CyclinB1;1::GUS is modulated within the 

vein orders so that cell cycling is prolonged in larger vein size 

classes, such as the midvein, but ceases first in smaller order 

veins. Modification of cell proliferation in leaves established 

that vein patterning is tightly coordinated with maintenance of 

meristematic competency of ground meristem cells to regulate 

(higher order) vein architecture (Kang et al. 2007). Although 

the direct association between the cell cycle and vascular 

patterning has yet to be determined, genes known to play a 

role in cell proliferation and/or stem cell maintenance may aid 

in elucidating this mechanistic pathway (Ji et al. 2010; Vanneste 

et al. 2011). 

Spatio-temporal regulation of root vascular 

development 

A number of comprehensive reviews exist that cover different 

aspects of root xylem development (Cano-Delgado et al. 2010; 

Scarpella and Helariutta 2010). In this section of the review, 

we will focus first on the morphological evidence for the timing 

of events that control vascular specification and differentiation 

within the root. We will then assess progress in elucidating 

the molecular markers and regulatory factors that govern 

spatio-temporal aspects of root vascular development. Spatial 

regulation involves cellular mechanisms that determine the 

arrangement of vascular cell types relative to each other. 

The temporal regulation of vascular development comprises 

mechanisms that determine cell specification of vascular cell 

type lineages beyond the quiescent center (QC), as well as differences 

in the timing of these differentiation events (Mähönen 

et al. 2000). The developmental time at which various cell types 

differentiate can be read according to the cell’s distance from 

the QC, or position relative to the root meristematic, elongation 

or maturation zone (Figure 11A). As the majority of research 

into the regulation of the spatial or temporal aspects of vascular 

development has been performed in the Arabidopsis root, we 

will use this model system to highlight progress in this area. 

Morphological markers of root vascular development 

Vasculature in the Arabidopsis root, as discussed earlier, is 

composed of the radially symmetric pericycle cell layer that 

surrounds the diarch vasculature. The pericycle is differentiated

Figure 11. Spatio-temporal regulation of root vascular patterning. 


(A) Spatial markers of vascular differentiation. Developmental time points at which morphological markers consistent with differentiation of 

vascular cell types are indicated relative to the position along the longitudinal axis of the root. Changes in differentiation are highlighted 

in a change in cell color. MeZ, meristematic zone; El, elongation zone; MaZ, maturation zone; PP, protophloem; MP, metaphloem; CC, 

companion cells; PX, protoxylem; MX, metaxylem. 

(B) Temporal regulation of vascular regulator gene expression. The distinct temporal patterns of different vascular regulators are demonstrated 

along with the cell type with which these markers are associated. If a gene has a much higher peak of gene expression, then only this peak 

is shown. 

(C) Examples of genes whose expression shows fluctuating peaks in developmental time (first row), or dynamic expression between roots 

(compare first and second rows). 

into two cell types, the xylem and phloem pole pericycle cells. 

The former are located at the poles of the xylem axis and 

are the only cells competent to become lateral root primordia, 

whereas the latter occupy the position between the xylem 

poles. There are no morphological markers for phloem pole 

pericycle differentiation, other than their position relative to 

xylem pole pericycle cells, and the function of these cells 

remains to be elucidated. 

Phloem tissue is positioned interior to the pericycle cell 

layer and is located at the opposing poles of the vascular 

cylinder, whereas the central xylem axis cells form a median 

line transecting the vascular cylinder, perpendicular to the two 

phloem poles. Procambial cells are positioned between the 

xylem and phloem tissues. Xylem tissue is composed of two 

different cell types: protoxylem and metaxylem vessels. In the 

Arabidopsis root, there are two outer protoxylem cells and three 

inner metaxylem cells that can be distinguished based on their 

secondary cell wall characteristics. Protoxylem cells have a 

helical or annular pattern of secondary cell wall deposition, 

whereas metaxylem cells have a pitted deposition pattern.


Protoxylem vessels in the root mature before the surrounding 

tissues elongate; during cell expansion of these surrounding 

cells, these protoxylem vessels are often destroyed. Thus, the 

metaxylem vessels act as the primary water conducting tissue 

throughout the main body of the plant (Esau 1965b). Metaxylem 

cell differentiation is temporally separated from protoxylem 

differentiation in that the outer metaxylem cells differentiate 

only after protoxylem cells differentiate and the surrounding 

tissues have completed their expansion. The inner metaxylem 

vessel differentiates later than the outer two metaxylem cells. 

Phloem tissue is composed of three cell types: protophloem 

SEs to the outside, and metaphloem SEs to the interior of 

the vascular cylinder, with CCs flanking the SEs. Protophloem 

SEs differentiate earlier than the metaphloem SEs and their 

associated CCs. 

Detailed anatomical studies of the Arabidopsis root tip have 

elucidated the earliest events in the timing and patterning of 

vascular initial cell divisions that give rise to all vascular cell 

types in the primary root (Mähönen et al. 2000). Just above 

the QC, asymmetric cell divisions of vascular initial cells give 

rise to the presumptive pericycle layer and protoxylem cells. 

At a position close to the QC (∼9 µm), five xylem cells are 

visible, and these will eventually differentiate into protoxylem 

and metaxylem vessels (Figure 11A). Two domains of vascular 

initials give rise to the phloem and procambial cell lineages, 

and they are located between 3 µm and 6 µm above the 

QC (Mähönen et al. 2000; Bonke et al. 2003). The number 

and exact pattern of future procambial cell divisions is variable 

between individual plants of the same species. 

The full set of phloem cells (protophloem, metaphloem 

and CC) can be observed at a distance above the QC 

(∼27 µm) (Mähönen et al. 2000) (Figure 11A). Protophloem 

and metaphloem SEs result from one tangential division of 

precursor cells, whereas CCs arise from one periclinal division 

of precursor cells (Bonke et al. 2003). At a further distance 

above the QC (∼70 µm), the first histological evidence of 

differentiation can be observed in protophloem SEs, as determined 

by staining with toluidine blue (Mähönen et al. 2000). 

Thus, protophloem SE differentiation occurs much earlier in 

developmental time compared to protoxylem vessel formation 

(Figure 11A). Metaphloem SEs and CCs differentiate at an 

approximately similar time to the outer metaxylem SEs. However, 

morphological analyses have determined that the spatial 

patterning of xylem cells occurs temporally prior to the spatial 

patterning of the phloem cells within the root. 

Vascular proliferation—cytokinin signaling 

Vascular initial cells or stem cells are the progenitor cell type for 

all vascular cells within the primary root. Regulation of vascular 

initial cell division is the first step in vascular development 

and is accomplished, in part, by the two-component cytokinin 

receptor WOL (Mähönen et al. 2000). WOL is expressed early 

in the Arabidopsis embryo during the globular stage and is 

present throughout the vascular cylinder during all subsequent 

stages of embryo and primary root development (Figure 11B). 

Interestingly, vascular defects within the embryonic root have 

not yet been reported. In the primary root of a wol mutant, 

there are fewer vascular initial cells, and the entire vascular 

bundle differentiates as protoxylem. Although this suggests 

that wol is deficient in procambial, metaxylem vessel and 

phloem cell specification, a double mutant between wol and 

fass (which results in supernumerary cell layers) produces 

phenotypically normal procambial and phloem cells, as well as 

both protoxylem and metaxylem vessels. This demonstrates 

that the role of WOL is in vascular initial cell proliferation, and 

that any influence on cell specification is secondary to this 

defect. 

Transcriptional master regulators 

and xylem development 

Xylem cell differentiation, as marked by secondary cell 

wall synthesis and deposition, occurs much later in root 

developmental time relative to protophloem cell differentiation 

(Figure 11A). However, cells destined to become xylem cells 

are morphologically identifiable immediately after division of 

vascular initial cells. Based on gene expression data, a downstream 

regulator of cytokinin signaling, the AHP6, an inhibitory 

pseudophosphotransfer protein, is likely one of the earliest 

regulators of protoxylem cell specification (Mähönen et al. 

2006), but is unlikely to be the sole regulator (Figure 11B). AHP6 

functions to negatively regulate cytokinin signaling through 

spatial restriction of signaling within protoxylem cells. In a 

wol mutant, therefore, there is a lack of cytokinin signaling, 

a decrease in the asymmetric division of vascular initial cells 

and ectopic protoxylem cell differentiation in the few remaining 

vascular cells. AHP6 acts in a negative feedback loop with 

cytokinin signaling – cytokinin represses AHP6 expression, 

while AHP6 represses and spatially restricts cytokinin signaling 

(Mähönen et al. 2006). Cytokinin regulates the spatial domain 

of AHP6 expression in embryogenesis prior to when primary 

root protoxylem differentiation occurs. Thus, it appears that 

this negative regulatory feedback between cytokinin and AHP6 

occurs upstream of protoxylem specification in the primary root 

(Mähönen et al. 2006). 

The earliest marker of protoxylem cell specification in the primary 

root is achieved through a TARGET OF MONOPTEROS 

5 (TMO5) promoter:GFP fusion, named S4 (Lee et al. 2006; 

Schlereth et al. 2010). TMO5 is required for embryonic root 

initiation, and expression of this bHLH transcription factor is 

turned on shortly after division of vascular initial cells in the 

primary root and is turned off prior to secondary cell wall 

differentiation in protoxylem cells. This marker then turns on

in metaxylem cells and subsequently turns off again prior 

to secondary cell wall differentiation. MYB46 is one of the 

transcription factors partially required for synthesis of various 

components of the secondary cell wall in the protoxylem and 

subsequent metaxylem (Lee et al. 2006; Zhong et al. 2008). 

This gene is expressed towards the end of the elongation 

zone in protoxylem cells and then later in metaxylem cells. 

Together, these findings suggest that, although there are no 

morphological markers of protoxylem cell specification early 

in developmental time, there are indeed molecular markers 

and two distinct developmental states for protoxylem and 

metaxylem cells: an “early” state and a “late” state. Gene 

expression data support this observation, as there are distinct 

gene expression profiles in cells marked by TMO5 as compared 

to MYB46 (Brady et al. 2007). 

As mentioned earlier, PHB, PHV, REV, COR and ATHB- 

8 act redundantly to regulate xylem cell fate differentiation 

and are sufficient to regulate xylem patterning. In a dominant 

mutant background of PHB, phb-1d, which its mRNA is resistant 

to miRNA degradation, ectopic protoxylem is observed 

(Carlsbecker et al. 2010). PHB is therefore sufficient to specify 

protoxylem differentiation. The developmental time point at 

which the patterning of xylem cells is first regulated by these 

transcription factors remains unknown. Upstream regulators 

of SHR have yet to be identified, although SHR is expressed 

throughout the vascular cylinder during vascular development. 

Class III HD-ZIP transcription factors are also expressed in 

various cell types of the vasculature, primarily early in root 

developmental time in the meristematic zone; miR165a and 

miR166b are also expressed with a peak in endodermis cells in 

this same zone (Carlsbecker et al. 2010). Based solely on the 

temporal nature of these expression patterns, the patterning 

of protoxylem and metaxylem cells appears to occur prior to 

the action of VND6 and VND7, although validation of this 

hypothesis requires further experimentation. 

Phloem cell patterning and differentiation 

Only a few factors are known to regulate phloem cell patterning 

and differentiation, despite protophloem being histologically 

evident quite early in development relative to xylem cell types 

(Mähönen et al. 2000). Mutations in OPS, ops-1 and ops-2, 

result in irregular phloem differentiation within the root. In 

WT roots, SE and CC differentiation is first marked by cell 

elongation followed by callose deposition and subsequent cell 

wall thickening in the longitudinal dimension. In the ops-1 

mutant, cell elongation, callose deposition and cell wall thickening 

fail to occur within the phloem cell lineage (Truernit 

et al. 2012). In addition, the ops-1 mutant has impaired longdistance 

phloem transport, likely because of gaps in phloem SE 

continuity. However, OPS is not sufficient to specify phloem cell 

differentiation. Overexpression of OPS results in precocious 


phloem cell differentiation within the root, but only within already 

specified protophloem and metaphloem lineages (Truernit et al. 

2012). 

In roots, APL is expressed later than OPS, that is, at a 

distance from the QC (Bonke et al. 2003; Truernit et al. 

2012) (Figure 11B). In an apl mutant, protophloem cells are 

misspecified as protoxylem cells, and there is a short root 

phenotype (Bonke et al. 2003). Contrary to an ops mutant 

phenotype, in an apl mutant background, no protophloem, 

metaphloem, or CCs are present anywhere in the phloem pole 

position within the vascular cylinder. APL plays a multifaceted 

role in phloem development. First, APL regulates the timing 

of the asymmetric cell divisions that would normally give rise 

to the SE and CC lineages. Second, APL is required for 

protophloem and metaphloem SE differentiation. Third, APL 

represses protoxylem differentiation within cells in the phloem 

pole position. However, despite all these roles, APL is not 

sufficient to specify phloem cell specification and differentiation. 

Clearly, additional factors remain to be identified in phloem 

cell development. The presence of a master regulator for 

phloem development like VND6/7 and the Class III HD-ZIP 

family in xylem development that is both necessary and 

sufficient has not been identified. Protophloem cells differentiate 

earlier than metaphloem cells and CCs. APL protein 

localization reflects this temporal difference in differentiation, 

but the factor(s) that determines this temporal delay has yet to 

be isolated. Finally, factors that determine the spatial patterning 

of protophloem, metaphloem and CCs are similarly unknown. 

Pericycle cell specification and differentiation 

Pericycle cells have been divided into two populations based 

on gene expression and function. Only xylem pole pericyle cells 

are competent to become lateral root primordia. One marker of 

xylem pole pericycle cell differentiation is the J0121 enhancer 

trap that marks xylem pole pericycle cells after they exit from the 

meristematic zone and pass through the elongation zone (Parizot 

et al. 2008). A marker of intervening cells within the pericycle 

tissue layer helped identify phloem pole pericycle cells. These 

cells are marked by expression of S17, a basic leucine zipper 

transcription factor. The function of phloem pole pericycle cells 

has not been determined, nor are there histological markers of 

phloem pole pericycle differentiation. However, phloem pole 

pericycle cells have a very distinct expression pattern and 

underlying transcriptional signature from that of xylem pole 

pericycle cells, as determined by expression profiling of marked 

populations of both of these cells relative to other cell types 

in the Arabidopsis root (Brady et al. 2007). Interestingly, the 

expression pattern of phloem pole pericycle cells more closely 

reflects that of cells in the developing phloem cell lineage. 

No early markers of either phloem or xylem pole pericycle 

cells have been identified, nor have regulatory factors been


found that are both necessary and sufficient for pericycle 

cell specification and differentiation. Cytokinin is sufficient to 

suppress xylem pole pericycle differentiation, as marked by 

the J0121 enhancer trap line, and this, in part, requires AHP6 

(Mähönen et al. 2006). MiR165/166 repression of PHB is also 

required for pericycle differentiation (Miyashima and Nakajima 

2011). In WT roots, AHP6 is expressed in protoxylem cells 

and the two abutting xylem pole pericycle cells (Figure 11B). 

In the scr-3 and phb-1d mutants, AHP6 expression was either 

completely lost or detected in only one of the aforementioned 

three cells. In addition, expression of an additional xylem pole 

pericycle marker gene, STELAR K + OUTWARD RECTIFIER 

(SKOR), was greatly reduced in these phb-1d and scr-3 mutant 

lines. Finally, in response to external auxin, which is sufficient to 

induce periclinal divisions in xylem pole pericyle cells, reduced 

periclinal divisions were observed in the phb-1d mutant. Expression 

of a mutant form of miR165, which is able to target the 

phb-1d miRNA-resistant PHB transcripts, was able to rescue 

the AHP6 and SKOR expression patterns in phb-1d. However, 

lateral root primordium development was somewhat delayed, 

suggesting that SHR/miR165-dependent regulation of PHB is 

required for pericycle function. 

Dynamic regulation of gene expression within the root 

vasculature 

Dynamic gene expression patterns have also been identified 

in the root vasculature, and thus far, oscillatory expression 

patterns have been linked to lateral root initiation. First, oscillations 

in auxin responsiveness, as measured by the DR5:GUS 

synthetic auxin response reporter in xylem pole pericycle cells 

within the root meristematic zone, were shown to temporally 

correlate with lateral root initiation at regular intervals in the maturation 

zone (De Smet et al. 2007). Further work demonstrated 

that this oscillatory auxin responsiveness likely determines 

competence of xylem pole pericycle cells to become lateral 

root primordia (Moreno-Risueno et al. 2010). 

To identify additional factors that play a role in lateral root development 

and which may act at the same time or downstream 

of this fluctuating auxin responsiveness, microarray analysis 

was used to identify genes whose expression oscillates in 

phase with DR5 auxin responsiveness as well as antiphase with 

auxin responsiveness. Many of these oscillating genes were 

shown to regulate prebranch initiation site formation and lateral 

root number (Moreno-Risueno et al. 2010). Numerous other 

genes have been identified that regulate dynamic expression in 

root developmental time. These were obtained by sectioning an 

individual Arabidopsis root into 12 successive sections, each 

representing a specific point in a developmental time (Brady 

et al. 2007). Sections of an independent root served as a 

biological replicate. 

Based on these data, genes were identified whose expression 

fluctuates over root developmental time; e.g., they showed 

peaks of expression in the meristematic zone and maturation 

zone, with downregulation of expression in the meristematic 

zone (Figure 11C). A rigorous statistical method was developed 

to identify cases of dynamic expression between roots 

(Figure 11C), and many of these were expressed specifically in 

root phloem cell types or in xylem pole pericycle cells. Their 

function was inferred to be associated with energy capture 

and lateral root initiation, respectively (Orlando et al. 2010). 

Together, these data indicate that oscillatory, rhythmic and 

fluctuating gene expression within roots and between roots in 

the root vasculature serve to regulate patterning of vascular 

cells, and likely other vascular biological functions. 

Spatio-temporal regulation of root and shoot vascular 

development and connectivity 

Studies on Arabidopsis root mutants defective in cell proliferation 

and cell differentiation may provide insight into possible 

common genetic regulatory mechanisms controlling radial 

vascular development in shoots and leaves. Mutants such as 

wol show decreased cell proliferation in root procambium and 

differentiate completely into protoxylem (Mähönen et al. 2000), 

whereas the apl mutant shows defects in phloem development 

(Bonke et al. 2003). Although these genes appear to affect 

root cells exclusively, recent studies revealed that SHR is also 

involved in regulating cell proliferation in leaves (Dhondt et al. 

2010). Furthermore, expression of SHR is tightly correlated with 

AtHB-8 expression during vascular strand formation (Gardiner 

et al. 2011). While the spatial and temporal patterns of cell 

proliferation during root vascular development are becoming 

clearer, understanding this mechanism in leaves has proved 

to be more challenging. It is likely that combinatorial control 

of both hormones and genetics are in effect, and that these 

components are tightly integrated in both tissue and organ 

(leaf) morphogenesis. Therefore, isolating these developmental 

components will be critical in order to thoroughly understand 

the spatial and temporal control of vascular development in 

leaves. 

Secondary Vascular Development 

The term “secondary growth” refers to the radial growth of 

stems, and is ultimately the result of cell division within a 

lateral meristem, the vascular cambium (Larson 1994). The 

vascular cambium produces daughter cells towards the center 

of the stem which become part of the secondary xylem, or 

wood (Figure 12). The cambium also produces daughter cells 

towards the outside of the stem which become part of the inner 

bark. There are two types of cambial initials: fusiform and ray.

Figure 12. Internal structure of a woody plant stem. 

The vascular cambium consists of a centrifugal layer of fusiform 

secondary phloem and a centripetal layer of secondary xylem cells 

surrounding a central zone comprising phloem and xylem transit 

amplifying cells with a central uniseriate layer of cambial stem cells. 

Most angiosperms and gymnosperm trees species also contain 

radial files of near isodiametric ray cells that play a role in nutrient 

transport and storage (reproduced from Matte Risopatron et al. 

(2010) with permission). 

Fusiform initials give rise to the vertically-oriented cells, including 

water-conducting tracheary elements of the secondary 

xylem, and nutrient- and molecular signaling-conducting SEs of 

the secondary phloem. Ray initials produce procumbent cells 

that serve to transport materials radially in the stem, and likely 

serve storage and other functions that are currently poorly 

defined. 

To produce a functional, woody stem, these and many 

other developmental processes must be coordinated (Du and 


Groover 2010). Importantly, these developmental processes 

are also highly influenced by environmental cues. This is 

evident when observing annual rings in a tree stump, where 

favorable environmental conditions in the spring can lead to 

rapid growth and production of wood with anatomical and 

chemical differences from wood produced under draught and 

less favorable conditions later in the growing season. Another 

notable example of how environmental cues influence secondary 

growth is the formation of reaction wood in response 

to gravity and mechanical stresses, with reaction wood serving 

to right bent stems or to support horizontal branches (Du and 

Yamamoto 2007). 

In this section of the review, we will highlight some of the more 

recent advances in the understanding of how secondary growth 

is regulated. This is an exciting period in the study of secondary 

growth, as genomic approaches applied to a number of species 

have yielded comprehensive lists of the genes expressed in 

the cambium and secondary vascular tissues. Additionally, the 

model forest tree genus Populus now has a fully sequenced 

genome (Tuskan et al. 2006), which, when paired with relatively 

efficient transformation systems for some Populus genotypes, 

has allowed detailed functional characterization of a modest 

number of regulatory genes. 

One emerging theme from these studies is that at least some 

of the major regulatory genes and mechanisms that regulate 

the cambium and secondary vascular development have been 

either directly co-opted from the shoot apical meristem, or else 

represent genes derived from duplication of an ancestral shoot 

apical meristem regulator (Spicer and Groover 2010). Thus, 

the study of the cambium and secondary growth also presents 

opportunities to understand the evolution of meristems and 

details as to how regulatory modules of genes can be reused 

and repurposed during plant evolution. Furthermore, since 

secondary growth in angiosperms, and perhaps in both angiosperms 

and gymnosperms, is likely homologous, advances 

in our understanding within model species like Populus can 

potentially greatly accelerate our understanding of secondary 

growth in less tractable species. 

The many values of secondary woody growth 

To better understand the importance of the fundamental processes 

involved in secondary growth, it is worthwhile to first 

gain an understanding of the practical reasons of how research 

of secondary growth is important to ecosystems, societies and 

industries. The wood produced by forest trees during secondary 

growth represents durable sequestration of the greenhouse gas 

CO2, which is ultimately incorporated into wood as byproducts 

of photosynthesis. Wood is primarily composed of fiber and 

tracheary element cells which undergo complex processes of 

differentiation in which they synthesize thickened secondary 

cell walls before undergoing PCD to produce cell corpses.


Fibers impart mechanical strength to wood, whereas tracheary 

elements provide both mechanical strength as well as water 

conduction. Lignin and cellulose are primary components of 

secondary cell walls, and thus of wood, and impart mechanical 

strength and resistance to degradation. Together, cellulose and 

lignin are the most abundant biopolymers on the planet. 

The secondary cell walls in wood ultimately reflect storage 

of energy and CO2 derived from photosynthesis. With regards 

to carbon sequestration, forests are second only to the oceans 

in the biological sequestration of carbon, and are thus central 

to the carbon cycle and to mediating the levels of atmospheric 

CO2. The energy stored in wood has played central roles in 

history by providing heating and cooking fuels, and continues 

to play these vital roles in developing countries today (Salim 

and Ullsten 1999; FAO 2008). Looking to the future, the woody 

tissues of trees are increasingly of interest as a net-carbon 

neutral source of bioenergy (FAO 2008). While wood wastes 

have long played roles in cogeneration plants to supplement 

coal or to provide energy to forest industry mills, more recently 

woody biomass is being utilized as a “next generation” biofuel. 

Wood from trees can be utilized as a feedstock to produce 

ethanol, syngas, or other biofuels. In addition, similar to a 

petroleum refinery, the utility and economics of biorefineries 

will be bolstered by production of value added products such 

as acetic acid and other chemicals (Naik et al. 2010). Fast 

growing woody perennial crops, including forest trees such 

Populus, are beginning to be utilized as a source of biofuels 

on industrial scales (Sannigrahi et al. 2010). 

Wood is also used to produce pulp, paper, lumber, and 

countless derived forest products. Forest products have been 

estimated to represent three percent of the total world trade 

(FAO 2009). Harvesting and processing of wood products, as 

well as related follow-on industries, represent vital components 

of rural economies in forested regions worldwide, where few 

alternative industries exist. Importantly, forests and the woody 

bodies of trees provide numerous ecosystem services to which 

it is difficult to ascribe an economic value (Salim and Ullsten 

1999; FAO 2009). Forests provide unique habitats, underpin 

crucial ecosystems, provide clean water, and are the focus of 

tourism industries worldwide. Perhaps the most difficult to value 

are the aesthetic, cultural, and spiritual aspects of forests, trees, 

and wood. In short, wood produced by forests is central to the 

health of our planet and society. 

The biology and regulation of secondary growth 

Secondary growth represents the culmination of a number 

of fascinating developmental processes. Currently, there are 

mechanisms identified and partially characterized that regulate 

specific developmental processes, including cambium initiation 

and maintenance, tissue patterning, and the balance of cell 

division and cell differentiation. While our understanding of 

secondary growth is far from complete, past and ongoing 

genomic studies have provided exhaustive lists of all the genes 

expressed during secondary growth. Molecular genetics studies 

have provided insights into the function of a modest number 

of regulatory genes, primarily encoding transcription factors, 

signaling peptides, and receptors. Plant growth regulators, 

including cytokinin, ethylene, gibberellic acid and most notably 

auxin, have all been implicated in influencing some aspect of 

secondary growth. 

Here, we provide a brief analysis of some of the mechanisms 

identified that regulate secondary growth. First, we will focus 

on auxin, as it has profound influences on secondary growth. 

Auxin has long been known to be a critical regulator of cambium 

functions and secondary growth. For example, exogenous 

auxin applied to decapitated shoots can stimulate cambial 

formation and activity (Snow 1935). It is generally assumed 

that auxin is produced in leaves and apical meristems, and 

transported down the cambium to stimulate growth. However, 

direct determinations of the actual routes of auxin transport in 

the stem and the relative amount of auxin synthesized in the 

stem are currently lacking. Indeed, looking at trees like the giant 

sequoia in which the canopy foliage can be a hundred feet from 

the active cambium at the base of the stem, bring into question 

models for auxin synthesis and transport that were developed 

in smaller and more tractable plant species. 

The role of auxin transport and auxin gradients during secondary 

growth have been researched directly in forest trees 

(Uggla et al. 1996; Schrader et al. 2003; Kramer et al. 2008; 

Nilsson et al. 2008), but remain inconclusive. A radial auxin gradient 

is present across secondary vascular tissues and peaks 

in the region of the cambium and nascent secondary xylem in 

both angiosperm and gymnosperm trees (Uggla et al. 1996; 

Tuominen et al. 1997). This observation spurred speculation 

that auxin could create a radial morphogen gradient, but that 

concept has been brought into question by studies showing 

that few genes that are auxin-responsive actually show peaks 

of expression in the cambial zone (Nilsson et al. 2008). 

Auxin has also been shown to be transported basipitally 

in the stem, and to be involved with polarity determination 

in secondary vascular tissues (Kramer et al. 2008). Genes 

encoding PIN-type auxin efflux carriers are preferentially expressed 

in the cambial zone and developing xylem in the 

radial gradient, and in the apical-basal gradient they show a 

sharp peak of expression in internodes that are transitioning to 

secondary growth (Schrader et al. 2003). Currently, the subcellular 

localization and function of PIN transporters is largely 

uncharacterized in stems, and the functional significance of 

the auxin gradients remains contentious (Nilsson et al. 2008). 

Thus, auxin is a central point for important new advances in the 

study of secondary growth. 

Indirect observations suggest that cell-cell communication 

might play important roles in secondary growth, and studies

are beginning to reveal important mechanisms in how the 

balance of cell differentiation and cell division is regulated, and 

how tissue identity is maintained across layers of secondary 

vascular tissues. Recent studies in Arabidopsis and Populus 

have identified mechanisms by which the balance of cell differentiation 

and tissue identity are established through cell-cell 

signaling. As previously discussed, in Arabidopsis, TDIFisa 

small peptide product of the phloem-expressed CLE41/44 gene 

(Ito et al. 2006). TDIF is secreted from the phloem, and acts to 

inhibit tracheary element differentiation and stimulate cambial 

activity (Ito et al. 2006; Hirakawa et al. 2008; Etchells and 

Turner 2010), as well as to regulate the orientation of cambial 

divisions (Etchells and Turner 2010). This peptide is perceived 

by the LRR-Receptor kinase Phloem Intercalated with Xylem 

(PXY), which is expressed in the procambium (Hirakawa et al. 

2008; Hirakawa et al. 2010). Loss-of-function pxy mutants 

show phloem cells intermixed in the xylem (Fisher and Turner 

2007). TRIF/PXY signaling activates WOX4 (Hirakawa et al. 

2010), which encodes a transcription factor that presumably 

influences gene expression associated with meristematic cell 

fate. Interestingly, stimulation of cambial activity by auxin 

requires functional WOX4 and PXY (Suer et al. 2011), providing 

insight into how auxin integrates into transcriptional regulation 

in secondary vascular tissues. 

Patterning and polarity in secondary vascular tissues 

Cross sections of a typical woody stem show that secondary 

vascular tissues are highly patterned (Figure 12), and that the 

proper position and patterning of the cambium, secondary 

xylem, and secondary phloem are crucial to the function of 

these tissues. Additionally, secondary vascular tissues can 

be described in terms of polarity, analogous to vasculature in 

leaves. Take for example the vasculature of a typical dicot tree, 

poplar, in which the vascular bundles in leaves always have 

xylem towards the adaxial and phloem towards the abaxial 

surface of the leaf. By following those vascular bundles through 

the leaf trace and into the stem, it becomes apparent that 

the same polarity relationships are found in both primary and 

secondary vascular tissues, with xylem towards the center and 

phloem towards the outside of the stem. 

Insights into how polarity is established and maintained 

in vascular tissues has been provided by pioneering studies 

of the Class III HD-ZIP and KANADI transcription factors in 

Arabidopsis. These Class III HD-ZIPs are highly conserved 

in plants, act antagonistically with KANADIs, and have been 

shown to regulate fundamental aspects of meristem function, 

polarity, and vascular development (Emery et al. 2003; Izhaki 

and Bowman 2007; Bowman and Floyd 2008). In Arabidopsis, 

REV is implicated in various developmental processes, 

including patterning of primary vascular bundles (Emery et al. 

2003; Bowman and Floyd 2008). A recent study showed that 


misexpression of a Populus REV ortholog results in formation 

of ectopic cambia in the cortex of the stem, and that these 

cambia can produce secondary xylem with reversed polarity 

(Robischon et al. 2011), indicating that the Class III HD-ZIPs 

also affect patterning and polarity in secondary vasculature. 

Genetics and genomics: critical tools for advancing 

knowledge on woody growth 

Research on secondary growth is at an exciting point, as genomic 

tools are now allowing the characterization of the genetic 

variation within species that is responsible for wood quality and 

growth traits. Association mapping is being taken to a wholegenome 

scale for some tree species, and will undoubtedly 

provide fascinating insights into macro- and micro-evolution 

of wood formation (Neale and Kremer 2011). Genomic tools 

are also now enabling the first generation of network biology 

approaches in the model tree genus Populus (Street et al. 

2011) that can be used in understanding woody growth. Such 

approaches utilize a variety of genomic data types to model 

the genetic networks that regulate specific aspects of woody 

growth, and can ultimately produce a “wiring diagram” of regulatory 

networks. This will be important for both better directing 

future research and for providing predictive models that can 

potentially be used to better direct breeding programs, identify 

regulatory genes for biotechnology, and provide insights into 

the complexities of biological processes fundamental to the 

future of forests worldwide. 

Physical and Physiological Constraints 

on Phloem Transport Function 

We now turn our attention to an examination of the constraints 

of photoassimilate transport in the most evolutionarily 

advanced plants, the angiosperms. Here, photoassimilate 

conducting units are comprised of SEs arranged end-to-end to 

form conduits that are referred to as sieve tubes. At maturity, 

SEs lack nuclei and vacuoles, and their parietal cytoplasm 

has a greatly reduced number of organelles. In contrast to 

xylem tracheary elements, SEs retain their semi-permeable 

plasma membrane. Their shared end walls contain interconnecting 

pores (sieve plate pores) formed from PD coalescing 

within pit fields (Evert 2006). In addition, each SE is highly 

interconnected symplasmically with a metabolically active CC, 

through specialized PD, to form a functional unit referred to as 

the SE-CC complex. 

In order to provide a framework on which to identify the 

physical and physiological constraints regulating phloem transport, 

we must first examine the physical mechanisms responsible 

for resource transport through the sieve tube system. 

As photoassimilate flow is polarized from source leaves (net 

exporters of resources) to heterotrophic sinks (net importers


of resources), phloem transport can be envisaged in terms 

of three key physiological components arranged in series. 

These components are: (a) loading of photosynthate (most 

commonly sucrose) in collection phloem (minor veins) within 

source leaves, (b) long-distance delivery in transport phloem, 

and (c) unloading from release phloem (Figure 13A). Principles 

of flux control analysis dictate that each component will confer 

an influence (constraint) on overall flow, from source to sink; 

thus, the question of constraint becomes one of degree. 

Bulk flow identifies the regulatory elements 

The phloem is generally buried deep within plant tissues, and 

this location, along with its sensitivity to mechanical perturbation, 

has made it technically challenging to observe flows 

through sieve tubes, non-invasively and in real-time. However, 

there is a critical body of experimental evidence showing that 

solutes and water move at similar velocities through sieve tubes 

(van Bel and Hafke 2005; Windt et al. 2006). These studies 

suggest that the phloem translocation stream moves by bulk 

flow, consistent with the now widely accepted pressure flow 

hypothesis put forward originally by Münch (1930). 

On the premise that transport through sieve tubes conforms 

to bulk flow, then the transport rate (Rf ) of a nutrient species is 

given by the product of transport velocity (V), sieve tube crosssectional 

area (A) and phloem sap concentration (C), whereby: 

Rf = V · A · C (1) 

Both A and C are finite elements, whereas, given that flow 

through sieve tubes approximates laminar flow through capillaries, 

the factors determining transport velocity (V), or solvent 

volume flux (Jv, having units of m 3 m −2 s −1 ,orms −1 ), are 

identified by the Hagen-Poiseuille Law as: 

Jv = Pπr 2 /8ηl (2) 

Here, P is the hydrostatic pressure difference between either 

end of each sieve tube of length (l), the translocation stream 

has a viscosity (η), and it moves though sieve tubes of known 

radii (r) that have sub-structural elements that serve to impede 

transport; i.e., sieve plate pores and parietal cytoplasm 

(Mullendore et al. 2010). 

The water potential (ψw) of any cell (e.g., a SE-CC complex) 

is given by: 

ψw = ψP + ψπ 

where ψP is the pressure potential and ψπ is the osmotic or 

solute potential within the cell of interest. The value of ψP within 

a cell is determined by the magnitude of ψπ and the ψw of the 

cell wall (apoplasmic potential). As ψπ in the wall is generally 

close to zero, and given that the cell is in quasi water potential 

equilibrium with its wall (i.e., the values of ψw for the cell and 

its surrounding wall [apoplasm] are close to being equal), then 

(3) 

the value of ψP in the cell (CC-SE) is given by: 

ψP = ψw − ψπ 

Armed with the above information, we can identify key elements 

that may constrain rates of phloem transport. 

Sieve element osmotic potentials are determined 

by phloem loading/retrieval 

The ψπ of the SE content (generally termed phloem sap) is 

primarily (60% to 75%) determined by one of several sugar 

species (sucrose, polyol or a raffinose family oligosaccharides 

– RFOs) with K + and the accompanying anions accounting for 

most of the remaining osmotic potential (Turgeon and Wolf 

2009). Thus, loading/retrieval of sugars plays a key role in 

setting the SE hydrostatic pressure (Equation 4), and, for this 

reason, we shall primarily focus attention on sugar transport, 

but where appropriate, we will comment on the involvement of 

other solutes. 

Proposed phloem-loading mechanisms are based on thermodynamic 

considerations and cellular pathways of loading. 

The most intensively studied is the apoplasmic loading mechanism 

in which sucrose (or polyol) loading requires the direct 

input of metabolic energy (Figure 13B, I), and is widespread 

amongst monocot and herbaceous eudicots species. An 

energy-dependent symplasmic loading mechanism also has 

been described. Here, sugar (sucrose or polyol) diffuses down 

its concentration gradient through a symplasmic route from 

mesophyll cells to specialized CCs, termed intermediary cells 

(ICs), where biochemical energy is required for sucrose/polyol 

conversion to large RFOs. This loading mechanism is referred 

to as the polymer trap mechanism (Figure 13B, II) and is thought 

to operate predominantly in herbaceous eudicots. 

Another loading system has been suggested to operate in 

woody plants (Davidson et al. 2011; Liesche and Schulz 2012) 

in which sugars are passively loaded through diffusion, driven 

by high sugar concentrations maintained in the mesophyll 

cell cytosol (Rennie and Turgeon 2009; Turgeon 2010a). This 

pathway is considered to be symplasmic, based on observed 

high PD densities between each cellular interface from mesophyll 

to SE-CC complexes (Figure 13B, III). It has also been 

suggested that delivery of sugars into SE-CC complexes could 

be achieved by bulk flow operating through interconnecting PD 

(Voitsekhovskaja et al. 2006). 

Several caveats must be considered for both the symplasmic 

diffusion and bulk flow models for passive phloem loading. 

If PD were to allow diffusion of sugars from mesophyll cells 

into sieve tubes, then all similarly-sized metabolites and ions 

should also pass into SE-CC complexes; i.e., the system would 

lack specificity. For situations in which bulk flow might transport 

sucrose, again all other soluble constituents present within the 

cytosol of cells forming the loading pathway should also gain 

(4)

Figure 13. Diagrammatic representation of resource flow through the phloem pathway. 


(A) Overall flow of resources through the phloem pathway. Within source leaves, resources (nutrients – green arrows and water – blue 

arrows) are loaded into sieve tubes (ST) of the collection phloem (chalk) formed by a network of minor veins. These loaded solutes lower ψπ 

in the ST that then causes water to move, by osmosis, across the ST semi-permeable plasma membrane resulting in a high ψPsl condition. 

This osmotically-driven increase in ψP serves as the thermodynamic driving force to drive bulk flow (green highlighted blue arrows) of ST sap 

throughout the phloem system. In this context, loaded resources flow from collection phloem STs into STs of lower order veins, functioning as 

transport phloem (light blue) for export from source leaves. Transport phloem supports long-distance axial transport of resources from source 

leaves to sinks through STs of exceptionally high hydraulic conductivities that homeostatically sustain their ψP by resource exchange with 

surrounding tissues (curved green superimposed on blue arrows). Upon reaching the release phloem (light mauve), resources are unloaded 

from STs by bulk flow through plasmodesmata (PD) that interconnect the surrounding cells. The difference in pressure potential between the 

source and sink (ψPsl – ψPs) represent the hydrostatic pressure differential that drives bulk flow through the phloem pathway from source to 

sink. 

(B) Phloem loading pathways and mechanisms within source leaves. Photosynthetically reduced carbon, generated in chloroplasts, is used 

to drive sucrose (Suc) or polyol (Poly) biosynthesis (green broken arrows) within the cytosol of mesophyll cells (MC). Excess Suc/Poly is 

transiently stored in vacuoles (V) of mesophyl cells and, along with carbon from remobilized chloroplastic starch grains (SG), buffers their 

cytosolic pool sizes available for phloem loading. Suc/poly (green arrows) moves from mesophyll cells along a phloem-loading pathway that 

includes bundle sheath cells (BSC), phloem parenchyma cells (PPC), companion cells (CC) or intermediary cells (IC) to finally enter sieve 

elements (SE) of the collection phloem. Three loading mechanisms are considered to function in different species. (I) Active apoplasmic 

loading: Suc (and/or Poly) is first released from phloem parenchyma cells (PPCs) by the action of a permease (Chen et al. 2012b), and 

subsequently retrieved into SE-CC complexes by symporters located along SE-CC plasma membranes. (II) Symplasmic loading: Raffinose 

oligosaccharides (RFOs) are synthesised in specialized CCs, termed ICs, from Suc delivered symplasmically from MCs. The larger molecular


entry into the phloem. When operating over a prolonged period, 

either of these proposed passive-loading systems would be 

anticipated to cause a perturbation to metabolism within the 

mesophyll cells. 

Rates of phloem loading depend upon the pool size of each 

transported solute available for loading, as well as the loading 

and retrieval mechanisms. For sugars, sucrose (Grodzinski 

et al. 1998) and polyol (Teo et al. 2006) pools are generated 

in mesophyll cells, whereas RFOs are synthesized from 

sucrose that enters the specialized ICs (Turgeon and Wolf 

2009)(Figure 13B, II). Irrespective of sugar species and phloem 

loading mechanism, during the photoperiod, transported sugars 

arise from current photosynthesis and export rates are 

linked positively with the sugar pool size (Grodzinski et al. 

1998; Leonardos et al. 2006; Lundmark et al. 2006). During 

the night, sugar pools are fed by starch reserves remobilized 

from chloroplasts (Smith and Stitt 2007) and sugars released 

from vacuolar storage in mesophyll cells (Eom et al. 2011) 

(Figure 13B). Depending upon carbon gain by leaf storage pools 

during the preceding photoperiod, remobilizing reserves during 

the night can sustain sugar pool sizes and, hence, export rates 

(Grimmer and Komor 1999). 

In situations where source leaves are operating at suboptimal 

photosynthetic activity, analyses of metabolic control 

have provided estimates that source leaf metabolism exercised 

approximately 80% of the control exerted over photoassimilates 

transported into developing potato tubers (Sweetlove et al. 

1998). However, the relationship between leaf metabolism and 

export rates also depends upon prevailing source/sink ratios. 

This can be illustrated by studies aimed at investigating effects 

associated with CO2 enrichment. Under conditions of source 

limitation, leaf photosynthetic rates are increased substantially 

by CO2 enrichment, and are matched proportionately by those 

of photoassimilate export (Farrar and Jones 2000). In contrast, 

more attenuated responses of leaf photosynthetic rates are 

elicited by CO2 enrichment under sink limitation, and these are 

not proportionately matched by export (Grodzinski et al. 1998; 

Grimmer and Komor 1999). The latter response suggests that, 

under sink limitation, predominant control of photoassimilate 

transport shifts to processes downstream of source leaf sugar 

metabolism. 

Estimates of membrane fluxes of sucrose loaded into SE- 

CC complexes in sugar beet leaves fall into the maximal 

range for plasma membrane transporter activity (Giaquinta 

1983). Therefore, if sucrose transporters are indeed operating 

at maximum capacity, then their overexpression might 

be expected to result in enhanced rates of phloem loading 

and photoassimilate export. However, overexpression of the 

spinach sucrose transporter (SoSUT1) in potato, while altering 

leaf metabolism, exerted no impact on biomass gain by the 

tubers (Leggewie et al. 2003). This finding indicates an absence 

of any constraint imposed by endogenous sucrose transporters 

on phloem loading. Indeed, phloem loading can respond quite 

rapidly (within minutes) to changes in sink demand (Lalonde 

et al. 2003). 

A striking example of the dynamic range available to the 

phloem loading system is shown by studies performed on 

Ricinus, a plant whose phloem sap will exude (bleed) from 

severed SE-CC complexes. Here, excisions made in Ricinus 

stems reduced ψP to zero in this region of the sieve tube 

system. This treatment resulted in exudation of phloem sap 

from the severed sieve tubes and an increase in translocation 

and, hence, phloem-loading rates of sucrose, by an order of 

magnitude (Smith and Milburn 1980a). This observed rapid 

response is envisaged to reflect signaling from the sink region 

(in this case, the site of SE-CC stem excision) to source leaves. 

Here, pressure-concentration waves transmitted through interconnecting 

sieve tubes (Mencuccini and Hölttä 2010) could act 

to regulate transporter activity mediating phloem loading (Smith 

and Milburn 1980b; Ransom-Hodgkins et al. 2003). 

←−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−− 

sizes of RFOs are thought to prevent their backward diffusion through PD interconnecting ICs with PPCs; however, the dilated PD that 

interconnect the IC-SE complexes permit forward diffusion of RFOs into the SEs (polymer trap model). (III) Passive – symplasmic loading: 

Suc or Poly is proposed to move by diffusion through the symplasm, via PD, down their concentration gradients from MCs to SEs. For all 

phloem-loading mechanisms, water enters (curved blue arrows) SE/CC or IC complexes through aquaporins (paired khaki ovals) (Fraysse 

et al. 2005). 

(C) Phloem unloading pathways from release phloem in sink organs. In all sinks, it is highly probable that imported resources are unloaded 

symplasmically by bulk flow from release phloem SE-CC complexes (green-highlighted blue arrows) into adjacent PPCs. Onward resource 

movement through the phloem-unloading pathway may occur by the following pathways. (I) Continuous symplasmic unloading: here, 

resources (green-highlighted blue arrows) likely continue to move by diffusion through PD into surrounding phloem parenchyma (PC) 

and ultimately sink cells (SC). (II) Apoplasmic step unloading: here, the phloem-unloading route involves resource transit through the sink 

apoplasm due to a symplasmic discontinuity in unloading pathways at either the PPC/SC or PC/SC interface. Membrane exchange of nutrients 

to, and from, the sink apoplasm occurs by transporter-mediated (khaki circles) membrane efflux and influx mechanisms, respectively. In both 

cases, water exiting SE-CC complexes can enter SCs (in the case of growing sinks) or, for non-expanding storage sinks, water returns to 

the xylem transpiration stream by exiting PPC/PCs (blue curved arrows) to the sink apoplasm through aquaporins (paired khaki ovals).

Other layers of post-translational control of sucrose transporter 

activity include protein-protein interactions, e.g., SUT1- 

SUT4 regulation of phloem loading (Chincinska et al. 2008), 

redox-induced dimerization of SUT1 (Krügel et al. 2008), and 

cytochrome b5 interaction with MdSUT1 and MdSOT6 (Fan 

et al. 2009). Interestingly, the question as to whether symplasmic 

loading species also have the capacity for short-term 

adjustments in phloem loading capacity is less certain (Amiard 

et al. 2005). 

Magnetic resonance imaging studies, conducted on longdistance 

transport of water through the vascular system in 

a range of species, have established that phloem transport 

remains unaffected by diurnal variations in transpiration-driven 

changes in apoplasmic leaf water potential (Windt et al. 2006). 

Thus, a regulatory mechanism must operate to maintain a 

constant pressure gradient (ψP) to drive bulk flow through 

sieve tubes. This likely involves osmoregulatory activities 

at the level of the SE-CC complex (Pommerrenig et al. 

2007). 

In general, it would appear that phloem loading of major 

osmotic species (sugars and K + ) does not constrain phloem 

transport under optimal growth conditions. During periods of 

abiotic stress, phloem loading can minimize the impact of 

water/salt stress through osmoregulatory activities of cells 

comprising the phloem-loading pathway(s) (Koroleva et al. 

2002; Pommerrenig et al. 2007). Interestingly, and perhaps 

surprisingly, both apoplasmic (Wardlaw and Bagnall 1981) and 

symplasmic (Hoffman-Thom et al. 2001) loaders undergo maintenance 

of phloem loading activities in cold-adapted plants. 

This indicates that changes in the viscosity of the phloem 

translocation stream may have little impact on bulk flow through 

sieve tubes. In contrast, elevated temperatures can slow 

translocation by callose occlusion of sieve pores (Milburn and 

Kallarackal 1989). In addition, deficiencies of K + and Mg 2+ can 

impact apoplasmic loading of sucrose into SE-CC complexes 

(Hermans et al. 2006). In the case of K + , this is thought to reflect 

a limitation in charge compensation across the SE-CC plasma 

membrane which could impede the operation of the sucrose-H + 

symport system (Deeken et al. 2002), whereas Mg 2+ deficiency 

could lower the availability of Mg 2+ -ATP which serves as 

substrate for the H + -ATPase that generates the proton motive 

force to power the sucrose H + symporter (Cakmak and Kirkby 

2008). 

In terms of minor osmotic species, direct control of their 

phloem translocation rates is determined entirely by the concentrations 

to which they accumulate in SE-CC complexes. 

This situation is nicely illustrated by studies performed on transgenic 

peas expressing a yeast S-methylmethione transporter 

under the control of the phloem-specific AtAAP1 promoter. 

Here, S-methylmethione levels in developing seeds were found 

to be proportional to their concentrations detected in phloem 

exudates (Tan et al. 2010). 


Mechanisms of phloem unloading 

The cellular pathway of phloem unloading may extend, functionally, 

from SE lumens of the release phloem to sites of 

nutrient utilization/storage in the particular sink organ/tissue 

(Lalonde et al. 2003; Figure 13C). Within these bounds, the 

cellular pathways followed circumscribe the physical conditions 

under which an unloading mechanism operates. Most sink 

systems investigated to date have PD interconnecting the 

SE-CC complex to cells of the surrounding ground tissues, 

and, thus, confer the potential for universal symplasmic unloading 

(Figure 13C, I). In general, such routes for symplasmic 

unloading have low densities of PD that interconnect 

SE-CCs with adjacent phloem parenchyma cells. Thus, a 

marked bottleneck for symplasmic nutrient delivery may exist 

at this cellular interface. 

Unloading routes in a variety of sink systems have 

been mapped by using membrane-impermeant fluorochromes 

loaded into phloem of source leaves. Upon import into the 

release phloem zone, fluorochrome movement can be retained 

within the vascular system of fleshy fruit during their major 

phase of sugar accumulation (e.g., apple, a sorbitol transporter 

(Zhang et al. 2004), grape berry, a sucrose transporter (Zhang 

et al. 2006), and cucumber, an RFO transporter (Hu et al. 

2011). However, more commonly, the fluorochrome moves 

symplasmically out from the phloem into surrounding ground 

tissues (Figure 13C, I) as found for root and shoot meristems 

(Stadler et al. 2005), expanding leaves (Stadler et al. 2005), 

young fruit prior to their major phase of sugar accumulation 

(Zhang et al. 2006), and developing seeds in which movement 

is restricted to maternal tissues (Zhang et al. 2007). 

In developing fruits, during the phase of sugar accumulation, 

SE-CC complexes are thought to be the site of sucrose release 

into the fruit apoplasm (Zhang et al. 2004, 2006; Hu et al. 2011). 

Studies conducted on tomato fruit have indicated that the 

cumulative membrane surface area of SE-CC complexes would 

be barely adequate to support sucrose unloading at maximal 

fluxes known to be associated with membrane transport. In 

contrast, using the range of reported PD-associated fluxes 

(Fisher 2000), it can be shown that PD densities could readily 

accommodate unloading of sucrose into surrounding phloem 

parenchyma cells (Figure 13C, II). Clearly, further studies are 

required to resolve whether or not phloem unloading universally 

includes a symplasmic passage from SE-CC complexes to 

phloem parenchyma cells in the release phloem zone, as found 

for developing seeds (Zhang et al. 2007) (Figure 13C, II). 

An obvious constraint on facilitated apoplasmic unloading 

from SE-CC complexes is a co-requirement for a hydrolysable 

transported sugar (e.g., sucrose or RFO) and an invertase 

present within the cell walls of the release phloem zone. This 

combination ensures maintenance of an outwardly directed diffusion 

gradient for the transported sugar, due to its conversion


into a different chemical species by the cell wall invertase. 

Some fleshy fruits (Zhang et al. 2006; Hu et al. 2011) and 

pre-storage phase seeds (Zhang et al. 2007) satisfy these 

requirements. However, these features do not apply to plants 

that transport polyol through their phloem, as exemplified by 

apple and temperate seeds during their storage phase; these 

systems may well rely on energy-coupled sugar release to the 

seed apoplasmic space (Zhang et al. 2004, 2007) (Figure 13C, 

II). In this context, maximal activities of symporters retrieving 

sucrose from seed apoplasmic spaces into pea cotyledons or 

wheat endosperm cells clearly constrain phloem unloading, 

as shown by increases in seed dry weight of transgenic 

plants overexpressing these transporters (Rosche et al. 2002; 

Weichert et al. 2010) (Figure 13C, II). 

A less obvious constraint, but an essential component of an 

unloading mechanism, is that unloading rates of all solutes (and 

particularly the major osmotic species) and water must match 

their rates of phloem import. Any mismatch will impact water relations 

of release phloem sieve tubes and, hence, translocation 

rates. This requirement is essentially irreconcilable if significant 

leakage were to occur by diffusion. To ensure matching rates 

of phloem import and membrane efflux, it is important to stress 

that membrane transport of all solutes and water needs to be 

facilitated. This allows activities of membrane transporters to be 

potentially coordinated to ensure that rates of resource import 

through, and unloading from, sieve tubes in the release phloem 

are matched (see Zhang et al. 2007). 

A simple resolution to this problem is for unloading from the 

SE-CC complex to follow a symplasmic route and occur by 

bulk flow. Currently, experimental support for bulk flow, as an 

unloading mechanism, is limited. Such evidence is derived from 

experiments in which hydrostatic pressure differences between 

SE-CC complexes and surrounding cells were manipulated at 

root tips (e.g., Gould et al. 2004b; Pritchard et al. 2004). However, 

given that PD may represent a low hydraulic conductivity 

pathway, a significant pressure differential would be required to 

ensure the operation of an effective bulk flow delivery system. 

Consistent with this prediction, large osmotic potential differences 

(−0.7 MPa to −1.3 MPa) between SEs of release 

phloem and downstream cells have been measured in symplasmic 

unloading pathways of root tips (Warmbrodt 1987; 

Pritchard 1996; Gould et al. 2004a) and developing seeds 

(Fisher and Cash-Clark 2000b). These differences translate 

into equally large differences in hydrostatic pressures, since 

sink apoplasmic ψP values approach zero (see Lalonde et al. 

2003). Expulsion of sieve tube contents by bulk flow into the 

much larger cell volumes of phloem parenchyma cells will 

dissipate the hydrostatic pressure of the expelled phloem sap 

within these cells. This, together with frictional drag imposed 

by a low hydraulic conductivity PD pathway, dictates that 

the large differentials in hydrostatic pressure between SE- 

CC complexes and phloem parenchyma cells are the result 

rather than the cause of bulk flow (Fisher and Cash-Clark 

2000b). 

The above considerations draw attention to the possibility 

that hydraulic conductivities of PD linking SE-CC complexes 

with phloem parenchyma cells play a significant role in regulating 

phloem unloading. Interestingly, size exclusion limits of 

PD at these cellular boundaries appear to be unusually high 

in roots (60 kDa; Stadler et al. 2005), sink leaves (50 kDa; 

Stadler et al. 2005) and developing seeds (400 kDa; Fisher and 

Cash-Clarke 2000a), compared to the frequently reported value 

of 0.8 kDa −1.0 kDa for PD linking various cell types in ground 

tissues (Fisher 2000). However, size exclusion limits based on 

molecular weight can be misleading, and Stokes radius is a 

preferable measure (Fisher and Cash-Clarke 2000a). Taking 

this into account, PD hydraulic conductivities computed on this 

basis appear to be sufficient to accommodate the required 

rates of bulk flow out from the SE-CC complex (Fisher and 

Cash-Clark 2000b). 

Large PD conductivities offer scope for considerable control 

over bulk flow across the SE-CC complex and adjoining phloem 

parenchyma cellular interfaces. A hint that such a system may 

operate is illustrated by studies on developing wheat grains. 

Here, imposition of a pharmacological block on sucrose uptake 

into the endosperm of an attached wheat grain was not accompanied 

by a change in sucrose concentration in cells forming 

the unloading pathway within maternal grain tissue (Fisher 

and Wang 1995). This finding points to a direct link between 

sucrose uptake by the endosperm and PD conductance at SE- 

CC-phloem parenchyma cell interfaces. How PD gating could 

be linked with sink demand has yet to be determined, but this 

would have significant implications for phloem translocation. 

Phloem-imported water drives cell expansion in growing 

sinks (Walter et al. 2009). However, in non-expanding storage 

sinks, phloem-imported water is recycled by the xylem back to 

the parent plant body (Choat et al. 2009). This necessitates 

water exit across plasma membranes, irrespective of the unloading 

pathway, and likely depends upon movement facilitated 

by aquaporins (Zhou et al. 2007). Thus, except for symplasmic 

unloading into growing sinks, aquaporins could play a vital role 

in constraining rates of phloem unloading and, hence, overall 

phloem transport (Figure 13C). 

Transport phloem – Far from being a passive conduit 

interconnecting sources and sinks 

Compared to collection and release phloem, transport phloem 

(Figure 13A) extends over considerable distances of up to 100 m 

in tall trees. Axial flows along sieve tubes occur at astonishingly 

high rates, as demonstrated by estimates of specific mass 

transfers of approximately 500 g biomass m −2 sieve tube crosssectional 

area s −1 (Canny 1975). For a time, these observations 

supported the notion that sieve tube cross-sectional areas

(Equation 1) were a major constraint over rates of phloem 

translocation (Canny 1975), a notion later dispelled by the 

finding that specific mass transfer rates could be elevated by 

an order of magnitude in modified plant systems (Passioura 

and Ashford 1974; Smith and Milburn 1980a; Kallarackal and 

Milburn 1984). More recently, a quest to obtain greater insights 

into sieve tube transport function (Thompson 2006) has reignited 

an interest in obtaining measures of sieve tube geometries 

to obtain meaningful estimates of sieve tube hydraulic 

conductivities (Thompson and Wolniak 2008; Mullendore et al. 

2010; Froelich et al. 2011). 

On the assumption that flows through sieve tube lumens 

and sieve pores are laminar, hydraulic conductivity (L) can be 

derived from the Hagen-Poiseulli Law as: 

L = πr 2 /8ηl (5) 

A technically innovative and thorough quantitative plant 

anatomical study, undertaken across a range of eudicot herbaceous 

life forms, yielded estimates of sieve tube hydraulic 

conductivities (Mullendore et al. 2010). Values of L were 

found to be dominated by sieve pore radii, as predicted 

by Equation 5, and were inversely related with independent 

measures of phloem transport velocities. Such an outcome 

is in contradiction to the Hagen-Poiseulli Law. Together 

with other phloem transport anomalies, such as gradients 

of sieve tube hydrostatic pressures not scaling with plant 

size (Turgeon 2010b), these studies point to a key feature 

in phloem translocation likely being overlooked. As outlined 

below, we contend that sieve tube properties that establish conduits 

of exceptionally high hydraulic conductivities, combined 

with their ability to osmoregulate, can account for transport 

phloem being capable of supporting high fluxes over long (m) 

distances. 

As indicated by Equation 5, sieve tube length (l), and most 

importantly, sieve pore radius (r), have a direct influence on 

hydraulic conductivity, along with sap viscosity (η). Sieve-tube 

sap viscosity varies approximately 2.5-fold across the range 

of measured sieve-tube sucrose concentrations (300 mM to 

1000 mM sucrose) and, hence, could influence sieve tube 

hydraulic conductivity. The viscosity of a 600 mM sucrose 

solution increases approximately 2-fold from 25 ◦ C to 0 ◦ C 

(Misra and Varshin 1961). A controlled experiment, in which an 

approximate doubling of phloem sap viscosity can be achieved 

without impacting source or sink activities, is to gradually (to 

avoid shock) cool a stem zone (from 25 ◦ C to just above 0 ◦ C). 

Absence of any slowing of transport rates through the cooled 

zone (Wardlaw 1974; Minchin and Thorpe 1983; Peuke et al. 

2006) argues that this range of phloem sap viscosities exerts 

little influence over phloem transport. 

To date, studies of hydraulic conductivities deduced from 

sieve tube geometries have yielded ambiguous results (Mullendore 

et al. 2010; Froelich et al. 2011). However, indirect 


observations suggest that sieve tube hydraulic conductivity 

is unlikely to constrain phloem transport. For instance, removal 

of substantial proportions of transport phloem crosssectional 

area from the stem had little impact on rates of 

translocation through the narrowed phloem zone (Wardlaw 

and Moncur 1976), thus indicating a considerable spare capacity 

for phloem transport. A spectacular example illustrating 

excess transport capacity is provided by a study of translocation 

rates through pedicels supporting developing apical 

fruits in racemes of Ricinus. Upon removal of apical fruits, 

and allowing exudation from their severed petiole stumps to 

proceed, translocation rates increased from 166 g to 3111 

g biomass m −2 sieve-tube area s −1 , a response suggesting 

that phloem transport was sink controlled, not phloem pathway 

controlled (Smith and Milburn 1980a; Kallarackal and Milburn 

1984). 

Experimental measurements conducted using a microfluidic 

system simulating phloem pressure flow as well as transport 

properties of ‘real’ plants (including tall trees) also yielded 

results that conformed with predictions of the Münch model 

(Jensen et al. 2011, 2012). Studies performed on an Arabidopsis 

mutant lacking P-protein agglomerations in sieve tubes, and 

hence conferring higher sieve tube conductivity, established 

that these plants had similar transport velocities (or volume 

flux – see Equation 2) to WT plants (Froelich et al. 2011). 

Collectively, these studies support the notion that sieve-tube 

hydraulic conductivities do not impose a significant limitation 

on transport fluxes along phloem pathways, even over considerable 

lengths of sieve tubes. Rather, as discussed above, the 

majority of control may well be exercised by bulk flow through 

PD linking SE-CC complexes of release phloem with adjacent 

phloem parenchyma cells (Figure 13C). 

Pressure-concentration waves generated by phloem unloading 

are transmitted over considerable distances (m) at velocities 

an order of magnitude higher than those of phloem translocation 

(Smith and Milburn 1980a; Mencuccini and Hölttä 2010). 

Such a signaling system is envisioned to underpin unified 

responses by all SE-CC complexes, comprising phloem paths 

from release to collection phloem, to altered resource demands 

by the various sinks (Thompson 2006). These responses are 

mediated by turgor-regulated membrane transport of sugars 

into SE-CC complexes; these sugars are supplied from mesophyll 

and axial pools for compensation within collection and 

transport phloem, respectively (Figure 13A). This mechanism 

results in homeostasis of hydrostatic pressure in sieve tubes 

along the phloem pathway (Gould et al. 2004a). The action 

of this pressure-concentration signaling system could account 

for differentials in hydrostatic pressures between collection and 

release phloem not scaling with transport distance, particularly 

in tall trees (Turgeon 2010b). In addition, such a mechanism 

could maintain sieve-tube sap concentrations of all solutes 

(Gould et al 2004a) and, hence, their rates of phloem transport.


The interconnected nature of the plant’s vascular system 

does not appear to exert a constraint over patterns of resource 

flow, as demonstrated by their plasticity in response to changes 

in source/sink ratios (Wardlaw 1990). This leads one to the 

notion that the elements of the phloem function, kinetically, as 

a single pool of resources. A ‘common’ kinetic pool of phloem 

sap depends upon hydraulic connectivity between all functional 

phloem conduits. Lateral sieve areas, phloem anastomoses 

(Evert 2006) and intervening phloem parenchyma cells (Oross 

and Lucas 1985) provide conduits for resource flows to function 

as a common kinetic pool. 

The above described transport behaviors led Don Fisher 

to propose a variant of the Münch pressure flow model in 

which he envisaged phloem systems functioning as highpressure 

manifolds (Fisher 2000). High hydrostatic pressures 

generated by phloem loading in source leaves (Gould et al. 

2005) are maintained throughout the transport phloem by 

osmoregulated loading in collection or re-loading by transport 

phloem (Figure 13). In the region of the release phloem, 

gradients in hydrostatic pressure across PD connecting SE- 

CC complexes to adjoining phloem parenchyma cells, in 

combination with PD hydraulic conductivity, control overall 

flows from source regions to each sink. As a corollary, 

relative magnitudes of PD conductivities between various 

sinks could control resource partitioning at a whole plant 

level. 

The high-pressure manifold model (Fisher 2000) accounts for 

all known aspects of phloem transport, except direct unloading 

across SE-CC plasma membranes. However, as we mentioned 

above, conclusive evidence for this pathway forming a major 

phloem-unloading route has not yet been established. Indeed, 

symplasmic flow into surrounding phloem parenchyma cells 

remains a real possibility in all sinks (Figure 13C) and, hence, 

the high-pressure manifold model appears to be universally 

applicable. 

Directing future studies to testing the phloem 

high-pressure manifold model 

In broad terms, there is strong evidence that the high-pressure 

manifold model (Fisher 2000) accounts for key elements underpinning 

phloem transport and resource partitioning at the 

whole plant level. The model highlights hydraulic conductivities 

of PD linking release phloem SE-CC complexes with 

phloem parenchyma cells as the pivotal point at which phloem 

transport is constrained both physically and physiologically. 

We consider the evidence sufficiently compelling to invest 

significant effort in future investigations to further test the 

general applicability of this model. Resolving the underpinning 

regulatory mechanisms could open up substantial biotechnological 

opportunities to divert biomass flows to enhance crop 

yields. 

Physical & Physiological Constraints on 

Xylem Function 

The xylem of the plant vascular system transports more fluid 

longer distances than any other vascular tissue. The collective 

flow of xylem sap summed over all the plants on a watershed 

can exceed the total runoff in streams (Schlesinger 1997). 

Typically, less than 5% of the xylem water is consumed 

by osmotically-driven cell expansion, and less than 1% is 

consumed by photosynthesis. The bulk of the transported 

water is lost to transpiration: the water evaporates from cell 

wall surfaces into the intercellular air spaces of the leaves, 

and diffuses out into the atmosphere through open stomata. 

Hence, the term “transpiration stream” is used to refer to xylem 

sap flow. Although the transpiration stream carries nutrients, 

molecular signals, and other compounds from roots to leaves, 

and evaporative cooling can minimize overheating of larger 

leaves, these benefits are usually regarded as secondary to the 

cost of having to lose such large quantities of water in exchange 

for stomatal CO2 uptake (Holtta et al. 2011). Under typical 

diffusion gradients, plants transpire hundreds of molecules of 

water for every CO2 molecule fixed by photosynthesis. If plants 

could evolve a way of obtaining CO2 without simultaneously 

losing water, their water consumption would be substantially 

reduced and water would presumably be much less of a limiting 

factor for their productivity. 

As expected for such a poor water-for-carbon exchange 

rate, plants have evolved a metabolically cheap mechanism for 

driving the transpiration stream; otherwise, the cost of moving 

water could easily exceed the meager energy return. According 

to the well-substantiated cohesion-tension mechanism summarized 

in Figure 14, water is pulled to the site of evaporation in 

the leaves by the tension established within the surface of the 

water at the top of the water column (capillary) (Pickard 1981). 

The plant functions more or less as a ‘water wick’. Once the 

‘wick’ is grown, the driving force for the transpiration stream 

is free of charge from the plant’s perspective. Most directly, 

the energy to drive the transpiration stream comes from the 

sun. However, despite its energetic efficiency, the cohesiontension 

mechanism has important limitations that constrain the 

productivity and survival of plants. Current research questions 

include the evolution, physiology, and ecology of these water 

transport constraints. 

The problem of frictional resistance to flow 

The basic wicking process (Figure 14A) presents a physical 

paradox. A narrower tube is better for generating capillary at 

the evaporating meniscus for pulling water up, but it is worse 

for creating high frictional resistance to the upward flow. The 

maximum drop in pressure (Pmin) created by an air-waterinterface 

across a cylindrical pore is inversely proportional to

Figure 14. The cohesion-tension mechanism for transpirationdriven 

xylem flow. 

(A) The basic hydraulic lift process: evaporation from a meniscus 

coupled to bulk liquid flow by capillary action. 

(B) In plants, surface tension created at the surface of narrow pores 

within the cell walls (w) acts as the energy gradient to drive longdistance 

bulk flow through a low-resistance network of dead xylem 

conduits (c). Conduits are connected by pits (p), which also protect 

against air-entry and embolism in the inevitable event of damage 

(see Figures 15, 16). Water is filtered through living cell membranes 

at the root endodermis (en) by reverse osmosis. 

(C) Living cells (rounded rectangles enclosed by hatched cell 

walls) do not generate transpirational flow (ep, leaf epidermis; m, 

mesophyll; cs, Casparian strip of the endodermis; rc, root cortex). 

Water flows through them to the site of evaporation via symplastic (s 

arrows) and apoplastic routes (a arrows). In the root, the apoplastic 

route (a’) is interrupted by the Casparian strip. Transpiration is 

actively regulated by stomatal opening through which the waterfor-carbon 

exchange occurs (from Sperry 2011). 

the cylinder radius (Pmin proportional to 1/radius), whereas the 

hydraulic resistivity (resistance per unit length) through the 

cylinder increases with 1/radius 4 . The higher the resistivity, 

the lower the flow rate at which P drops to Pmin, pulling 

the meniscus down the tube and drying out the wick. The 

evaporating menisci of the plant are held in the nanometerscale 

pores of primary walls facing internal (intercellular) air 

spaces. Although ideal for generating a potentially substantial 

driving force for pulling up the transpiration stream, the high 

resistivity of these pores to bulk flow limits the distance and rate 

of water flow. Hence, high flow resistivity of parenchymatous 

plant ground tissue was a major factor limiting the size of nonvascular 

(pre-tracheophyte) plants. 


With the evolution of xylem conduits, the wick paradox 

was solved: maximum capillary action at nano-scale cell wall 

pores is coupled to minimum bulk flow resistivity in micro-scale 

xylem conduit lumens for carrying the transpiration stream over 

most of the soil-to-leaf distance (Figure 14B). The conduits are 

dead cell wall structures that resist collapse by having lignified 

secondary walls. They form a continuous apoplasmic pipeline 

from terminal leaf vein to root tip, propagating the tensional 

component to the supply of water in the soil. As water is pulled 

into the stele of the absorbing root tissues, it is forced across 

the endodermal membrane by reverse osmosis, providing a 

mechanism for filtration and selective uptake (Figure 14B, en). 

Like cell wall water, the soil water is held by capillary and 

absorptive forces. In short, the cohesion-tension mechanism is 

a “tug-of-war” on a rope of liquid water between capillary forces 

in soil vs. plant apoplasm. Living protoplasts do not participate 

in driving the transpiration stream, but draw from it by osmosis 

during cell expansion growth and to stay hydrated and turgid 

(Figure 14C). 

Low resistivity xylem facilitates larger plants and higher flow 

rates (equals greater photosynthetic productivity), and xylem 

evolution is a history of innovations for presumably moving 

water more efficiently. The increasing literature on the topic 

is beyond the more physiological emphasis of this section 

of the current review, but one example will suffice. The rise 

of high-productivity angiosperms appears to coincide with the 

evolution of greater vein density within leaves, which minimizes 

the distance the transpiration stream flows in high-resistance 

parenchymatous ground tissue (Boyce et al. 2009). Presumably, 

the evolutionary pressure for maximizing hydraulic efficiency 

varied over geological time scales, being less during 

periods of higher atmospheric CO2 and arid (dry) conditions, 

and increasing during periods of low CO2 and mesic (wet) 

conditions (Boyce and Zwieniecki 2012). 

The reason that lower frictional resistance correlates with 

greater photosynthetic rate is because it also correlates with 

greater diffusive conductance of the stomatal pores (Meinzer 

et al. 1995; Hubbard et al. 2001). The physics of the xylem 

conduit does not account for this coupling between low flow 

resistance and high diffusive conductance (equates to high 

evaporation rate). As long as the integrity of the xylem conduit 

remains constant, its evaporation rate is essentially independent 

of its internal liquid phase flow resistance. The observed 

coupling must result from a physiological response of the plant. 

The simplest explanation for the coordination of low hydraulic 

resistance and high diffusive conductances is that stomata are 

responding in a feedback manner to some measure of leaf 

or plant water status (Sperry 2000; Brodribb 2009). Increased 

plant hydraulic resistances result in a greater tensional component 

(i.e., more negative values of ψP) at a given transpiration 

rate and soil water status. If the ψP falls below some regulatory 

set-point (which need not be constant), hydraulic or chemical


signals are sent to the stomatal complex, reducing the stomatal 

aperture and transpiration rate, which causes the ψP to rise 

back above the set-point. This ψP feedback is at least broadly 

consistent with most observed stomatal behavior in response to 

hydraulic resistance, as well as to evaporative demand and soil 

moisture stress (Oren et al. 1999; Brodribb 2009; Pieruschka 

et al. 2010). 

Recent evidence indicates that the ancestral feedback was 

entirely passive, as indeed it appears to still be in many seedless 

vascular plants (Brodribb and McAdam 2011). Accordingly, 

as ψP becomes more negative, guard cell turgor drops without 

any chemical signaling or active osmotic adjustment. Ensuing 

stomatal closure reduces the transpiration rate, which causes 

ψP to stop falling and rise back up. Only with the divergence 

of the seed plants 360 Mya did apparently active signaling 

evolve, which involves ψP sensing mechanisms, triggering of 

abscisic acid and other chemical signaling molecules, and 

active osmotic adjustment via ion pumps at the stomatal 

complex (McAdam and Brodribb 2012). The details of how this 

active feedback is achieved, the extent of active vs. passive 

mechanisms, the actual sites of evaporation within the leaf, 

the sites of water potential sensing, and the extent of hydraulic 

coupling between the stomatal complex and xylem are basic 

questions that are still poorly understood, and are the subject 

of considerable research and debate (Pieruschka et al. 2010; 

Mott and Peak 2011). 

Regardless of the feedback details, plants appear to have 

evolved to regulate ψP at the expense of sacrificing photosynthesis 

via reduced stomatal diffusive conductance. Presumably, 

this response avoids deleterious consequences of 

excessively negative ψP. Certainly most physiological processes 

are more energy-demanding at more negative ψP: the 

xylem conduit has to be stronger, and protoplasmic osmotic 

concentrations have to be greater. The implication is that 

there is some optimal midday ψP, in so far as it maximizes 

the cost/benefit margin of water transport vs. CO2 processing 

(Holtta et al. 2011). A priori, the optimal ψP would differ across 

habitats. For example, it would need to be more negative in drier 

habitats where plants have to pull harder to extract soil water as 

compared to wetter habitats. A full understanding of the costs 

associated with xylem transport requires detailed consideration 

of how xylem structure relates to its role in the water conducting 

process. 

Confining embolism with inter-conduit pitting 

While the evolution of xylem conduits solved the so called 

“wick” paradox, a new problem was created: the low-resistivity 

xylem conduits are necessarily too wide to generate, in and 

of themselves, much of a tension, i.e., a negative ψP value. 

In the inevitable event that a conduit becomes damaged and 

exposed to air, the surface tension present in the new meniscus 

spanning the conduit lumen is too weak to resist retreat (Pmin is 

not negative enough), and so water is pulled from this specific 

conduit into neighboring conduits and becomes embolized; i.e., 

it eventually is filled with N2, O2, CO2 and water vapor gases. 

Thus, capillary rise within the xylem conduit lumen cannot do 

the job of pulling up the transpiration stream. For the system 

to function, the xylem conduits must be primed by being water 

filled from inception, as they are, having developed from living 

cells. Nevertheless, there must be a means of limiting the 

spread of embolism when inevitably a conduit is damaged, 

even if by normal developmental events such as abscission of 

parts or protoxylem rupture. 

The problem of embolism is mitigated most fundamentally 

by dividing the fluid conducting space into thousands 

of overlapping and inter-connected conduits (Figure 14B, C). 

Each one embolizes as a unit because the inter-connections 

consist of porous partitions (inter-conduit pits) fine enough 

to trap and hold an air-water meniscus against a sufficiently 

negative ψP to minimize further gas propagation (Figure 14B,p). 

This multi-conduit system necessarily compromises hydraulic 

conductance because of the added resistance to flow through 

the inter-conduit pitting. Presumably, the lowest hydraulic resistance 

would be achieved by a single branching tube akin 

to the animal positive-pressure cardiovascular system. But a 

tensional system of such design would fail completely from a 

single point of air entry without any partitions to check the influx 

of gas. 

The presence of inter-conduit pitting is of great consequence 

for xylem functioning. The distribution of pitting and the structure 

and chemistry of individual pits influence both the flow 

resistance through the xylem and the Pmin for the xylem system 

(Choat et al. 2008). Although there is tremendous variation in 

inter-conduit pit structure across lineages, their basic structure 

has three elements held in common (Figure 15). As water flows 

from one conduit lumen to another, it passes through a pit 

aperture in the secondary wall, which opens into a usually 

wider pit chamber. Spanning the chamber is a porous pit 

membrane through which the water filters before passing out 

the downstream aperture. The pit membrane is the modified 

primary cell walls plus middle lamella of the adjacent conduits. 

There is no cell membrane or protoplasm in the dead but 

functioning conduit. 

Hydrolysis during xylem cell death is thought to remove all 

hemicelluloses and a debatable portion of pectins from the 

pit membrane, leaving a porous cellulosic mesh of microfibrils 

(Butterfield 1995). However, atomic force microscopy suggests 

that the microfibrillar structure lies beneath a non-porous 

coating of amorphous non-fibrillar material (Pesacreta et al. 

2005). Pit membrane chemistry, structure, and development 

are crucial to understanding the frictional resistance to flow as 

well as the ability of the xylem to sustain significant tensions in 

the water column (Choat et al. 2008; Lee et al. 2012).

Figure 15. Inter-vessel pit structure in angiosperm wood. 

Upper-right insert shows brightfield image of two overlapping vessels. 

Their common wall is studded with inter-vessel pits as shown in 

the main scanning electron microscopy (SEM) image where an intervessel 

wall has been sectioned to show individual pit structure. Pits 

consist of openings in the secondary wall (apertures) leading to pit 

chambers that are spanned by a pit membrane which is the modified 

primary cell wall and middle lamella of the adjacent vessel elements. 

The lower-right insert shows an SEM face view of the micro-porous 

pit membrane. Scale bars: 5 µm and 30 µm for the upper inset. 

Micrographs courtesy of Fredrick Lens, Jarmila Pittermann, and 

Brendan Choat. 

Inter-conduit pits add substantial flow resistance to the xylem 

conduit lumen. An unobstructed lumen conducts water as 

efficiently as an ideal cylindrical capillary tube of the same 

diameter (Zwieniecki et al. 2001a; Christman and Sperry 2010). 

The most extensive survey indicates that adding inter-conduit 

pits increases flow resistivity over that of an unobstructed lumen 

by an average factor of 2.8 in conifers with unicellular tracheids 

and 2.3 in angiosperms with multicellular vessels (Hacke et al. 

2006; Pittermann et al. 2006a). The lower number for vessels 

is not surprising given that they are roughly 10 times longer 

than a tracheid of the same diameter (Pittermann et al. 2005), 

thereby spacing high resistance pits further apart and reducing 

the length-normalized resistance (resistivity). What is surprising 

is that the greater length of vessels does not have more of 

an effect: if inter-vessel pits have the same area-specific pit 

resistance as inter-tracheid pits, placing the end-walls 10 times 

further apart should increase lumen resistivity by less than a 

factor of 1.18. The higher observed factor of 2.3 indicates that 

inter-vessel pits have higher flow resistance than inter-tracheid 

ones, a difference consistent with anatomy and estimations 

based on modeling (Pittermann et al. 2005). 

Inter-vessel pits have nano-porous “homogeneous” pit membranes 

(pores usually < 100 nm) (Choat et al. 2008), or 


Figure 16. Structure and function of inter-conduit pits in 

conifer tracheids (left) and angiosperm vessels (right). 

(A) Pit membranes, face view. 

(B) Side view schematic of membranes within the pit chamber 

formed by the secondary walls. Pits open and functioning in water 

transport. 

(C) Schematic of pit location within conduit network. 

(D) Side view of pits in sealed position showing proposed airseeding 

process (from Pittermann et al. 2005). 

often with no pores detectable, whereas inter-tracheid pits 

of conifers have a highly porous margo (pores ≫ 500 nm) 

(Petty and Preston 1969) peripheral to a central thickened torus 

(Figure 16A, left). The greater porosity of the margo decreases 

the area-specific pit resistance by an estimated 59-fold relative 

to inter-vessel pits of angiosperms (Pittermann et al. 2005). 

The homogenous-type pit membrane is presumably ancestral, 

and the implication is that the evolution of efficient torus-margo 

pitting, within in the gymnosperm lineage, was as hydraulically 

advantageous as the evolution of vessels in angiosperms.


Pit membrane chemistry interacts with xylem sap chemistry 

to influence xylem flow resistance in a very complex and poorly 

understood manner. According to the “ionic effect”, increasing 

the concentration of KCl up to 50 mM (encompassing the 

physiological range) can decrease resistivity anywhere from 

2% to 37% relative to pure water, depending on the angiosperm 

species and even the season (Nardini et al. 2011b). The 

KCl effect is much less or even negative in the already 

low-resistance torus-margo pits of conifer species (Cochard 

et al. 2010b). Furthermore, this KCl effect can be reduced or 

eliminated in the presence of as little as 1 mM Ca 2+ in some 

species and conditions (Van Iepeeren and Van Gelder 2006), 

but not in others (Nardini et al. 2011b). Skepticism about the 

importance of the phenomenon in planta (Van Iepeeren 2007) 

has been answered with observations indicating KCl-mediated 

decreases in resistivity associated with embolism and exposure 

of branches to sunlight (Nardini et al. 2011b). Interestingly, 

adaptive adjustments in KCl concentration may be mediated 

by xylem-phloem exchange (Zwieniecki et al. 2004). 

The ionic effect has been localized to the pit membranes, 

but the mechanism remains unknown. A “hydrogel” model implicates 

ionic shrinkage of pit membrane pectins or equivalent 

hydrogel polymers, and, hence, a widening of membrane pores 

(Zwieniecki et al. 2001b). However, recent observations with 

atomic force microscopy do not support a pore-widening effect. 

Although KCl was observed to thin the membrane, pores were 

not observed, suggesting that the decrease in resistance resulted 

from membrane thinning and perhaps increased permeability 

of non-porous gel material (Lee et al. 2012). The extent of 

pectins or similar gel materials in pit membranes appears to be 

highly variable across species, perhaps underlying the extreme 

variation in the ionic effect (Nardini et al. 2011b). Uncertainty 

about the extent of hydrogel components of pit membranes has 

led to an alternative (and perhaps complementary) hypothesis 

that ions increase permeability by reducing the diffuse-double 

layer of cations lining negatively charged nano-scale pores 

in the membrane (Van Doorn et al. 2011b). All of these 

hypotheses are consistent with a minor effect in torus-margo 

pit membranes, with their large micro-scale pores between 

cellulosic strands having presumably minimal pectin content. 

Although inter-conduit pits have the disadvantage of adding 

substantial flow resistance, they perform the highly advantageous 

function of trapping an air-water meniscus and minimizing 

the embolism event such that it does not compromise 

the conducting system (Figure 16B, C). The homogenous pits 

of angiosperm vessels have pores narrow enough to trap 

themeniscuswithaPmin negative enough to hold against a 

substantial range of negative ψP values (Figure 16D). The torusmargo 

pits function somewhat differently. The wider margo 

pores cannot sustain a very negative Pmin, but they can 

generate just enough pressure difference to aspirate the solid 

torus against the pit aperture on the water-filled side (Petty 

1972). In this way, the torus can seal off the pit with a sufficiently 

negative Pmin to minimize air passage (Figure 16D). 

While inter-conduit pits minimize the propagation of embolism, 

as the next section indicates, they nevertheless play 

a major role in limiting the tensional gradient that can be 

generated by the cohesion-tension mechanism. 

Limits to negative ψ P values: the problem of cavitation 

Periodically, the cohesion-tension mechanism comes under 

question for its prediction of liquid pressures that fall below 

the vapor pressure of water, and also below pure vacuum for 

a gas (Canny 1998; Zimmermann et al. 2004). A tree 30 m tall 

requires a ψP of −0.3 MPa on its stationary water column just 

to balance the gravity component. To this we need to add, 

say, −0.3 MPa to balance a favorable soil water potential 

of −0.3 MPa. Finally, we need to add the typical ψP of 

−1 MPa needed to overcome frictional resistance under midday 

transpiration rates. The required ψP totals −1.6 MPa. At sea 

level and 20 ◦ C, a vapor pressure of only −0.098 MPa will 

bring water to its boiling point, and −0.1013 MPa corresponds 

to pure vacuum for a gas. Clearly, for the cohesion-tension 

mechanism to operate, transition from the liquid phase to the 

vapor phase must be suppressed, and the xylem sap must 

remain in a metastable liquid state. The xylem sap is in effect 

super-heated, although “super-tensioned” is more descriptive. 

The liquid water column becomes analogous to a solid whose 

strong atomic and intermolecular bonds allow it to be placed 

under tension; i.e., water is a tensile liquid! 

The concept of metastable water is foreign to the macroscopic 

world of normal human experience, hence the cohesiontension 

skeptics. Water boils at 100 ◦ C, and vacuum pumps become 

gas-locked at or above −0.098 MPa. But in these familiar 

cases, the phase change to vapor (cavitation) is nucleated by 

contact with foreign agents that destabilize the inter-molecular 

hydrogen-bonding of liquid water (Pickard 1981). Such “heterogeneous 

nucleation” of cavitation is typically triggered by 

minute and ubiquitous gas bubbles in the system. When care 

is taken to minimize such heterogeneous nucleation, liquid 

water can develop substantially metastable negative ψP values. 

Theoretical calculations, based on equations of state for water, 

put the limiting ψP at homogeneous cavitation (where energy 

of the water molecules themselves is sufficient to trigger the 

phase change) below −200 MPa at 20 ◦ C(Mercury and Tardy 

2001). Experiments with a variety of systems ranging from 

centrifuged capillary tubes to water-filled quartz crystals have 

reached values well below −25 MPa, with some as low as −180 

MPa and approaching the theoretical limit (Briggs 1950; Zheng 

et al. 1991). Such values dwarf even the most negative ψP in 

plants, which is about −13 MPa (Jacobson et al. 2007); more 

typical plant ψP values are less negative than −3 MPa.

There is abundant evidence that cavitation “pressures” in 

plants are negative enough for the cohesion-tension mechanism 

to operate. Such pressures are determined from a “vulnerability 

curve” which usually plots the hydraulic conductivity 

(reciprocal of resistivity) of the xylem (often as a percentage 

loss from maximum) as a function of the ψP value in the xylem 

sap. Curves are generated in several ways, but the centrifuge 

method is commonly used because of its rapidity (Alder et al. 

1997; Cochard et al. 2005). Stems or roots are spun in a custom 

centrifuge rotor that places their xylem under a known tension 

at the center of rotation. The conductivity is measured either 

during or between spinning, and the experiment is continued 

until the conductivity has dropped to negligible values, thus 

indicating complete blockage of flow by cavitation. Typical 

species-specific variation is shown in Figure 17A, and Figure 17B 

compares the ψP value causing complete loss of xylem conductivity 

with the minimum ψP values measured in nature for 

102 species (Sperry 2000). Clearly, the xylem of some species 

cavitates much more readily than others, but these vulnerable 

species also do not develop very negative ψP values in nature. 

Across the board, the minimum ψP is generally less negative 

than the value of ψP at zero conductivity, as is required by the 

cohesion-tension mechanism. 

Although the centrifuge method is widely accepted for 

conifers and for short-vesseled angiosperms, there is some 

controversy about its ability to measure the vulnerability of longvesseled 

taxa where many conduits can exceed the length of 

the spinning conductivity segment. Response curves in these 

taxa indicate that a significant number of these large vessels 

cavitate at very modest negative ψP values. This pattern is 

seen in two of the curves shown in Figure 17A (open symbols). 

Comparisons with other methods have verified this type of 

curve in many (Christman et al. 2012; Jacobson and Pratt 

2012; Sperry et al. 2012), but not in all cases (Choat et al. 

2010; Cochard et al. 2010a), and resolving the matter requires 

further research. 

A major cause of the cavitation in plant xylem is air-seeding 

through the inter-conduit pits that normally are responsible 

for confining gas embolism (Crombie et al. 1985; Sperry and 

Tyree 1988; Sperry and Tyree 1990; Jarbeau et al. 1995). 

The Pmin of inter-conduit connections is easily measured from 

the positively applied gas pressure that just breaches their 

seal (Christman et al. 2012). From a variety of techniques 

employed across a wide range of species, the Pmin range 

of inter-conduit pitting is generally indistinguishable from the 

range of ψP values that cause cavitation. In consequence, 

vulnerability curves can usually be reproduced using positive 

gas pressures rather than placing the xylem sap under tension 

(Cochard et al. 1992). The prevailing model of cavitation 

nucleation is that when the ψP in the transpiration stream 

drops to the Pmin of the inter-conduit seal against adjoining 

embolized vessels, a gas bubble is pulled into the transpi- 


ration stream which then nucleates cavitation. The sap ψP 

in the cavitated conduit immediately rises to 0, and the sap 

is very quickly drained out of the conduit by the surrounding 

transpiration stream until the entire conduit becomes filled with 

water vapor and air, the exact composition of the embolism 

depending on diffusive exchange with the surrounding tissue 

(Tyree and Sperry 1989). According to this model, cavitation 

and embolism can propagate from conduit-to-conduit via pit 

failure. 

Considerable attention has been given to how the structure 

of inter-conduit pitting relates to its function in cavitation resistance. 

In conifer tracheids, where torus aspiration seals the 

pits, it has been proposed that air-seeding occurs when the pit 

membrane is stretched sufficiently to displace the torus from 

the aperture, exposing part of the margo through which air can 

readily pass (Figure 16D). Displacement has been observed microscopically, 

and conifers that are more resistant to cavitation 

generally have less flexible pit membranes, and can have a 

torus that covers the aperture with greater overlap (Sperry and 

Tyree 1990; Domec et al. 2006; Hacke and Jansen 2009). An 

alternative model is that air seeding occurs by displacement of 

the air-water-interface between the torus and the pit chamber 

wall. In some conifers, air-seeding could also occur through 

pores in the torus. In support of these mechanisms is a dependence 

of Pmin on the sap surface tension (Cochard et al. 2009; 

Holtta et al. 2012; Jansen et al. 2012). However, it is not clear 

whether sap solutions that lower the surface tension also alter 

pit membrane flexibility and, hence, the ease of torus displacement. 

It is also unclear whether the torus-margo membrane can 

de-aspirate to function a second time if the embolized tracheids 

refill. 

In angiosperm inter-vessel pits, air seeding probably occurs 

by displacement of the air-water meniscus from pores in 

their homogenously nano-porous pit membranes (Figure 16D), 

but beyond this, the details are surprisingly complex and 

ambiguous. Air seeding at pores is implied by the predicted 

sensitivity to surfactants and correspondence between pore 

dimensions, sized by particle penetration, and air-seeding 

pressures (Crombie et al. 1985; Sperry and Tyree 1988; 

Jarbeau et al. 1995). The pores may be pre-existing in the 

membrane, or perhaps created or enlarged by the partial 

or complete aspiration of the membrane that likely precedes 

the air-seeding (Thomas 1972). A major role of membrane 

strength is indicated by the effects of removing Ca 2+ from 

the membrane using various chelators such as oxalic acid. 

These treatments do not alter surface tension or hydraulic 

conductivity, but they can dramatically increase membrane 

flexibility and vulnerability to cavitation (Sperry and Tyree 

1988; Sperry and Tyree 1990; Herbette and Cochard 2010). 

Further support for the importance of membrane mechanics is 

the cavitation “fatigue” phenomenon wherein xylem becomes 

more vulnerable to cavitation after having been cavitated and


Figure 17. Vulnerability of xylem to cavitation by negative pressure. 

(A) Vulnerability curves for five species showing the drop in xylem hydraulic conductivity (normalized by stem cross-sectional area) as the 

xylem pressure becomes more negative. Curves were generated using the centrifugal force method. Species can differ considerably in their 

maximum hydraulic conductivity (x axis intercept) and how readily they lose it to cavitation (from Hacke et al. 2006). 

(B) Xylem pressure at zero hydraulic conductivity from cavitation (the y intercept of a, above) vs. the minimum pressure observed in nature 

for 102 species (from Sperry 2000). 

re-filled. The implication is that mechanical stress has plastically 

deformed the membrane to make it more porous. 

Cavitation fatigue is potentially reversible in vivo and with 

artificial xylem saps, suggesting the stressed and air-seeded 

pit membrane can be restored back to its normal form (Hacke 

et al. 2001b; Stiller and Sperry 2002). 

The possibility that pores are created by mechanical stress 

on the pit membrane is also consistent with the typical rarity of

observable pores of predicted air-seeding size in non-stressed 

membranes (Choat et al. 2008), as well as the tendency for 

pit membranes to be thicker, pit chambers shallower (less 

deflection), and apertures proportionally smaller in cavitationresistant 

species (Choat et al. 2008; Jansen et al. 2009; 

Lens et al. 2010). Furthermore, the KCl-induced decrease in 

membrane flow resistance does not translate into increased 

vulnerability to cavitation (Cochard et al. 2010b). This could 

mean there are few to no pores in the non-stressed water conducting 

pit membrane, with water penetrating the hydrated gel 

phase. Alternatively, this could support the diffuse-double layer 

hypothesis for the KCl effect which does not require changes 

in pore dimensions (Van Doorn et al. 2011b). The apparent 

interaction between membrane stress and air-seeding would 

be largely eliminated for “vestured” pits where, in some plant 

lineages, the membrane is supported by outgrowths from the pit 

chamber wall, a factor that must lie behind the elusive adaptive 

significance of these structures (Jansen et al. 2001; Choat et al. 

2004). 

An additional complexity in linking cavitation resistance to 

pit structure is a seemingly inescapable role for probability. A 

single inter-conduit seal consists of hundreds if not thousands 

of individual pits. Not all of these pits will be created equal, and 

it only takes one leaky pit to air-seed the cavitation. According 

to the “rare pit” hypothesis, the more pits that constitute the 

seal, the weaker the seal will be, because by chance the 

leakier will be the leakiest pit (Wheeler et al. 2005). This will 

be true whether the air-seeding pores are pre-existing, or are 

created by mechanical stress. This hypothesis is supported 

by the general trend for larger vessels to be more vulnerable 

to cavitation (because they would have more pits), and by a 

correlation between the extent of inter-conduit pit area and 

increasing vulnerability (Hacke et al. 2006; Christman et al. 

2009; Christman et al. 2012). It seems safe to conclude that 

both pit quantity and pit quality interact to set the ψP value at 

which air-seeding occurs and, hence, the cavitation resistance 

(Lens et al. 2010). 

Because cavitation appears to spread from conduit to conduit, 

the three dimensional connectivity of the conduit network 

should have an additional impact on cavitation resistance. 

Woody species differ considerably in the extent of overlap 

or connectivity between individual conduits (Carlquist 1984). 

Modeling studies suggest that greater connectivity would tend 

to facilitate the spread of embolism, whereas more limited connectivity 

would tend to confine it (Loepfe et al. 2007). However, 

limited comparative data do not always support this prediction. 

A greater vessel grouping index (higher connectivity) in some 

lineages correlates with higher cavitation resistance rather than 

enhanced vulnerability (Lens et al. 2010). This trend supports 

the earlier notion that the greater redundancy provided by high 

connectivity would be advantageous for minimizing the effect 

of embolism in xeric habitats (Carlquist 1984). 


The evident complexity that links xylem conduit structure to 

cavitation resistance comes from the multitude of variables 

involved: features at the individual pit level (e.g. membrane 

porosity, thickness, Ca 2+ content, mechanical properties) can 

be balanced by features at the inter-conduit wall scale (e.g., 

pit number), which in turn can be balanced by features at 

the conduit network level (e.g., conduit connectivity). Thus, the 

same cavitation resistance is likely to be achieved by multiple 

structural combinations. 

Freeze-thaw associated cavitation 

Cavitation can also be induced by freeze-thaw cycles and likely 

by a “thaw expansion” nucleating mechanism (Pittermann and 

Sperry 2006) (Figure 18A). Freezing of the xylem sap in nature 

usually occurs under conditions where transpiration is minimal. 

Thus, xylem blockage by ice formation would normally not 

result in ψP becoming more negative. Instead, ψP would likely 

become less negative, or even become positive because of 

the expansion of ice. However, dissolved gases in the sap are 

insoluble in ice, and under typical freezing conditions will form 

bubbles in the middle of the conduit. On thawing, if these bubbles 

do not dissolve fast enough they can nucleate cavitation 

if the thawed sap ψP is sufficiently negative. Hence, cavitation 

would occur during the thawing rather than the freezing phase, 

a prediction supported by experimental observation (Mayr and 

Sperry 2010). 

Just as for cavitation by water stress, there is considerable 

variation between species in their vulnerability to freeze-thaw 

cavitation. It is scarcely detectible in some species regardless 

of the negative sap ψP values, and others are blocked 

completely by a single freeze-thaw event at ψP values close 

to zero (Davis et al. 1999) (Figure 18B). However, unlike the 

water stress situation, there appears to be a single structural 

variable of over-riding importance for determining vulnerability 

to freeze-thaw cavitation. That variable is the xylem conduit 

lumen diameter: wider conduits are uniformly more susceptible 

to cavitation than narrower ones, regardless of whether the 

conduit is a conifer tracheid or angiosperm vessel (Figure 18B). 

The simplest explanation is that wider vessels form larger 

bubbles during freezing because of their greater water volume, 

and large bubbles take longer to re-dissolve and, hence, are 

more likely to nucleate cavitation, post-thaw. As the thawexpansion 

model predicts, a more negative ψP post-thaw, or 

a more rapid thawing rate, will also induce more cavitation at 

a given conduit diameter (Langan et al. 1997; Pittermann and 

Sperry 2003, 2006). Similarly, the amount of embolism should 

decrease with a greater rate of freezing, which reduces gas 

bubble size (Sevanto et al. 2012). 

In some species, the amount of embolism increases with the 

minimum freezing temperature, an observation not necessarily 

predicted from the thaw-expansion mechanism (Pockman and


Figure 18. Vulnerability of xylem to cavitation by freeze-thaw 

events. 

(A) The “thaw expansion” mechanism for cavitation by freezing 

and thawing. Freezing of a sap-filled functional vessel creates 

gas bubbles in the ice-filled frozen conduit. If bubbles persist long 

enough after thawing, and negative pressures are low enough, they 

will trigger cavitation and result in an embolized conduit (from Sperry 

1993). 

(B) Loss of hydraulic conductivity caused by a single freeze-thaw 

cycle at -0.5 MPa versus the average conduit lumen diameter. Data 

are species averages for angiosperm vessels (black) and conifer 

tracheids (blue) (from Pittermann and Sperry 2003). 

Sperry 1997; Ball et al. 2006; Pittermann and Sperry 2006). It is 

possible that the lower ice temperature and consequent tissue 

dehydration creates, locally, a more negative sap ψP, postthaw 

(Ball et al. 2006), or perhaps causes tissue damage that 

nucleates cavitation, post-thaw. Acoustic emissions are often 

detected during the freezing phase, which could indicate that at 

least some cavitation occurs prior to the thawing of tissue (Mayr 

and Zublasing 2010). However, experiments indicate no loss of 

conductivity when stems frozen under negative ψP conditions 

are thawed at atmospheric pressure; the conductivity only 

drops when the thaw occurs under negative ψP values (Mayr 

and Sperry 2010). It is possible that other phenomena besides 

conduit cavitation are causing freezing-associated acoustic 

emissions. Importantly, not all embolism events during winter 

are necessarily caused by freeze-thaw cycles. Sublimation and 

cavitation by water stress in thawed and transpiring crowns 

with frozen boles or soil represent other potential causes 

(Peguero-Pina et al. 2011). 

Negative xylem sap ψ P and conduit collapse 

The cohesion-tension mechanism requires conduit walls that 

are sufficiently rigid to withstand collapse by the required 

negative ψP values. Hence, the evolution of secondary walls 

and lignification necessarily paralleled the evolution of xylem 

tissues. While many factors contribute to the strength of conduit 

walls to prevent implosion, a dominant variable is the ratio 

of wall thickness to conduit lumen radius. This ratio tends to 

increase with cavitation resistance, as expected from concomitantly 

more negative sap ψP values. A higher thickness-to-span 

ratio also increases wood density, consistent with the tendency 

for greater wood density in more cavitation-resistant trees that 

generally experience more negative ψP values (Hacke et al. 

2001a; Domec et al. 2009). 

Estimates of wall strength give an average safety factor 

from implosion of 1.9 in woody angiosperm vessels and 6.8 

in conifer stem tracheids (Hacke et al. 2001a). The lower value 

for vessels presumably reflects their minimal role in mechanical 

support of the tree, a function performed by wood fibers. 

However, conifer tracheids must be additionally reinforced 

because they not only have to hold up against negative sap 

ψP, but they also support the tree itself. Interestingly, not all 

conduits avoid implosion, as it has been observed in the axial 

tracheids of pine needles (Cochard et al. 2004), transfusion 

tracheids of podocarps (Brodribb and Holbrook 2005), and 

metaxylem vessels in maize (Kaufman et al. 2009). In each 

case, the collapse was apparently reversible. Not unexpectedly, 

implosion is also observed in conduits of lignin-deficient 

mutants (Piquemal et al. 1998). 

Trade-offs between efficiency and safety 

The cohesion-tension mechanism, and its limitation by cavitation 

and conduit collapse, suggest potential trade-offs in 

the xylem conduit structure for minimizing flow resistance on 

the one hand (efficiency), and sustaining greater negative ψP 

without cavitation or conduit collapse on the other hand (safety). 

With respect to greater resistance to collapse, large increases

in the thickness-to-span ratio would be more readily achieved 

by narrowing the lumen, because walls arguably have a limited 

maximum thickness (Pittermann et al. 2006b). Tradeoffs arise 

because narrower lumens would have exponentially greater 

flow resistance, and higher thickness-to-span increases construction 

costs. Greater resistance to cavitation by freeze-thaw 

cycles also comes at the expense of narrower lumens, and 

consequently, higher flow resistance per conduit (Davis et al. 

1999). Resistance to freeze-thaw embolism can, in turn, tradeoff 

with photosynthetic capacity in evergreen species (Choat 

et al. 2011). 

Enhanced resistance to cavitation by water stress also tends 

to be associated with higher flow resistances, as seen in the 

collection of vulnerability curves shown in Figure 17A. The 

more vulnerable species tend to have higher initial hydraulic 

conductivities than the more cavitation-resistant ones. The 

basis for this association is not straightforward because so 

many variables interact to determine a species’ vulnerability 

curve. While intuitively one might expect that pits which are 

more resistant to air-seeding would also have a greater flow 

resistance (Holtta et al. 2011), the data suggest otherwise: 

membrane flow resistance is uncoupled from membrane airseeding 

resistance. Estimates of pit flow resistances across 

several angiosperm and conifer species showed no relationship 

with cavitation resistance (Pittermann et al. 2005; Hacke et al. 

2006). Furthermore, the drop in flow resistance via the ionic 

effect on pit membranes had no effect on cavitation resistance 

(Cochard et al. 2010b). Instead, the efficiency-safety trade-off 

may arise at the whole-conduit and conduit-network scale. If the 

rare pit hypothesis were correct, greater cavitation resistance 

would require fewer pits per conduit, which would generally 

correspond to narrower and shorter conduits of consequently 

greater flow resistance (Wheeler et al. 2005). And if reducing 

the connectivity of a conduit (the number of conduits it contacts) 

limits the spread of embolism, as expected, the lower 

connectivity may translate to lower conductivity at the network 

scale (Loepfe et al. 2007). 

The flow resistance penalty of narrower and shorter conduits 

can be compensated by increasing their number per wood 

area (Hacke et al. 2006). This “packing” strategy is exemplified 

by conifers which devote over 95% of their wood volume 

to tracheids and, thus, achieve similar whole stem hydraulic 

conductivity as angiosperms whose generally wider and longer 

vessels occupy only a small fraction of wood space, most of 

which is devoted to structural fibers and storage parenchyma. 

The packing strategy exemplifies how trade-offs at one level 

of structure can be compensated for at another scale, vastly 

complicating the adaptive interpretation of wood structure and 

function. 

Trade-offs of one sort or another presumably underlie the 

observed correlation between cavitation resistance and the 

range of native sap ψP values (Figure 17B). Accordingly, mesic- 


adapted species that experience less negative xylem ψP are 

vulnerable to cavitation because being overly resistant would 

cost them in terms of greater flow resistance and vascular 

construction costs. Conversely, xeric-adapted species that 

periodically endure more negative ψP values must be more 

resistant to cavitation, and their consequently greater flow 

resistance and construction costs competitively exclude them 

from more mesic habitats. The result of this adaptive scenario 

is that species tend to be only as resistant to cavitation as they 

have to be for the native ψP range they experience over their 

lifespan (Maherali et al. 2003; Markesteijn et al. 2011). 

The limiting process of cavitation naturally constrains the 

xylem ψP over which productivity can be sustained. Indeed, 

an important adaptive advantage of stomatal regulation of ψP 

is to keep it from reaching such negative values that would 

induce excessive cavitation (Tyree and Sperry 1988). “Runaway 

cavitation,” which is the loss of all hydraulic conductivity 

caused by unregulated ψP, can be induced experimentally, and 

it is a dramatic and quick cause of mortality (Holtta et al. 

2012). Not surprisingly, plants have evolved the necessary 

cavitation resistance and stomatal control mechanisms to avoid 

such an efficient suicidal scenario. But stomatal control cannot 

directly prevent the gradual accumulation of cavitation as the 

xylem ψP becomes more negative due to limited soil water 

availability. In consequence, extreme stress events or climatic 

shifts push plants towards excessive cavitation, resulting in 

partial or complete dieback from chronic reductions (70% or 

greater) in hydraulic conductivity from cavitation (Anderegg 

et al. 2012; Plaut et al. 2012). Soil-plant-atmosphere models 

that incorporate cavitation resistance can be successful in 

predicting responses of vegetation to drought (Sperry et al. 

2002), allowing effects of climate change on plant water and 

carbon flux to be anticipated. 

Refilling of embolized conduits 

The refilling of embolized xylem conduits has been documented 

in numerous studies. Embolisms accumulating over the winter 

from freeze-thaw cycles and other causes can be reversed in 

the spring in many species (Sperry 1993; Hacke and Sauter 

1996). Diurnal embolism and refilling cycles have also been 

documented (Stiller et al. 2005; Yang et al. 2012), as well as 

refilling after relief from prolonged drought (West et al. 2008). 

Two kinds of refilling have been observed. In “bulk” refilling, 

the sap ψP of the entire bulk xylem stream rises close to or 

above zero to force sap back into the embolized conduits. For 

this to happen, at a minimum, transpiration must be negligible 

and the soil ψw must be close to zero. Xylem ψP would thus 

decrease only to what is necessary to counter-act gravity, 

dropping by approximately 0.01 MPa per meter height. Less 

negative ψP values, even positive values, could develop from 

foliar uptake of rain or dew, and especially from osmotically


generated root pressures. A few species, Acer in particular, 

also develop positive stem ψP values in response to freezethaw 

cycles in early spring (Tyree and Zimmermann 2002). 

Root pressures can exceed 0.5 MPa and are strongly associated 

with bulk refilling in a variety of woody and herbaceous 

plants (Sperry 1993; Stiller et al. 2005; Yang et al. 2012). 

Experimental suppression of root pressure has been shown to 

block refilling in some species (Sperry 1993). The natural failure 

of root pressure and spring refilling, owing to freezing-related 

mortality of shallow roots, has been linked to birch dieback 

episodes (Cox and Malcolm 1997). Diminishing root pressure 

with plant height has also been invoked as a limit to the stature 

of refilling bamboos (Yang et al. 2012). 

In a second type of “novel refilling,” the bulk xylem ψP is much 

too negative to allow sap to move back into the embolized conduits 

(Salleo et al. 1996). There must be a pumping mechanism 

that brings sap into the embolized conduit and keeps it there 

until the gas is dissolved or escapes. The pumping mechanism 

is unknown, but several hypotheses have been proposed (see 

Nardini et al. 2011a for a recent review). Two basic driving 

forces are suggested: forward osmosis associated with solute 

accumulation in the thin water film along the embolized conduit 

wall, or reverse osmosis driven by either tissue pressure, or 

perhaps more likely, Münch pressure flow redirected from the 

phloem to the embolism, via ray tissue. In the latter case, 

refilling becomes a special case of phloem unloading. The data 

suggest the mechanism is: (a) triggered by the presence of a 

gas-filled conduit (rather than a particular plant water potential 

(Salleo et al. 1996)), (b) associated with starch hydrolysis 

(Bucci et al. 2003; Salleo et al. 2009), (c) upregulation of 

certain aquaporins (Secchi and Zwieniecki 2010), and (d) active 

phloem transport in the vicinity of the embolism (Salleo et al. 

2006). 

Xylem conduit wall sculpturing and chemistry may also be 

important (Kohonen and Helland 2009) with wettable areas 

assisting water uptake and gas dissolution, and hydrophobic 

areas perhaps allowing gas escape through minute wall pores 

(Zwieniecki and Holbrook 2009). Two very different roles have 

been proposed for inter-conduit pits in the refilling process. 

In one model, air pockets in the pit chamber and “wicking” 

forces at the aperture serve to isolate the pressurized sap in 

the embolized vessel from the negative ψP in the adjacent transpiration 

stream (Zwieniecki and Holbrook 2000). Alternatively, 

it has been proposed that pit membranes can act as osmotic 

membranes, with sap being pulled from the transpiration stream 

into the refilling conduit by an osmotic gradient, analogous 

to the generation of positive turgor pressures in protoplasts 

(Hacke and Sperry 2003). 

A particularly informative study is the imaging work of Brodersen 

et al. (2010). Embolized vessels in grapevine were observed 

to refill while the ψP of the surrounding xylem was more 

negative than −0.7 MPa, confirming the need for a pumping 

process. Water entered empty vessels from the direction of 

the rays, a pattern consistent with phloem-directed water influx 

rather than either pit membrane osmosis from the transpiration 

stream or root pressure. There was no obvious mechanism 

to prevent drainage of the accumulating water back into the 

surrounding sap stream, contradicting a role of inter-vessel 

pits in hydraulic isolation. Whether the vessel refilled or stayed 

partially embolized depended on the difference between the 

rate of water influx from the rays, versus drainage to the 

surrounding transpiration stream. There was considerable variation 

in the onset, rate, and eventual success or failure, in the 

plants imaged. Unfortunately, the ψπ of the refilling sap was not 

determined, so forward- versus reverse-osmosis mechanisms 

could not be distinguished. Nevertheless, the results lend 

strong support to a phloem-coupled refilling mechanism that 

refills by pumping water into the embolized vessels faster than 

it is withdrawn. 

Engineering xylem properties: A path to increased 

plant productivity 

The cohesion-tension mechanism constrains the productivity 

and survival of plants, arguably constituting the “functional 

backbone of terrestrial plant productivity” (Brodribb 2009). Because 

of the stomatal regulation of canopy xylem ψP, frictional 

resistance to water flow through the plant is coupled to the 

maximum potential photosynthetic rate and, hence, to productivity 

in general. The coupling in turn is necessary for avoiding 

hydraulic failure by cavitation, which limits plant survival 

in extremis. The cavitation limit presumably evolved in response 

to complex trade-offs with frictional resistance, with 

competition selecting for minimal flow resistance at the expense 

of excessive cavitation safety margins. Although the driving 

force for the transpiration stream is passive, flow resistance 

(via the ionic effect) and conduit refilling is modulated by 

active metabolic processes. Probably the single most important 

structures in the pipeline are the inter-conduit pits: their distribution, 

chemistry, structure, and mechanical properties greatly 

influence both frictional resistance to flow and vulnerability to 

cavitation by water stress. 

The tools of molecular biology have the potential to greatly 

advance our knowledge of the flow resistance, cavitation, and 

refilling phenotypes, as well as the nature of trade-offs among 

them. As the genetic and developmental controls of xylem 

anatomical traits become better understood (Demari-Weissler 

et al. 2009), they can be manipulated to untangle structurefunction 

relationships that can otherwise only be inferred from 

comparative studies. Crucial to advancement in this area 

are model organisms in which the hydraulic physiology can 

be phenotyped and manipulated. Among woody plants, the 

Populus system is perhaps most promising, and much 

has already been learned from it (Secchi and Zwieniecki

2010; Schreiber et al. 2011). The next decade should 

bring rapid progress as molecular biology continues to 

merge with comparative and evolutionary whole plant 

physiology. 

Long-distance Signaling Through 

the Phloem 

Over the past several decades, considerable attention has 

been paid to unraveling the mechanics of phloem loading. 

Genetic and molecular studies have identified the major players 

that mediate in the loading of sugars, predominantly sucrose, 

into the CC-SE complex. Interestingly, in terms of the apoplasmic 

loaders, the recent identification of the permease that 

controls release of sucrose from the phloem parenchyma cells 

into the CC apoplasm (Figure 13B, I) served to complete the 

molecular characterization of this important pathway (Chen 

et al. 2012b). Based on such studies and extensive physiological 

experiments, the nature of the photosynthates (sugars 

and amino acids) loaded into the phloem translocation stream 

is well established. 

The phloem has also been shown to carry additional cargo, 

including the phytohormones auxin, gibberellins, cytokines and 

abscisic acid (Hoad 1995), signaling agents involved in plant 

defense (discussed later in the review), as well as certain 

proteins and various forms of RNA (Lough and Lucas 2006; 

Buhtz et al. 2010). That specific proteins are present in the 

phloem has been recognized for some time (Fisher et al. 

1992; Bostwick et al. 1992), and furthermore, some such 

proteins have been shown to move within the translocation 

stream (Golecki et al. 1998, 1999; Xoconostle-Cázares et al. 

1999). 

Phloem proteins: Potential roles in enucleate SE 

maintenance and long-distance signaling 

Phloem exudate can be collected from a number of plant 

species, and this feature has been used to develop proteomic 

databases for these species (Barnes et al. 2004; Giavalisco 

et al. 2006; Lin et al. 2009; Rodriguez-Medina et al. 2011). 

This collection process requires that an incision be made 

into the petiole or stem in order to allow the phloem to 

“bleed.” Thus, due care is required to minimize the level 

of protein contamination from surrounding (CCs and phloem 

parenchyma) tissues. As excision results in an abrupt pressure 

drop between the sieve tube system and the surrounding cells, 

it is generally appreciated that some level of contamination 

is unavoidable (Atkins et al. 2011). Here, use of molecular 

markers such as Rubisco (Doering-Saad et al. 2006; Giavalisco 

et al. 2006; Lin et al. 2009), can help in assessing the extent 

to which contamination may have occurred. Generally, 


contamination does not appear to be an important issue, 

especially for the prominent proteins, but proteins present in 

very low abundance need to be viewed with a degree of 

caution. 

Other methods, including cutting aphid stylets (Aki et al. 

2008; Gaupels et al. 2008a), EDTA-induced phloem exudation 

(Gaupels et al. 2008b; Batailler et al. 2012) and laser microdissection 

of phloem tissues (Deeken et al. 2008) have also 

been employed to develop phloem databases. Collectively, 

these studies have established phloem proteome databases 

containing more than 1,000 proteins, with activities encompassing 

a very broad range of activities, including enzymes involved 

in metabolic networks, amino acid synthesis, protein turnover, 

RNA binding, transcriptional regulation, stress responses, defense, 

and more. 

The next step will be to partition these proteins into those 

involved in local maintenance of the functional enucleate sieve 

tube system and long-distance signaling. For these studies, 

a combination of hetero-grafting experiments conducted between 

species from different genera or families, and advanced 

mass spectroscopy methods, will prove most useful. The cucurbits, 

such as pumpkin, cucumber, melon and watermelon, 

from which analytical quantities of phloem exudate can generally 

be collected, may prove ideal for this purpose. The 

recent completion of annotated genomes for three of these 

cucurbits (Huang et al. 2009; Garcia-Mas et al. 2012; Guo 

et al. 2012) adds to the utility of these species for such critical 

experiments. 

The complexity of the phloem proteome raises the question 

as to the stability of these proteins and the mechanism by 

which they might be turned over within the sieve tube system. 

The large population of proteinase inhibitors probably 

prevents turnover by simple proteolysis (Dinant and Lucas 

2012). However, identification in the phloem sap of ubiquitin 

and numerous enzymes involved in protein ubiquitination and 

turnover, including all the components of the 26S proteasome 

(Figure 19), indicates that enucleate SEs likely have retained the 

ability to engage in protein sorting and turnover (Lin et al. 2009). 

Thus, once they have performed their function(s), phloem 

proteins can be degraded either through export into neighboring 

CCs, or in loco via the ubiquitin-26S proteasome pathway. 

The mature, enucleate sieve tube system also has been 

shown to contain all the enzymes and associated activities 

required for a complete antioxidant defense system (Walz et al. 

2002; Lin et al. 2009; Batailler et al. 2012). Interestingly, these 

enzyme activities appear to increase in response to imposed 

drought stress (Walz et al. 2002). This complement of enzymes 

would appear to function, locally, to afford protection against 

oxidative stresses, thereby preventing damage to essential 

components of the SEs. Such local maintenance functions will 

likely be performed by a specific subset of the proteins detected 

in phloem exudates.


Figure 19. Pumpkin phloem sap contains the machinery for ubiquitin-mediated proteolysis. 

(A) Schematic representation of the 26S proteasome indicating that all components except for Rpn4 were identified from the pumpkin phloem 

sap. Orange boxes indicate components identified by Lin et al. (2009). 

(B) Identification of phloem proteins associated with ubiquitin-mediated proteolysis. Note that green boxes represent proteins present in the 

Arabidopsis genome, and red lettering indicates identification in pumpkin phloem proteins. White boxes represent Saccharomyces cerevisiaespecific 

proteins. Ub, ubiquitin; CHIP, carboxyl terminus of Hsc 70-interacting protein; APC/C, anaphase promoting complex/cyclosome; 

DCAF, DD B1-CUL4-associated factor; SCF, Skp1-Cul1-F-box protein; ECS, Elongin C-Cul2-SOCS box; ECV, SCF-like E3 ubiquitin ligase 

complex (from Lin et al. 2009).

FLOWERING LOCUS T (FT) as the phloem-mobile 

florigenic signal 

It has long been known that, for plants whose flowering is 

controlled by day length, the phloem is involved in the transmission 

of a photoperiod-induced signal that moves from the 

mature/source leaves to the shoot apex where it induces the 

onset of flowering (Zeevaart 1976). The nature of this grafttransmissible 

signal, termed florigen (Zeevaart 2006), was 

recently identified as FT, a member of the CETS protein 

family (consisting of CENTRORADIALIS [CEN], TERMINAL 

FLOWER 1 [TFL1] and FT). FT expression is confined to CCs 

in source leaves, and this small protein enters the sieve tube 

system by passage through the CC-SE PD. 

Direct evidence for the presence of FT in the phloem translocation 

stream was provided by studies performed on a pumpkin 

(Cucurbita moschata) accession in which flowering occurs 

only under short-day (SD) conditions (Lin et al. 2007). Mass 

spectroscopy studies conducted on phloem sap collected from 

plants grown under long-day (LD) and SD conditions provided 

unequivocal evidence that the C. moschata FT orthologue 

was present only in exudate collected from SD-grown plants 

in which flowering was induced. Supporting evidence for FT 

as a component of the long-distance florigenic signal was 

provided by studies on Arabidopsis (Corbesier et al. 2007) and 

rice (Tamaki et al. 2007). Here, FT and Hd3a, the rice FT 

orthologue, were expressed as GFP fusions driven by a CCspecific 

promoter. Detection of a GFP signal in the meristem 

of these transgenic plants was consistent with movement from 

the phloem into the meristem where floral induction was taking 

place. 

An absolute quantitation peptide approach was used to 

determine the level of FT in the pumpkin phloem translocation 

stream. Recorded values were in the low femtomolar range (Lin 

et al. 2007), clearly placing FT in a protein hormone category 

(Shalit et al. 2009). Here, it is noteworthy that FT peptides 

could not be detected in these pumpkin phloem exudates 

when analyzed using the proteomics approach reported by Lin 

et al. (2009). This is important, as it indicates that not all low 

abundance proteins detected in phloem exudates represent 

contaminants from surrounding tissues. 

As a number of examples exist in which both mRNA and 

the encoded protein have been detected in phloem exudates 

(Lough and Lucas 2006), Lin et al. (2007) carried out extensive 

tests for the presence of the pumpkin FT transcripts using the 

same phloem exudates in which FT peptides were identified. 

As FT transcripts could not be amplified, it appeared that, 

in these plants, FT protein, but not its mRNA, serves as 

the florigenic signal. This finding was consistent with earlier 

studies conducted on the tomato FT orthologue, SINGLE- 

FLOWER TRUSS (SFT). SFT-dependent graft-transmissible 

signals were found to induce flowering in day length neutral 


tomato and tobacco plants; however, no evidence was obtained 

for the presence of SFT transcripts within the scion meristem 

tissues (Lifschitz et al. 2006). Extensive mutagenesis of the 

FT gene, in which sequence and structural modifications 

were made to the mRNA whilst leaving the FT protein unaltered, 

had little or no effect on long-distance floral induction 

(Notaguchi et al. 2008). Again, these findings are consistent 

with FT protein, not its mRNA serving as the phloem-mobile 

signal. 

Experimental support for FT mRNA as a component of 

the florigenic signaling mechanism has been suggested from 

studies based on movement-defective viral expression systems 

(Li et al. 2009). Here, the first 100 nucleotides of the FT RNA 

sequence were shown to function in cis to allow systemic 

movement of heterologous viral sequences. Similar findings 

with Arabidopsis have been reported in terms of FT sequences 

acting in cis to mediate in the long-distance transport of 

otherwise cell-autonomous transcripts (Lu et al. 2012). Further 

support for the role of endogenous FT mRNA, as a component 

of the florigenic signal, was claimed from studies with transgenic 

Arabidopsis lines in which flowering was delayed through 

expression, in the apex, of RNAi/artificial miRNA-FT (Lu et al. 

2012). Unfortunately, these results are contradictory to findings 

from a similar FT silencing experiment in which expression of 

amiRNA-FT in the phloem caused a significant delay in floral 

induction, whereas its expression in the apex failed to delay 

flowering (Mathieu et al. 2007). Although the controversy over 

whether or not FT mRNA contributes to floral induction remains 

to be resolved, there is no a priori reason why, for any gene, its 

mRNA and protein cannot both serve as signaling agents via 

the phloem. 

ATC as a phloem-mobile anti-florigenic signal 

Systemic floral inhibitors or anti-florigens have been proposed 

to participate in down regulating floral induction under noninducing 

conditions (Zeevaart 2006). Evidence in support of 

this concept was recently provided by studies performed on 

Arabidopsis plants carrying mutations in ATC, aCEN/TFL1 

homologue. Flowering in Arabidopsis is promoted under LDand 

inhibited under SD-conditions. Expression of ATC is enhanced 

during short days and, based on the effect of an atc-2 

mutant, the WT gene appears to contribute to the suppression 

of flowering (Huang et al. 2012). 

At the tissue level, ATC was found to be expressed in the 

vascular system, and the phloem in particular, but not in the 

apex. This suggested a non-cell-autonomous function for either 

the ATC transcripts or protein. A range of grafting experiments 

were performed with Arabidopsis stock and scions connected 

above the hypocotyl region (with the stock containing several 

mature source leaves). Analysis of RNA extracted from atc-2


scions grafted onto WT stocks provided clear evidence for 

the movement of ATC transcripts across the graft union, and 

compared to WT:WT grafts, flowering time was delayed (Huang 

et al. 2012). Parallel experiments were performed to address 

whether ATC protein moves across the graft union. In these 

Western blot experiments, a clear ATC signal was detected in 

atc-2 scions grafted onto WT stocks. These findings support the 

possibility that, in Arabidopsis, both ATC transcripts and protein 

are phloem-mobile; i.e., together, they may enter the shoot 

apex to compete with FT for FD, thereby inhibiting the transition 

to flowering. However, it is also possible that the phloemmobile 

ATC transcripts enter CCs located in the atc-2 scion 

tissues where they then produce ATC protein. Irrespective of 

this potential complication, identification of ATC as a negative 

regulator of flowering time in Arabidopsis constitutes an important 

step forward in understanding the role of the phloem in the 

overall regulation of plant growth and development. 

Phloem-mediated long-distance lipid-based signaling? 

Lipids and lipid-binding proteins have been detected in phloem 

exudates collected from a number of plant species. Some 14 

putative lipid-binding proteins were detected in Arabidopsis 

phloem exudates collected from excised petioles that were 

incubated in EDTA to facilitate the bleeding process (Guelette 

et al. 2012). Bioinformatics analysis of these proteins indicated 

potential roles in membrane synthesis and/or turnover, prevention 

of lipid aggregation, participation in synthesis of the 

glycosyphosphatidylinositol (GPI) anchor, and biotic and abiotic 

stress. A range of lipids have also been reported in phloem 

exudates, including simple lipids to complex glycolipids and 

phytosterols such as cholesterol (Behmer et al. 2011; Guelette 

et al. 2012). 

An interesting study recently conducted on an Arabidopsis 

small (20 kDa) phloem lipid-associated family protein (PLAFP) 

revealed that it displayed specific bind properties for phosphatidic 

acid (PA) (Benning et al. 2012). As both PA and 

PLAFP were detected in Arabidopsis exudate, these results 

suggest that PA may well be either trafficked into or translocated 

through the sieve tube system by PLAFP. In any event, 

detection of lipids and lipid-binding proteins within phloem 

exudates certainly raises the question as to whether they function 

in membrane maintenance and/or long-distance signaling 

events. 

Messenger RNA: A smart way to send a “message”! 

A number of recent studies have identified specific mRNA 

populations within the phloem sap of various plant species 

(Sasaki et al. 1998; Doering-Saad et al. 2006; Lough and 

Lucas 2006; Omid et al. 2007; Deeken et al. 2008; Gaupels 

et al. 2008a; Rodriguez-Medina et al. 2011; Guo et al. 2012). 

These databases indicate that the phloem translocation stream 

of the angiosperms likely contains in excess of 1,000 mRNA 

species that encode for proteins involved in a very wide range 

of processes. While many of these transcripts are held in 

common between plant species, specific differences have been 

reported. For example, a comprehensive analysis carried out 

using the phloem transcriptomes prepared from cucumber 

(1,012 transcripts) and watermelon (1,519 transcripts) phloem 

exudate indicated that 55% were held in common (Guo et al. 

2012). In contrast, the vascular transcriptomes (13,775 and 

14,242 mRNA species in watermelon and cucumber, respectively) 

were 97% identical. Thus, differences in phloem transcripts 

most likely reflect unique functions specific to these 

species. 

A comparative analysis of the vascular and phloem transcriptomes 

for cucumber and watermelon identified populations of 

transcripts that are highly enriched in phloem exudates over 

the level detected in excised vascular bundles. The numbers 

given above represent the transcripts that were present at 

≥2-fold higher than the level detected in vascular bundles. 

Concerning cucumber, more than 30% of the phloem transcripts 

were enriched >10-fold above the level in the vascular 

bundles. Importantly, some transcripts were enriched above 

500-fold, with another 210 displaying >20-fold enrichment. A 

similar situation was observed for watermelon, with some 120 

transcripts displaying >10-fold enrichment and 320 having 5fold 

or greater enrichment in the phloem sap. These data indicate 

that, following transcription in the CCs, many transcripts 

must undergo sequestration in the sieve tube system through 

trafficking mediated by the CC-SE PD. 

To date, only a limited number of these phloem mRNAs have 

been characterized in terms of whether they act locally or traffic 

long-distance to specific target sites. Excellent examples where 

translocation through the phloem has been established include 

NACP (Ruiz-Medrano et al. 1999), PP16 (Xoconostle-Cázares 

et al. 1999), the PFP-LeT6 fusion gene (Kim et al. 2001), GAIP 

(Haywood et al. 2005), BEL5 (Benerjee et al. 2006; Hannapel 

2010), POTH1 (a KNOTTED1-Like transcription factor) (Mahajan 

et al. 2012) and Aux/IAA18 and Aux/IAA28 (Notaguchi et al. 

2012). The stability of these phloem-mobile transcripts is made 

possible by the fact that phloem exudates have been shown 

to lack RNase activity (Xoconostle-Cázares et al. 1999), and 

thus, by extension, the phloem translocation stream is likely 

also devoid of this activity. 

Phloem delivery of GAIP transcripts modifies 

development in tomato sink organs 

The pumpkin phloem sap was found to contain transcripts for 

two members of the DELLA subfamily of GRAS transcription 

factors, CmGAIP and CmGAIPB, known to function in 

the GA signaling pathway (Ruiz-Medrano et al. 1999). The

function of CmGAIP and GAI from Arabidopsis was investigated 

using transgenic Arabidopsis and tomato lines expressing 

engineered dominant gain-of-function Cmgaip and 

DELLA-gai genes. Importantly, these transgenic plants exhibit 

clear morphological changes in leaf development, and this 

characteristic was used to test whether phloem delivery of the 

Cmgaip/DELLA-gai transcripts into sink tissues could induce 

this mutant phenotype. These engineered gai transcripts were 

found to move long-distance through the phloem, and could 

then exit from the terminal phloem and subsequently traffic 

into the apex, where they accumulated in developing leaf 

primordial (Haywood et al. 2005). Parallel studies conducted 

with transgenic plants expressing EGFP revealed the inability 

of these transcripts to enter the phloem. This finding suggested 

that phloem entry of Cmgaip/DELLA-gai transcripts must 

occur by a selective process. 

Analysis of WT tomato scions grafted onto Cmgaip and 

DELLA-gai stocks indicated that import of these transcripts 

caused highly reproducible morphological changes in leaf phenotype 

(Figure 20). Unexpectedly, tomato leaflet morphology 

was found to be influenced quite late in development. Of 

equal importance, the presence of Cmgaip and DELLAgai 

transcripts, within the various tissues of the scion, was 

not correlated with overall sink strength. Strong signals were 

detected in young developing flowers and the apex, but signal 

could not be amplified from fruit stalks or rapidly expanding 

fruit (Figure 20C). These findings indicated an unexpected 

complexity in the events underlying phloem delivery 

of these transcripts, suggesting a high degree of regulation 

over such trafficking of macromolecules. Furthermore, these 

studies revealed that phloem long-distance delivery of RNA 

can afford flexibility in adjusting developmental programs to 

ensure that newly emerging leaves are optimized for performance 

under existing environmental conditions (Haywood et al. 

2005). 

A model has been proposed that selective entry of transcripts 

from the CC into the sieve tube system involves specific 

sequences within the RNA (Lucas et al. 2001). As both Cmgaip 

and DELLA-gai transcripts were able to move within the 

heterologous plant, tomato, this finding suggested that such 

sequence motifs, termed “zip codes,” must be conserved and, 

further, the molecular machinery required for this recognition 

and trafficking must similarly be conserved between pumpkin, 

tomato and Arabidopsis. 

Support for this hypothesis was provided by mutational 

analysis of GAI in which it was clearly established that mRNA 

entry into the phloem is facilitated by a motif located within the 3 ′ 

region of the transcript (Huang and Yu 2009). Furthermore, this 

motif was specific to GAI, as parallel experiments conducted 

with the four additional members of the DELLA family failed 

to detect their transcripts in heterografting assays. By testing 

an extensive series of GAI mutants, it was found that two zip 


Figure 20. Phloem delivery of Cmgaip/DELLA-gai transcripts 

modifies leaf development. 

(A) Schematic illustrating the V-grafting method used to test the 

influence of phloem-mobile transcripts, derived from the stock and 

imported across the graft union (gu) into the scion, on developmental 

processes occurring in the scion. 

(B) Schematic showing that import of Cmgaip/DELLA-gai transcripts 

into wild-type tomato scions results in the development of 

scion leaves with characteristic morphological features associated 

with transgenic stock plants carrying the dominant gain-of-function 

Cmgaip or DELLA-gai gene (Haywood et al. 2005). Numbers 

represent tomato leaflet position along the leaf axis. Leaflet L0 does 

not exhibit this change in morphology as it had passed through this 

developmental stage prior to formation of the graft union. 

(C) Detection of Cmgaip/DELLA-gai transcripts in wild-type tomato 

scions grafted onto Cmgaip/DELLA-gai transgenic tomato stocks; 

presence (+), absence (-). Absence of phloem-mobile transcripts 

from the rapidly growing fruit indicates the operation of a regulatory 

system controlling delivery of phloem mobile transcripts to specific 

tissues/organs (from Haywood et al. 2005). 

codes appear to be required for phloem entry and translocation, 

one being located in the coding sequence and the other in the 

3 ′ untranslated region. Finally, GFP transcripts carrying these 

two zip codes were detected in the scion, confirming that these 

sequence motifs are necessary and sufficient for targeting GAI 

transcripts to the phloem (Huang and Yu 2009).


Role of phloem RNP complexes in RNA delivery 

to target tissues 

Considering that the phloem translocation stream contains in 

excess of 1,000 transcripts, it is perhaps not surprising that the 

pumpkin phloem proteome was found to contain in excess of 

80 recognized RNA binding proteins (RBPs) (Lin et al. 2009). 

Several of these RBPs have been characterized, with the first 

being CmPP16-1 and CmPP16-2 from pumpkin (Xoconostle-Cázares 

et al. 1999). These two proteins display properties 

equivalent to those of viral movement proteins (Lucas 2006) 

in that they bind RNA in a sequence non-specific manner and 

mediate the cell-to-cell trafficking of transcripts through PD. 

Entry of CmPP16-1/2 into the sieve tube system appears to 

be controlled by post-translational modifications. Interestingly, 

both CmPP16 and its PD receptor, NCAPP1 (Lee et al. 2003), 

require serine residues to be phosphorylated and glycosylated 

for effective interaction and delivery of CmPP16 to and through 

PD (Taoka et al. 2007). In an elegant experiment, Aoki et al. 

(2005) used severed brown leafhopper stylets to introduce 

CmPP16-1 and CmPP16-2 directly into the sieve tube system 

of rice. Analysis of the long-distance movement of these two 

pumpkin proteins, within the rice plant, clearly revealed that 

they did not simply follow the direction of bulk flow. Destinationselective 

movement was shown to be controlled by proteins 

from the pumpkin phloem sap that interact with CmPP16- 

1/2. Collectively, these studies on CmPP16 provide important 

insights into the complexity of the processes that underlie 

macromolecular trafficking within the phloem translocation 

system. 

The most extensively characterized phloem RBP is Cm- 

RBP50, a polypyrimidine tract-binding protein that accumulates 

to high levels in pumpkin phloem sap (Ham et al. 2009). Pull 

down assays, using a polyclonal antibody directed against 

CmRBP50 and pumpkin phloem exudates, led to the identification 

of the proteins and mRNA species contained within 

a CmRBP50-associated ribonucleoprotein (RNP) complex 

(Figure 21A). Interestingly, CmGAIP transcripts were contained 

within this CmRBP50 RNP complex. Binding specificity between 

CmRBP50 and these CmGAIP transcripts is imparted 

by a series of polypyrimidine tracts located within the mRNA. 

As these sites differ from those involved in mediating CmGAIP 

transcript entry into the phloem (Huang and Yu 2009), it is likely 

that assembly of the CmRBP50 RNP complex occurs within the 

sieve tube system. 

Heterografting studies conducted between pumpkin (stock) 

and cucumber (scion) established that this RNP complex is 

engaged in the long-distance delivery of CmGAIP mRNA to 

developing tissues. Important insights into the basis for the stability 

of this CmRBP50-CmGAIP mRNA complex were provided 

by reconstitution experiments. These studies identified a series 

of serine residues within the CmRBP50 C-terminus that, when 

phosphoryated, allow for the assembly of the RNP complex. 

Sequential binding of CmPP16 and the other proteins that form 

the complex results in an increase in its overall stability (Li et al. 

2011) (Figure 21B). 

In addition to CmGAIP, CmSCARECROW-LIKE, CmSHOOT 

MERISTEMLESS, CmETHYLENE RESPONSE FACTOR and 

CmMybP transcripts were also isolated from CmRBP50 RNP 

complexes. Given that the watermelon phloem exudate was 

found to contain transcripts for some 118 transcription factors, 

there remains much to be done in terms of identifying and characterizing 

the associated RNP complexes involved in mediating 

their entry into, and presumed long-distance transport through, 

the phloem. 

Phloem transcripts and protein synthesis 

in the enucleate sieve tube system 

Analysis of cucumber phloem proteome and transcriptome 

databases identified some 169 proteins for which transcripts 

were also present in phloem exudates. This represents around 

15% of the phloem transcripts and raises the question as to why 

there would be the need for such transcripts when, presumably, 

the proteins can enter the sieve tube system by trafficking 

through CC-SE PD. The possibility exists that some proteins 

required for SE maintenance are cell-autonomous. If this were 

the case, synthesis within the enucleate sieve tube system 

would be required. It has long been assumed that the mature 

SE does not have the capacity for protein synthesis. However, 

the pumpkin phloem proteome contains numerous proteins 

involved in translation (Lin et al. 2009). Furthermore, gel 

filtration chromatography experiments performed on pumpkin 

phloem exudates identified complexes of proteins containing 

CmeIF5A and elongation factor 2, both known to be involved 

in protein synthesis (Ma et al. 2010). Thus, synthesis of a 

discrete set of essential proteins may well occur within mature 

SEs. 

Phloem-based delivery of small RNA and systemic 

gene silencing 

In recent years, post-transcriptional gene silencing has 

emerged as an important component of the regulatory networks 

that control a broad array of developmental and physiological 

processes (Brodersen and Voinnet 2006). These events can 

occur in local tissues, and the phloem also functions as a 

conduit for the systemic spread of gene silencing (Melnyk et al. 

2011). The pioneering work of Palauqui and coworkers laid a 

solid foundation for this concept. Transgenic tobacco plants 

expressing additional copies of a nitrate reducase gene (Nia) 

were found to undergo a perplexing process in which small 

clusters of cells within mature source leaves were observed to


Figure 21. Delivery of phloem-mobile transcripts to sink tissues requires formation of stable ribonucleoprotein complexes. 

(A) Model illustrating the protein composition of the pumpkin CmRBP50-based ribonucleoprotein (RNP) complex that binds specifically to 

five phloem transcripts that encode transcription factors which are delivered into sink tissues (from Ham et al. 2009). 

(B) Phosphorylation of four serine residues at the C-terminus of CmRBP50 is essential for assembly of a stabilized RNP complex (left image). 

Mutating these serine residues prevents RNP complex assembly in the sieve tube system (right image) (from Li et al. 2011). 

turn white (Palauqui et al. 1996). Subsequently, this process 

moved up the body of the plant in a source-to-sink pattern 

reflective of phloem translocation. 

An insightful follow-up series of experiments revealed that a 

graft-transmissible signal moved into the scion where it caused 

the turnover of Nia transcripts. A deficiency in fixed nitrogen 

then caused the leaves of the scion to turn white (Palauqui 

et al. 1997) (Figure 22A). Cell-to-cell movement of the Nia 

silencing signal was tested by placement of a WT stem seg- 

ment between the silenced stock and the non-silenced scion. 

That white leaves still developed in these scions confirmed 

the involvement of the phloem (Figure 22B). This conclusion 

was further supported by grafting Nia-silenced scions onto 

non-silenced root stocks. As the direction of the phloem is 

from the stock to the scion, this graft combination did not 

result in the generation of white leaves (Figure 22C), again 

consistent with transmission of the silencing signal through the 

phloem.


Figure 22. Grafting experiments illustrating that transmission 

of a phloem long-distance signal can induce posttranscriptional 

gene silencing within developing scion tissues. 

(A–C) Transgenic tobacco plants expressing a nitrate reductase 

gene (Nia) segregated into silenced (S) and non-silenced (NS) 

phenotypes were employed in grafting studies to test for the longdistance 

propagation of the silencing condition through the phloem. 

(A) Grafting of an NS scion onto an S stock resulted in silencing 

of Nia within the scion leaves. (B) Placement of a wild-type (WT) 

stem segment between the NS scion and S stock did not prevent 

the transmission of the silencing signal. (C) Grafting of a silenced 

scion (S) onto a NS stock does not activate silencing in the NS stock 

tissues. 

(D) Analysis of RNA samples collected from specific grafted tissues 

( ∗ ) confirmed that the observed silencing phenotype reflected 

sequence-specific targeting of the Nia transcripts (redrawn from 

Palauqui et al. 1997). 

Sequence-specificity of the silencing signal was established 

by grafting studies performed with nitrite reductase (Nii) transgenic 

tobacco stocks and non-silenced Nia scions (Figure 22D). 

A hypothesis was advanced that phloem-mobile silencing 

signals involved the translocation of antisense RNA, whose 

entry into the developing scion tissues caused an enzymemediated 

cleavage of the double-stranded form of the target 

RNA (Jorgensen et al. 1998). Detection, in silenced tissues, 

of small (20–25 nucleotide [nt]) antisense RNA complementary 

to the silenced gene (sRNA) (Hamilton and Baulcombe 1999) 

provided strong support for the general features of this model. 

Analysis of RNA extracted from pumpkin phloem sap identified 

a population of 21 nt – 24 nt sRNA. Sequencing and 

bioinformatics analysis indicated that these sRNAs belong to 

both the micro(mi)RNA and small interfering (si)RNA silencing 

pathways (Yoo et al. 2004). Interestingly, although equal signal 

strength was detected for sense and antisense sRNA probes, 

they did not appear to exist in the phloem sap as duplexes. 

The involvement of these phloem sRNAs in systemic silencing 

was explored using silencing (stock) and non-silencing (scion) 

transgenic squash (Cucurbita pepo) plants expressing a viral 

coat protein gene. Phloem sap from both the stock and scion 

tested positive for CP siRNA, and analysis of apical tissues 

from these scions confirmed that the level of CP mRNA had 

been greatly reduced. Interestingly, a low level signal for 

the antisense CP transcript could also be detected in the 

phloem sap of both stock and scion plants. Collectively, these 

findings offered support for the hypothesis that both siRNA and 

antisense RNA are likely components of the systemic silencing 

machinery. 

Limited information is available concerning the mechanism 

by which these sRNA molecules enter and move longdistance 

through the phloem. Biochemical studies performed 

on pumpkin and cucumber phloem exudate identified a 20 kDa 

PHLOEM SMALL RNA BINDING PROTEIN1 (PSRP1) that 

bound specifically to sRNA (Yoo et al. 2004). This protein has 

the capacity to traffic its sRNA cargo through PD, and in the 

cucurbits, PSRP1 may be involved in shuttling sRNA from CCs 

into the sieve tube system. Interestingly, PSRP1 homologues 

have yet to be identified in the genomes of other plant species. 

This raises the possibility that additional proteins have evolved 

to carry out these same functions. 

Role of phloem-mobile sRNA in directing 

transcriptional gene silencing in target tissues 

The phloem sap collected from pumpkin and oilseed rape 

contains a significant population of 24-nt sRNA (Yoo et al. 

2004; Buhtz et al. 2008), indicating a likely involvement in 

transcriptional gene silencing (TGS) within sink tissues (Mosher 

et al. 2008). Grafting experiments performed with various 

combinations of GFP transgenic and DICER-LIKE mutant 

Arabidopsis lines provided further confirmation that a significant 

population of exogenous/endogenous 23 nt – 24 nt sRNA can 

cross the graft union (Molnar et al. 2010). Methylation analysis 

of DNA extracted from these grafted target tissues provided

compelling evidence for the hypothesis that phloem-mobile 24nt 

sRNA can mediate in epigenetic TGS of specific genomic 

loci. 

Similar findings were reported for studies on endogenous 

inverted repeats that generate double-stranded RNA molecules 

(Dunoyer et al. 2010), as well as for transgenic plants expressing 

a hairpin-structured gene under the control of a viral 

companion-cell-specific promoter (Bai et al. 2011). In this latter 

case, this phloem-transmissible TGS event was shown to be 

inherited by subsequent progeny. Collectively, these findings 

support the notion that phloem-mobile sRNA can serve to regulate 

gene expression within developing tissues epigenetically 

to allow for adaptation to environmental inputs. 

Phloem-mobile miRNA 

Although generally considered to act cell-autonomously 

(Voinnet 2009), as mentioned above, numerous miRNAs have 

been detected in phloem exudates from various plant species, 

and the roles played by some of these have recently been 

established. In plants, adaptation to changing nutrient availability 

in the soil involves both root-to-shoot (see next section) 

and shoot-to-root signaling. In the case of phosphate (Pi), 

changes in availability within leaves leads to an upregulation in 

miR399 production and, subsequently, its entry into the phloem 

translocation stream (Lin et al. 2008; Pant et al. 2008). Delivery 

of miR399 into the roots results in the cleavage of the target 

mRNA encoding for PHO2, a ubiquitin-conjugating E2 enzyme 

(UBC24). This gives rise to increased uptake of Pi into these 

roots and restoration of Pi levels within the body of the plant. 

Loss of PHO2 activity probably allows for an increase in Pi 

transporter capacity; i.e., of influx carriers located in the outer 

region of the root and xylem parenchyma-located efflux carriers 

that function in Pi loading into the transpiration stream (Chiou 

and Lin 2011). 

Tuber induction in potato is regulated by phloem delivery 

of BEL5 transcripts and miR172 (Martin et al. 2009). Vascular 

expression of miR172 and its upregulation under tuber-inducing 

SD conditions suggested that this miRNA may act as a longdistance 

signaling component in the control of potato tuber 

induction. Support for this notion was provided by grafting studies 

involving P35S:MIR172 stocks grafted to WT potato scions. 

Here, tuberization occurred as early as in P35S:MIR172 potato 

lines. In contrast, when P35S:MIR172 scions were grafted to 

WT stocks, early tuber induction did not occur. Although these 

findings are consistent with miR172 serving as a phloem-mobile 

signal, it is also possible that it might act through regulation, in 

the CCs, of an independent mobile signal. 

Phloem-mobile sRNA control over host infection 

by parasitic plants 

Parasitic plants cause major losses in some regions of the 

world (Ejeta 2007). Recent studies have established that host 


transcripts can enter into parasitic plants (Roney et al. 2007; 

David-Schwartz et al. 2008). The pathway for this trafficking 

is through the haustoria of the parasite that interconnects its 

vascular system to that of the host. This suggested that host 

invasion by parasitic plants might be controlled by phloem 

delivery of sRNA species designed specifically to target critical 

genes involved in the physiology or development of the 

parasitic weed (Yoder et al. 2009). 

Based on the observation that parasitic broomrape 

(Orobanche aegyptiaca) accumulates large quantities of mannitol, 

Aly et al. (2009) engineered transgenic tomatoes to 

express a hairpin construct to target the mannose 6-phosphate 

reductase (M6PR) that functions as a key enzyme in mannitol 

biosynthesis. Analysis of tissue from broomrape growing on 

these transgenic tomato plants indicated a significant reduction 

in both M6PR transcript and mannitol levels. This strategy 

gave rise to a level of tomato protection against this plant 

parasite. 

An alternate control approach based on targeting a parasitic 

developmental program involved the development of transgenic 

tobacco plants expressing hairpin constructs for two 

dodder (Cuscuta pentagona) haustoria-expressed KNOTTEDlike 

HOMEOBOX1 (KNOX1) genes. These constructs were 

driven by the CC-specific SUC2 promoter and were based 

on 3 ′ UTRs that did not display sequence homology to the 

related tobacco orthologues, STM and KNAT1–3 (Alakonya 

et al. 2012). Defects in haustoria development and connection 

to the transgenic tobacco plants were highly correlated with 

the presence of KNOX1 siRNA, delivered most likely through 

the vascular system, and down-regulation of the C. pentagona 

KNOX1 transcript levels. Importantly, dodder plants growing 

on these transgenic tobacco plants exhibited greatly reduced 

vigor. Collectively, these studies indicate that an effective 

control of plant parasitism may be achieved by targeting a 

pyramided combination of parasite genes involved in various 

aspects of growth and development. 

Root-to-shoot Signaling 

Response to abiotic stress 

Signals arising within the root system can provide shoots with 

an early warning of root conditions, such as water deficiency, 

nutrient availability/deficiency, and so forth (Figure 23). The 

xylem transports hormones, such as abscisic acid (ABA) 

(Bahrun et al. 2002; Jiang and Hartung 2008), ethylene and 

cytokinin (CK) (Takei et al. 2002; Hirose et al. 2008; Kudo 

et al. 2010; Ghanem et al. 2011), as well as strigolactones 

(SLs) (Gomez-Roldan et al. 2008; Umehara et al. 2008; Brewer 

et al. 2013; Ruyter-Spira et al. 2012) from roots to aboveground 

tissues. In this section of the review, we will address the role of 

these xylem-borne signaling agents.


Figure 23. The plant vascular system serves as an effective 

inter-organ communication system. 

In response to a wide range of environmental and endogenous 

inputs, the xylem (blue lines) transmits root-to-shoot signals (blue 

circles), including hormones such as abscisic acid, ACC (ethylene 

precursor) and cytokinin, as well as strigolactones (SLs). These 

xylem-borne signaling agents serve to communicate the prevailing 

conditions within the soil. The phloem (pink lines) transports a wide 

array of shoot-to-root signaling molecules (pink circles), including 

auxin, cytokinin, proteins and RNA species, including mRNA and 

sRNA. These phloem-borne signaling agents complete the longdistance 

communication circuit that serves to integrate developmental 

and physiological events, occurring within shoot and root 

tissues, in order to optimize the plant performance under the existing 

growth/environmental conditions. 

When soils become dry, root-derived signals are transported 

through the xylem to leaves in order to effect a reduction in both 

leaf transpiration and vegetative growth. Tight control between 

water uptake by the root system and the xylem transpiration 

stream is achieved through regulation of leaf stomatal aperture. 

Production of ABA within roots and its transport to the leaves 

could contribute to preventing excess water loss, as it is has 

long been known that ABA is a key regulator of stomatal conductance 

(Mittelheuser and Van Steveninck 1969; Schachtman 

and Goodger 2008). 

The ABA content in roots is well correlated with both soil 

moisture and root relative water content (Davis and Zhang 

1991; Thompson et al. 2007). Although large increases in ABA 

are detected in the xylem sap, when plants are exposed to 

drought conditions (Christmann et al. 2007), grafting studies 

have indicated that root-derived ABA is not necessary for 

drought-induced stomatal closure (Holbrook et al. 2002). Furthermore, 

recent studies have shown that leaf-derived synthesis 

of ABA contributes to water-stress-induced down-regulation 

of stomatal conductance (Holbrook et al. 2002; Thompson 

et al. 2007). Thus, further studies are required to evaluate the 

relative contribution of root-derived versus shoot-synthesized 

ABA in terms of the overall efficacy of stomatal control over the 

transpiration stream. 

There is some indication that ethylene-based signaling may 

also contribute to root-to-shoot communication under abiotic 

stress conditions. For example, the anaerobic environment 

caused by soil flooding can increase the level of aminocyclopropane 

carboxylic acid (ACC, the immediate precursor of 

ethylene) in plant roots. ACC has been detected in the xylem 

from both flooded and drought-stressed plants (Tudela and 

Primo-Millo 1992; Belimov et al. 2009). This root-derived ACC 

is transported to the shoot where it then gives rise to increased 

ethylene production which can play a role in regulating shoot 

growth and development under these stress conditions (Voesenek 

et al. 2003; Pérez-Alfocea et al. 2011). 

Changes in xylem sap pH have also been reported for 

plants exposed to drought conditions. Alkalinization of the 

xylem sap appears to be correlated with enhanced stomatal 

closure (Jia and Davies 2008; Sharp and Davies 2009). These 

pH changes may act synergistically with ABA and ACC to 

generate an effective root-to-shoot signaling system for water 

stress. The involvement of other known xylem-based root-toshoot 

signals, such as CK, etc., remains to be established in 

terms of contributing to water stress signaling. In any event, 

advancing our understanding of the mechanisms of root-toshoot 

signaling associated with water stress should lead the 

way for the development of crops with improved water use 

efficiencies. 

Xylem signals associated with nutrient stress 

The phenotypic plasticity that plants display in response to 

changes in their nutrient supply requires the operation of rootto-shoot 

signaling. Such signals from roots can provide shoots 

with an early warning of decreases in nutrient supply, while 

signals from shoots can ensure that the nutrient acquisition 

by roots is integrated to match the nutrient demand of shoots 

(Lough and Lucas 2006; Liu et al. 2009). 

CK plays an important role in plant growth and development 

and its involvement as a xylem-mobile signal in regulating the 

nutrient starvation response, such as occurs under nitrogen and 

phosphorus deficiency conditions, is well established (Takei 

et al. 2002; Hirose et al. 2008; Ghanem et al. 2011). Nitrate 

deprivation leads to a reduction in the level of mobile CK in the 

xylem sap, whereas upon resupply of nitrate to these stress

oots, CK again increases in the xylem transpiration stream 

(Rahayu et al. 2005; Ruffel et al. 2011). Interestingly, transzeatin-type 

CK moves in the xylem, and isopentenyl-type CK 

is present in the phloem translocation stream. This suggests 

that these structural variations carry specific information from 

the root-to-shoot and shoot-to-root, respectively (Hirose et al. 

2008; Werner and Schmülling 2009). 

As discussed above, phosphate acquisition by the root system 

involves phloem-mobile signals from the shoot (Figure 24). 

In terms of the root-to-shoot component of this phosphate 

signaling network, it has been suggested that the level of 

phosphate in the xylem transpiration stream may serve as one 

component in this signaling pathway (Bieleski 1973, Poirier 

et al. 1991; Burleigh and Harrison 1999; Hamburger et al. 2002; 

Lai et al. 2007; Stefanovic et al. 2007; Chiou and Lin 2011; 

Thibaud et al. 2010). Studies on the growth of Arabidopsis roots 

being exposed to low phosphate conditions identified the tip of 

the primary root, including the meristem region and root cap, 

as the site that may sense local phosphate availability (Linkohr 

et al. 2002; Svistoonoff 2007). However, currently, there is no 

evidence for the existence of a phosphate sensor or receptor. 

Both CK and SLs have also been considered to function in 

xylem transmission of root phosphate status (Martin et al. 2000; 

Franco-Zorrilla et al. 2005; Kohlen et al. 2011). Plants grown 

under limiting phosphate conditions have repressed levels of 

trans-zeatin-type CK in their xylem sap (Martin et al. 2000) and, 

under these conditions, expression of the CK receptor CRE1 

is similarly decreased (Franco-Zorrilla et al. 2002, 2005). In 

many plant species, the SLs are up-regulated upon exposure 

to phosphate deficiency conditions. Grafting studies have indicated 

that SLs produced in the root can move to the shoot 

(Beveridge et al. 1994; Napoli 1996; Turnbull et al. 2002). In 

such studies, WT rootstocks grafted to mutant scions lacking 

the ability to produce SLs were able to restore WT branching 

patterns in these scions. Thus, xylem-transported SLs can 

contribute to the regulation of shoot architectural responses 

to phosphate-limiting conditions (Kohlen et al. 2011). Collectively, 

these findings suggest that the levels of phosphate, CK 

and SLs in the xylem transpiration stream play an important 

role in coordinating vegetative growth with phosphate nutrient 

availability (Rouached et al. 2011). 

Xylem signaling in plant-symbiotic associations 

The interaction of nitrogen-fixing bacteria (Rhizobia) is generally 

confined to legumes, whereas most flowering plants 

establish symbiotic associations with arbuscular mycorrhizal 

(AM) fungi for phosphate acquisition. In both types of plantsymbiont 

association, there is a significant metabolic cost to 

the plant host. Thus, there is a need for the plant to ensure 

that the cost-benefit ratio remains favorable. To this end, 


Figure 24. Long-distance signaling in response to phosphate 

deficiency conditions. 

Phosphate (Pi) availability in the soil solution is transmitted through 

the xylem transpiration stream (blue lines) which passes predominantly 

to the mature source leaves. Pi per se, and/or other root-toshoot 

signals (1), including cytokinin and strigolactones, are thought 

to be involved in this nutrient signal transduction pathway. When 

roots encounter low levels of available Pi, changes in these xylemborne 

signaling components are decoded in the leaves (2), resulting 

in the activation of Pi deficiency responsive pathways. Outputs from 

this Pi-stress response pathway, including miR399 21-nucleotide 

silencing agents and other potential signaling components (3) are 

loaded into the phloem (red lines). Phloem-mobile signals move 

down to the root where they enter different target receiver cells 

to mediate an increase in Pi uptake (4) and alter root architecture 

(4’). The miR399 signal targets PHOSPHATE2 (PHO2) transcripts 

to derepress Pi transporter activity. A different set of phloemmobile 

signals are likely delivered to developing leaves (5) and 

the shoot apex (6) to regulate growth and development in order to 

survive under the Pi-stress condition. This long-distance signaling 

network operates to ensure that the root system integrates its 

physiological activities to optimize growth conditions within the 

shoot.


Figure 25. Autoregulation of symbiosis between plants and 

bacteria/fungi involves long-distance signaling through the 

vascular system. 

During plant-symbiont associations a significant metabolic cost is 

incurred by the plant host. To ensure that the cost-benefit ratio 

remains favorable to the plant, systemic autoregulation systems 

evolved to control the level of root nodulation associated with 

nitrogen-fixing Rhizobium and growth on the roots of arbuscular 

mycorrhizal (AM) fungi. In this system, root-derived signals (blue 

circles) which are still unknown but may involve CLE peptides in 

the case of nodulation, are transported through the xylem (blue 

lines) to the shoot. In mature source leaves, these autoregulation 

of nodulation signals appear to be recognized by a LRR-RLK. 

Although grafting studies have established that signals (pink circles) 

enter and move through the phloem (pink lines) to the roots, their 

identities remain to be elucidated. These shoot-to-root signals are 

involved in down-regulating the root – Rhizobium/AM fungi symbiotic 

association. 

plants have evolved a systemic feedback regulatory system 

termed “autoregulation” in which nodule formation and the 

ongoing development of the association with the AM fungus is 

negatively controlled by long-distance signaling (Catford et al. 

2003; Staehelin et al. 2011) (Figure 25). 

The process of autoregulation in legumes in which the 

number of symbiotic root nodules is controlled by long-distance 

communication between the root and shoot has been well 

studied. Two component pathways are thought to operate 

involving root-derived signals through the xylem and shootderived 

signals through the phloem. Following rhizobial infection, 

a root-derived signal is generated that is then translocated 

to the shoot. This xylem-borne signal is perceived in the 

shoot and subsequently leads to the production of a shootderived 

signal whose movement through the phloem to the 

roots causes a block to further nodulation. Consistent with this 

notion, mutant legumes have been identified that are defective 

in autoregulation of nodulation, and grafting experiments established 

that these mutants were not capable of producing the 

requisite shoot-derived signals. 

Genetic studies suggest that CLE peptides, induced in 

response to rhizibial nodulation signals in roots, serve as 

signaling agents that travel through the xylem to the shoots. 

A leucine-rich repeat (LRR) CLAVATA-like receptor kinase, 

located in the leaves, appears to function in this signaling 

pathway (Searle et al. 2003). Although not yet proven, the 

CLE peptides imported from roots are likely perceived by the 

LRR autoregulation receptor kinase in the shoots (Okamoto 

et al. 2009; Miyazawa et al. 2010; Osipova et al. 2012). In any 

event, it will be of considerable interest to identify the feedback 

signal(s) that enters the phloem to down-regulate nodulation 

back in the roots (Oka-Kira et al. 2005). 

In addition to CLE – LRR-RLK signals from the root, soil available 

nitrogen also appears to participate in this long-distance 

signaling system. Consistent with this notion, in legumes, high 

levels of soil nitrate cause strong up-regulation of root CLE 

gene expression (Okamoto et al. 2009; Reid et al. 2011). 

Furthermore, high nitrate or ammonia levels abolish nodulation, 

and autoregulation-defective mutants exhibit more or fewer 

nitrate-insensitive phenotypes. Thus, a combination of CLE 

peptides and nitrate/ammonia levels in the xylem transpiration 

stream could function as the root-to-shoot signals that allow 

legumes to integrate autoregulation of nodulation with environmental 

nitrogen conditions. 

With respect to phosphatesignaling, studies using cucumber 

revealed that application of root extracts from mycorrhizal 

plants reduced the degree of root colonization by AM fungi 

(Vierheilig et al. 2003). In contrast, root extracts from noninfected 

plants stimulated successful AM fungal colonization. 

This study supports the hypothesis that an equivalent autoregulation 

system operates to control the plant-AM fungal 

association involved in plant phosphate homeostasis. Given 

the attributes of the cucurbits for analysis of phloem and xylem 

sap, this system might prove invaluable for studies aimed 

at identifying the agents that serve to coordinate phosphate 

acquisition by the roots with utilization in the shoots.

Role of xylem signals in coordination of shoot 

architecture 

A number of studies have indicated that root-derived signals 

play a role in the regulation of vegetative growth (Van Norman 

et al. 2004; Van Norman and Sieburth 2007; Sieburth and 

Lee 2010). These signals contribute to water and nutrient use 

efficiency through the control over shoot branching and growth. 

The BYPASS1, 2, 3 (BPS1, 2, 3) genes are required to prevent 

the synthesis of a novel substance, a bps signal that moves 

from root-to-shoot, where it modifies shoot growth (Van Norman 

et al. 2004; Van Norman and Sieburth 2007; Lee et al. 2012; 

Lee and Sieburth 2012). 

Although the functions of the BPS proteins remain to be 

elucidated, studies based on bps mutants clearly established 

that the roots of these mutants cause arrested shoot growth, 

likely due to the over-production of an inhibitor of shoot growth. 

Grafting experiments established that the root system of the 

bps mutants was both necessary and sufficient to induce shoot 

arrest, and revealed that BPS proteins can work to generate 

mobile root-to-shoot signals that can inhibit shoot growth (Van 

Norman et al. 2004; Lee et al. 2012). It will be of great interest 

to unravel the underlying mechanism by which shoot growth is 

controlled by this signaling pathway. 

Recent studies reported that bps mutants show normal 

responses to both exogenous auxin and polar auxin transport 

inhibitors, suggesting that the primary target of the bps signal 

is independent of auxin. Furthermore, this root-to-shoot signal 

appears to act in parallel with auxin to regulate patterning 

and growth in various tissues and at multiple developmental 

stages (Lee et al. 2012). Clearly, further characterization of 

the BPS signaling pathway could well open the door to novel 

approaches towards controlling shoot architecture in specialty 

crops. 

The SLs have been referred to as rhizosphere signaling 

molecules (Nagahashi and Douds 2000; Akiyama et al. 2005) 

that also participate in the regulation of shoot architecture by 

suppressing lateral shoot development. Biosynthetic SL mutants 

exhibit a highly branching phenotype and, interestingly, 

phosphate starvation in these plants causes a reduction in 

shoot branching (Umehara et al. 2008, 2010). As previously 

reported, plants grown under phosphate deficiency conditions 

have fewer shoots and an increase in lateral roots (Umehara 

et al. 2010; Kohlen et al. 2011). Several plant hormones, 

such as auxin and ethylene, also appear to be involved in 

linking phosphate signaling with plant growth responses (Chiou 

and Lin 2011). Auxin signaling was shown to be associated 

with changes in root system architecture under phosphate 

deficiency conditions (López-Bucio et al. 2002). Recent studies 

suggest that an auxin receptor TRI and the auxin signaling 

pathway are involved in this SL-regulated root-sensing of low 

phosphate conditions (Mayzlish-Gati et al. 2012). It is likely 


that auxin and SLs function, cooperatively, to control shoot 

branching (Brewer et al. 2009; Domagalska and Leyser 2011). 

These hormones move through the xylem and phloem, respectively, 

to form a network of systemic signals to orchestrate plant 

architecture at the whole plant level. 

Vascular Transport of Microelement 

Minerals 

In the last decade, significant advances have been made in the 

understanding of the mechanisms that control the intracellular 

homeostasis of microelement minerals (Takano et al. 2008; 

Curie et al. 2009; Williams and Pittman 2010; Conte and 

Walker 2011; Waters and Sankaran 2011; Ivanov et al. 2012; 

Sperotto et al. 2012). However, relatively little is known about 

the processes governing their long-distance transport. Major 

questions remaining relate to the mechanisms of vascular 

loading/unloading, as well as the chemical speciation of these 

elements during their transport. Furthermore, transport is not 

a static process and, therefore, may differ not only with the 

nutrient and plant species but also with other factors, such as 

developmental stage, circadian cycle, and nutritional status. In 

this section of the review, we assess the current knowledge 

on microelement vascular transport focusing on these open 

questions. 

Microelement trafficking and speciation in xylem sap 

In the xylem sap, the non-proteinogenic amino acid nicotianamine 

(NA), histidine, and organic acids are usually associated 

with cationic microelements (Figure 26, Table 2). NA 

binds several transition metals with very high affinity, including, 

in order of stability, Fe(III), Cu(II), Ni(II), Co(II), Zn(II), Fe(II) 

and Mn(II) (von Wirén et al. 1999; Rellán-Álvarez et al. 2008; 

Curie et al. 2009). Insights into the role of NA complexation 

and trafficking have been provided by studies of NA-deficient 

mutants. Here, the chloronerva tomato mutant is interesting 

as it has high root Cu concentrations, but the concentration 

in xylem sap is low, indicating a failure in Cu transport into 

mature leaves. This finding indicates that Cu(II)-NA likely 

serves as a key complex in the xylem sap (Herbik et al. 

1996; Pich and Scholz 1996). With regard to other cationic 

microelements, analysis of an A. thaliana nicotianamine synthase 

(NAS) quadruple mutant (which has low levels of NA) 

showed that long-distance transport of Fe through the xylem 

was not affected, and Fe accumulated in the leaves (Klatte 

et al. 2009). Other studies performed on NAS-overexpressing 

tobacco and A. thaliana plants reported elevated Ni tolerance 

and high Zn levels in young leaves. Finally, studies conducted 

on several metal hyperaccumulator species (Krämer 2010) 

identified Cu(II)-NA, Zn(II)-NA and Ni(II)-NA complexes in 

the roots, xylem sap and leaves (Schaumlöffel et al. 2003;


Figure 26. Schematic depicting membrane transporters involved 

in loading and unloading of micronutrient elements in 

the vascular systems. 

Homologues from different plant species (At, Arabidopsis thaliana; 

Tc, Thlaspi caerulescens; Os,Oryza sativa) are given as examples. 

P-type ATPase (HMA), ferroportin (IREG) and and MATE. 

(FRD) families are involved in loading Zn and Cu, Fe, and citrate, 

respectively, into the xylem. Borate is loaded into the xylem by 

the anion efflux system, AtBOR1. The chemical species present 

in the xylem and phloem sap are indicated; several micronutrient 

species may occur in xylem sap. Histidine (His), Nicotianamine 

(NA), and organic acids are the most likely chelating agents of these 

mineral micronutrients. The complex Fe3Cit3 has been detected in 

xylem sap from tomato. Unloading of Ni, Fe and Zn from the xylem 

takes place via members of the Yellow Stripe-Like family of metal 

transporters (YSL). Phloem loading and unloading of Fe, Mn, Cu 

and Zn is also mediated by several members of the YSL family 

in rice and Arabidopsis. AtOPT3, a member of the oligopeptide 

transporter family, is involved in Fe and Mn loading into the sieve 

tube system. Chemical species of micronutrient minerals in the 

phloem sap include complexes of Ni, Cu, Zn and Fe with NA. The 

complexes Zn-NA and Fe (III)-2 ′ DMA have been recently detected 

in phloem sap from rice. Iron Transporter Protein (ITP) and Copper 

Chaperone (CCH) may have a role in Fe and Cu transport within 

the phloem, respectively, whereas, Mn and Ni have been detected 

Vacchina et al. 2003; Ouerdane et al. 2006; Mijovilovich et al. 

2009, Trampczynska et al. 2010). 

Histidine (His) can function to chelate Zn, Cu and Ni in the 

xylem sap (Krämer et al. 1996; Salt et al. 1999; Liao et al. 

2000; Küpper et al. 2004). An extended X-ray absorption fine 

structure (EXAFS) study demonstrated that most of the Zn 

in petioles and stems of Noccacea caerulescens existed as 

a complex with His (Küpper et al. 2004). However, a recent 

study performed on the same species proposed His as a Zn 

ligand within cells, and NA as the Zn chelator involved in longdistance 

transport (Trampczynska et al. 2010). For Cu, as 

commented above, NA plays a key role in xylem transport. 

However, xylem transport of Cu in tomato and chicory is 

efficient even in the absence of NA, provided that His is present, 

thus offering support for the existence of both mechanisms 

for Cu complexation in xylem sap (Liao et al. 2000). Based 

on these findings, Irtelli et al. (2009) proposed that, under Cu 

deficiency conditions, NA is responsible for Cu chelation in 

xylem sap, whereas His and Pro serve as the major chelators 

in excess Cu conditions. 

The involvement of His in Ni chelation in the xylem sap 

has been proposed based on studies in Ni-hyperaccumulator 

species (Krämer et al. 1996; Kerkeb and Krämer 2003; Mari 

et al. 2006; Krämer 2010; McNear et al. 2010). In these plants, 

there is an enhanced expression of the first enzyme in the 

His biosynthetic pathway and higher concentrations of His in 

xylem sap (Krämer et al. 1996; Ingle et al. 2005). On the other 

hand, His-overproducing transgenic A. thaliana lines displayed 

enhanced Ni tolerance, but did not exhibit increased Ni concentrations 

in xylem sap or leaves (Wycisk et al. 2004; Ingle et al. 

2005). These studies suggest that, in non-hyperaccumulator 

plants, other chelating agents such as NA and organic acids, 

may also play important roles (Verbruggen et al. 2009; Hassan 

and Aarts 2011). Accordingly, studies on natural variation 

among Arabidopsis accessions indicated that a Ni(II)-malic acid 

complex may be involved in translocation of Ni from roots to 

shoots (Agrawal et al. 2012). 

As mentioned above, organic acids have also been hypothesized 

to serve as chelators for Fe, Zn, Ni and Mn in xylem sap, 

based on in silico calculations using xylem sap composition 

(von Wirén et al. 1999; López-Millán et al. 2000; Rellán-Alvárez 

et al. 2008). For instance, in silico speciation studies in xylem 

sap of the hyperaccumulator Alyssum serpyllifolium found 

approximately 18% of Ni bound to organic acids, mainly malate 

and citrate (Alves et al. 2011), and in tomato, Mn was predicted 

←−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−− 

in association with low molecular (LMW) peptides and organic 

compounds. The molybdate anion has been detected in both xylem 

and phloem sap. Boron is present as borate and boric acid in xylem 

sap and as complexes with sugar alcohols in phloem sap.


Table 2. Chemical speciation of cationic microelements present within the xylem transpiration stream and the phloem translocation 

stream 

Xylem sap Phloem sap 

Nicotianamine Histidine Organic acids Nicotianamine Peptides/proteins 

Fe Rellán-Álvárez et al. Klatte et al. 2009 ITP 

2010 

Takahashi et al. 2003 

a 

Krüger et al. 2002 

Zn Klatte et al. 2009 Küpper et al. 2004 Salt et al. 1999 

Nishiyama et al. 2012 

Trampczynska et al. 

2010 

Takahashi et al. 2003 

Cu Pich and Scholz 1996 

Liao et al. 2000 

Pich et al. 1994 CCH 

Herbik et al. 1996 

Irtelli et al. 2009 

Mira et al. 2001 

Liao et al. 2000 

(metallothioneins) 

Guo et al. 2003, 2008 

Mn White et al. 1981 LMW peptides 

Van Goor and Wiersma 

1976 

Ni Ouerdane et al. 2006 Krämer et al. 1996 Agrawal et al. 2012 Schaumlöffel et al. 2003 LMW compounds 

Kerkeb and Krämer Alves et al. 2011 

Wiersma and Van Goor 

2003 

Krämer 2010 

McNear et al. 2010 

1979 

a ITP, Iron Transport Protein; CCH, Copper Chaperone Protein; LMW, Low Molecular Weight. 

as a citrate complex (White et al. 1981). However, these 

models did not include other possible chelating agents such 

as amino acids or NA. Recently, a tri-Fe(III), tri-citrate complex 

(Fe3Cit3) was identified in the xylem sap of tomato plants, using 

an integrated mass spectrometry approach (Rellán-Álvárez 

et al. 2010). Also, by means of X-ray absorption spectroscopy, 

organic acids have been shown to complex Zn in xylem sap of 

Noccacea caerulescens (Salt et al. 1999). 

With regard to anionic microelements, the soluble molybdate 

anion, which is the predominant aqueous species at pH values 

above 4.0, has been detected in both xylem and phloem 

sap, and is assumed to be the major chemical species of 

Mo delivered by these two long-distance transport systems 

(Marschner 1995). The fact that the molybdate anion is not 

very biologically active may allow for its transport as a free 

anion. For boron (B), in addition to boric acid and borate, at 

least one other yet unidentified B-containing compound has 

been described in the xylem sap of squash roots (Iwai et al. 

2003). This compound has a lower molecular weight than the 

rhamnogalacturan II-B complex, which contributes to cell wall 

strengthening (Takano et al. 2008). 

Microelement trafficking and speciation in the phloem 

sap 

Metal mobility in the phloem sap depends on the individual 

microelement, its chemical species, and in some cases on 

the nutritional status of the plant. Zn and Ni are considered 

highly mobile in the phloem translocation stream; for instance, 

the loading of these metals into the developing wheat grain 

occurs mostly via the phloem, with transfer from xylem to 

phloem occurring in the rachis and the peduncle (Riesen and 

Feller 2005). In contrast, manganese (Mn) appears to be poorly 

mobile in the phloem; it can be translocated out of source 

leaves, but the loading of Mn into the developing grain is 

poor in most crop species (Riesen and Feller 2005; Williams 

and Pittman 2010). For Cu, its mobility in the phloem sap is 

intermediate. For instance, in wheat, translocation from mature 

to younger developing leaves does not occur, and it has been 

proposed that when Cu enters the cell it becomes bound 

by chaperones and, therefore, is not immediately available 

for retranslocation. Later remobilization of Cu appears to be 

possible during leaf senescence when proteins are hydrolyzed, 

thereby releasing Cu (Puig and Penarrubia 2009). Fe is also 

considered intermediately mobile in plants, and studies have 

shown that it is translocated to young barley leaves mainly via 

phloem transport (Tsukamoto et al. 2009). Grusak (1994) also 

reported phloem transport of Fe from various source tissues to 

developing Pisum sativum seeds. And, as occurs with Cu, an 

increased remobilization of Fe occurs during leaf senescence 

(Waters et al. 2009; Sperotto et al. 2010). 

With regard to the chemical species in which these metals 

are transported through the phloem (Figure 26), Fe, Cu and 

Zn are considered to move as either NA- or metal-mugineic


acid complexes (mugineic acids are synthesized from NA), 

especially as the neutral-to-basic pH of the phloem sap is 

suitable for metal-NA formation (Pich et al. 1994; von Wirén 

et al. 1999; Takahashi et al. 2003). Accordingly, Zn(II)-NA and 

Fe(III)-2 ′ -deoxymugineic acid complexes have been detected 

in phloem sap from rice (Nishiyama et al. 2012). Furthermore, 

mutants defective in NA production display lower Mn, Zn, 

Fe and Cu concentrations in reproductive organs (Takahashi 

et al. 2003; Curie et al. 2009). However, it is still unclear 

whether this impediment to metal delivery into sink organs 

is due to alterations in phloem loading and transport, per se, 

or to changes in the intracellular concentrations of metals in 

source tissues where NA also plays an important role in metal 

chelation. 

The phloem may also transport other forms of Fe; e.g., an 

iron transport protein (ITP) is present in the phloem sap of 

Ricinus communis (Krüger et al. 2002), but it has not yet been 

found in other species. A copper chaperone protein (CCH) has 

also been proposed to play a role in phloem transport of Cu 

from senescing to young leaves (Mira et al. 2001). In addition, 

metallothioneins (types 1, 2 and 3), proteins predominantly 

regulated by Cu, also appear to function in Cu accumulation 

and phloem transport during senescence (Guo et al. 2003, 

2008). These proteins are also associated with Cu tolerance 

(Murphy and Taiz 1997; van Hoof et al. 2001; Jack et al. 2007). 

Currently, little information is available concerning the chemical 

forms of Ni and Mn in the phloem sap. In R. communis, Mn 

has been detected in association with low molecular weight 

peptides (van Goor and Wiersma 1976), whereas Ni can be 

complexed with negatively charged organic compounds with 

a molecular weight in the range of 1,000–5,000 Da (Wiersma 

and van Goor 1979). 

The mobility of molybdenum (Mo) in the phloem varies 

depending on its concentration and on plant age. Interestingly, 

in wheat, Mo has been associated with the existence of “Mobinding 

sites” in the phloem that, until saturated, appear to 

prevent its long-distance translocation (Yu et al. 2002). B mobility 

in the phloem is highly dependent on the plant species. In 

plants that transport sugar alcohols, B appears to be complexed 

with diols and polyols (Brown and Hu 1996; Hu et al. 1997; 

Takano et al. 2008). Complexes of sorbitol-B-sorbitol, fructose- 

B-fructose, sorbitol-B-fructose and mannitol-B-mannitol have 

been identified in peach and celery phloem sap (Hu et al. 1997). 

Enhancement of sorbitol production results in an increase of B 

translocation from mature leaves to sink tissues as well as 

tolerance to B deficiency. In plant species that do not produce 

significant amounts of sugar alcohols, B is thought to be phloem 

immobile, or only slightly mobile, and its distribution in shoots 

seems primarily to follow the xylem transpiration stream (Oertli 

1993; Bolaños et al. 2004; Lehto et al. 2004; Takano et al. 

2008). 

Vascular loading and unloading of cationic 

micronutrients 

Yellow Stripe-Like (YSL) proteins play important roles in the 

short- and long-distance transport of microelements and their 

delivery to sink tissues (Curie et al. 2009). Members of the YSL 

family, AtYSL1, AtYSL2, AtYSL3, OsYSL2 and OsYSL18, are 

expressed in vascular tissues (Table 3) and may have a role 

in the lateral movement of Fe within the veins and in phloem 

transport (DiDonato et al. 2004; Koike et al. 2004; Le Jean et al. 

2005; Schaaf et al. 2005; Aoyama et al. 2009). The rice OsYSL2 

can transport Fe(II)-NA and Mn(II)-NA to an equal extent (Koike 

et al. 2004). OsYSL18 transports Fe(III)-deoxymugineic acid 

(Aoyama et al. 2009), whereas there are contradictory reports 

concerning the ability of AtYSL2 to transport Fe(II)-NA and 

Cu(II)-NA (DiDonato et al. 2004; Schaaf et al. 2005). AtYSL1 

seems to play a role in Fe(II)-NA translocation to seeds (Le 

Jean et al. 2005). A study on the Arabidopsis double mutant 

ysl1ysl3 reported reduced accumulation of Fe, Cu and Zn in 

seeds, consistent with involvement of the YSL1 and YSL3 

transporters in remobilization from leaves (Waters et al. 2006). 

There is also evidence for a role of YSLs in the Zn and 

Ni hyperaccumulation of Thlaspi caerulescens, especially for 

TcYSL3 and TcYSL7, which are highly expressed around 

vascular tissues particularly in shoots when compared with 

their A. thaliana orthologs (Gendre et al. 2007). TcYSL3 is an 

Fe(II)-NA and Ni(II)-NA influx transporter that is suggested to 

facilitate the movement of these metal-NA complexes from the 

xylem into leaf cells. 

A number of other transporters involved in vascular loading 

and unloading of microelements have also been identified 

(Figure 26, Table 3). The Arabidopsis OPT3 transporter 

(OligoPeptide Transporter) appears to be essential for embryo 

development (Stacey et al. 2008). This protein transports Mn, 

and its expression in the vascular tissue suggests a role 

in Mn long-distance transport. Although yeast studies have 

suggested that it can also transport Cu, OPT3 does not appear 

to play a role in Zn or Cu loading, as seeds of opt3-2 plants 

actually accumulate increased levels of these two metals 

(Stacey et al. 2008). The opt3-2 mutant also has reduced 

Fe concentrations in its seeds as well as impaired seedling 

growth under Fe-deficient conditions, thus suggesting a role in 

Fe loading into the seed (and perhaps even phloem-mediated 

redistribution). 

Another Fe efflux transporter, IREG1/FPN1 (Iron Regulated1/Ferroportin1), 

is considered to function in Fe loading 

into the xylem within the roots (Morrissey et al. 2009). Loss 

of FPN1 function results in chlorosis, and FPN1-GUS plants 

show staining at the plasma membrane of the root vascular 

system. However, yeast complementation studies using FPN1 

have failed, and information on the chemical form of Fe

Table 3. Transporters involved in vascular loading and unloading of cationic microelements 


Family Gene Microelement Function Reference 

YSL AtYSL1 Fe-NA Seed loading, phloem loading Le Jean et al. 2005 

AtYSL2 Fe, Zn Xylem loading/unloading DiDonato et al. 2004 

Schaaf et al. 2005 

AtYSL3 Fe, Cu, Zn Phloem loading Waters et al. 2006 

OsYSL2 Fe-NA Mn-NA Phloem loading/unloading to grains Koike et al. 2004 

OsYSL18 Fe(III)-DMA Phloem loading/unloading 

to reproductive organs 

Aoyama et al. 2009 

TcYSL3 Fe-NA, Ni-NA Xylem unloading Gendre et al. 2007 

TcTSL7 Ni-NA Xylem unloading Gendre et al. 2007 

OPT AtOPT3 Mn Fe Seed loading Stacey et al. 2008 

Ferroportin IREG1/FPN1 Fe Xylem loading Morrissey et al. 2009 

MATE FRD3 Citrate Xylem loading Durrett et al. 2007 

Yokosho et al. 2009 

Yokosho et al. 2010 

P-type ATPases AtHMA5 Cu Xylem loading André-Colás et al. 2006 

Kobayashi et al. 2008 

AtHMA2/4 Zn Xylem loading Hussain et al. 2004 

Mills et al. 2005 

Wong and Cobbett 2009 

Verret et al. 2004, 2005 

AhHMA4 Zn Xylem loading Hanikenne et al. 2008 

transported by FPN1 has yet to be established (Morrissey et al. 

2009). 

The Arabidopsis P-type ATPases, AtHMA5 and AtHMA2/4, 

have been implicated in Cu and Zn efflux, respectively, into 

the xylem at the root level, for long-distance transport to 

the shoots (Hussain et al. 2004; Mills et al. 2005; Andrés- 

Colas et al. 2006). Consistent with this model, both hma5 

and hma2hma4 loss-of-function mutants accumulate increased 

levels of the corresponding metal within the root, and show 

lower levels in their shoots (Hanikenne et al. 2008; Wong and 

Cobbett 2009). HMA5 is predominantly expressed in the root 

and is specifically induced by excess Cu. Mutants of HMA5 

overaccumulate Cu in the root, suggesting a compromised 

efflux system. Further evidence in support of the role of HMA5 

in xylem transport of Cu from the roots to the shoots comes from 

a study of natural variation in Cu tolerance among Arabidopsis 

accessions, which identified HMA5 as a major QTL associated 

with Cu translocation capacity and sensitivity (Kobayashi et al. 

2008). HMA2 and 4 are present in the plasma membrane of 

root and shoot vascular tissues (Mills et al. 2003; Hussain 

et al. 2004; Verret et al. 2004; Mills et al. 2005; Verret 

et al. 2005; Williams and Mills 2005; Sinclair et al. 2007; 

Blindauer and Schmid 2010). In addition, functional analysis 

of HMA4 in A. halleri and A. thaliana showed that silencing 

of AhHMA4, by RNA interference, completely suppressed Zn 

hyperaccumulation. These studies provided a clear demonstration 

that HMA4 plays a key role in xylem loading and, 

consequently, in root-to-shoot transport of Zn (Hanikenne et al. 

2008). 

Organic acids may also have a role in xylem Fe loading. 

Citrate has been described as an Fe(III) chelator in the xylem 

sap (Rellán-Álvarez et al. 2010) and FRD3 (Ferric Reductase 

Defective), a transporter of the MATE family, is localized to 

the plasma membrane of the pericycle and vascular cylinder. 

FRD3 proteins facilitate citrate efflux into the xylem of the root 

vasculature and have been described in Arabidopsis (Durrett 

et al. 2007), rice (Yokosho et al. 2009) and rye (Yokosho 

et al. 2010). Mutant frd3 plants are chlorotic, show reduced 

citrate and Fe concentrations in the xylem and the shoot, 

accumulate Fe in the root, and exhibit constitutive expression 

of the Fe uptake components, thus suggesting that FRD3 is 

necessary for efficient Fe transport to the shoot through the 

transpiration stream. Also, independent Fe-citrate and Fe-NA 

xylem loading systems may complement each other, as in the 

frd3 mutant, the nicotianamine synthase NAS4 gene is induced, 

and the double mutant nas4x-2/frd3 shows impaired growth 

and low Fe levels in the shoot (Schuler et al. 2010). FRD3 

is constitutively expressed in the hyperaccumulators A. halleri 

and N. caerulescens compared to A. thaliana and N. arvensis, 

and may also play a role in Zn transport (Talke et al. 2006; 

van de Mortel et al. 2006). However, this overexpression may 

be related to an altered Fe homeostasis leading to high Zn 

concentrations in the hyperaccumulators (Roschzttardtz et al. 

2011).


Vascular loading and unloading of anionic 

micronutrients 

Xylem loading of B is mediated by BOR1 (Takano et al. 2001; 

Miwa and Fujiwara 2010). BOR1 is an anion efflux system that 

is strongly expressed in the root pericycle cells surrounding the 

xylem vessels. The bor1-1 mutant is defective in xylem loading 

of B (Takano et al. 2002). The specific chemical form of the 

substrate for BOR1 remains unknown, but electrophysiological 

analyses in the human homolog, NaBC1, suggest borate anion 

as the likely candidate (Park et al. 2004). There are six BOR1 

paralogs in the A. thaliana genome which may also have roles 

in xylem-phloem loading and unloading (Miwa and Fujiwara 

2010). A second B transporter, NIP6,1, is a channel protein 

required for proper distribution of boric acid, particularly to 

young developing shoot tissues (Tanaka et al. 2008). This 

transporter is predominantly expressed in nodal regions of 

shoots, especially the phloem region of vascular tissues, where 

it is likely involved in xylem-phloem transfer of boric acid. 

The mechanisms for Mo loading and unloading in the vascular 

tissues remain to be elucidated. To date, only one Mo 

transporter from Arabidopsis, MOT1, has been identified in 

plants. MOT1 is a high-affinity molybdate transporter localized 

to the plasma membrane or mitochondrial membranes, and 

plays an important role in efficient Mo uptake from soils and 

accumulation within the plant (Tomatsu et al. 2007; Baxter 

et al. 2008). MOT1 belongs to the family of sulphate transporters, 

SULTR (Hawkesford 2003), which has 14 members 

in A. thaliana. It is tempting to speculate that some of these 

transporters may be involved in vascular tissue loading or 

unloading. 

Systemic Signaling: Pathogen 

Resistance 

Like all living organisms, plants have to constantly resist 

pathogenic microbes. The absence of a circulatory vascular 

system and their sessile nature can pose particular problems. 

Plants have therefore evolved unique defense mechanisms to 

ensure survival. The multiple modes of plant defense include 

both passive and active mechanisms that provide defense 

against a wide variety of pathogens. Active defense includes 

the production of antimicrobial compounds, cell wall reinforcement 

via the synthesis of lignin and callose, and the specific induction 

of elaborate defense signaling pathways. These include 

species level (non-host) resistance, race-specific resistance 

expressed both locally and systemically, and basal resistance. 

Race-specific resistance is induced when strain-specific 

avirulent (Avr) proteins from the pathogen associate directly/indirectly 

with cognate plant resistance (R) proteins (reviewed 

in Jones and Dangl 2006; Caplan et al. 2008). Induction 

of R-mediated signaling is often accompanied by the onset of 

a hypersensitive response (HR), a form of PCD resulting in 

necrotic lesions, at the site of pathogen entry (Dangl et al. 

1996). HR is one of the first visible manifestations of pathogeninduced 

host defenses, and is thought to help confine the 

pathogen to the dead cells. R-mediated signaling is also often 

accompanied by the induction of a robust form of resistance 

against secondary pathogens in the systemic parts of the 

plants, termed systemic acquired resistance (SAR) (Durrant 

and Dong 2004; Vlot et al. 2008; Spoel and Dong 2012). 

Identified as a form of plant immunity nearly 100 years ago, 

SAR is a highly desirable form of resistance that protects 

against a broad-spectrum of pathogens. SAR involves the generation 

of a mobile signal at the site of primary infection, which 

moves to and arms distal portions of a plant against subsequent 

secondary infections (Figure 27). The identification of this signal 

could greatly facilitate the use of SAR in protecting agriculturally 

important plants against a wide range of pathogens. Because of 

its unique mechanistic properties and its exciting potential applications 

in developing sustainable crop protection strategies, 

SAR has been one of the most intensely researched areas 

of plant biology. The last decade has witnessed considerable 

progress, and a number of signals contributing to SAR have 

been isolated and characterized. Despite concerted efforts to 

harness this mode of plant immunity, the plant defense field 

lacks a consensus regarding the identity of the SAR signal, 

whether this signal constitutes multiple molecular components, 

and how these component(s) might coordinate the systemic 

induction of broad-spectrum resistance. 

Among the signals contributing to SAR are salicylic acid (SA) 

and several components that feed into the SA pathway, including 

the methylated derivative of SA (MeSA; Park et al. 2007), 

the diterpenoid dehdryoabietinal (DA; Chaturvedi et al. 2012), 

the nine carbon (C9) dicarboxylic acid azelaic acid (AA; Jung 

et al. 2009), auxin (Truman et al. 2010), the phosphorylated 

sugar glycerol-3-phosphate (G3P; Chanda et al. 2011; Mandal 

et al. 2011), and two lipid transfer proteins (LTPs), Defective 

in Induced Resistance (DIR1; Maldonado et al. 2002) and AA 

insensitive (AZI1; Jung et al. 2009). Jasmonic acid (JA) has also 

been suggested to participate in SAR (Truman et al. 2007), but 

its precise role remains contentious (Attaran et al. 2009). The 

diverse chemical natures of the SAR-inducing molecules have 

led to the growing belief that SAR might involve the interplay of 

multiple diverse and independent signals. In this final section 

of the review, we will evaluate the role of SA and the recently 

identified mobile inducers of SAR. 

SA and SAR 

SA is a central and critical component of SAR. The biosynthesis 

of SA occurs via the shikimic acid pathway, which 

bifurcates into two branches after the biosynthesis of chorismic 

acid. In one branch, chorismic acid is converted to SA via


Figure 27. A simplified model summarizing the known mobile inducers of systemic acquired resistance (SAR). 

Pathogen infection induces an increase in the levels of glycerol-3-phosphate (G3P), azelaic acid (AA), dehdryoabietinal (DA), salicylic acid 

(SA) and methyl SA (MeSA). Some of the SA is converted to MeSA, whereas G3P is converted to an unknown derivative (indicated by 

asterisk). Owing to its volatile nature, a significant proportion of the MeSA is thought to escape by emissions. Although both SA and MeSA 

are phloem mobile, SA likely functions downstream of mobile signal generation. This is based on grafting experiments that indicate SA is not 

the SAR signal, yet basal SA is essential for G3P-, DA-, and AA-mediated SAR. DA induces SA accumulation in infected and distal tissues, 

whereas AA primes for SA biosynthesis in response to secondary pathogen stimulus. Neither G3P nor AA induces SA accumulation. G3P-, 

AA-, and DA-mediated SAR require the endoplasmic reticulum-localized lipid transfer-like protein, DIR1. Systemic movement of G3P and 

DIR1 is mutually inter-dependent. Upon transport, complex(es) comprising DIR1 and the G3P-derivative induce de novo G3P biosynthesis 

in the distal tissues. 

phenylalanine and cinnamic acid intermediates, and in the other 

branch chorismic acid is converted to SA via isochorismic acid. 

Two well characterized enzymes in these branches include 

PHENYLALANINE AMMONIA LYASE (PAL), which converts 

phenylalanine to cinnamic acid, and ISOCHORISMATE SYN- 

THASE (ICS), which catalyzes the conversion of chorismic 

acid to isochorismic acid (Wildermuth et al. 2001; Strawn et al. 

2007). 

Transcriptional profiling has shown that expression of hundreds 

of genes is altered during the development of SAR 

(Schenk et al. 2000; Wang et al. 2006; Truman et al. 2007; 

Chanda et al. 2011). This is likely to have wide-ranging effects, 

including strengthening of the cell wall and production of 

reactive oxygen species and SA. A hallmark of plants that have 

manifested SAR is the induction of pathogenesis-related (PR) 

proteins (Carr et al. 1987; Loon et al. 1987; Ward et al. 1991). 

These observations and the fact that exogenous SA induces 

PR expression led to the suggestion that SA was involved in 

SAR signaling. 

Exogenous application of SA or its synthetic functional 

analogues such as BTH (1,2,3-benzothiadiazole-7-carbothioic 

acid, S-methyl ester) also induce generalized defense against a 

variety of pathogens. Evidence supporting a role for SA in plant 

defense came from analysis of transgenic plants expressing 

the bacterial gene encoding salicylate hydroxylase, an enzyme 

that catalyzes conversion of SA to catechol. These transgenic 

plants were unable to accumulate free SA, showed compromised 

defense, and were unable to induce SAR (Gaffney et al. 

1993; Friedrich et al. 1995; Lawton et al. 1995). The fact that 

pathogen inoculation induces SA accumulation in both local 

and distal uninoculated tissues led to the hypothesis that SA 

might well be the phloem-mobile signal (Vernooij et al. 1994;


Shulaev et al. 1995). However, the timeframe of SAR signal 

movement precedes that of SA accumulation in the distal 

tissues. Moreover, SA-deficient rootstocks of plants expressing 

SA hydroxylase or plants suppressed in PAL expression are 

capable of activating SAR in the leaves of WT scions. These 

data argued against a role for SA as the phloem-mobile SAR 

signal (Vernooij et al. 1994; Pallas et al. 1996). Regardless, 

SA is required for the proper induction of SAR, and mutations 

in either the ICS or PAL pathway are sufficient to compromise 

SAR (Vernooij et al. 1994; Pallas et al. 1996; Wildermuth et al. 

2001). 

Radiolabel feeding experiments suggest that over 50% of 

SA in distal leaves is transported from the inoculated leaves 

with the remainder being synthesized de novo (Shulaev et al. 

1995; Molders et al. 1996). Whether the induced synthesis of 

SA in the distal tissues is required for SAR remains unclear. 

A marginal difference (∼10 ng/g FW) in SA levels between 

SAR-competent WT versus SAR-compromised SA-deficient 

scions grafted onto SA-deficient rootstocks suggests that an 

increase in SA accumulation may not be a prerequisite for the 

normal induction of SAR (Vernooij et al. 1994; Chanda et al. 

2011). 

In general, most of the endogenous SA is metabolized 

to a glucose conjugate, SA 2-O-β-D-glucose (SAG) or SA 

glucose ester (SGE), and this reaction is catalyzed by SA 

glucosyltransferases (Enyedi et al. 1992; Edwards 1994; Lee 

and Raskin 1998; Lee and Raskin 1999; Dean and Delaney 

2008; Song et al. 2008). Other derivates of SA include its 

methylated ester and methyl SA. (MeSA), and hydroxylated 

form gentisic acid (GA), both of which are also present as 

glucose conjugates. Exogenous GA induces a specific set of 

PR proteins in tomato that are not induced by SA, suggesting 

that SA and GA differ in their mode of action (Bellés et al. 

1999). 

Unlike SA, MeSA is biologically inactive and only functions 

when converted back to SA. MeSA is a well-characterized 

volatile organic compound that can function as an airborne 

defense signal and can mediate plant-plant communication 

(Shulaev et al. 1997; Koo et al. 2007). Conversion of SA to 

MeSA is mediated by SA methyltransferases (SAMT), also 

designated BA (benzoic acid)-/SA-MT because it can utilize 

either SA or BA as substrates (Chen et al. 2003; Effmert et al. 

2005; Koo et al. 2007). Overexpression of BSMT leads to the 

depletion of endogenous SA and SAG, as most of the available 

SA is converted to MeSA (Koo et al. 2007). This in turn is 

associated with increased susceptibility to bacterial and fungal 

pathogens, suggesting that levels of free SA, but not MeSA, 

are critical for plant immunity. Likewise, overexpression of the 

Arabidopsis SA glucosyltransferase (AtSGT1) also results 

in the depletion of SA and an increase in MeSA levels, 

which again correlates with increased susceptibility to bacterial 

pathogens (Song et al. 2008). 

MeSA accumulates in the phloem following induction of SAR, 

and this requires SAMT activity. Upon translocation to the distal 

tissues, MeSA is converted back to SA via MeSA esterase 

(Figure 27). Most of the MeSA accumulating in response to 

pathogen inoculation was shown to escape by volatile emissions 

(Attaran et al. 2009). Furthermore, Arabidopsis BSMT 

mutant plants do not accumulate MeSA, but remain SAR 

competent. This discrepancy was attributed to the dependency 

of MeSA-derived signaling on light (Liu et al. 2011), which is 

well-known to play an important role in plant defense (Karpinski 

et al. 2003; Roberts and Park 2006). Notably, the phloem 

translocation time of the SAR signal to distal tissues precedes 

the time of MeSA requirement; i.e., 48 h and 72 h post primary 

infection, respectively (Park et al. 2009; Chanda et al. 2011; 

Chaturvedi et al. 2012). This suggests that MeSA is unlikely 

to be the primary mobile signal, and possibly might act as a 

downstream contributor to SAR. 

Recent studies also suggest that defective SAR in dir1 plants 

is associated with increased expression of BSMT1, which correlates 

with increased accumulation of MeSA and a reduction 

in SA and SAG levels (Liu et al. 2011). However, this is in 

contrast with two other independent studies that showed normal 

SA levels in pathogen inoculated dir1 plants (Maldonado et al. 

2002; Chaturvedi et al. 2012). Some possibilities that might 

account for these discrepancies are disparate regulation of 

BSMT1 expression and the associated changes in MeSA and 

SA levels in different ecotypic backgrounds, and/or plant growth 

conditions, such as light, humidity, temperature, and wind. 

For example, light intensities could affect SA levels/defense 

responses since photoreceptors are well known to regulate 

both SA- and R-mediated signaling (Genoud et al. 2002; Jeong 

et al. 2010). 

Components that affect SAR by regulating SA levels 

Many proteins known to mediate SA-derived signaling have 

been identified as contributors to SAR. These include proteins 

involved in SA biosynthesis (including ICS and PAL), transport, 

and/or SA-dependent R-mediated signaling (ENHANCED 

DISEASE SUSCEPTIBILITY 1 (EDS1), EDS5, PHYTOALEXIN 

DEFICIENT 4 (PAD4), and SENESCENCE-ASSOCIATED 

gene 101 (SAG101)). The Arabidopsis EDS5 (also called 

SA INDUCTION-DEFICIENT 1) encodes a plastid-localized 

protein that shows homology to the bacterial multidrug and 

toxin extrusion transporter (MATE) proteins. EDS5 is required 

for the accumulation of SA after pathogen inoculation (Nawrath 

et al. 2002; Ishihara et al. 2008) and, consequently, a mutation 

in EDS5 causes enhanced susceptibility against oomycete, 

bacterial, and viral pathogens (Rogers and Ausubel 1997; 

Nawrath et al. 2002; Chandra-Shekara et al. 2004). Mutations 

in ICS1 and EDS5 lead to similar phenotypes (Venugopal 

et al. 2009), suggesting that EDS5 might be involved in

the transport of SA and/or its precursors across the plastid 

membrane. 

Currently, EDS5 is thought to act downstream of three other 

signaling components, EDS1 and PAD4, and NON-RACE 

SPECIFIC DISEASE RESISTANCE 1 (NDR1), which are 

required for basal R protein-mediated signaling and the SAR 

response (Glazebrook and Ausubel 1994; Century et al. 1995; 

Glazebrook et al. 1996; Parker et al. 1996; Century et al. 1997; 

Aarts et al. 1998; Zhou et al. 1998; Shapiro and Zhang 2001; 

Liu et al. 2002; Coppinger et al. 2004; Hu et al. 2005; Truman 

et al. 2007). Some of the Arabidopsis ecotypes express two 

functionally redundant isoforms of EDS1 which interact with 

each other as well as with the structurally similar PAD4 and 

SAG101 proteins (Feys et al. 2001; He and Gan 2002; Feys 

et al. 2005; García et al. 2010; Zhu et al. 2011). EDS1, PAD4, 

and SAG101 proteins also exist as a ternary complex (Zhu et al. 

2011). 

EDS1 interacts with several R proteins, suggesting that 

EDS1 and, by extension, PAD4 and SAG101, likely act at 

the R protein level (Bhattacharjee et al. 2011; Heidrich et al. 

2011; Zhu et al. 2011). Notably, mutations in EDS1, PAD4, 

and SAG101 lead to overlapping as well as independent 

phenotypes, suggesting that these proteins might function as 

complex(s) as well as individual proteins. Mutations in EDS1 

or PAD4 attenuate the expression of FLAVIN-DEPENDENT 

MONOOXYGENASE 1 (FMO1), which is required for SA accumulation 

in the distal tissues and, thereby, SAR (Mishina 

and Zeier 2006). The pathogen-induced SA levels are also 

regulated by AGD2-LIKE DEFENSE 1 (ALD1), which is induced 

in distal tissues after avirulent inoculation in a PAD4-dependent 

manner (Song et al. 2004a). The ALD1 encoded protein shows 

aminotransferase activity in vitro, suggesting that an amino 

acid-derived signal might participate in the regulation of SA 

levels and, thereby, SAR (Song et al. 2004b). 

SA signaling components that affect SAR 

The NON-EXPRESSOR OF PR1 (NPR1), an ankyrin repeat 

containing protein also called NON-INDUCIBLE IMMUNITY 1 

(NIM1; Delaney et al. 1995; Ryals et al. 1997) or SA INSEN- 

SITIVE 1) (SAI1; Shah et al. 1997), is considered a central 

regulator of SA-derived signaling. A mutation in Arabidopsis 

NPR1 abolishes SAR, suggesting that it is a positive regulator 

of SAR (Cao et al. 1994; Cao et al. 1997). Besides SA signaling 

and SAR, NPR1 also functions in induced systemic resistance 

(ISR) and, possibly, in regulating cross-talk between the SA 

and jasmonic acid (JA) pathways (van Wees et al. 2000; 

Kunkel and Brooks 2002; Lavicoli et al. 2003; Spoel and Dong 

2008). SA and NPR1 negatively affect the symbiotic interaction 

between Medicago and Rhizobium (Peleg-Grossman et al. 

2009), suggesting that SA and NPR1 are essential components 

of multiple signaling pathway(s). 


In the absence of SA, NPR1 exists as an oligomer via intermolecular 

disulfide bonding, and remains in the cytoplasm (Mou 

et al. 2003). Reducing conditions triggered upon activation of 

defense responses and the accumulation of SA result in the 

dissociation of the NPR1 oligomer into monomers, which are 

transported into the nucleus (Mou et al. 2003; Tada et al. 

2008). Within the nucleus, these NPR1 monomers interact 

with members of the TGACG motif binding transcription factors 

belonging to the basic leucine zipper (bZIP) protein family 

(Zhang et al. 1999; Després et al. 2000; Niggeweg et al. 

2000; Zhou et al. 2000; Chern et al. 2001; Fan and Dong 

2002; Kim and Delaney 2002). SA also induces the reduction 

of the disulfide bridges in TGA proteins, thereby allowing the 

proteins to interact with NPR1 with subsequent activation of 

gene expression (Després et al. 2003). 

Genetic evidence supporting a role for TGA factors in SAR 

was provided by the analysis of the tga2 tga5 tga6 triple mutant, 

which was unable to induce PR gene expression in response 

to SA and was defective in the onset of SAR (Zhang et al. 

2003). Recent studies have also shown that, like NPR1, TGA1 

also undergoes S-nitrosylation, which promotes the nuclear 

translocation of NPR1 and increases the DNA binding activity 

of TGA1 (Tada et al. 2008; Lindermayr et al. 2010). 

The monomerization of NPR1 also appears to be important 

for the activation of the NPR1 regulated members of the 

WRKY transcription factor family (Mou et al. 2003; Wang 

et al. 2006). In addition, NPR1 controls the expression of the 

protein secretory pathway genes in a TGA2-, TGA5- and TGA6independent 

manner (Wang et al. 2006). The nuclear NPR1 

is phosphorylated and recycled in a proteasome-dependent 

manner (Spoel et al. 2009). This turnover is required for 

the establishment of SAR. The Arabidopsis genome contains 

five paralogs of NPR1 (Liu et al. 2005). Like NPR1, 

NPR3 and NPR4 also interact with TGA proteins (Zhang 

et al. 2006). The npr3 npr4 mutant plants accumulate higher 

levels of NPR1 and, consequently, are unable to induce 

SAR. 

In a recent study, NPR3 and NPR4 were shown to bind SA 

and to function as adaptors of the Cullin 3 ubiquitin E3 ligase 

to mediate NPR1 degradation in an SA-dependent manner (Fu 

et al. 2012). NPR3 and NPR4 are neither the first nor the only 

known SA binding proteins. However, much like most of the 

plant hormone receptors, NPR3 and NPR4 are the only known 

proteins which regulate the proteasome-dependent recycling 

of a master regulator of the SA-signaling pathway. For this 

reason, these proteins have been suggested to serve as the 

long-sought-after SA receptors. In yet another study, NPR1 

was also proposed to function as a SA receptor (Wu et al. 

2012). Like NPR3/NPR4, NPR1 bound SA and the kinetics of 

this binding were similar to those of other receptor-hormone 

interactions. Thus, SA might well bind to multiple NPRs and 

differentially modulate their function(s).


Other proteins that bind SA include the MeSA esterase 

SABP2 (Kumar and Klessig 2003). Binding of SA to SABP2 

inhibits its esterase activity, resulting in the accumulation of 

MeSA. In addition, SA binds catalase (Chen et al. 1993) 

and carbonic anhydrase (Slaymaker et al. 2002), and inhibits 

the activities of the heme-iron-containing enzymes catalase, 

ascorbate peroxidase, and aconitase (Durner and Klessig 

1995). The ability of SA to chelate iron has been suggested 

as one of the mechanisms for SA-mediated inhibition of these 

enzymes (Rueffer et al. 1995). Tobacco plants silenced for 

carbonic anhydrase or aconitase show increased pathogen 

susceptibility, suggesting that these proteins are required for 

plant defense (Slaymaker et al. 2002; Moeder et al. 2007). 

Whereas NPR1 is an essential regulator of SA-derived signaling, 

many protein-induced pathways are known to activate 

SA-signaling in an NPR1-independent manner (Kachroo et al. 

2000; Takahashi et al. 2002; Raridan and Delaney 2002; Van 

der Biezen et al. 2002). Furthermore, a number of mutants have 

been isolated that induce defense SA signaling in an NPR1independent 

manner (Kachroo and Kachroo 2006). A screen 

for npr1 suppressors resulted in the identification of SNI1 (SUP- 

PRESSOR OF npr1, INDUCIBLE), a mutation which restores 

SAR in npr1 plants by de-repressing NPR1-dependent SA 

responsive genes (Li et al. 1999; Mosher et al. 2006). SNI1 has 

been suggested to regulate recombination rates through chromatin 

remodeling (Durrant et al. 2007). A subsequent screen 

for sni1 suppressors identified BRCA2 (BREAST CANCER) 

and RAD51D, which when mutated abolish the sni1-induced 

de-repression of NPR1-dependent gene expression (Durrant 

et al. 2007; Wang et al. 2010). SNI1 is therefore thought to 

act as a negative regulator that prevents recombination in 

the uninduced state. A role for SNI1, BRCA2 and RAD51D in 

recombination and defense suggests a possible link between 

these processes. Collectively, such findings support a key role 

for chromatin modification in the activation of plant defense and 

SAR (March-Díaz et al. 2008; Walley et al. 2008; Dhawan et al. 

2009; Ma et al. 2011). 

Mobile inducers of SAR 

Recent advances in the SAR field have led to the identification 

of four mobile inducers of SAR, including MeSA, AA, DA 

and G3P. All of these inducers accumulate in the inoculated 

leaves after pathogen inoculation and translocate systemically 

(Figure 27). The role of MeSA, a methylated derivative of 

SA, was discussed above. The dicarboxylic acid AA and the 

diterpenoid DA induce SAR in an ICS1-, NPR1-, DIR1-, and 

FMO1-dependent manner (Jung et al. 2009; Chaturvedi et al. 

2012). Their common requirements for these components suggest 

that AA- and DA-mediated SAR may represent different 

branches of a common signaling pathway. Indeed, exogenous 

application of low concentrations of DA and AA, that do not 

activate SAR, do so when applied together. However, AA and 

DA differ in their mechanism of SAR activation: DA increases 

SA levels in local and distal tissues, whereas AA primes for 

pathogen-induced biosynthesis of SA in the distal tissues. DA 

application also induces local accumulation of MeSA. Unlike 

DA, AA does not induce SA biosynthesis when applied by itself. 

This is intriguing, considering their common requirements for 

downstream factors. At present, the biosynthetic pathways for 

AA and DA and the biochemical basis of AA- and DA-induced 

SAR remain unclear. Furthermore, firm establishment of AA 

or DA as mobile SAR inducers awaits the demonstration that 

plants unable to synthesize these compounds are defective in 

SAR. 

G3P is a phosphorylated three-carbon sugar that serves 

as an obligatory component of glycolysis and glycerolipid 

biosynthesis. In the plant, G3P levels are regulated by enzymes 

directly/indirectly involved in G3P biosynthesis, as well as those 

involved in G3P catabolism. Recent results have demonstrated 

a role for G3P in R-mediated defense leading to SAR and 

defense against the hemibiotrophic fungus Colletotrichum higginsianum 

(Chanda et al. 2008). Arabidopsis plants containing 

the RPS2 gene rapidly accumulate G3P when infected with an 

avirulent (Avr) strain of the bacterial pathogen Pseudomonas 

syringae (avrRpt2); G3P levels peak within 6 h post-inoculation 

(Chanda et al. 2011). Strikingly, accumulation of G3P in the 

infected and systemic tissues precedes the accumulation of 

other metabolites known to be essential for SAR (SA, JA). 

Mutants defective in G3P synthesis are compromised in 

SAR, and this defect can be restored by the exogenous 

application of G3P (Chanda et al. 2011). Exogenous G3P 

also induces SAR in the absence of primary pathogen, albeit 

only in the presence of the LTP-like protein DIR1, which is a 

well-known positive regulator of SAR (Maldonado et al. 2002; 

Champigny et al. 2011; Chanda et al. 2011; Liu et al. 2011; 

Chaturvedi et al. 2012). DIR1 is also required for AA- and 

DA-mediated SAR, suggesting that DIR1 might be a common 

node for several SAR signals. Interestingly, G3P and DIR1 

are interdependent on each other for their translocation to the 

distal tissues. However, G3P does not interact directly with 

DIR1. Moreover, 14 C-G3P-feeding experiments have shown 

that G3P is translocated as a modified derivative during SAR. 

These results suggest that DIR1 likely associates with a G3Pderivative 

and, upon translocation to the distal tissues this 

complex, then induces the de novo synthesis of G3P and 

consequently SAR (Figure 27). 

This defense-related function of G3P is conserved because 

exogenous G3P can also induce SAR in soybean (Chanda 

et al. 2011). Exogenous application of G3P on local leaves 

induces transcriptional reprogramming in the distal tissues, 

which among other changes leads to the induction of the gene 

encoding a SABP2-like protein and repression of BSMT1. Thus, 

it is possible that G3P-mediated signaling functions to prime

the system for SA biosynthesis in the presence of an invading 

pathogen. However, exogenous G3P alone is not associated 

with increased SA biosynthesis, in either local or distal leaves. 

In this regard, it is interesting that similar to G3P, AA does 

not induce the expression of the genes normally associated 

with SA signaling, or those induced in response to exogenous 

SA. Induced SA accumulation diverts carbon, nitrogen and 

energy away from the plant’s primary metabolic pathways, 

which negatively impacts growth and development (Heil and 

Baldwin 2002; Heidel et al. 2004). Thus, chemicals like AA and 

G3P, which induce SAR without increasing SA levels, could be 

tremendously beneficial in improving crop resistance without 

affecting plant growth, development and ultimately yield. 

Fatty acids, lipids, cuticle and plant defense 

The primary role of G3P in plant metabolism is that of an 

obligatory precursor for glycerolipid biosynthesis. G3P enters 

lipid biosynthesis upon acylation with the fatty acid (FA) oleic 

acid (18:1) to form lyso-phosphatidic acid (lyso-PA), via the 

activity of the soluble plastidial G3P acyltransferase (GPAT). 

Genetically-based reductions in 18:1 levels induce constitutive 

defense signaling via the SA pathway (Kachroo et al. 2003, 

2004, 2005; Venugopal et al. 2009). Consequently, low 18:1containing 

plants exhibit enhanced resistance to bacterial and 

oomycete pathogens (Shah et al. 2001; Kachroo et al. 2001). 

Low 18:1 levels also specifically induce the expression of 

several R genes, which in turn induces defense signaling. 

SA and EDS1 regulate this low 18:1-dependent induction of 

defense responses in a redundant manner (Chandra-Shekara 

et al. 2007; Venugopal et al. 2009; Xia et al. 2009). Interestingly, 

it has also been shown that 18:1 levels regulate the NOA1 

(NITRIC OXIDE ASSOCIATED) protein and thereby nitric oxide 

levels. Thus, the increased NO in low 18:1-containing plants 

is responsible for their altered defense related phenotypes 

(Mandal et al. 2012). 

A number of cuticle-defective mutants are compromised 

in SAR (Xia et al. 2009, 2010). Whereas acp4 plants can 

generate the signal required for inducing SAR, they are unable 

to respond to it. This loss of ability to “perceive” the SAR signal 

appears to be related to the defective cuticle of acp4 plants, 

because mechanical abrasion of the cuticle disrupts SAR in 

WT plants. This SAR-disruptive effect of cuticle abrasion is 

highly specific because it hinders SAR only during the timeframe 

of mobile signal generation and translocation to distal 

tissues; it does not alter local defenses. These observations 

suggest that cuticle-derived component(s) likely participate in 

processing/perception of the SAR signal(s). The requirement 

for the plant cuticle in SAR development, the presence of 

lipids and FAs in petiole exudates (Madey et al. 2002; Behmer 

et al. 2011; Guelette et al. 2012), and the derivatization of 

G3P (a glycerolipid precursor) into an unknown compound 


that translocates with the LTP DIR1, all suggest a role for 

lipids/FAs/sugars in SAR. 

Clearly, more work is required to dissect the relationships 

between these chemically diverse signals. For example, what 

factors govern the transport and movement of these signals 

through the vascular system, and their subsequent unloading 

into distal tissues? How are these signals processed at 

their systemic destinations? What reprogramming of metabolic 

events is required to activate defense and subsequently depress 

the tissues to the resting phase? 

Future Perspectives 

The emergence of the tracheophyte-based vascular system 

had major impacts on the evolution of terrestrial biology, 

in general, through its role in facilitating the development 

of plants with increased stature, photosynthetic output, 

and ability to colonize a greatly expanded range of environmental 

habitats. Significant insights have been gained 

concerning the genetic and hormonal networks that cooperate 

to orchestrate vascular development in the angiosperms, 

and progress is currently being made for the 

gymnosperms. However, much remains to be learned in 

terms of the early molecular events that led to the co-opting 

of pre-tracheophyte transcription factors and hormone signaling 

pathways, in order to establish the developmental 

programs that underlay the emergence of the tracheids 

as an effective/superior system for water conduction over 

the WCCs/hydroids. The same situation holds for the 

FCCs/leptoids to sieve cell/SE transition. Certainly, future 

application of genomic and molecular tools should offer 

important insights into the relationships between these preand 

post-tracheophyte/SE programs. 

Cost-effective, high-throughput sequencing technologies 

are opening the door to studies that integrate plant functional 

genomics with physiology and ecology. Such studies 

will likely provide important insights into novel strategies, 

achieved by different plant families, to refine the operational 

characteristics of their xylem/phloem transport systems to 

meet the challenges imposed by their specific ecological 

niches. Much of our current knowledge of vascular development 

is built upon studies conducted on “model” systems 

such as Arabidopsis. Although the general principles are 

likely to apply to most, if not all, advanced tracheophytes, 

many surprises are likely to be unearthed as research 

expands to cover plants with increased stature and concomitant 

challenges in terms of environmental inputs. 

Fundamental details are now established in terms of 

the mechanics underlying the thermodynamics of bulk flow 

though both the xylem and phloem. For the xylem, important


questions remain to be resolved, including the mechanism 

by which a single cavitation event can propagate within 

the tracheid/vessel system, the processes involved in refilling 

of embolized tracheary elements, especially when 

the transpiration stream is under tension, and the degree 

to which pit architecture between species contributes to 

ecological fitness. With regard to the phloem, one of the 

most fundamental questions that remains to be resolved 

relates to the mechanism(s) by which the plant integrates 

sink demand with source capacity to optimize growth under 

prevailing environmental conditions. The phloem manifold 

hypothesis and the concept of delivery to various sink 

tissues being controlled by local PD properties warrants 

close attention. 

The role of the plant vascular system as a long-distance 

signaling system for integration of abiotic and biotic inputs 

is also firmly established. However, much remains to be 

learned concerning the nature of the xylem- and phloemmobile 

signals that function in nutrient homeostasis, environmental 

signaling to control stomatal density in emerging 

leaves, pathogen-host plant interactions, etc. In addition, 

the discovery that angiosperm phloem sap collected from 

the enucleate sieve tube system contains a broad spectrum 

of proteins and RNA species is consistent with the phloem 

functioning as a sophisticated communication system. A 

number of pioneering studies have demonstrated the role 

of protein and RNA as long-distance signaling agents. 

Future studies are required to both to expand the number 

of proteins/RNA investigated as well as to focus on the 

molecular mechanisms involved in determining how these 

signaling agents are targeted to specific sink tissues. 

Commercial applications of knowledge gained on the 

development and functions of the plant vascular system 

are likely to be boundless. Access to methods to control 

source-sink relationships would have profound effects over 

yield potential and biomass production for the biofuels 

industry. Modifications to secondary xylem development 

will likely allow for engineering of wood that has unique 

properties for industrial applications. Engineering of novel 

traits for agriculture will likely be achieved by acquiring a 

better understanding of the root-to-shoot and shoot-to-root 

signaling networks. Thus, the future for research on plant 

vascular biology is very bright indeed! 

Acknowledgements 

Work in the authors’ laboratories was supported in part by the 

National Science Foundation (grants IOS-0752997 and IOS- 

0918433 to WJL; grants IOS#0749731, IOS#051909 to PK), the 

Department of Energy, Division of Energy Biosciences (grant 

DE-FG02-94ER20134 to WJL), the US Department of Agriculture, 

Agricultural Research Service (under Agreement number 

58-6250-0-008 to MAG), the Spanish Ministry of Science and 

Innovation (MICINN) (grants AGL2007-61948 and AGL2009- 

09018 to AFLM), the Ministry of Education, Science, Sports 

and Culture of Japan (grant 19060009 to HF), from the Japan 

Society for the Promotion of Science (JSPS grant 23227001 

to HF), and from the NC-CARP project (to HF), the National 

Key Basic Research Program of China (grant 2012CB114500 

to XH), the National Natural Science Foundation of China (grant 

31070156 to XH), and the NSFC-JSPS cooperation project 

(grant 31011140070 to HF and XH). 

Received 20 Jan. 2013 Accepted 19 Feb. 2013 

References 

Aarts N, Metz M, Holub E, Staskawicz BJ, Daniels MJ, Parker 

JE (1998) Different requirements for EDS1 and NDR1 by disease 

resistance genes define at least two R gene-mediated signaling 

pathways in Arabidopsis. Proc. Natl. Acad. Sci. USA 95, 10306– 

10311. 

Agrawal B, Lakshmanan V, Kaushik S, Bais HP (2012) Natural 

variation among Arabidopsis accessions reveals malic acid 

as a key mediator of Nickel (Ni) tolerance. Planta 236, 477– 

489. 

Aki T, Shigyo M, Nakano R, Yoneyama T, Yanagisawa S (2008) 

Nano scale proteomics revealed the presence of regulatory proteins 

including three FT-Like proteins in phloem and xylem saps from rice. 

Plant Cell Physiol. 49, 767–790. 

Akiyama K, Matsuzaki K, Hayashi H (2005) Plant sesquiterpenes 

induce hyphal branching in arbuscular mycorrhizal fungi. Nature 

435, 824–827. 

Alakonya A, Kumar R, Koenig D, Kimura S, Townsley B, Runo 

S, Garces HM, Kang J, Yanez A, David-Schwartz R, Machuka 

J, Sinha N (2012) Interspecific RNA interference of SHOOT 

MERISTEMLESS-Like disrupts Cuscuta pentagona plant parasitism. 

Plant Cell 4, 3153–3166. 

Alder NN, Pockman WT, Sperry JS, Nuismer S (1997) Use of 

centrifugal force in the study of xylem cavitation. J. Exp. Bot. 48, 

665–674. 

Aloni R (1987) Differentiation of vascular tissues. Plant Physiol. 38, 

179–204. 

Aloni R, Schwalm K, Langhans M, Ullrich CI (2003) Gradual shifts in 

sites of free-auxin production during leaf-primordium development 

and their role in vascular differentiation and leaf morphogenesis in 

Arabidopsis. Planta 216, 841–853. 

Alves S, Nabais C, Simões Gonçalves MDL, Correia dos Santos MM 

(2011) Nickel speciation in the xylem sap of the hyperaccumulator 

Alyssum serpyllifolium ssp. lusitanicum growing on serpentine soils 

of northeast Portugal. Plant Physiol. 168, 1715–1722.

Aly R, Cholakh H, Joel DM, Leibman D, Steinitz B, Zelcer A, Naglis 

A, Yarden O, Gal-On A (2009) Gene silencing of mannose 6phosphate 

reductase in the parasitic weed Orobanche aegyptiaca 

through the production of homologous dsRNA sequences in the 

host plant. Plant Biotechnol. J. 7, 487–498. 

Amiard V, Mueh K, Demming-Adams B, Ebbert V, Turgeon R (2005) 

Anatomical and photosynthetic acclimation to the light environment 

in species with differing mechanisms of phloem loading. Proc. Natl. 

Acad. Sci. USA 102, 12968–12973. 

Anderegg WRL, Berry JA, Smith DD, Sperry JS, Anderegg LDL, 

Field CB (2012) The roles of hydraulic and carbon stress in a 

widespread climate-induced forest die-off. Proc. Am. Acad. Arts Sci. 

109, 233–237. 

Andrés-Colás N, Sancenón V, Rodríguez-Navarro S, Mayo S, Thiele 

DJ, Ecker JR, Puig S, Peñarrubia L (2006) The Arabidopsis 

heavy metal P-type ATPase HMA5 interacts with metallochaperones 

and functions in copper detoxification of roots. Plant J. 45, 

225–236. 

Aoki K, Suzui N, Fujimaki S, Dohmae N, Yonekura-Sakakibara 

K, Fujiwara T, Hayashi H, Yamaya T, Sakakibara H (2005) 

Destination-selective long-distance movement of phloem proteins. 


Aoyama T, Kobayashi T, Takahashi M, Nagasaka S, Usuda K, 

Kakei Y, Ishimaru Y, Nakanishi H, Mori S, Nishizawa NK (2009) 

OsYSL18 is a rice iron(III)-deoxymugineic acid transporter specifically 

expressed in reproductive organs and phloem of lamina joints. 

Plant Mol. Biol. 70, 681–692. 

Atkins CA, Smith PMC, Rodriguez-Medina C (2011) Macromolecules 

in phloem exudates – A review. Protoplasma 248, 165–172. 

Attaran E, Zeier TE, Griebel T, Zeier J (2009) Methyl salicylate 

production and jasmonate signaling are not essential for systemic 

acquired resistance in Arabidopsis. Plant Cell 21, 954–971. 

Avci U, Petzold HE, Ismail IO, Beers EP, Haigler CH (2008) Cysteine 

proteases XCP1 and XCP2 aid micro-autolysis within the intact 

central vacuole during xylogenesis in Arabidopsis roots. Plant J. 

56, 303–315. 

Bai SL, Kasai A, Yamada K, Li TZ, Harada T (2011) A mobile 

signal transported over a long distance induces systemic transcriptional 

gene silencing in a grafted partner. J. Exp. Bot. 62, 

4561–4570. 

Baima S, Nobili F, Sessa G, Lucchetti S, Ruberti I, Morelli G 

(1995) The expression of the athb-8 homeobox gene is restricted to 

provascular cells in Arabidopsis thaliana. Development 121, 4171– 

4182. 

Bahrun A, Jensen CR, Asch F, Mogensen VO (2002) Droughtinduced 

changes in xylem pH, ionic composition, and ABA concentration 

act as early signals in field-grown maize (Zea mays L.). 

J. Exp. Bot. 53, 251–263. 

Ball MC, Canny MJ, Huang CX, Egerton JJG, Wolfe J (2006) 

Freeze/thaw-induced embolism depends on nadir temperature: The 

heterogeneous hydration hypothesis. Plant Cell Environ. 29, 729– 

745. 


Banerjee AK, Chatterjee M, Yu YY, Suh SG, Miller WA, Hannapel 

DJ (2006) Dynamics of a mobile RNA of potato involved in a longdistance 

signaling pathway. Plant Cell 18, 3443–3457. 

Barnes A, Bale J, Constantinidou C, Ashton P, Jones A, Pritchard 

J (2004) Determining protein identity from sieve element sap in 

Ricinus communis L. by quadrupole time of flight (Q-TOF) mass 

spectrometry. J. Exp. Bot. 55, 1473–1481. 

Barnett JR (1982) Plasmodesmata and pit development in secondary 

xylem elements. Planta. 155, 251–260. 

Batailler B, Lemaitre T, Vilaine F, Sanchez C, Renard D, Cayla T, 

Beneteau J, Dinant S (2012) Soluble and filamentous proteins in 

Arabidopsis sieve elements. Plant Cell Environ. 35, 1258–1273. 

Bauby H, Divol F, Truernit E, Grandjean O, Palauqui JC (2007) Protophloem 

differentiation in early Arabidopsis thaliana development. 


Baxter I, Muthukumar B, Hyeong CP, Buchner P, Lahner B, Danku 

J, Zhao K, Lee J, Hawkesford MJ, Guerinot ML, Salt DE (2008) 

Variation in molybdenum content across broadly distributed populations 

of Arabidopsis thaliana is controlled by a mitochondrial 

molybdenum transporter (MOT1). PLoS Genetics 4, e1000004. 

Behmer ST, Grebenok RJ, Douglas AE (2011) Plant sterols and host 

plant suitability for a phloem-feeding insect. Funct. Ecol. 25, 484– 

491. 

Belimov AA, Dodd IC, Hontzeas N, Theobald JC, Safronova 

VI, Davies WJ (2009) Rhizosphere bacteria containing 

1-aminocyclopropane-1-carboxylate deaminase increase yield 

of plants grown in drying soil via both local and systemic hormone 

signalling. New Phytol. 181, 413–423. 

Bellés JM, Garro R, Fayos J, Navarro P, Primo J, Conejero V (1999) 

Gentisic acid as a pathogen-inducible signal, additional to salicylic 

acid for activation of plant defenses in tomato. Mol. Plant-Microbe 

Interact. 12, 227–235. 

Benschop JJ, Mohammed S, O’Flaherty M, Heck AJR, Slijper 

M, Menke FLH (2007) Quantitative phosphoproteomics of 

early elicitor signaling in Arabidopsis. Molec. Cell. Proteomics 6, 

1198–1214. 

Benkova E, Michniewicz M, Sauer M, Teichmann T, Seifertova 

D, Jurgens G, Friml J (2003) Local, efflux-dependent auxin 

gradients as a common module for plant organ formation. Cell 115, 

591–602. 

Benning UF, Tamot B, Guelette BS, Hoffmann-Benning S (2012) 

New aspects of phloem-mediated long-distance lipid signaling in 

plants. Front. Plant Sci. 3, 53. 

Berleth T, Mattsson J, Hardtke CS (2000) Vascular continuity and 

auxin signals. Trends Plant Sci. 5, 387–393. 

Betsuyaku S, Takahashi F, Kinoshita A, Miwa H, Shinozaki K, 

Fukuda H, Sawa S (2011) Mitogen-activated protein kinase regulated 

by the CLAVATA receptors contributes to shoot apical 

meristem homeostasis. Plant Cell Physiol. 52, 14–29. 

Beveridge CA, Ross JJ, Murfet IC (1994) Branching mutant rms-2 

in Pisum sativum (grafting studies and endogenous indole-3-acetic 

acid levels). Plant Physiol. 104, 953–959.


Bhattacharjee S, Halane MK, Kim SH, Gassmann W (2011) 

Pathogen effectors target Arabidopsis EDS1 and alter its interaction 

with immune regulators. Science 334, 1405–1408. 

Bieleski RL (1973) Phosphate pools, phosphate transport, and phosphate 

availability. Ann. Rev. Plant Physiol. 24, 225–252. 

Bishopp A, Help H, El-Showk S, Weijers D, Scheres B, Friml 

J, Benková E,Mähönen AP, Helariutta Y (2011a) A mutually 

inhibitory interaction between auxin and cytokinin specifies vascular 

pattern in roots. Curr. Biol. 21, 917–926. 

Bishopp A, Lehesranta S, Vatén A, Help H, El-Showk S, Scheres B, 

Helariutta K, Mähönen AP, Sakakibara H, Helariutta Y (2011b) 

Phloem-transported cytokinin regulates polar auxin transport and 

maintains vascular pattern in the root meristem. Curr. Biol. 21, 927– 

932. 

Blindauer CA, Schmid R (2010) Cytosolic metal handling in plants: 

Determinants for zinc specificity in metal transporters and metallothioneins. 

Metallomics 2, 510–529. 

Bolaños L, Lukaszewski K, Bonilla I, Blevins D (2004) Why boron? 

Plant Physiol. Biochem. 42, 907–912. 

Bollhoner B, Prestele J, Tuominen H (2012) Xylem cell death: 

Emerging understanding of regulation and function. J. Exp. Bot. 

63, 1081–1094. 

Bonke M, Thitamadee S, Mähönen AP, Hauser MT, Helariutta Y 

(2003) APL regulates vascular tissue identity in Arabidopsis. Nature 

426, 181–186. 

Boot KJM, Libbenga KR, Hille SC, Offringa R, van Duijn B (2012) 

Polar auxin transport: An early invention. J. Exp. Bot. 63, 4213– 

4218. 

Bostwick DE, Dannenhoffer JM, Skaggs MI, Lister RM, Larkins BA, 

Thompson GA (1992) Pumpkin phloem lectin genes are specifically 

expressed in companion cells. Plant Cell 4, 1539–1548. 

Bowman JL, Floyd SK (2008) Patterning and polarity in seed plant 

shoots. Annu. Rev. Plant Biol. 59, 67–88. 

Boyce KC, Brodribb TJ, Feild TS, Zwieniecki MA (2009) Angiosperm 

leaf vein evolution was physiologically and environmentally transformative. 

Proc. R. Soc. Series B 276, 1771–1776. 

Boyce KC, Zwieniecki MA (2012) Leaf fossil record suggests limited 

influence of atmospheric CO2 on terrestrial productivity prior to 

angiosperm evolution. Proc. Natl. Acad. Sci. USA 109, 10403– 

10408. 

Brady SM, Orlando DA, Lee JY, Wang JY, Koch J, Dinneny 

JR, Mace D, Ohler U, Benfey PN (2007) A high-resolution root 

spatiotemporal map reveals dominant expression patterns. Science 

318, 801–806. 

Brady SM, Zhang L, Megraw M, Martinez NJ, Jiang E, Yi CS, 

Liu W, Zeng A, Taylor-Teeples M, Kim D, Ahnert S, Ohler U, 

Ware D, Walhout AJ, Benfey PN (2011) A stele-enriched gene 

regulatory network in the Arabidopsis root. Mol. Syst. Biol. 7, 

459. 

Brewer PB, Dun EA, Ferguson BJ, Rameau C, Beveridge CA (2009) 

Strigolactone acts downstream of auxin to regulate bud outgrowth 

in pea and Arabidopsis. Plant Physiol. 150, 482–493. 

Brewer PB, Koltai H, Beveridge CA (2013) Diverse roles of strigolactones 

in plant development. Mol. Plant 6, 18–28. 

Briggs LJ (1950) Limiting negative pressure of water. J. Appl. Phys. 

21, 721–722. 

Brodersen C, McElrone AJ, Choat B, Matthews MA, Shackel KA 

(2010) The dynamics of embolism repair in xylem: in vivo visualizations 

using high-resolution computed tomography. Plant Physiol. 

154, 1088–1095. 

Brodersen P, Voinnet O (2006) The diversity of RNA silencing pathways 

in plants. Trends Genet. 22, 268–280. 

Brodribb TJ (2009) Xylem hydraulic physiology: The functional backbone 

of terrestrial plant productivity. Plant Sci. 177, 245–251. 

Brodribb TJ, Holbrook NM (2005) Water stress deforms tracheids 

peripheral peripheral to the leaf vein of a tropical conifer. Plant 

Physiol. 137, 1139–1146. 

Brodribb TJ, McAdam SAM (2011) Passive origins of stomatal control 

in vascular plants. Science 331, 582–585. 

Brown PH, Hu H (1996) Phloem mobility of boron is species dependent: 

Evidence for phloem mobility in sorbitol-rich species. Ann. Bot. 77, 

497–505. 

Bucci SJ, Scholz FG, Goldstein G, Meinzer FC, Sternberg LDSL 

(2003) Dynamic changes in hydraulic conductivity in petioles of 

two savanna species: Factors and mechanisms contributing to the 

refilling of embolized vessels. Plant Cell Environ. 26, 1633–1645. 

Buhtz A, Pieritz J, Springer F, Kehr J (2010) Phloem small RNAs, 

nutrient stress responses, and systemic mobility. BMC Plant Biol. 

10, 64. 

Buhtz A, Springer F, Chappell L, Baulcombe DC, Kehr J (2008) 

Identification and characterization of small RNAs from the phloem 

of Brassica napus. Plant J. 53, 739–749. 

Burleigh SH, Harrison MJ (1999) The down-regulation of Mt4-like 

genes by phosphate fertilization occurs systemically and involves 

phosphate translocation to the shoots. Plant Physiol. 119, 241–248. 

Butenko MA, Vie AK, Brembu T, Aalen RB, Bones AM (2009) Plant 

peptides in signalling: Looking for new partners. Trends Plant Sci. 

14, 255–263. 

Butterfield BG (1995) Vessel element differentiation. In: Iqbal M, ed. 

The Cambial Derivatives. Encyclopedia of Plant Anatomy. Gebruder 

Borntraeger, Berlin. pp. 93–106. 

Cakmak I, Kirkby EA (2008) Role of magnesium in carbon partitioning 

and alleviating photo-oxidative damage. Physiol. Plant 133, 692– 

704. 

Canny MJ (1975) Mass Transfer. In: MH Zimermann, JA Milburn, eds. 

Transport in Plants I. Phloem Transport. Springer-Verlag, Berlin. 

pp. 139–153. 

Canny MJ (1998) Transporting water in plants. Amer. Sci. 86, 152–159. 

Cano-Delgado A, Lee JY, Demura T (2010) Regulatory mechanisms 

for specification and patterning of plant vascular tissues. Annu. Rev. 

Cell Dev. Biol. 26, 605–637. 

Cao H, Bowling SA, Gordon S, Dong X (1994) Characterization of an 

Arabidopsis mutant that is nonresponsive to inducers of systemic 

acquired resistance. Plant Cell 6, 1583–1592.

Cao H, Glazebrook J, Clark JD, Volko S, Dong X (1997) The 

Arabidopsis NPR1 gene that controls systemic acquired resistance 

encodes a novel protein containing ankyrin repeats. Cell 88, 

57–64. 

Caplan J, Padmanabhan M, Dinesh-Kumar SP (2008) Plant NB-LRR 

immune receptors: From recognition to transcriptional reprogramming. 

Cell Host Microbe 3, 126–135. 

Carlquist S (1984) Vessel grouping in dicotyledon wood: Significance 

and relationship to imperforate tracheary elements. Aliso 10, 505– 

525. 

Carlsbecker A, Lee JY, Roberts CJ, Dettmer J, Lehesranta S, Zhou 

J, Lindgren O, Moreno-Risueno MA, Vatén A, Thitamadee S, 

Campilho A, Sebastian J, Bowman JL, Helariutta Y, Benfey 

PN (2010) Cell signalling by microRNA165/6 directs gene dosedependent 

root cell fate. Nature 465, 316–321. 

Carr JP, Dixon DC, Nikolau BJ, Voelkerding KV, Klessig DF (1987) 

Synthesis and localization of pathogenesis-related proteins in tobacco. 

Mol. Cell Biol. 7, 1580–1583. 

Catford JG, Staehelin C, Lerat S, Piché Y, Vierheilig H (2003) 

Suppression of arbuscular mycorrhizal colonization and nodulation 

in split-root systems of alfalfa after pre-inoculation and treatment 

with Nod factors. J. Exp. Bot. 54, 1481–1487. 

Century KS, Holub EB, Staskawicz BJ (1995) NDR1, a locus of 

Arabidopsis thaliana that is required for disease resistance to both 

a bacterial and a fungal pathogen. Proc. Natl. Acad. Sci. USA 92, 

6597–6601. 

Century KS, Shapiro AD, Repetti PP, Dahlbeck D, Holub E, 

Staskawicz BJ (1997) NDR1, a pathogen-induced component 

required for Arabidopsis disease resistance. Science 278, 1963– 

1965. 

Champigny MJ, Shearer H, Mohammad A, Haines K, Neumann M, 

Thilmony R, He SY, Fobert P, Dengler N, Cameron RK (2011) 

Localization of DIR1 at the tissue, cellular and subcellular levels 

during systemic acquired resistance in Arabidopsis using DIR1:GUS 

and DIR1:EGFP reporters. BMC Plant Biol. 11, 125. 

Chanda B, Venugopal SC, Kulshrestha S, Navarre D, Downie 

B, Vaillancourt L, Kachroo A, Kachroo P (2008) Glycerol-3phosphate 

levels are associated with basal resistance to the 

hemibiotrophic fungus Colletotrichum higginsianum in Arabidopsis. 

Plant Physiol. 147, 2017–2029. 

Chanda B, Ye X, Mandal M, Yu K, Sekine KT, Gao QM, Selote 

D, Hu Y, Stromberg A, Navarre D, Kachroo A, Kachroo P 

(2011) Glycerol-3-phosphate is a critical mobile inducer of systemic 

immunity in plants. Nat. Genet. 43, 421–427. 

Chandra-Shekara AC, Navarre D, Kachroo A, Kang HG, Klessig 

D, Kachroo P (2004) Signaling requirement and role of salicylic 

acid in HRT- and rrt-mediated resistance to turnip crinkle virus in 

Arabidopsis. Plant J. 40, 647–659. 

Chandra-Shekara AC, Venugopal SC, Kachroo A, Kachroo P (2007) 

Plastidal fatty acid levels regulate resistance gene-dependent defense 

signaling in Arabidopsis. Proc. Natl. Acad. Sci. USA 104, 

7277–7282. 


Chaturvedi R, Krothapalli K, Makandar R, Nandi A, Sparks A, Roth 

M,WeltiR,ShahJ(2008) Plastid omega3-fatty acid desaturasedependent 

accumulation of a systemic acquired resistance inducing 

activity in petiole exudates of Arabidopsis thaliana is independent 

of jasmonic acid. Plant J. 54, 106–117. 

Chaturvedi R, Venables B, Petros R, Nalam V, Li M, Wang 

X, Takemoto LJ, Shah J (2012) An abietane diterpenoid is a 

potent activator of systemic acquired resistance. Plant J. 71, 

161–172. 

Chen F, D’Auria JC, Tholl D, Ross JR, Gershenzon J, Noel JP, Pichersky 

E (2003) An Arabidopsis thaliana gene for methylsalicylate 

biosynthesis, identified by a biochemical genomics approach, has 

a role in defense. Plant J. 36, 577–588. 

Chen HM, Pang Y, Zeng J, Ding Q, Yin SY, Liu C, Lu MZ, Cui KM, He 

XQ (2012a) The Ca2+ -dependent DNases are involved in secondary 

xylem development in Eucommia ulmoides. J. Integr. Plant Biol. 54, 

456–470. 

Chen LQ, Qu XQ, Hou BH, Sosso D, Osorio S, Fernie AR, Frommer 

WB (2012b) Sucrose efflux mediated by SWEET proteins as a key 

step for phloem transport. Science 335, 207–211. 

Chen Z, Silva H, Klessig DF (1993) Involvement of reactive oxygen 

species in the induction of systemic acquired resistance by salicylic 

acid in plants. Science 242, 883–886. 

Chern MS, Fitzgerald HA, Yadav RC, Canlas PE, Dong X, Ronald 

PC (2001) Evidence for a disease-resistance pathway in rice similar 

to the NPR1-mediated signaling pathway in Arabidopsis. Plant J. 27, 

101–113. 

Chincinska IA, Liesche J, Krügel U, Michalska J, Geigenberger P, 

Grimm B, Kühn C (2008) Sucrose transporter StSUT4 from potato 

affects flowering, tuberization, and shade avoidance response. 


Chiou TJ, Lin SI (2011) Signaling network in sensing phosphate 

availability in plants. Annu. Rev. Plant Biol. 62, 185–206. 

Choat B, Cobb AR, Jansen S (2008) Structure and function of bordered 

pits: New discoveries and impacts on whole-plant hydraulic 

function. New Phytol. 177, 608–626. 

Choat B, Drayton WM, Brodersen C, Matthews MA, Schackel KA, 

Wada H, McElrone AJ (2010) Measurement of vulnerability to 

water stress-induced cavitation in grapevine: A comparison of four 

techniques applied to a long-vesseled species. Plant Cell Environ. 

33, 1502–1512. 

Choat B, Gambetta GA, Shakel KA, Matthews MA (2009) Vascular 

function in grape berries across development and its relevance to 

apparent hydraulic isolation. Plant Physiol. 151, 1677–1687. 

Choat B, Jansen S, Zwieniecki MA, Smets E, Holbrook NM (2004) 

Changes in pit membrane porosity due to deflection and stretching: 

The role of vestured pits. J. Exp. Bot. 55, 1569–1575. 

Choat B, Medek DE, Stuart SA, Pasquet-Kok J, Egerton JJG, 

Salari H, Sack L, Ball MC (2011) Xylem traits mediate a trade-off 

between resistance to freeze-thaw-induced embolism and photosynthetic 

capacity in overwintering evergreens. New Phytol. 191, 

996–1005.


Christman MA, Sperry JS, Adler FR (2009) Testing the rare pit 

hypothesis in three species of Acer. New Phytol. 182, 664–674. 

Christman MA, Sperry JS (2010) Single vessel flow measurements 

indicate scalariform perforation plates confer higher resistance to 

flow than previously estimated. Plant Cell Environ. 33, 431–443. 

Christman MA, Sperry JS, Smith DD (2012) Rare pits, large vessels, 

and extreme vulnerability to cavitation in a ring-porous tree species. 

New Phytol. 193, 713–720. 

Christmann A, Weiler EW, Steudle E, Grill E (2007) A hydraulic signal 

in root-to-shoot signalling of water shortage. Plant J. 52, 167–174. 

Clay NK, Nelson T (2005) Arabidopsis thickvein mutation affects vein 

thickness and organ vascularization, and resides in a provascular 

cell-specific spermine synthase involved in vein definition and in 

polar auxin transport. Plant Physiol. 138, 767–777. 

Cochard H, Cruiziat P, Tyree MT (1992) Use of positive pressures to 

establish vulnerability curves: Further support for the air-seeding 

hypothesis and implications for pressure-volume analysis. Plant 

Physiol. 100, 205–209. 

Cochard H, Froux F, Mayr S, Coutand C (2004) Xylem wall collapse 

in water-stressed pine needles. Plant Physiol. 134, 401–408. 

Cochard H, Gaelle D, Bodet C, Tharwat I, Poirier M, Ameglio T 

(2005) Evaluation of a new centrifuge technique for rapid generation 

of xylem vulnerability curves. Physiol. Plant. 124, 410–418. 

Cochard H, Herbette S, Barigah T, Vilagrosa A (2010a) Does sample 

length influence the shape of vulnerability to cavitation curves? A 

test with the Cavitron spinning technique. Plant Cell Environ. 33, 

1543–1552. 

Cochard H, Herbette S, Hernandez E, Holtta T, Mencuccini M 

(2010b) The effects of sap ionic composition on xylem vulnerability 

to cavitation. J. Exp. Bot. 61, 275–285. 

Cochard H, Holtta T, Herbette S, Delzon S, Mencuccini M (2009) 

New insights into the mechanisms of water-stress-induced cavitation 

in conifers. Plant Physiol. 151, 949–954. 

Colombani A, Djerbi S, Bessueille L, Blomqvist K, Ohlsson A, 

Berglund T, Teeri TT, Bulone V (2004) In vitro synthesis of 

(1 → 3)-β-D-glucan (callose) and cellulose by detergent extracts of 

membranes from cell suspension cultures of hybrid aspen. Cellulose 

11, 313–327. 

Conte SS, Walker EL (2011) Transporters contributing to iron trafficking 

in plants. Mol. Plant 4, 464–476. 

Coppinger P, Pepetti PP, Day B, Dahlbeck D, Mehlert A, Staskawicz 

BJ (2004) Overexpression of the plasma membrane-localized 

NDR1 protein results in enhanced bacterial resistance in Arabidopsis 

thaliana. Plant J. 40, 225–237. 

Corbesier L, Vincent C, Jang SH, Fornara F, Fan QZ, Searle I, 

Giakountis A, Farrona S, Gissot L, Turnbull C, Coupland G 

(2007) FT protein movement contributes to long-distance signaling 

in floral induction of Arabidopsis. Science 316, 1030–1033. 

Courtois-Moreau CL, Pesquet E, Sjodin A, Muniz L, Bollhoner B, 

Kaneda M, Samuels L, Jansson S, Tuominen H (2009) A unique 

program for cell death in xylem fibers of Populus stem. Plant J. 58, 

260–274. 

Cox RM, Malcolm JM (1997) Effects of duration of a simulated winter 

thaw on dieback and xylem conductivity of Betula papyrifera. Tree 

Physiol. 17, 389–396. 

Crombie DS, Hipkins MF, Milburn JA (1985) Gas penetration of 

pit membranes in the xylem of Rhododendron as the cause of 

acoustically detectable sap cavitation. Aust. J. Plant Physiol. 12, 

445–454. 

Cronshaw J (1981) Phloem structure and function. Annu Rev. Plant 

Physiol. Plant Mol. Biol. 32, 465–484. 

Cui H, Hao YM, Stolc V, Deng XW, Sakakibara H, Kojima M (2011) 

Genome-wide direct target analysis reveals a role for SHORT- 

ROOT in root vascular patterning through cytokinin homeostasis. 


Cui H, Levesque MP, Vernoux T, Jung JW, Paquette AJ, Gallagher 

KL, Wang JY, Blilou I, Scheres B, Benfey PN (2007) 

An evolutionarily conserved mechanism delimiting SHR movement 

defines a single layer of endodermis in plants. Science 316, 

421–425. 

Cui KM, He XQ (2012) The Ca2+ -dependent DNases are involved in 

secondary xylem development in Eucommia ulmoides. J. Integr. 

Plant Biol. 54, 456–470. 

Curie C, Cassin G, Couch D, Divol F, Higuchi K, Le Jean M, Misson 

J, Schikora A, Czernic P, Mari S (2009) Metal movement within 

the plant: Contribution of nicotianamine and yellow stripe 1-like 

transporters. Ann. Bot. 103, 1–11. 

Dangl JL, Dietrich RA, Richberg MH (1996) Death don’t have no 

mercy: Cell death programs in plant-microbe interactions. Plant Cell 

8, 1793–1807. 

David-Schwartz R, Runo S, Townsley B, Machuka J, Sinha N (2008) 

Long-distance transport of mRNA via parenchyma cells and phloem 

across the host-parasite junction in Cuscuta. New Phytol. 179, 

1133–1141. 

Davidson A, Keller F, Turgeon R (2011) Phloem loading, plant growth 

form, climate. Protoplasma 248, 153–163. 

Davis SD, Sperry JS, Hacke UG (1999) The relationship between 

xylem conduit diameter and cavitation caused by freeze-thaw 

events. Am. J. Bot. 86, 1367–1372. 

Davis WJ, Zhang J (1991) Root signals and the regulation of growth 

and development of plants in drying soil. Annu. Rev. Plant Physiol. 

Mol. Biol. 42, 55–76. 

Dean JV, Delaney SP (2008) Metabolism of salicylic acid in wild-type, 

ugt74f1 and ugt74f2 glucosyltransferase mutants of Arabidopsis 

thaliana. Physiol. Plant. 132, 417–425. 

Deeken R, Ache P, Kajahan I, Klinkenberg J, Bringmann G, Hedrich 

R (2008) Identification of Arabidopsis thaliana phloem RNAs 

provides a search criterion for phloem-based transcripts hidden 

in complex datasets of microarray experiments. Plant J. 55, 

746–759. 

Deeken R, Geiger D, Fromm J, Koroleva O, Ache P, Langenfeld- 

Heyser R, Sauer N, May ST, Hedrich R (2002) Loss of AKT2/3 

potassium channel affects sugar loading into the phloem of Arabidopsis. 

Planta 216, 334–344.

Delaney TP, Friedrich L, Ryals JA (1995) Arabidopsis signal transduction 

mutant defective in chemically and biologically induced disease 

resistance. Proc. Natl. Acad. Sci. USA 92, 6602–6606. 

Demari-Weissler H, Rachamilavech S, Aloni R, German MA, Cohen 

S, Zwieniecki MA, Holbrook NM, Granot D (2009) LeFRK2 is 

required for phloem and xylem differentiation and the transport of 

both sugar and water. Planta 230, 795–805. 

Demura T, Tashiro G, Horiguchi G, Kishimoto N, Kubo M, Matsuoka 

N, Minami A, Nagata-Hiwatashi M, Nakamura K, Okamura Y, 

Sassa N, Suzuki S, Yazaki J, Kikuchi S, Fukuda H (2002) Visualization 

by comprehensive microarray analysis of gene expression 

programs during transdifferentiation of mesophyll cells into xylem 

cells. Proc. Natl. Acad. Sci. USA 99, 15794–15799. 

Demura T, Fukuda H (2007) Transcriptional regulation in wood formation. 

Trends Plant Sci. 12, 64–70. 

Dengler N, Kang J (2001) Vascular patterning and leaf shape. Curr. 

Opin. Plant Biol. 4, 50–56. 

De Smet I, Tetsumura T, De Rybel B, Frey NF, Laplaze L, Casimiro I, 

Swarup R, Naudts M, Vanneste S, Audenaert D, Inze D, Bennett 

MJ, Beeckman T (2007) Auxin-dependent regulation of lateral root 

positioning in the basal meristem of Arabidopsis. Development 134, 

681–690. 

Després C, DeLong C, Glaze S, Liu E, Fobert PR (2000) The 

Arabidopsis NPR1/NIM1 protein enhances the DNA binding activity 

of a subgroup of the TGA family of bZIP transcription factors. Plant 

Cell 12, 279–290. 

Després C, Chubak C, Rochon A, Clark R, Bethune T, Desveaux 

D, Fobert PR (2003) The Arabidopsis NPR1 disease resistance 

protein is a novel cofactor that confers redox regulation of DNA 

binding activity to the basic domain/leucine zipper transcription 

factor TGA1. Plant Cell 15, 2181–2191. 

Dhawan W, Luo H, Foerster AM, AbuQamar S, Du HN, Briggs SC, 

Scheid OM, Mengiste T (2009) HISTONE MONOUBIQUITINA- 

TION1 interacts with a subunit of the mediator complex and regulates 

defense against necrotrophic fungal pathogens in Arabidopsis. 


Dhondt S, Coppens F, De Winter F, Swarup K, Merks RMH, 

Inze D, Bennett MJ, Beemster GTS (2010) SHORT-ROOT and 

SCARECROW regulate leaf growth in Arabidopsis by stimulating 

S-phase progression of the cell cycle. Plant Physiol. 154, 1183– 

1195. 

DiDonato R, Roberts L, Sanderson T, Eisley R, Walker E (2004) 

Arabidopsis yellow stripe-like2 (YSL2): A metal-regulated gene 

encoding a plasma membrane transporter of nicotianamine-metal 

complexes. Plant J. 39, 403–414. 

Diévart A, Clark SE (2004) LRR-containing receptors regulating plant 

development and defense. Development 131, 251–261. 

Dinant S, Lucas WJ (2012) Sieve elements: Puzzling activities 

deciphered by proteomics studies. In: GA Thompson, 

AJE van Bel, eds. Phloem: Molecular Cell Biology, Systemic 

Communication, Biotic Interactions. Wiley-Blackwell, Ames. pp. 

157–185. 


Doering-Saad C, Newbury HJ, Couldridge CE, Bale JS, Pritchard J 

(2006) A phloem-enriched cDNA library from Ricinus: Insights into 

phloem function. J. Exp. Bot. 57, 3183–3193. 

Dolan L, Janmaat K, Willemsen V, Linstead P, Poethig S, Roberts K, 

Scheres B (1993) Cellular organisation of the Arabidopsis thaliana 

root. Development 119, 71–84. 

Domagalska MA, Leyser O (2011) Signal integration in the control of 

shoot branching. Nat. Rev. Mol. Cell Biol. 12, 211–221. 

Domec JC, Lachenbruch B, Meinzer FC (2006) Bordered pit structure 

and function determine spatial patterns of air-seeding thresholds in 

xylem of douglas-fir (Psuedotsuga menziesii; Pinaceae) trees. Am. 

J. Bot. 93, 1588–1600. 

Domec JC, Warren JM, Meinzer FC, Lachenbruch B (2009) Safety 

factors for xylem failure by implosion and air-seeding with roots, 

trunks, and branches of young and old conifer trees. IAWA J. 30, 

100–120. 

Donner TJ, Sherr I, Scarpella E (2009) Regulation of preprocambial 

cell state acquisition by auxin signaling in Arabidopsis leaves. 

Development 136, 3235–3246. 

Du J, Groover A (2010) Transcriptional regulation of secondary growth 

and wood formation. J. Integr. Plant Biol 52, 17–27. 

Du S, Yamamoto F (2007) An overview of the biology of reaction wood 

formation. J. Integr. Plant Biol. 49, 131–143. 

Duckett JG, Pressel S, P’ng KMY, Renzaglia, KS (2009) Exploding 

a myth: The capsule dehiscence mechanism and the function of 

pseudostomata in Sphagnum. New Phytol. 183, 1053–1063. 

Dunoyer P, Brosnan CA, Schott G, Wang Y, Jay F, Alioua A, Himber 

C, Voinnet O (2010) An endogenous, systemic RNAi pathway in 

plants. EMBO J. 29, 1699–1712. 

Durner J, Klessig DF (1995) Inhibition of ascorbate peroxidase by 

salicylic acid and 2.6-dichloroisonicdinic acid, two inducers of plant 

defense responses. Proc. Natl. Acad. Sci. USA 92, 11312–11316. 

Durrant WE, Wang S, Dong X (2007) Arabidopsis SNI1 and RAD51D 

regulate both gene transcription and DNA recombination during the 

defense response. Proc. Natl. Acad. Sci. USA 104, 4223–4227. 

Durrant W, Dong X (2004) Systemic acquired resistance. Annu. Rev. 

Phytopathol. 42, 185–209. 

Durrett TP, Gassmann W, Rogers EE (2007) The FRD3-Mediated 

efflux of citrate into the root vasculature is necessary for efficient 

iron translocation. Plant Physiol. 144, 197–205. 

Edwards D, Davies KL, AXE L (1992) A vascular conducting strand in 

the early plant Cooksonia. Nature 357, 683–685. 

Edwards R (1994) Conjugation and metabolism of salicylic acid in 

tobacco. J. Plant Physiol. 143, 609–614. 

Effmert U, Saschenbrecker S, Ross J, Negre F, Fraser CM, Noel JP, 

Dudareva N, Piechulla B (2005) Floral benzenoid carboxyl methyltransferases: 

From in vitro to in planta function. Phytochemistry 66, 

1211–1230. 

Ejeta G (2007) Breeding for Striga resistance in sorghum: Exploitation 

of an intricate host-parasite biology. Crop Sci. 47, S216–S227. 

Emery JF, Floyd SK, Alvarez J, Eshed Y, Hawker NP, Izhaki A, 

Baum SF, Bowman JL (2003) Radial patterning of Arabidopsis


shoots by Class III HD-ZIP and KANADI genes. Curr. Biol. 13, 1768– 

1774. 

Endo S, Demura T, Fukuda H (2001) Inhibition of proteasome activity 

by the TED4 protein in extracellular space: A novel mechanism for 

protection of living cells from injury caused by dying cells. Plant Cell 

Physiol. 42, 9–19. 

Enyedi AJ, Yalpani N, Silverman P, Raskin I (1992) Localization, 

conjugation, and function of salicylic acid in tobacco during the 

hypersensitive reaction to tobacco mosaic virus. Proc. Natl. Acad. 

Sci. USA 89, 2480–2484. 

Eom JS, Cho JI, Reinder A, Lee SW, Yoo Y, Tuan PQ, Choi SB, 

Bang G, Park YI, Cho MH, Bhoo SH, An G, Hahn TR, Ward JM, 

Jeon JS (2011) Impaired function of the tonoplast-localized sucrose 

transporter in rice (Oryza sativa), OsSUT2, limits the transport of 

vacuolar reserve sucrose and affects plant growth. Plant Physiol. 

157, 109–119. 

Esau K (1965a) Plant Anatomy. John Wiley & Sons, Inc., New York. 

Esau K (1965b) Vascular Differentiation in Plants. Holt, Rinehart and 

Winston, New York. 

Esau K (1969) The Phloem. Encyclopedia of Plant Anatomy. Borntaeger, 

Berlin. 

Esau K, Cheadle VI, Gifford EM (1953) Comparative structure and 

possible trends of specialization of the phloem. Am.J.Bot.40, 9– 

19. 

Eshed Y, Baum SF, Perea JV, Bowman JL (2001) Establishment of 

polarity in lateral organs of plants. Curr. Biol. 11, 1251–1260. 

Etchells JP, Provost CM, Turner SR (2012) Plant vascular cell 

division is maintained by an interaction between PXY and ethylene 

signalling. PLoS Genet. 8, e1002997. 

Etchells JP, Turner SR (2010) The pxy-cle41 receptor ligand pair defines 

a multifunctional pathway that controls the rate and orientation 

of vascular cell division. Development 137, 767–774. 

Evert RF (2006) Esau’s Plant Anatomy: Meristems, Cells, and Tissues 

of the Plant Body: Their Structure, Function, and Development. John 

Wiley & Sons, Hoboken, NJ. 

Evert RF, Warmbrodt RD, Eichhorn SE (1989) Sieve-pore development 

in some leptosporangiate ferns. Am. J. Bot. 76, 1404–1413. 

Fan RC, Peng CC, Xu YH, Wang XF, Li Y, Shang Y, Du SY, Zhao R, 

Zhang XY, Zhang LY, Zhang DP (2009) Apple sucrose transporter 

SUT1 and sorbitol transporter SOT6 interact with cytochrome b5 

to regulate their affinity for substrate sugars. Plant Physiol. 150, 

1880–1901. 

Fan W, Dong X (2002) In vivo interaction between NPR1 and transcrip- 

tion factor TGA2 leads to salicylic acid-mediated gene activation in 

Arabidopsis. Plant Cell 14, 1377–1389. 

FAO (2008) Forest and energy. Food and Agriculture Organization of 

the United Nations. FAO Forestry Paper 154. 

FAO (2009) State of the world’s forests. FAO Report, Food and 

Agriculture Organization of the United Nations. ISBN 978-92-5- 

106057-5. 

Farrar JF, Jones DL (2000) The control of carbon acquisition by roots. 

New Phytol. 147, 43–53. 

Feys BJ, Moisan LJ, Newman MA, Parker JE (2001) Direct interaction 

between the Arabidopsis disease resistance signaling proteins, 

EDS1 and PAD4. EMBO J. 20, 5400–5411. 

Feys BJ, Wiermer M, Bhat RA, Moisan LJ, Medina-Escobar N, 

Neu C, Cabral A, Parker JE (2005) Arabidopsis SENESCENCE- 

ASSOCIATED GENE101 stablizes and signals within an EN- 

HANCED DISEASE SUSCEPTIBILITY1 complex in plant innate 

immunity. Plant Cell 17, 2601–2613. 

Fiers M, Ku KL, Liu CM (2007) CLE peptide ligands and their roles in 

establishing meristems. Curr. Opin. Plant Biol. 10, 39–43. 

Fisher DB (2000) Long-Distance Transport. In: B Buchanan, W Grissem, 

R Jones, eds. Biochemistry & Molecular Biology of Plants. 

American Soc. of Plant Biologists, Maryland, pp. 730–785. 

Fisher DB, Cash-Clarke CE (2000a) Sieve tube unloading, postphloem 

transport of fluorescent tracers, proteins injected into sieve 

tubes severed aphid stylets. Plant Physiol. 123, 125–137. 

Fisher DB, Cash-Clarke CE (2000b) Gradients in water potential, 

turgor pressure along the translocation pathway during grain filling 

in normally watered, water-stressed wheat plants. Plant Physiol. 

123, 139–147. 

Fisher DB, Wang N (1995) Sucrose concentration gradients along the 

post-phloem transport pathway in the maternal tissue of developing 

wheat grains. Plant Physiol. 109, 587–592. 

Fisher DB, Wu K, Ku MSB (1992) Turnover of soluble proteins in the 

wheat sieve tube. Plant Physiol. 100, 1433–1441. 

Fisher K, Turner S (2007) Pxy, a receptor-like kinase essential for 

maintaining polarity during plant vascular-tissue development. Curr. 

Biol. 17, 1061–1066. 

Floyd SK, Bowman JL (2004) Gene regulation: Ancient microrna 

target sequences in plants. Nature 428, 485–486. 

Floyd SK, Zalewski CS, Bowman JL (2006) Evolution of class iii 

homeodomain-leucine zipper genes in streptophytes. Genetics 173, 

373–388. 

Franco-Zorrilla JM, Martín AC, Leyva A, Paz-Ares J (2005) Interaction 

between phosphate-starvation, sugar, and cytokinin signaling 

in Arabidopsis and the roles of cytokinin receptors CRE1/AHK4 and 

AHK3. Plant Physiol. 38, 847–57. 

Franco-Zorrilla JM, Martin AC, Solano R, Rubio V, Leyva A, Paz- 

Ares J (2002) Mutations at CRE1 impair cytokinin-induced repression 

of phosphate starvation responses in Arabidopsis. Plant J. 32, 

353–360. 

Franks P, Brodribb T (2005) Stomatal control and water transport in 

stems. In: Holbrook NM, Zwieniecki MA, eds. Vascular Transport in 

Plants. Elsevier, Amsterdam. pp. 69–89. 

Fraysse LC, Wells B, McCann MC, Kjellbom P (2005) Specific plasma 

membrane aquaporins of the PIP1 subfamily are expressed in sieve 

elements and guard cells. Biol. Cell 97, 519–534. 

Friedrich L, Vernooij E, Gaffney T, Morse A, Ryals J (1995) 

Characterization of tobacco plants expressing a bacterial salicylate 

hydroxylase gene. Plant Mol. Biol. 29, 959–968. 

Froelich DR, Mullendore DL, Jensen KH, Ross-Elliot TJ, Anstead 

JA, Thompson GA, Pélissier HC, Knoblauch M (2011) Phloem

ultrastructure and pressure flow: Sieve-element-occlusion-related 

agglomerations do not affect translocation. Plant Cell 23, 4428– 

4445. 

Fu ZQ, Yan S, Saleh A, Wang W, Ruble J, Oka N, Mohan R, 

Spoel SH, Tada Y, Zheng Y, Dong X (2012) NPR3 and NPR4 

are receptors for the immune signal salicylic acid in plants. Nature 

16, 228–232. 

Fukuda H (2000) Programmed cell death of tracheary elements as a 

paradigm in plants. Plant Mol. Biol. 44, 245–253. 

Fukuda H (2004) Signals that control plant vascular cell differentiation. 

Nat. Rev. Mol. Cell Biol. 5, 379–391. 

Fukuda H, Hirakawa Y, Sawa S (2007) Peptide signaling in vascular 

development. Curr. Opin. Plant Biol. 10, 477–482. 

Funk V, Kositsup B, Zhao C, Beers EP (2002) The Arabidopsis xylem 

peptidase XCP1 is a tracheary element vacuolar protein that may 

be a papain ortholog. Plant Physiol. 128, 84–94. 

Gabaldon C, Ros LVG, Pedreno MA, Barcelo AR (2005) Nitric oxide 

production by the differentiating xylem of Zinnia elegans. New 

Phytol. 165, 121–130. 

Gaffney T, Friedrich L, Vernooij B, Negmtto D, Nye G, Uknes S, 

Ward E, Kessmann H, Ryals J (1993) Requirement of salicylic 

acid for the induction of systemic acquired resistance. Science 261, 

754–756. 

Gallagher KL, Benfey PN (2009) Both the conserved GRAS domain 

and nuclear localization are required for SHORT-ROOT movement. 

Plant J. 57, 785–797. 

García AV, Blanvillain-Baufumé S,HuibersRP,WiermerM,LiG, 

Gobbato E, Rietz S, Parker JE (2010) Balanced nuclear and 

cytoplasmic activities of EDS1 are required for a complete plant 

innate immune response. PLoS Pathogens 6, e1000970 

Garcia-Mas J, Benjak A, Sanseverino W, Bourgeois M, Mir G, 

González VM, Hénaff E, Câmara F, Cozzuto L, Lowy E, Alioto 

T, Capella-Gutiérrez S, Blanca J, Cañizares J, Ziarsolo P, 

Gonzalez-Ibeas D, Rodríguez-Moreno L, Droege M, Du L, 

Alvarez-Tejado M, Lorente-Galdos B, Melé M, Yang L, Weng 

YQ, Navarro A, Marques-Bonet T, Aranda MA, Nuez F, Picó 

B, Gabaldón T, Roma G, Guigó R, Casacuberta JM, Arús P, 

Puigdomènech P (2012) The genome of melon (Cucumis melo L.) 

Proc. Natl. Acad. Sci. USA 109, 11872–11877. 

Gardiner J, Donner TJ, Scarpella E (2011) Simultaneous activation 

of SHR and ATHB8 expression defines switch to preprocambial cell 

state in Arabidopsis leaf development. Dev. Dynam. 240, 261–270. 

Gaupels F, Buhtz A, Knauer T, Deshmurkh S, Waller F, van Bel 

AJE, Kogel KH, Kehr J (2008a) Adaption of aphid stylectomy for 

analyses of proteins and mRNA in barley phloem sap. J. Exp. Bot. 

59, 3297–3306. 

Gaupels F, Knauer T, van Bel AJE (2008b) A combinatory approach 

for analysis of protein sets in barley sieve-tube samples using 

EDTA-facilitated exudation and aphid stylectomy. J. Plant Physiol. 

165, 95–103. 

Gendre D, Czernic P, Conejero G, Pianelli K, Briat J, Lebrun M, Mari 

S (2007) TcYSL3, a member of the YSL gene family from the hyper- 


accumulator Thlaspi caerulescens, encodes a nicotianamine-Ni/Fe 

transporter. Plant J. 49, 1–15. 

Genoud T, Buchala AJ, Chua NH, Metraux JP (2002) Phytochrome 

signalling modulates the SA-perceptive pathway in Arabidopsis. 


Ghanem ME, Albacete A, Smigocki AC, Frébort I, Pospísilová 

H, Martínez-Andújar C, Acosta M, Sánchez-Bravo J, Lutts S, 

Dodd IC, Pérez-Alfocea F (2011) Root-synthesized cytokinins 

improve shoot growth and fruit yield in salinized tomato (Solanum 

lycopersicum L.) plants. J. Exp. Bot. 62, 125–140. 

Giaquinta RT (1983) Phloem loading of sucrose. Annu. Rev. Plant 

Physiol. 34, 347–387. 

Giavalisco P, Kapitza K, Kolasa A, Buhtz A, Kehr J (2006) Towards 

theproteomeofBrassica napus phloem sap. Proteomics 6, 896– 

909. 

Glazebrook J, Ausubel FM (1994) Isolation of phytoalexin-deficient 

mutants of Arabidopsis thaliana and characterization of their interactions 

with bacterial pathogens. Proc. Natl. Acad. Sci. USA 91, 

8955–8959. 

Glazebrook J, Rogers EE, Ausubel FM (1996) Isolation of Arabidopsis 

mutants with enhanced disease susceptibility by direct screening. 

Genetics 143, 973–982. 

Golecki B, Schulz A, Carstens-Behrens U, Kollmann R (1998) 

Evidence for graft transmission of structural phloem proteins or 

their precursors in heterografts of Cucurbitaceae. Planta 206, 

630–640. 

Golecki B, Schulz A, Thompson GA (1999) Translocation of structural 

P proteins in the phloem. Plant Cell 11, 127–140. 

Gomez-Roldan V, Fermas S, Brewer PB, Puech-Pagès V, Dun 

EA, Pillot JP, Letisse F, Matusova R, Danoun S, Portais 

JC, Bouwmeester H, Bécard G, Beveridge CA, Rameau C, 

Rochange SF (2008) Strigolactone inhibition of shoot branching. 

Nature 455, 189–194. 

Gould N, Minchin PEH, Thorpe MR (2004a) Direct measurements of 

sieve element hydrostatic pressure reveal strong regulation after 

pathway blockage. Funct. Plant Biol. 31, 987–993. 

Gould N, Thorpe MR, Koroleva O, Minchin PEH (2005) Phloem 

hydrostatic pressure relates to solute loading rate: A direct test of 

the Münch hypothesis. Funct. Plant Biol. 32, 1019–1026. 

Gould N, Thorpe MR, Minchin PEH, Pritchard J, White PJ (2004b) 

Solute is imported to elongating root cells of barley as a pressure 

driven-flow of solution. Funct. Plant Biol. 31, 391–397. 

Grimmer C, Komor E (1999) Assimilate export by leaves of Ricinus 

communis L. growing under normal and elevated carbon dioxide 

concentrations: The same rate during the day, a different rate at 

night. Planta 209, 275–281. 

Grodzinski B, Jiao J, Leonardos ED (1998) Estimating photosynthesis 

and concurrent export rates in C3 and C4 species at ambient 

and elevated CO2. Plant Physiol. 117, 207–215. 

Grusak MA (1994) Iron transport to developing ovules of Pisum 

sativum. (I. seed import characteristics and phloem iron-loading 

capacity of source regions). Plant Physiol. 104, 649–655.


Guelette BS, Benning UF, Hoffmann-Benning S (2012) Identification 

of lipids and lipid-binding proteins in phloem exudates from 

Arabidopsis thaliana. J. Exp. Bot. 63, 3603–3616. 

Guo S, Zhang J, Sun H, Salse J, Lucas WJ, Zhang H, Zheng Y, 

Mao L, Ren Y, Wang Z, Min J, Guo X, Murat F, Ham BK, Zhang 

Z, Gao S, Huang M, Xu Y, Zhong S, Bombarely A, Mueller LA, 

Zhao H, He H, Zhang Y, Zhang Z, Huang S, Tan T, Pang E, Lin 

K, Hu Q, Kuang H, Ni P, Wang B, Liu J, Kou Q, Hou W, Zou X, 

Jiang J, Gong G, Klee K, Schoof H, Huang Y, Hu X, Dong S, 

Liang D, Wang J, Wu K, Xia Y, Zhao X, Zheng Z, Xing M, Liang 

X, Huang B, Lv T, Wang J, Yin Y, Yi H, Li R, Wu M, Levi A, 

Zhang X, Giovannoni JJ, Wang J, Li Y, Fei Z, Xu Y (2012) The 

draft genome of watermelon (Citrullus lunatus) and resequencing of 

20 diverse accessions. Nat. Genet. 45, 51–58. 

Guo WJ, Bundithya W, Goldsbrough PB (2003) Characterization 

of the Arabidopsis metallothionein gene family: Tissue-specific 

expression and induction during senescence and in response to 

copper. New Phytol. 159, 369–381. 

Guo WJ, Meetam M, Goldsbrough PB (2008) Examining the specific 

contributions of individual Arabidopsis metallothioneins to copper 

distribution and metal tolerance. Plant Physiol. 146, 1697– 

1706. 

Hacke U, Sauter JJ (1996) Xylem dysfunction during winter and 

recovery of hydraulic conductivity in diffuse-porous and ring-porous 

trees. Oecologia 105, 435–439. 

Hacke UG, Sperry JS (2003) Limits to xylem refilling under negative 

pressure in Laurus nobilis and Acer negundo. Plant Cell Environ. 

26, 303–311. 

Hacke UG, Sperry JS, Pockman WP, Davis SD, McCulloh KA 

(2001a) Trends in wood density and structure are linked to prevention 

of xylem implosion by negative pressure. Oecologia 126, 

457–461. 

Hacke UG, Sperry JS, Wheeler JK, Castro L (2006) Scaling of 

angiosperm xylem structure with safety and efficiency. Tree Physiol. 

26, 689–701. 

Hacke UG, Stiller V, Sperry JS, Pittermann J, McCulloh KA 

(2001b) Cavitation fatigue: Embolism and refilling cycles can 

weaken cavitation resistance of xylem. Plant Physiol. 125, 779– 

786. 

Hacke UG, Jansen S (2009) Embolism resistance of three boreal 

conifers varies with pit structure. New Phytol. 182, 675–686. 

Ham B-K, Brandom J, Xoconostle-Cázares, B, Ringgold V, Lough 

TJ, Lucas WJ (2009) Polypyrimidine tract binding protein, Cm- 

RBP50, forms the basis of a pumpkin phloem ribonucleoprotein 

complex. Plant Cell 21, 197–215. 

Hamburger D, Rezzonico E, MacDonald-Comber Petétot J, 

Somerville C, Poirier Y (2002) Identification and characterization 

of the Arabidopsis PHO1 gene involved in phosphate loading to the 

xylem. Plant Cell 14, 889–902. 

Hamilton AJ, Baulcombe DC (1999) A species of small antisense 

RNA in posttranscriptional gene silencing in plants. Science 286, 

950–952. 

Han JJ, Lin W, Oda Y, Cui KM, Fukuda H, He XQ (2012) The 

proteasome is responsible for caspase-3-like activity during xylem 

development. Plant J. 72, 129–141. 

Hanikenne M, Talke IN, Haydon MJ, Lanz C, Nolte A, Motte P, 

Kroymann J, Weigel D, Krämer U (2008) Evolution of metal 

hyperaccumulation required cis-regulatory changes and triplication 

of HMA4. Nature 453, 391–395. 

Hannapel DJ (2010) A model system of development regulated by the 

long-distance transport of mRNA. J. Integr. Plant Biol. 52, 40–52. 

Hanzawa Y, Takahashi T, Komeda Y (1997) ACL5: AnArabidopsis 

gene required for internodal elongation after flowering. Plant J. 12, 

863–874. 

Hardtke CS, Berleth T (1998) The Arabidopsis gene MONOPTEROS 

encodes a transcription factor mediating embryo axis formation and 

vascular development. EMBO J. 17, 1405–1411. 

Hardtke CS, Ckurshumova W, Vidaurre DP, Singh SA, Stamatiou G, 

Tiwari SB, Hagen G, Guilfoyle TJ, Berleth T (2004) Overlapping 

and non-redundant functions of the Arabidopsis auxin response 

factors MONOPTEROS and NONPHOTOTROPIC HYPOCOTYL 

4. Development 131, 1089–1100. 

Haritatos E, Medville R, Turgeon R (2000) Minor vein structure and 

sugar transport in Arabidopsis thaliana. Planta 211, 105– 111. 

Hassan Z, Aarts MGM (2011) Opportunities and feasibilities for 

biotechnological improvement of Zn, Cd or Ni tolerance and accumulation 

in plants. Environ. Exp. Bot. 72, 53–63. 

Hawkesford MJ (2003) Transporter gene families in plants: The 

sulphate transporter gene family – redundancy or specialization? 

Physiol. Plant. 117, 155–163. 

Haywood V, Yu TS, Huang NC, Lucas WJ (2005) Phloem longdistance 

trafficking of GIBBERELLIC ACID INSENSITIVE RNA 

regulates leaf development. Plant J. 42, 49–68. 

He Y, Gan S (2002) A gene encoding an acyl hydrolase is involved in 

leaf senescence in Arabidopsis. Plant Cell 14, 805–815. 

Heidel AJ, Clarke JD, Antonovics J, Dong X (2004) Fitness costs 

of mutations affecting the systemic acquired resistance pathway in 

Arabidopsis thaliana. Genetics 168, 2197–2206. 

Heidrich K, Wirthmueller L, Tasset C, Pouzet C, Deslandes L, 

Parker JE (2011) Arabidopsis EDS1 connects pathogen effector 

recognition to cell compartment-specific immune responses. 

Science 334, 1401–1404. 

Heil M, Baldwin IT (2002) Fitness costs of induced resistance: Emerging 

experimental support for a slippery concept. Trends Plant Sci. 

7, 61–66. 

Herbette S, Cochard H (2010) Calcium is a major determinant of xylem 

vulnerability to cavitation. Plant Physiol. 153, 1932–1939. 

Herbik A, Giritch A, Horstmann C, Becker R, Balzer HJ, Bäumlein H, 

Stephan UW (1996) Iron and copper nutrition-dependent changes 

in protein expression in a tomato wild type and the nicotianaminefree 

mutant chloronerva. Plant Physiol. 111, 533–540. 

Hermans C, Hammond JP, White PJ, Verbuggen N (2006) How do 

plants respond to nutrient shortage by biomass allocation? Trends 

Plant Sci. 11, 610–617.

Hickey L (1973) Classification of the architecture of dicotyledonous 

leaves. Amer. J. Bot. 60, 17–33. 

Hirakawa Y, Kondo Y, Fukuda H (2010) Regulation of vascular 

development by CLE peptide-receptor systems. J. Integr. Plant Biol. 

52, 8–16. 

Hirakawa Y, Kondo Y, Fukuda H (2010) TDIF peptide signaling 

regulates vascular stem cell proliferation via the WOX4 homeobox 

gene in Arabidopsis. Plant Cell 22, 2618–2629. 

Hirakawa Y, Kondo Y, Fukuda H (2011) Establishment and maintenance 

of vascular cell communities through local signaling. Curr. 

Opin. Plant Biol. 14, 17–23. 

Hirakawa Y, Shinohara H, Kondo Y, Inoue A, Nakanomyo I, Ogawa 

M, Sawa S, Ohashi-Ito K, Matsubayashi Y, Fukuda H (2008) 

Non-cell-autonomous control of vascular stem cell fate by a CLE 

peptide/receptor system. Proc. Natl. Acad. Sci. USA 105, 15208– 

15213. 

Hirose N, Takei K, Kuroha T, Kamada-Nobusada T, Hayashi H, 

Sakakibara H (2008) Regulation of cytokinin biosynthesis, compartmentalization 

and translocation. J. Exp. Bot. 59, 75–83. 

Hoad GV (1995) Transport of hormones in the phloem of higher plants. 

Plant Growth Reg. 16, 173–182. 

Hoffman-Thoma G, van Bel AJE, Ehlers K (2001) Ultrastructure of 

minor-vein phloem and assimilate export in summer and winter 

leaves of the symplasmically loading evergreens Ajuga reptans L., 

Aucuba japonica Thumb. and Hedrix helix L. Planta 212, 231–242. 

Holbrook NM, Shashidhar VR, James RA, Munns R (2002) Stomatal 

control in tomato with ABA-deficient roots: Response of grafted 

plants to soil drying. J. Exp. Bot. 53, 1503–1514. 

Holtta T, Mencuccini M, Nikinmaa E (2011) A carbon cost-gain model 

explains the observed patterns of xylem safety and efficiency. Plant 

Cell Environ. 34, 1819–1834. 

Holtta T, Juurola E, Lindfors L, Porcar-Castell A (2012) Cavitation 

induced by a surfactant leads to transient release of water stress 

and subsequent “run away” embolism in scots pine (Pinus sylvestris) 

seedlings. J. Exp. Bot. 63, 1057–1067. 

Hu G, deHart AKA, Li Y, Ustach C, Handley V, Navarre R, Hwang 

CF, Aegerter BJ, Williamson VM, Baker B (2005) EDS1 in tomato 

is required for resistance mediated by TIR-class R genes and the 

receptor-like R gene Ve. Plant J. 42, 376–391. 

Hu H, Penn SG, Lebrilla CB, Brown PH (1997) Isolation and characterization 

of soluble boron complexes in higher plants: The mechanism 

of phloem mobility of boron. Plant Physiol. 113, 649–655. 

Hu L, Sun H, Li R, Zhang L, Wang S, Sui X, Zhang Z (2011) Phloem 

unloading follows an extensive apoplasmic pathway in cucumber 

(Cucumis sativa L.) fruit from anthesis to marketable maturing stage. 

Plant Cell Environ. 40, 743–748. 

Huang SW, Li RQ, Zhang ZH, Li L, Gu XF, Fan W, Lucas WJ, 

Wang XW, Xie BY, Ni PX, Ren YY, Zhu HM, Li J, Lin K, Jin 

WW, Fei ZJ, Li GC, Staub J, Kilian A, van der Vossen EAG, 

Wu Y, Guo J, He J, Jia ZQ, Ren Y, Tian G, Lu Y, Ruan J, Qian 

WB, Wang MW, Huang QF, Li B, Xuan ZL, Cao JJ, Asan, Wu 

ZG, Zhang JB, Cai QL, Bai YQ, Zhao BW, Han YH, Li Y, Li 


XF, Wang SH, Shi QX, Liu SQ, Cho WK, Kim JY, Xu Y, Heller- 

Uszynska K, Miao H, Cheng ZC, Zhang SP, Wu J, Yang YH, 

KangHX,LiM,LiangHQ,RenXL,ShiZB,WenM,JianM, 

Yang HL, Zhang GJ, Yang ZT, Chen R, Liu SF, Li JW, Ma LJ, 

Liu H, Zhou Y, Zhao J, Fang XD, Li GQ, Fang L, Li YR, Liu DY, 

Zheng HK, Zhang Y, Qin N, Li Z, Yang GH, Yang S, Bolund L, 

Kristiansen K, Zheng HC, Li SC, Zhang XQ, Yang HM, Wang J, 

Sun RF, Zhang BX, Jiang SZ, Wang J, Du YC, Li SG (2009) The 

genome of the cucumber, Cucumis sativus L. Nat. Genet. 41, 1275– 

1281. 

Huang NC, Yu TS (2009) The sequences of Arabidopsis GA- 

INSENSITIVE RNA constitute the motifs that are necessary and 

sufficient for RNA long-distance trafficking. Plant J. 59, 921–929. 

Huang NC, Jane WN, Chen J, Yu TS (2012) Arabidopsis thaliana 

CENTRORADIALIS homologue (ATC) acts systemically to inhibit 

floral initiation in Arabidopsis. Plant J. 72, 175–184. 

Hubbard RM, Stiller V, Ryan MG, Sperry JS (2001) Stomatal conductance 

and photosynthesis vary linearly with plant hydraulic 

conductance in ponderosa pine. Plant Cell Environ. 24, 113– 

121. 

Husbands AY, Chitwood DH, Plavskin Y, Timmermans MC (2009) 

Signals and prepatterns: New insights into organ polarity in plants. 

Genes Dev. 23, 1986–1997. 

Hussain D, Haydon MJ, Wang Y, Wong E, Sherson SM, Young 

J, Camakaris J, Harper JF, Cobbett CS (2004) P-type ATPase 

heavy metal transporters with roles in essential zinc homeostasis in 


Ilegems M, Douet V, Meylan-Bettex M, Uyttewaal M, Brand L, 

Bowman JL, Stieger PA (2010) Interplay of auxin, kanadi and 

class iii hd-zip transcription factors in vascular tissue formation. 


Imai A, Hanzawa Y, Komura M, Yamamoto KT, Komeda Y, Takahashi 

T (2006) The dwarf phenotype of the Arabidopsis acl5 mutant 

is suppressed by a mutation in an upstream ORF of a bHLH gene. 


Imai A, Komura M, Kawano E, Kuwashiro Y Takahashi T (2008) 

A semi-dominant mutation in the ribosomal protein L10 gene 

suppresses the dwarf phenotype of the acl5 mutant in Arabidopsis 

thaliana. Plant J. 56, 881–890. 

Ingle RA, Mugford ST, Rees JD, Campbell MM, Smith JAC (2005) 

Constitutively high expression of the histidine biosynthetic pathway 

contributes to nickel tolerance in hyperaccumulator plants. Plant 

Cell 17, 2089–2106. 

Ingram P, Dettmer J, Helariutta Y, Malamy JE (2011) Arabidopsis 

lateral root development 3 is essential for early phloem development 

and function, and hence for normal root system development. Plant 

J. 68, 455–467. 

Irtelli B, Petrucci WA, Navari-Izzo F (2009) Nicotianamine and 

histidine/proline are, respectively, the most important copper 

chelators in xylem sap of Brassica carinata under conditions 

of copper deficiency and excess. J. Exp. Bot. 60, 269– 

277.


Ishihara T, Sekine KT, Hase S, Kanayama Y, Seo S, Ohashi Y, 

Kusano T, Shibata D, Shah J, Takahashi H (2008) Overexpression 

of the Arabidopsis thaliana EDS5 gene enhances resistance to 

viruses. Plant Biol. 10, 451–461. 

Ito J, Fukuda H (2002) ZEN1 is a key enzyme in the degradation of 

nuclear DNA during programmed cell death of tracheary elements. 


Ito Y, Nakanomyo I, Motose H, Iwamoto K, Sawa S, Dohmae N, 

Fukuda H (2006) Dodeca-cle peptides as suppressors of plant stem 

cell differentiation. Science 313, 842–845. 

Ivanov R, Brumbarova T, Bauer P (2012) Fitting into the harsh reality: 

Regulation of iron-deficiency responses in dicotyledonous plants. 

Mol. Plant 5, 27–42. 

Iwai H, Usui M, Hoshino H, Kamada H, Matsunaga T, Kakegawa K, 

IshiiT,SatohS(2003) Analysis of sugars in squash xylem sap. 


Izhaki A, Bowman JL (2007) KANADI and Class III HD-ZIP gene 

families regulate embryo patterning and modulate auxin flow during 

embryogenesis in Arabidopsis. Plant Cell 19, 495–508. 

Jack E, Hakvoort HWJ, Reumer A, Verkleij JAC, Schat H, 

Ernst WHO (2007) Real-time PCR analysis of metallothionein- 

2b expression in metallicolous and non-metallicolous populations 

of Silene vulgaris (Moench) Garcke. Environ. Exp. Bot. 59, 

84–91. 

Jacobson AL, Pratt RB (2012) No evidence for an open vessel effect in 

centrifuge-based vulnerability curves of a long-vesseled liana (Vitis 

vinifera). New Phytol. 194, 982–990. 

Jacobson AL, Pratt RB, Ewers FW, Davis SD (2007) Cavitation 

resistance among 26 chaparral species of southern California. Ecol. 

Mon. 77, 99–115. 

Jansen S, Baas P, Smets E (2001) Vestured pits: Their occurrence 

and systematic importance in eudicots. Taxon 50, 135–167. 

Jansen S, Choat B, Pletsers A (2009) Morphological variation in 

intervessel pit membranes and implications to xylem function in 

angiosperms. Am. J. Bot. 96, 409–419. 

Jansen S, Lamy JB, Burlett R, Cochard H, Gasson P, Delzon S 

(2012) Plasmodesmatal pores in the torus of bordered pit membranes 

affect cavitation resistance of conifer xylem. Plant Cell 

Environ. 35, 1109–1120. 

Jarbeau JA, Ewers FW, Davis SD (1995) The mechanism of waterstress-induced 

embolism in two species of chaparral shrubs. Plant 

Cell Environ. 18, 189–196. 

Jensen KH, Lee J, Bohr T, Bruus H, Holbrook NM, Zwieniecki 

MA (2011) Optimality of the Münch mechanism for translocation 

of sugars in plants. J. Royal Soc. Interface 8, 1155–1165. 

Jensen KH, Liesche J, Bohr T, Schulz A (2012) Universality of phloem 

transport in seed plants. Plant Cell Environ. 35, 1065–1076. 

Jeong R-D, Chandra-Shekara AC, Barman SR, Navarre DA, Klessig 

D, Kachroo A, Kachroo P (2010) CRYPTOCHROME 2 and PHO- 

TOTROPIN 2 regulate resistance protein mediated viral defense by 

negatively regulating a E3 ubiquitin ligase. Proc. Natl. Acad. Sci. 

USA 107, 13538–13543. 

Ji J, Strable J, Shimizu R, Koenig D, Sinha N, Scanlon MJ (2010) 

WOX4 promotes procambial development. Plant Physiol. 152, 

1346–1356. 

Jia W, Davies WJ (2008) Modification of leaf apoplastic pH in relation 

to stomatal sensitivity to root-sourced abscisic acid signals. Plant 

Physiol. 143, 68–77. 

Jiang F, Hartung W (2008) Long-distance signalling of abscisic acid 

(ABA): The factors regulating the intensity of the ABA signal. J. Exp. 

Bot. 59, 37–43. 

Johri M (2008) Hormonal regulation in green plant lineage families. 

Physiol. Mol. Biol. Plants 14, 23–38. 

Jones JD, Dangl JL (2006) The plant immune system. Nature 444, 

323–329. 

Jorgensen RA, Atkinson RG, Forster RLS, Lucas WJ (1998) 

An RNA-based information superhighway in plants. Science 279, 

1486–1487. 

Jun JH, Fiume E, Fletcher JC (2008) The CLE family of plant 

polypeptide signaling molecules. Cell Mol. Life Sci. 65, 743–755. 

Jung HW, Tschaplinkski TJ, Wang L, Glazebrook J, Greenberg JT 

(2009) Priming in systemic plant immunity. Science 324, 89–91. 

Kachroo A, Kachroo P (2006) Salicylic acid-, jasmonic acid- and 

ethylene-mediated regulation of plant defense signaling. In: Setlow 

J, ed. Genetic Engineering, Principles and Methods. 28, 55–83. 

Kachroo A, Lapchyk L, Fukushigae H, Hildebrand D, Klessig D, 

Kachroo P (2003) Plastidial fatty acid signaling modulates salicylic 

acid- and jasmonic acid-mediated defense pathways in the Arabidopsis 

ssi2 mutant. Plant Cell 12, 2952–2965. 

Kachroo A, Venugopal SC, Lapchyk L, Falcone D, Hildebrand 

D, Kachroo P (2004) Oleic acid levels regulated by glycerolipid 

metabolism modulate defense gene expression in Arabidopsis. 


Kachroo P, Kachroo A, Lapchyk L, Hildebrand D, Klessig D (2003) 

Restoration of defective cross talk in ssi2 mutant: Role of salicylic 

acid, jasmonic acid, and fatty acids in SSI2-mediated signaling. Mol. 

Plant-Microbe Interact. 11, 1022–1029. 

Kachroo P, Shanklin J, Shah J, Whittle E, Klessig D (2001) 

A fatty acid desaturase modulates the activation of defense 

signaling pathways in plants. Proc. Natl. Acad. Sci. USA 98, 

9448–9453. 

Kachroo P, Srivathsa CV, Navarre DA, Lapchyk L, Kachroo A (2005) 

Role of salicylic acid and fatty acid desaturation pathways in ssi2mediated 

signaling. Plant Physiol. 139, 1717–1735. 

Kachroo P, Yoshioka K, Shah J, Dooner H, Klessig DF (2000) 

Resistance to turnip crinkle virus in Arabidopsis requires two host 

genes and is salicylic acid dependent but NPR1, ethylene and 

jasmonate independent. Plant Cell 12, 677–690. 

Kakehi J, Kuwashiro Y, Motose H, Igarashi K, Takahashi T (2010) 

Norspermine substitutes for thermospermine in the control of stem 

elongation in Arabidopsis thaliana. FEBS Lett. 584, 3042–3046. 

Kakehi JI, Kuwashiro Y, Niitsu Y, Takahashi T (2008) Thermospermine 

is required for stem elongation in Arabidopsis thaliana. Plant 

Cell Physiol. 49, 1342–1349.

Kallarackal J, Milburn JA (1984) Specific mass transfer, sinkcontrolled 

phloem translocation in castor bean. Aust. J. Plant 

Physiol. 10, 561–568. 

Kang J, Dengler N (2002) Cell cycling frequency and expression 

of the homeobox gene ATHB-8 during leaf vein development in 

Arabidopsis. Planta 216, 212–219. 

Kang J, Dengler N (2004) Vein pattern development in adult leaves of 

Arabidopsis thaliana. Int. J. Plant Sci. 165, 231–242. 

Kang J, Mizukami Y, Wang H, Fowke L, Dengler NG (2007) Modification 

of cell proliferation patterns alters leaf vein architecture in 

Arabidopsis thaliana. Planta 226, 1207–1218. 

Karpinski S, Gabrys H, Mateo A, Karpinska B, Mullineaux P (2003) 

Light perception in plant disease defense signaling. Curr. Opin. 


Kaufman I, Schulze-Till T, Schneider H, Zimmermann U, Jakob P, 

Wegner L (2009) Function repair of embolized vessels in maize 

roots after temporal drought stress, as demonstrated by magnetic 

resonance imaging. New Phytol. 184, 245–256. 

Kempers R, van Bel AJE (1997) Symplasmic connections between 

sieve element and companion cell in the stem of Vicia fava L. 

have a molecular exclusion limit of at least 10 kD. Planta 201, 

195–201. 

Kenrick P, Crane PR (1997) The origin and early evolution of plants 

on land. Nature 389, 33–39. 

Kerkeb L, Krämer U (2003) The role of free histidine in xylem loading 

of nickel in Alyssum lesbiacum and Brassica juncea. Plant Physiol. 

131, 716–724. 

Kerstetter RA, Bollman K, Taylor RA, Bomblies K, Poethig RS 

(2001) KANADI regulates organ polarity in Arabidopsis. Nature 411, 

706–709. 

Kiba T, Yamada H, Sato S, Kato T, Tabata S, Yamashino T, 

Mizuno T (2003) The type-A response regulator, ARR15, acts as a 

negative regulator in the cytokinin-mediated signal transduction in 

Arabidopsis thaliana. Plant Cell Physiol. 44, 868–874. 

Kim HS, Delaney TP (2002) Over-expression of TGA5, which encodes 

a bZIP transcription factor that interacts with NIM1/NPR1, 

confers SAR-independent resistance in Arabidopsis thaliana to 

Peronospora parasitica. Plant J. 32, 151–163. 

Kim M, Canio W, Kessler S, Sinha N (2001) Developmental changes 

due to long-distance movement of a homeobox fusion transcript in 

tomato. Science 293, 287–289. 

Klatte M, Schuler M, Wirtz M, Fink-Straube C, Hell R, Bauer P 

(2009) The analysis of Arabidopsis nicotianamine synthase mutants 

reveals functions for nicotianamine in seed iron loading and iron 

deficiency responses. Plant Physiol. 150, 257–271. 

Kobayashi Y, Kuroda K, Kimura K, Southron-Francis JL, Furuzawa 

A, Iuchi S, Kobayashi M, Taylor GJ, Koyama H (2008) Amino acid 

polymorphisms in strictly conserved domains of a P-type ATPase 

HMA5 are involved in the mechanism of copper tolerance variation 

in Arabidopsis. Plant Physiol. 148, 969–980. 

Kohlen W, Charnikhova T, Liu Q, Bours R, Domagalska MA, 

Beguerie S, Verstappen F, Leyser O, Bouwmeester H, Ruyter- 


Spira C (2011) Strigolactones are transported through the xylem 

and play a key role in shoot architectural response to phosphate 

deficiency in nonarbuscular mycorrhizal host Arabidopsis. Plant 

Physiol. 155, 974–987. 

Kohonen M, Helland A (2009) On the function of wall sculpturing in 

xylem conduits. J. Bionics Eng. 6, 324–329. 

Koike S, Inoue H, Mizuno D, Takahashi M, Nakanishi H, Mori 

S, Nishizawa N (2004) OsYSL2 is a rice metal-nicotianamine 

transporter that is regulated by iron and expressed in the phloem. 


Kondo Y, Hirakawa Y, Fukuda H (2011) CLE peptides can negatively 

regulate protoxylem vessel formation via cytokinin signaling. Plant 

Cell Physiol. 52, 37–48. 

Koo YJ, Kim MA, Kim EH, Song JT, Jung C, Moon JK, Kim JH, 

Seo HS, Song SI, Kim JK, Lee JS, Cheong JJ, Choi YD (2007) 

Overexpression of salicylic acid carboxyl methyltransferase reduces 

salicylic acid-mediated pathogen resistance in Arabidopsis thaliana. 

Plant Mol. Biol. 64, 1–15. 

Koroleva OA, Tomos AD, Farrar J, Pollock CJ (2002) Changes in 

osmotic and turgor pressure in response to sugar accumulation in 

barley source leaves. Planta 215, 210–219. 

Kramer EM, Lewandowski M, Beri S, Bernard J, Borkowski M, 

Borkowski MH, Burchfield LA, Mathisen B, Normanly J (2008) 

Auxin gradients are associated with polarity changes in trees. 

Science 320, 1610. 

Krämer U (2010) Metal hyperaccumulation in plants. Annu. Rev. Plant 

Biol. 61, 517–534. 

Krämer U, Cotter-Howells JD, Charnock JM, Baker AJM, Smith JAC 

(1996) Free histidine as a metal chelator in plants that accumulate 

nickel. Nature 379, 635–638. 

Krecek P, Skupa P, Libus J, Naramoto S, Tejos R, Friml J, 

Zazimalova E (2009) The pin-formed (pin) protein family of auxin 

transporters. Genome Biol. 10, 249. 

Krügel U, Veenhoff LM, Langbein J, Wiederhold E, Liesche J, 

Friedrich T, Grimm B, Martinoia E, Poolman B, Kühn C (2008) 

Transport and sorting of Solanum tuberosum sucrose transporter 

SUT1 is affected by posttranslational modification. Plant Cell 20, 

2497–2513. 

Krüger C, Berkowitz O, Stephan U, Hell R (2002) A metal-binding 

member of the late embryogenesis abundant protein family transports 

iron in the phloem of Ricinus communis L. J. Biol. Chem. 277, 

25062–25069. 

Kubo M, Udagawa M, Nishikubo N, Horiguchi G, Yamaguchi M, Ito 

J, Mimura T, Fukuda H, Demura T (2005) Transcription switches 

for protoxylem and metaxylem vessel formation. Genes Dev. 19, 

1855–1860. 

Kudo T, Kiba T, Sakakibara H (2010) Metabolism and long-distance 

translocation of cytokinins. J. Integr. Plant Biol. 52, 53–60. 

Kumar D, Klessig DF (2003) High-affinity salicylic acid-binding protein 

2 is required for plant innate immunity and has salicylic acid 

stimulated lipase activity. Proc. Natl. Acad. Sci. USA 100, 16101– 

16106.


Kunkel BN, Brooks DM (2002) Cross talk between signaling pathways. 

Curr. Opin. Plant Biol. 5, 325–331. 

Küpper H, Mijovilovich A, Meyer-Klaucke W, Kroneck PMH (2004) 

Tissue- and age-dependent differences in the complexation of 

cadmium and zinc in the cadmium/zinc hyperaccumulator Thlaspi 

caerulescens (Ganges Ecotype) revealed by X-ray absorption spectroscopy. 


Kuriyama H, Fukuda H (2002) Developmental programmed cell death 

in plants. Curr. Opin. Plant Biol. 5, 568–573. 

Kwon SI, Cho HJ, Jung JH, Yoshimoto K, Park OK (2010) The 

RabGTPase RabG3b functions in autophagy and contributes to 

tracheary element differentiation in Arabidopsis. Plant J. 64, 151– 

164. 

Lachaud S, Maurousset L (1996) Occurrence of plasmodesmata 

between differentiating vessels and other xylem cells in Sorbus 

torminalis L. Crantz and their fate during xylem maturation. Protoplasma 

191, 220–226. 

Lai F, Thacker J, Li Y, Doerner P (2007) Cell division activity 

determines the magnitude of phosphate starvation responses in 


Lalonde S, Tegeder M, Throne-Holst M, Frommer WB, Patrick JW 

(2003) Phloem loading, unloading of amino acids, sugars. Plant Cell 

Environ. 26, 37–56. 

Langan S, Ewers FW, Davis SD (1997) Xylem dysfunction caused by 

water stress and freezing in two species of co-occurring chaparral 

shrubs. Plant Cell Environ. 20, 425–437. 

Lau S, Shao N, Bock R, Jürgens G, De Smet I (2009) Auxin 

signaling in algal lineages: Fact or myth? Trends Plant Sci. 14, 182– 

188. 

Lavicoli A, Boutet E, Buchala A, Metraux JP (2003) Induced resistance 

in Arabidopsis thaliana in response to root inoculation with 

Pseudomonas fluorescens CHA0. Mol. Plant-Microbe Interact. 16, 

851–858. 

Lawton K, Weymann K, Friedrich L, Vernooij B, Uknes S, Ryals 

J (1995) Systemic acquired resistance in Arabidopsis requires 

salicylic acid but not ethylene. Mol. Plant-Microbe Interact. 8, 863– 

870. 

Le Jean ML, Schikora A, Mari S, Briat JF, Curie C (2005) A loss-offunction 

mutation in AtYSL1 reveals its role in iron and nicotianamine 

seed loading. Plant J. 44, 769–782. 

Lee DK, Sieburth LE (2012) The bps signal: Embryonic arrest from 

an auxin-independent mechanism in bypass triple mutants. Plant 

Signal. Behav. 7, 698–700. 

Lee DK, Van Norman JM, Murphy C, Adhikari E, Reed JW, Sieburth 

LE (2012) In the absence of BYPASS1-related gene function, 

the bps signal disrupts embryogenesis by an auxin-independent 

mechanism. Development 139, 805–815. 

Lee HI, Raskin I (1998) Glucosylation of salicylic acid in Nicotiana 

tabacum cv. Xanthi-nc. Phytopathol. 88, 692–697. 

Lee HI, Raskin I (1999) Purification, cloning, and expression of a 

pathogen inducible UDP-glucose:salicylic acid glucosuyltransferase 

from tobacco. J. Biol. Chem. 274, 36637–36642. 

Lee JY, Colinas J, Wang JY, Mace D, Ohler U, Benfey PN (2006) 

Transcriptional and posttranscriptional regulation of transcription 

factor expression in Arabidopsis roots. Proc. Natl. Acad. Sci. USA 

103, 6055–6060. 

Lee J, Holbrook NM, Zwieniecki MA (2012) Ion-induced changes 

in the structure of bordered pit membranes. Front. Plant Sci. 3, 

1–4. 

Lee JY, Yoo BC, Rojas M, Gomez Ospina N, Staehelin LA, Lucas 

WJ (2003) Selective trafficking of non-cell-autonomous proteins 

mediated by NtNCAPP1. Science 299, 392–396. 

Leggewie G, Kolbe A, Lemoine R, Roessner U, Lytovchenko 

A, Zuther E, Kehr J, Frommer WB, Riemeier JW, Willmitzer 

L, Fernie AR (2003) Overexpression of the sucrose transporter 

SoSUT1 in potato results in alterations in leaf carbon partitioning 

and in tuber metabolism but has little impact on tuber morphology. 

Planta 217, 158–167. 

Lehesranta SJ, Lichtenberger R, Helariutta Y (2010) Cell-to-cell 

communication in vascular morphogenesis. Curr. Opin. Plant Biol. 

13, 59–65. 

Lehmann K, Hause B, Altmann D, Kock M (2001) Tomato ribonuclease 

LX with the functional endoplasmic reticulum retention motif 

HDEF is expressed during programmed cell death processes, 

including xylem differentiation, germination, and senescence. Plant 

Physiol. 127, 436–449. 

Lehto T, Lavola A, Julkunen-Tiitto R, Aphalo PJ (2004) Boron 

retranslocation in scots pine and norway spruce. Tree Physiol. 24, 

1011–1017. 

Lens F, Sperry JS, Christman MA, Choat B, Rabaey D, Jansen 

S (2010) Testing hypotheses that link wood anatomy to cavitation 

resistance and hydraulic conductivity in the genus Acer. New Phytol. 

190, 709–723. 

Leonardos ED, Micallef BJ, Micallef MC, Grodzinski B (2006) Diel 

patterns of leaf export and of main shoot growth for Flaveria linearis 

with altered leaf sucrose-starch partitioning. J. Exp. Bot. 57, 801– 

814. 

Levesque MP, Vernoux T, Busch W, Cui H, Wang JY, Blilou I, 

Hassan H, Nakajima K, Matsumoto N, Lohmann JU, Scheres B, 

Benfey PN (2006) Whole-genome analysis of the SHORT-ROOT 

developmental pathway in Arabidopsis. PLoS Biol. 4, e143. 

Li CY, Zhang K, Zeng XW, Jackson S, Zhou Y, Hong YG (2009) A 

cis element within Flowering Locus T mRNA determines its mobility 

and facilitates trafficking of heterologous viral RNA. J. Virol. 83, 

3540–3548. 

Li P, Ham BK, Lucas WJ (2011) CmRBP50 phosphorylation is 

essential for assembly of a stable phloem-mobile high-affinity ribonucleoprotein 

complex. J. Biol. Chem. 286, 23142–23149. 

Li X, Zhang Y, Clarke JD, Li Y, Dong X (1999) Identification 

and cloning of a negative regulator of systemic acquired 

resistance, SNI1, through a screen for supressors of npr1-1. Cell 

3, 329–339. 

Li Y, Beisson F, Koo AJ, Molina I, Pollard M, Ohlrogge J (2007) 

Identification of acyltransferases required for cutin biosynthesis and

production of cutin with suberin-like monomers. Proc. Natl. Acad. 

Sci. USA 104, 18339–18344. 

Liao MT, Hedley MJ, Woolley DJ, Brooks RR, Nichols MA (2000) 

Copper uptake and translocation in chicory (Cichorium intybus L. 

cv Grasslands Puna) and tomato (Lycopersicon esculentum Mill. cv 

Rondy) plants grown in NFT system. II. The role of nicotianamine 

and histidine in xylem sap copper transport. Plant Soil 223, 243– 

252. 

Liesche J, Schulz A (2012) In vivo quantification of cell coupling in 

plants with different phloem loading strategies. Plant Physiol. 159, 

355–365. 

Lifschitz E, Eviatar T, Rozman A, Shalit A, Goldshmidt A, Amsellem 

Z, Alvarez JP, Eshed Y (2006) The tomato FT ortholog triggers 

systemic signals that regulate growth and flowering and substitute 

for diverse environmental stimuli. Proc. Natl. Acad. Sci. USA 103, 

6398–6403. 

Ligrone R, Duckett JG, Renzaglia KS (2000) Conducting tissues and 

phyletic relationships of bryophytes. Phil. Trans. R. Soc. Lond. B. 

Biol. Sci. 355, 795–813. 

Ligrone R, Duckett JG, Renzaglia KS (2012) Major transitions in the 

evolution of early land plants: A bryological perspective. Ann. Bot. 

109, 851–871. 

Lin MK, Belanger H, Lee YJ, Varkonyi-Gasic E, Taoka KI, Miura 

E, Xoconostle-Cázares B, Gendler K, Jorgensen RA, Phinney 

B, Lough TJ, Lucas WJ (2007) FT protein may act as the longdistance 

florigenic signal in the cucurbits. Plant Cell 19, 1488–1506. 

Lin MK, Lee YJ, Lough TJ, Phinney BS, Lucas WJ (2009) Analysis 

of the pumpkin phloem proteome provides insights into angiosperm 

sieve tube function. Mol. Cell. Proteomics 8, 343–356. 

Lin SI, Chiang SF, Lin WY, Chen JW, Tseng CY, Wu PC, Chiou TJ 

(2008) Regulatory network of microRNA399 and PHO2 by systemic 

signaling. Plant Physiol. 147, 732–746. 

Lindermayr C, Sell S, Müller B, Leister D, Durner J (2010) Redox 

regulation of the NPR1-TGA1 system of Arabidopsis thaliana by 

nitric oxide. Plant Cell 22, 2894–2907. 

Linkohr BI, Williamson LC, Fitter AH, Leyser HMO (2002) Nitrate 

and phosphate availability and distribution have different effects on 

root system architecture of Arabidopsis. Plant J. 29, 751–760. 

Liu G, Holub EB, Alonso JM, Ecker JR, Fobert PR (2005) An Arabidopsis 

NPR1-like gene, NPR4, is required for disease resistance. 


Liu PP, Dahl CC, Klessig DF (2011) The extent to which methyl 

salicylate is required for systemic acquired resistance is dependent 

on exposure to light after infection. Plant Physiol. 157, 2216–2226. 

LiuTY,ChangCY,ChiouTJ(2009) The long-distance signaling of 

mineral macronutrients. Curr. Opin. Plant Biol. 12, 312–319. 

Liu Y, Schiff M, Marathe R, Dinesh-Kumar SP (2002) Tobacco Rar1, 

EDS1 and NPR1/NIM1 like genes are required for N-mediated 

resistance to tobacco mosaic virus. Plant J. 30, 415–429. 

Loepfe L, Martinez-Vilalta J, Pinol J, Mencuccini M (2007) The 

relevance of xylem network structure for plant hydraulic efficiency 

and safety. J. Theor. Biol. 247, 788–803. 


Lomax TL, Hicks GR (1992) Specific auxin-binding proteins in the 

plasma membrane: Receptors or transporters? Biochem. Soc. 

Trans. 20, 64–69. 

Loon LCV, Gerritsen YAM, Ritter CE (1987) Identification, purification, 

and characterization of pathogenesis-related proteins from virusinfected 

Samsun NN tobacco leaves. Plant Mol. Biol. 9, 593–609. 

López-Bucio J, Hernández-Abreu E, Sánchez-Calderón L, Nieto- 

Jacobo MF, Simpson J, Herrera-Estrella L (2002) Phosphate 

availability alters architecture and causes changes in hormone 

sensitivity in the Arabidopsis root system. Plant Physiol. 129, 244– 

256. 

López-Millán AF, Morales F, Abadía A, Abadía J (2000) Effects of 

iron deficiency on the composition of the leaf apoplastic fluid and 

xylem sap in sugar beet. Implications for iron and carbon transport. 


Lough TJ, Lucas WJ (2006) Integrative plant biology: Role of phloem 

long-distance macromolecular trafficking. Annu. Rev. Plant Biol. 57, 

203–232. 

Lu KJ, Huang NC, Liu YS, Lu CA, Yu TS (2012) Long-distance 

movement of Arabidopsis FLOWERING LOCUS T RNA participates 

in systemic floral regulation. RNA Biology 9, 653–662. 

Lucas WJ (2006) Plant viral movement proteins: Agents for cell-to-cell 

trafficking of viral genomes. Virology 344, 169–184. 

Lucas WJ, Ding B, van der Schoot C (1993) Plasmodesmata and the 

supracellular nature of plants. New Phytol. 125, 435–476. 

Lucas WJ, Yoo BC, Kragler F (2001) RNA as a long-distance 

information macromolecule in plants. Nat. Rev. Molec. Cell Biol. 

2, 849–857. 

Lundmark M, Cavaco AM, Trevanior S, Hurry V (2006) Carbon 

partitioning and export in transgenic Arabidopsis thaliana with 

altered capacity for sucrose synthesis grown at low temperature: 

A role for metabolite transporters. Plant Cell Environ. 29, 1703– 

1714. 

MaKW,FloresC,MaW(2011) Chromatin configuration as a battlefield 

in plant-bacteria interactions. Plant Physiol. 157, 535–543. 

Ma Y, Miura E, Ham BK, Cheng HW, Lee YJ, Lucas WJ (2010) 

Pumpkin eIF5A isoforms interact with components of the translational 

machinery in the cucurbit sieve tube system. Plant J. 64, 

536–550. 

Madey E, Nowack LM, Thompson JE (2002) Isolation and characterization 

of lipid in phloem sap of canola. Planta 214, 625–634. 

Mähönen AP, Bishopp A, Higuchi M, Nieminen KM, Kinoshita K, 

Törmäkangas K, Ikeda Y, Oka A, Kakimoto T, Helariutta Y (2006) 

Cytokinin signaling and its inhibitor AHP6 regulate cell fate during 

vascular development. Science 311, 94–98. 

Mähönen AP, Bonke M, Kauppinen L, Riikonen M, Benfey PN, 

Helariutta Y (2000) A novel two-component hybrid molecule regulates 

vascular morphogenesis of the Arabidopsis root. Genes Dev. 

14, 2938–2943. 

Mahajan A, Bhogale S, Kang IH, Hannapel DJ, Banerjee AK (2012) 

The mRNA of a knotted1-like transcription factor of potato is phloem 

mobile. Plant Mol. Biol. 79, 595–608.


Maherali H, Pockman WT, Jackson RB (2003) Adaptive variation in 

the vulnerability of woody plants to xylem cavitation. Ecology 85, 

2184–2199. 

Maldonado AM, Doerner P, Dixon RA, Lamb CJ, Cameron RKA 

(2002) Putative lipid transfer protein involved in systemic resistance 

signaling in Arabidopsis. Nature 419, 399–403. 

Mandal M, Chanda B, Xia Y, Yu K, Sekine K, Gao QM, Selote D, 

Kachroo A, Kachroo P (2011) Glycerol-3-phosphate and systemic 

immunity. Plant Signal. Behav. 6, 1871–1874. 

Mandal MK, Chandra-Shekara AC, Jeong RD, Yu K, Zhu S, Chanda 

B, Navarre D, Kachroo A, Kachroo P (2012) Oleic acid-dependent 

modulation of NITRIC OXIDE ASSOCIATED 1 protein levels 

regulates nitric oxide-mediated defense signaling in Arabidopsis. 


March-Díaz R, García-Domínguez M, Lozano-Juste J, León J, 

Florencio FJ, Reyes JC (2008) Histone H2A.Z and homologues of 

components of the SWR1 complex are required to control immunity 

in Arabidopsis. Plant J. 53, 475–487. 

Mari S, Gendre D, Pianelli K, Ouerdane L, Lobinski R, Briat JF, 

Lebrun M, Czernic P (2006) Root-to-shoot long-distance circulation 

of nicotianamine and nicotianamine-nickel chelates in the metal 

hyperaccumulator Thlaspi caerulescens. J. Exp. Bot. 57, 4111– 

4122. 

Markesteijn L, Poorter L, Paz H, Sack L, Bongers F (2011) Ecological 

differentiation in xylem cavitation is associated with stem and leaf 

structural traits. Plant Cell Environ. 34, 137–148. 

Marschner H (1995) Mineral Nutrition of Higher Plants. Academic 

Press, London. 

Martin A, Adam H, Diaz-Mendoza M, Zurczak M, Gonzalez-Schain 

ND, Suarez-Lopez P (2009) Graft-transmissible induction of potato 

tuberization by the microRNA miR172. Development 136, 2873– 

2881. 

Martin AC, del Pozo J, Iglesias J, Rubio V, Solano R, de La Pena 

A, Leyva A, Paz-Ares J (2000) Influence of cytokinins on the expression 

of phosphate starvation responsive genes in Arabidopsis. 


Mathieu J, Warthmann N, Kuttner F, Schmid M (2007) Export of FT 

protein from phloem companion cells is sufficient for floral induction 

in Arabidopsis. Curr. Biol. 17, 1055–1060. 

Matsubayashi Y, Sakagami Y (2006) Peptide hormones in plants. 

Annu Rev. Plant Biol. 57, 649–674. 

Matsumoto-Kitano M, Kusumoto T, Tarkowski P, Kinoshita- 

Tsujimura K, Václavíková K, Miyawaki K, Kakimoto T (2008) 

Cytokinins are central regulators of cambial activity. Proc. Natl. 

Acad. Sci. USA 105, 20027–20031. 

Matte Risopatron JP, Sun Y, Jones BJ (2010) The vascular cambium: 

Molecular control of cellular structure. Protoplasma 247, 145–161. 

Mattsson J, Ckurshumova W, Berleth T (2003) Auxin signaling in 

Arabidopsis leaf vascular development. Plant Physiol. 131, 1327– 

1339. 

Mayr S, Sperry JS (2010) Freeze-thaw induced embolism in Pinus 

contorta: Centrifuge experiments validate the “thaw-expansion” 

hypothesis but conflict with ultrasonic data. New Phytol. 18, 1016– 

1024. 

Mayr S, Zublasing V (2010) Ultrasonic emissions from conifer xylem 

exposed to repeated freezing. Plant Physiol. 167, 34–40. 

Mayzlish-Gati E, De-Cuyper C, Goormachtig S, Beeckman T, Vuylsteke 

M, Brewer PB, Beveridge CA, Yermiyahu U, Kaplan 

Y, Enzer Y, Wininger S, Resnick N, Cohen M, Kapulnik Y, 

Koltai H (2012) Strigolactones are involved in root response to 

low phosphate conditions in Arabidopsis. Plant Physiol. 160, 1329– 

1341. 

McAdam SAM, Brodribb TJ (2012) Stomatal innovation and the rise 

of seed plants. Ecol. Lett. 15, 1–8. 

McCarthy RL, Zhong R, Ye ZH (2009) MYB83 is a direct target of 

SND1 and acts redundantly with MYB46 in the regulation of secondary 

cell wall biosynthesis in Arabidopsis. Plant Cell Physiol. 50, 

1950–1964. 

McConnell JR, Barton MK (1998) Leaf polarity and meristem formation 

in Arabidopsis. Development 125, 2935–2942. 

McConnell JR, Emery J, Eshed Y, Bao N, Bowman J, Barton 

MK (2001) Role of PHABULOSA and PHAVOLUTA in determining 

radial patterning in shoots. Nature 411, 709–713. 

McNear DH Jr, Chaney RL, Sparks DL (2010) The hyperaccumulator 

Alyssum murale uses complexation with nitrogen and oxygen 

donor ligands for Ni transport and storage. Phytochemistry 71, 

188–200. 

Meinzer FC, Goldstein G, Jackson P, Holbrook NM, Gutierrez 

MV, Cavelier J (1995) Environmental and physiological regulation 

of transpiration in tropical forest gap species: The influence 

of boundary layer and hydraulic properties. Oecologia 101, 

514–522. 

Melnyk CW, Molnar A, Baulcombe DC (2011) Intercellular and 

systemic movement of RNA silencing signals. EMBO J. 30, 3553– 

3563. 

Mencuccini M, Hölttä T(2010) The significance of phloem transport 

for the speed with which canopy photosynthesis and belowground 

respiration are linked. New Phytol. 185, 189–203. 

Mercury L, Tardy Y (2001) Negative pressure of stretched liquid water. 

Geochemistry of soil capillaries. Geochem. Cosmochem. Acta 65, 

3391–3408. 

Mijovilovich A, Leitenmaier B, Meyer-Klaucke W, Kroneck PMH, 

Gotz B, Küpper H (2009) Complexation and toxicity of copper in 

higher plants. II. Different mechanisms for copper versus cadmium 

detoxification in the copper-sensitive cadmium/zinc hyperaccumulator 

Thlaspi caerulescens (Ganges ecotype). Plant Physiol. 151, 

715–731. 

Milburn JA, Kallarackal J (1989) Physiological aspects of phloem 

translocation. In: Baker DA, Milburn JA, eds. Transport of Photoassimilates. 

Lonhman Scientific & Technical, Essex. pp. 262–305. 

Milioni D, Sado PE, Stacey NJ, Roberts K, McCann MC (2002) Early 

gene expression associated with the commitment and differentiation 

of a plant tracheary element is revealed by cDNA-amplified fragment 

length polymorphism analysis. Plant Cell 14, 2813–2824.

Mills RF, Krijgert GC, Baccarini PJ, Hall JL, Williams LE (2003) 

Functional expression of AtHMA4, a P1B-type ATPase of the 

Zn/Co/Cd/Pb subclass. Plant J. 35, 164–176. 

Mills RF, Francini A, Ferreira Da Rocha PSC, Baccarini PJ, Aylett 

M, Krijger GC, Williams LE (2005) The plant P1B-type ATPase 

AtHMA4 transports Zn and Cd and plays a role in detoxification 

of transition metals supplied at elevated levels. FEBS Lett. 579, 

783–791. 

Minchin PEH, Thorpe MR (1983) A rate of cooling response in phloem 

translocation. J. Exp. Bot. 34, 529–536. 

Mira H, Martínez-García F, Peñarrubia L (2001) Evidence for the 

plant-specific intercellular transport of the Arabidopsis copper chaperone 

CCH. Plant J. 25, 521–528. 

Mishina TE, Zeier J (2006) The Arabidopsis flavin-dependent 

monooxygenase FMO1 is an essential component of biologically 

induced systemic acquired resistance. Plant Physiol. 141, 1666– 

1675. 

Mishler BD, Churchill SP (1984) A cladistic approach to the phylogeny 

of the “bryophytes.” Brittonia 36, 406–424. 

Misra NB, Varshini YP (1961) Viscosity-temperature relation to solutions. 

J. Chem. Eng. Data 6, 194–196. 

Mitsuda N, Seki M, Shinozaki K, Ohme-Takagi M (2005) The NAC 

transcription factors NST1 and NST2 of Arabidopsis regulate secondary 

wall thickenings and are required for anther dehiscence. 


Mitsuda N, Iwas A, Yamamoto H, Yoshida M, Seki M, Shinozaki 

K, Ohme-Takagi M (2007) NAC transcription factors, NST1 and 

NST3, are key regulators of the formation of secondary walls in 

woody tissues of Arabidopsis. Plant Cell 19, 270–280. 

Mittelheuser CJ, Van Steveninck RFM (1969) Stomatal closure and 

inhibition of transpiration induced by (RS)-abscisic acid. Nature 221, 

281–282. 

Miwa H, Betsuyaku S, Iwamoto K, Kinoshita A, Fukuda H, Sawa 

S (2008) The receptor-like kinase SOL2 mediates CLE signaling in 

Arabidopsis. Plant Cell Physiol. 49, 1752–1757. 

Miwa K, Fujiwara T (2010) Boron transport in plants: Co-ordinated 

regulation of transporters. Ann. Bot. 105, 1103–1108. 

Miyashima S, Nakajima K (2011) The root endodermis: A hub of 

developmental signals and nutrient flow. Plant Signal. Behav. 6, 

1954–1958. 

Miyashima S, Koi S, Hashimoto T, Nakajima K (2011) Non-cellautonomous 

microRNA165 acts in a dose-dependent manner to 

regulate multiple differentiation status in the Arabidopsis root. Development 

138, 2303–2313. 

Miyashima S, Sebastian J, Lee JY, Helariutta Y (2012) Stem cell 

function during plant vascular development. EMBO J. 32, 178–193. 

Miyazawa H, Oka-Kira E, Sato N, Takahashi H, Wu GJ, Sato S, 

Hayashi M, Betsuyaku S, Nakazono M, Tabata S, Harada K, 

Sawa S, Fukuda H, Kawaguchi M (2010) The receptor-like kinase 

KLAVIER mediates systemic regulation of nodulation and nonsymbiotic 

shoot development in Lotus japonicus. Development 137, 

4317–4325. 


Moeder W, Pozo OD, Navarre DA, Martin GB, Klessig DF (2007) 

Aconitase plays a role in regulating resistance to oxidative stress 

and cell death in Arabidopsis and Nicotiana benthamiana. Plant Mol. 

Biol. 63, 273–287. 

Mok DW, Mok MC (2001) Cytokinin metabolism and action. Annu. Rev. 

Plant Physiol. Plant Mol. Biol. 52, 89–118. 

Molders W, Buchala A, Metraux J-P (1996) Transport of salicylic acid 

in tobacco necrosis virus-infected cucumber plants. Plant Physiol. 

112, 787–792. 

Molnar A, Melnyk CW, Bassett A, Hardcastle TJ, Dunn R, 

Baulcombe DC (2010) Small silencing RNAs in plants are mobile 

and direct epigenetic modification in recipient cells. Science 328, 

872–875. 

Moreno-Risueno MA, Van Norman JM, Moreno A, Zhang JY, Ahnert 

SE, Benfey PN (2010) Oscillating gene expression determines 

competence for periodic Arabidopsis root branching. Science 329, 

1306–1311. 

Morrissey J, Baxter IR, Lee J, Li L, Lahner B, Grotz N, Kaplan J, 

Salt DE, Guerinot ML (2009) The ferroportin metal efflux proteins 

function in iron and cobalt homeostasis in Arabidopsis. Plant Cell 

21, 3326–3338. 

Mosher RA, Durrant WE, Wang D, Song J, Dong X (2006) A comprehensive 

structure-function analysis of Arabidopsis SNI1 defines 

essential regions and transcriptional supppressor activity. Plant Cell 

18, 1750–1765. 

Mosher RA, Schwach F, Studhollme D, Baulcombe DC (2008) 

PolIVb influences RNA-directed DNA-methylation independently of 

its role in siRNA biogenesis. Proc. Natl. Acad. Sci USA 105, 3145– 

3150. 

Motose H, Sugiyama M, Fukuda H (2004) A proteoglycan mediates 

inductive interaction during plant vascular development. Nature 429, 

873–878. 

Mott KA, Peak D (2011) Alternative perspective on the control of 

transpiration by radiation. Proc. Natl. Acad. Sci. USA 108, 19820– 

19823. 

Mou Z, Fan W, Dong X (2003) Inducers of plant systemic acquired 

resistance regulate NPR1 function through redox changes. Cell 113, 

935–944. 

Mullendore DL, Windt CW, Van As H, Knoblauch M (2010) Sieve 

tube geometry in relation to phloem flow. Plant Cell 22, 579– 

593. 

Müller R, Bleckmann A, Simon R (2008) The receptor kinase 

CORYNE of Arabidopsis transmits the stem cell-limiting signal 

CLAVATA3 independently of CLAVATA1. Plant Cell 20, 934–946. 

Münch E (1930) Material Flow in Plants. Translated 2003 by Milburn 

JA and Kreeb KH, University of Bremen, Germany. Gustav Fischer 

Verlag, Jena Germany. 

Muñiz L, Minguet EG, Singh SK, Pesquet E, Vera-Sirera F, Moreau- 

Courtois CL, Carbonell J, Blázquez MA, Tuominen H (2008) 

ACAULIS5 controls Arabidopsis xylem specification through the 

prevention of premature cell death. Development 135, 2573– 

2582.


Murphy A, Taiz L (1997) Correlation between potassium efflux and 

copper sensitivity in 10 Arabidopsis ecotypes. New Phytol. 136, 

211–222. 

NagahashiG,DoudsDD(2000) Partial separation of root exudate 

components and their effects upon the growth of germinated spores 

of AM fungi. Mycol Res. 104, 1453–1464. 

Nagawa S, Sawa S, Sato S, Kato T, Tabata S, Fukuda H (2006) 

Gene trapping in Arabidopsis reveals genes involved in vascular 

development. Plant Cell Physiol. 47, 1394–1405. 

Naik SN, Goud VV, Rout PK, Dalai AK (2010) Production of first and 

second generation biofuels: A comprehensive review. Renew. Sust. 

Energ. Rev. 14, 578–597. 

Napoli C (1996) Highly branched phenotype of the petunia dad1-1 

mutant is reversed by grafting. Plant Physiol. 111, 27–37. 

Nardini A, LoGullo MA, Salleo S (2011a) Refilling embolized xylem 

conduits: Is it a matter of phloem unloading? Plant Sci. 180, 604– 

611. 

Nardini A, Salleo S, Jansen S (2011b) More than just a vulnerable 

pipeline: Xylem physiology in the light of ion-mediated regulation of 

plant water transport. J. Exp. Bot. 62, 4701–4718. 

Nawrath C, Heck S, Parinthawong N, Métraux JP (2002) EDS5, 

an essential component of salicylic acid-dependent signaling for 

disease resistance in Arabidopsis, is a member of the MATE 

transporter family. Plant Cell 14, 275–286. 

Neale DB, Kremer A (2011) Forest tree genomics: Growing resources 

and applications. Nat. Rev. Genet. 12, 111–122. 

Nelson T, Dengler N (1997) Leaf vascular pattern formation. Plant Cell 

9, 1121–1135. 

Niggeweg R, Thurow C, Weigel R, Pfitzner U, Gatz C (2000) 

Tobacco TGA factors differ with respect to interaction with NPR1, 

activation potential and DNA-binding properties. Plant Mol. Biol. 42, 

775–788. 

Nilsson J, Karlberg A, Antti H, Lopez-Vernaza M, Mellerowicz 

E, Perrot-Rechenmann C, Sandberg G, Bhalerao RP (2008) 

Dissecting the molecular basis of the regulation of wood formation 

by auxin in hybrid aspen. Plant Cell 20, 843–855. 

Nishiyama R, Kato M, Nagata S, Yanagisawa S, Yoneyama T (2012) 

Identification of Zn-nicotianamine and Fe-2 ′ -deoxymugineic acid in 

the phloem sap from rice plants (Oryza sativa L.). Plant Cell Physiol. 

53, 381–390. 

Notaguchi M, Abe M, Kimura T, Daimon Y, Kobayashi T, Yamaguchi 

A, Tomita Y, Dohi K, Mori M, Araki T (2008) Long-distance, grafttransmissible 

action of Arabidopsis FLOWERING LOCUS T protein 

to promote flowering. Plant Cell Physiol. 49, 1645–1658. 

Notaguchi M, Wolf S, Lucas WJ (2012) Phloem-mobile Aux/IAA 

transcripts target to the root tip and modify root architecture. J. 

Integr. Plant Biol. 54, 760–772. 

Oda Y, Fukuda H (2012a) How xylem cells design wood cell walls: 

Secondary cell wall patterning by microtubule-associated proteins. 


Oda Y, Fukuda H (2012b) Initiation of cell wall pattern by a Rho- and 

microtubule-driven symmetry breaking. Science 337, 11333–11336. 

Oda Y, Hasezawa S (2006) Cytoskeletal organization during xylem cell 

differentiation. J. Plant Res. 119, 167–177. 

Oda Y, Iida Y, Kondo Y, Fukuda H (2010) Wood cell-wall structure 

requires local 2D-microtubule disassembly by a novel plasma 

membrane-anchored protein. Curr. Biol. 20, 1197–1202. 

Oda Y, Mimura T, Hasezawa S (2005) Regulation of secondary cell 

wall development by cortical microtubules during tracheary element 

differentiation in Arabidopsis cell suspensions. Plant Physiol. 137, 

1027–1036. 

Oertli JJ (1993) The mobility of boron in plants. Plant Soil 155, 301– 

304. 

Ohashi-Ito K, Fukuda H (2003) HD-zip III homeobox genes that include 

a novel member, ZeHB-13 (Zinnia)/ATHB-15 (Arabidopsis), are 

involved in procambium and xylem cell differentiation. Plant Cell 

Physiol. 44, 1350–1358. 

Ohashi-Ito K, Oda Y, Fukuda H (2010) Arabidopsis VASCULAR- 

RELATED NAC-DOMAIN6 directly regulates the genes that govern 

programmed cell death and secondary wall formation during xylem 

differentiation. Plant Cell 22, 3461–3473. 

Oka-Kira E, Tateno K, Miura K-I, Haga T, Hayashi M, Harada 

K, Sato S, Tabata S, Shikazono N, Tanaka A, Watanabe Y, 

Fukuhara I, Nagata T, Kawaguchi M (2005) klavier (klv), A novel 

hypernodulation mutant of Lotus japonicus affected in vascular 

tissue organization and floral induction. Plant J. 44, 505–515. 

Okamoto S, Ohnishi E, Sato S, Takahashi H, Nakazono M, Tabata 

S, Kawaguchi M (2009) Nod factor/nitrate-induced CLE genes that 

drive HAR1-mediated systemic regulation of nodulation. Plant Cell 

Physiol. 50, 67–77. 

Omid A, Keilin T, Glass A, Leshkowitz D, Wolf S (2007) Characterization 

of phloem-sap transcription profile in melon plants. J. Exp. 

Bot. 58, 3645–3656. 

Oren R, Sperry JS, Katul GG, Pataki DE, Ewers BE, Phillips N, 

Schafer KVR (1999) Survey and synthesis of intra- and interspecific 

variation in stomatal sensitivity to vapour pressure deficit. Plant Cell 

Environ. 22, 1515–1526. 

Orlando DA, Brady SM, Fink TM, Benfey PN, Ahnert SE (2010) 

Detecting separate time scales in genetic expression data. BMC 

Genomics 11, 381. 

Oross JW, Lucas WJ (1985) Sugar-beet petiole structure – vascular 

anastomoses and phloem ultrastructure. Can. J. Bot. 63, 2295– 

2304. 

Osipova MA, Mortier V, Demchenko KN, Tsyganov VE, 

Tikhonovich IA, Lutova LA, Dolgikh EA, Goormachtig S (2012) 

Wuschel-related homeobox5 gene expression and interaction of 

CLE peptides with components of the systemic control add two 

pieces to the puzzle of autoregulation of nodulation. Plant Physiol. 

158, 1329–1341. 

Ouerdane L, Mari S, Czernic P, Lebrun M, Łobiński R (2006) 

Speciation of non-covalent nickel species in plant tissue extracts 

by electrospray Q-TOFMS/MS after their isolation by 2D size 

exclusion-hydrophilic interaction LC (SEC-HILIC) monitored by ICP- 

MS. J. Anal. At. Spec. 21, 676–683.

Palauqui JC, Elmayan T, deBorne FD, Crete P, Charles C, 

Vaucheret H (1996) Frequencies, timing, and spatial patterns of cosuppression 

of nitrate reductase and nitrite reductase in transgenic 

tobacco plants. Plant Physiol. 112, 1447–1456. 

Palauqui JC, Elmayan T, Pollien JM, Vaucheret H (1997) Systemic 

acquired silencing: Transgene-specific post-transcriptional silencing 

is transmitted by grafting from silenced stocks to non-silenced 

scions. EMBO J. 16, 4738–4745. 

Pallas JA, Paiva NL, Lamb C, Dixon RA (1996) Tobacco plants epigenetically 

suppressed in phenylalanine ammonia-lyase expression 

do not develop systemic acquired resistance in response to infection 

by tobacco mosaic virus. Plant J. 10, 281–293. 

Pant BD, Buhtz A, Kehr J, Scheible WR (2008) MicroRNA399 is a 

long-distance signal for the regulation of plant phosphate homeostasis. 


Parizot B, Laplaze L, Ricaud L, Boucheron-Dubuisson E, Bayle 

V, Bonke M, De Smet I, Poethig SR, Helariutta Y, Haseloff J, 

Chriqui D, Beeckman T, Nussaume L (2008) Diarch symmetry of 

the vascular bundle in Arabidopsis root encompasses the pericycle 

and is reflected in distich lateral root initiation. Plant Physiol. 146, 

140–148. 

Park M, Li Q, Shcheynikov N, Zeng W, Muallem S (2004) NaBC1 is 

a ubiquitous electrogenic Na + -coupled borate transporter essential 

for cellular boron homeostasis and cell growth and proliferation. Mol. 

Cell 16, 331–341. 

Park SW, Kaimoyo E, Kumar D, Mosher S, Klessig D (2007) Methyl 

salicylate is a critical mobile signal for plant systemic acquired 

resistance. Science 318, 113–116. 

Park SW, Liu PP, Forouhar F, Vlot AC, Tong L, Tietjen K, Klessig D 

(2009) Use of a synthetic salicylic acid analog to investigate the roles 

of methyl salicylate and its esterases in plant disease resistance. J. 

Biol. Chem. 284, 7307–7317. 

Parker J, Holub E, Frost L, Falk A, Gunn N, Daniels M (1996) 

Characterization of eds1, a mutation in Arabidopsis suppressing 

resistance to Peronospora parasitica specified by several different 

RPP genes. Plant Cell 8, 2033–2046. 

Passioura JB, Ashford AE (1974) Rapid translocation in the phloem 

of wheat roots. Aust. J. Plant Physiol. 1, 521–527. 

Peguero-Pina JS, Alquezar-Alquezar JM, Mayr S, Cochard H, Gil- 

Pelegrin D (2011) Embolism induced by winter drought in Pinus 

sylvestris L.: A possible reason for dieback near its southern 

distribution limit? Ann. For. Sci. 38, 565–574. 

Peleg-Grossman S, Golani Y, Kaye Y, Melamed-Book N, Levine 

A (2009) NPR1 protein regulates pathogenic and symbiotic interactions 

between Rhizobium and legumes and non-legumes. PLoS 

ONE 4, e8399. 

Pérez-Alfocea F, Ghanem ME, Gómez-Cadenas A, Dodd IC (2011) 

Omics of root-to-shoot signaling under salt stress and water deficit. 

OMIC 15, 893–901. 

Pesacreta TC, Groom LH, Rials TG (2005) Atomic force microscopy 

of the intervessel pit membrane in the stem of Sapium sebiferum 

(Euphorbiaceae). IAWA J. 26, 397–426. 


Pesquet E, Ranocha P, Legay S, Digonnet C, Barbier O, Pichon 

M, Goffner D (2005) Novel markers of xylogenesis in zinnia are 

differentially regulated by auxin and cytokinin. Plant Physiol. 139, 

1821–1839. 

Pesquet E, Korolev AV, Calder G, Lloyd CW (2010) The microtubuleassociated 

protein AtMAP70–5 regulates secondary wall patterning 

in Arabidopsis wood cells. Curr. Biol. 20, 744–749. 

Petráˇsek J, Mravec J, Bouchard R, Blakeslee JJ, Abas M, Seifertova 

D, Wisniewska J, Tadele Z, Kubes M, Covanova M, 

Dhonukshe P, Skupa P, Benkova E, Perry L, Krecek P, Lee 

OR, Fink GR, Geisler M, Murphy AS, Luschnig C, Zazimalova 

E, Friml J (2006) PIN proteins perform a rate-limiting function in 

cellular auxin efflux. Science 312, 914–918. 

Petráˇsek J, Friml J (2009) Auxin transport routes in plant development. 


Petty JA (1972) The aspiration of bordered pits in conifer wood. Proc. 

R. Soc. Ser. B. Biol. Sci. 181, 395–406. 

Petty JA, Preston RD (1969) The dimensions and number of pit 

membrane pores in conifer wood. Proc. R. Soc. Ser. B. Biol. Sci. 

172, 137–151. 

Peuke AD, Windt C, van As H (2006) Effects of cold girdling on flows in 

the transport phloem in Ricinus communis: Is mass flow inhibited? 


Pich A, Scholz G (1996) Translocation of copper and other micronutrients 

in tomato plants (Lycopersicon esculentum Mill.): 

Nicotianamine-stimulated copper transport in the xylem. J. Exp. Bot. 

47, 41–47. 

Pich A, Scholz G, Stephan U (1994) Iron-dependent changes of 

heavy-metals, nicotianamine, and citrate in different plant organs 

and in the xylem exudate of 2 tomato genotypes. Nicotianamine 

as possible copper translocator. Plant Soil 165, 189– 

196. 

Pickard WF (1981) The ascent of sap in plants. Progr. Biophys. Mol. 

Biol. 37, 181–229. 

Pieruschka R, Huber G, Berry JA (2010) Control of transpiration by 

radiation. Proc. Natl. Acad. Sci. USA 107, 13372–13377. 

Piquemal J, LaPierre C, Myton K, O’Connel A, Schuch W, Grima- 

Pettenati J, Boudet AM (1998) Down-regulation of cinnamoyl-CoA 

reductase induces significant changes of lignin profiles in transgenic 

tobacco plants. Plant J. 13, 71–83. 

Pires ND, Dolan L (2012) Morphological evolution in land plants: New 

designs with old genes. Phil Trans. R. Soc. 367, 508–518. 

Pittermann J (2010) The evolution of water transport in plants: An 

integrated approach. Goebiology 8, 112–139. 

Pittermann J, Sperry JS (2003) Tracheid diameter is the key trait 

determining extent of freezing-induced cavitation in conifers. Tree 

Physiol. 23, 907–914. 

Pittermann J, Sperry JS (2006) Analysis of freeze-thaw embolism in 

conifers: The interaction between cavitation pressure and tracheid 

size. Plant Physiol. 140, 374–382. 

Pittermann J, Sperry JS, Hacke UG, Wheeler JK, Sikkema EH (2005) 

Torus-margo pits help conifers compete with angiosperms. Science


310, 1924. 

Pittermann J, Sperry JS, Hacke UG, Wheeler JK, Sikkema EH 

(2006a) Inter-tracheid pitting and the hydraulic efficiency of conifer 

wood: The role of tracheid allometry and cavitation protection. Am. 

J. Bot. 93, 1105–1113. 

Pittermann J, Sperry JS, Wheeler JK, Hacke UG, Sikkema EH 

(2006b) Mechanical reinforcement of tracheids compromises the 

hydraulic efficiency of conifer xylem. Plant Cell Environ. 29, 1618– 

1628. 

Plaut JA, Yepez EA, Hill J, Pangle R, Sperry JS, Pockman WT, 

McDowell NG (2012) Hydraulic limits preceeding mortality in a 

pinon-juniper woodland under experimental drought. Plant Cell 

Environ. 35, 1601–1617. 

Pockman WT, Sperry JS (1997) Freezing-induced xylem cavitation 

and the northern limit of Larrea tridentata. Oecologia 109, 19–27. 

Poirier Y, Thoma S, Somerville C, Schiefelbein J (1991) Mutant of 

Arabidopsis deficient in xylem loading of phosphate. Plant Physiol. 

97, 1087–1093. 

Pommerrenig B, Papini-Terzi FS, Sauer N (2007) Differential regulation 

of sorbitol and sucrose loading into the phloem of Plantago 

major in response to salt stress. Plant Physiol. 144, 1029–1038. 

Pressel S, Ligrone R, Duckett JG (2006) Effects of de- and rehydration 

on food-conducting cells in the moss Polytrichum formosum: A 

cytological study. Ann. Bot. 98, 67–76. 

Prigge MJ, Clark SE (2006) Evolution of the Class III HDd-ZIP gene 

family in land plants. Evol. Dev. 8, 350–361. 

Prigge MJ, Otsuga D, Alonso JM, Ecker JR, Drews GN, Clark SE 

(2005) Class III homeodomain-leucine zipper gene family members 

have overlapping, antagonistic, and distinct roles in Arabidopsis 

development. Plant Cell 17, 61–76. 

Pritchard J (1996) Aphid stylectomy reveals an osmotic step between 

sieve tube, cortical cells in barley roots. J. Exp. Bot. 47, 1519– 

1524. 

Pritchard J, Tomos AD, Farrar JF, Minchin PEH, Gould N, Paul MJ, 

MacRae EA, Ferrier RA, Gray DW, Thorpe MR (2004) Turgor, 

solute import, growth in maize roots treated with galactose. Funct. 


Puig S, Penarrubia L (2009) Placing metal micronutrients in context: 

Transport and distribution in plants. Curr. Op. Plant Biol. 12, 299– 

306. 

Pyo H, Demura T, Fukuda H (2007) TERE; a novel cis-element responsible 

for a coordinated expression of genes related to programmed 

cell death and secondary wall formation during differentiation of 

tracheary elements. Plant J. 51, 955–965. 

Rahayu YS, Walch-Liu P, Neumann G, Römheld V, von Wirén N, 

Bangerth F (2005) Root-derived cytokinins as long-distance signals 

for NO − 

3 -induced stimulation of leaf growth. J. Exp. Bot. 56, 1143– 

1152. 

Ransom-Hodgkins WD, Vaughn MW, Bush DR (2003) Protein phosphorylation 

plays a key role in sucrose-mediated transcriptional 

regulation of a phloem-specific proton-sucrose symporter. Planta 

217, 483–489. 

Raridan GJ, Delaney TP (2002) Role of salicylic acid and NIM1/NPR1 

in race-specific resistance in Arabidopsis. Genetics 161, 803–811. 

Raven JA (1991) Long-term functioning of enucleate sieve elements 

- possible mechanisms of damage avoidance and damage repair. 


Raven JA (2003) Long-distance transport in non-vascular plants. Plant 

Cell Environ. 26, 73–85. 

Reid DE, Ferguson BJ, Gresshoff PM (2011) Inoculation- and nitrateinduced 

CLE peptides of soybean control NARK-dependent nodule 

formation. Mol. Plant-Microbe Interact. 24, 606–618. 

Reinhardt D, Mandel T, Kuhlemeier C (2000) Auxin regulates the 

initiation and radial position of plant lateral organs. Plant Cell 12, 

507–518. 

Reinhardt D, Pesce ER, Stieger P, Mandel T, Baltensperger K, 

Bennett M, Traas J, Friml J, Kuhlemeier C (2003) Regulation 

of phyllotaxis by polar auxin transport. Nature 426, 255–260. 

Rellán-Álvarez R, Abadía J, Álvarez-Fernández A (2008) Formation 

of metal-nicotianamine complexes as affected by pH, ligand exchange 

with citrate and metal exchange. A study by electrospray 

ionization time-of-flight mass spectrometry. Rapid Commun. Mass 

Sp. 22, 1553–1562. 

Rellán-Álvarez R, Giner-Martínez-Sierra J, Orduna J, Orera I, 

Rodríguez-Castrillón JA, García-Alonso JI, Abadía J, Álvarez- 

Fernández A (2010) Identification of a tri-iron(III), tri-citrate complex 

in the xylem sap of iron-deficient tomato resupplied with iron: New 

insights into plant iron long-distance transport. Plant Cell Physiol. 

51, 91–102. 

Rennie EA, Turgeon R (2009) A comprehensive picture of phloem 

loading strategies. Proc. Natl. Acad. Sci. USA 106, 14162–14167. 

Riesen O, Feller U (2005) Redistribution of nickel, cobalt, manganese, 

zinc, and cadmium via the phloem in young and maturing wheat. 

J. Plant Nutr. 28, 421–430. 

Roberts MR, Paul ND (2006) Seduced by the dark side: Integrating 

molecular and ecological perspectives on the influence of light on 

plant defense against pests and pathogens. New Phytol. 170, 677– 

699. 

Robischon M, Du J, Miura E, Groover A (2011) The Populus 

Class III HD ZIP, popREVOLUTA, influences cambium initiation 

and patterning of woody stems. Plant Physiol. 155, 1214– 

1225. 

Rodriguez-Medina C, Atkins CA, Mann AJ, Jordan ME, Smith PMC 

(2011) Macromolecular composition of phloem exudate from white 

lupin (Lupinus albus L.). BMC Plant Biol. 11, 36. 

Rogers EE, Ausubel FM (1997) Arabidopsis enhanced disease susceptibility 

mutants exhibit enhanced susceptibility to several bacterial 

pathogens and alterations in PR-1 gene expression. Plant Cell 

9, 305–316. 

Roney JK, Khatibi PA, Westwood JH (2007) Cross-species translocation 

of mRNA from host plants into the parasitic plant dodder. 


Rosche E, Blackmore D, Tegeder M, Richardson T, Schroeder 

H, Higgins TJV, Frommer WB, Offler CE, Patrick JW (2002)

Seed-specific expression of a potato sucrose transporter increases 

sucrose uptake, growth rates of developing pea cotyledons. Plant 

J. 30, 165–175. 

Roschzttardtz H, Séguéla-Arnaud M, Briat J, Vert G, Curie C (2011) 

The FRD3 citrate effluxer promotes iron nutrition between symplastically 

disconnected tissues throughout Arabidopsis development. 


Rothwell GW, Lev-Yadun S (2005) Evidence of polar auxin flow in 375 

million-year-old fossil wood. Am. J. Bot. 92, 903–906. 

Rouached H, Stefanovic A, Secco D, Bulak Arpat A, Gout E, 

Bligny R, Poirier Y (2011) Uncoupling phosphate deficiency from 

its major effects on growth and transcriptome via PHO1 expression 

in Arabidopsis. Plant J. 65, 557–570. 

Rueffer M, Steipe B, Zenk MH (1995) Evidence against specific 

binding of salicylic acid to plant catalase. FEBS Lett. 377, 175–180. 

Ruffel S, Krouk G, Ristova D, Shasha D, Birnbaum KD, Coruzzi 

GM (2011) Nitrogen economics of root foraging: Transitive closure 

of the nitrate-cytokinin relay and distinct systemic signaling for N 

supply vs. demand. Proc. Natl. Acad. Sci. USA 108, 18524–18529. 

Ruiz-Medrano R, Xoconostle-Cázares B, Lucas WJ (1999) Phloem 

long-distance transport of CmNACP mRNA: Implications for supracellular 

regulation in plants. Development 126, 4405–4419. 

Ruszala EM, Beerling DJ, Franks PJ, Chater C, Casson SA, Gray 

JE, Hetherington AM (2011) Land plants acquired active stomatal 

control early in their evolutionary history. Curr. Biol. 21, 1030–1035. 

Ruyter-Spira C, Al-Babili S, van der Krol S, Bouwmeester H (2012) 

The biology of strigolactones. Trends Plant Sci. 12, 1360–1385. 

Ryals J, Weymann K, Lawton K, Friedrich L, Ellis D, Steiner H- 

Y, Johnson J, Delaney TP, Jesse T, Vos P, Uknes S (1997) 

The Arabidopsis NIM1 protein shows homology to the mammalian 

transcription factor inhibitor IκB. Plant Cell 9, 425–439. 

Sachs T (1981) The control of patterned differentiation of vascular 

tissues. Adv. Bot. Res. 9, 152–262. 

Sachs T (1991) Pattern Formation in Plants. Cambridge University 

Press, Cambridge UK. 

Salim E, Ullsten O (1999) Our forests our future. Report of the World 

Commission on Forests and Sustainable Development, Cambridge 

University Press, Cambridge UK. 

Salleo S, Lo Gullo MA, De Paoli D, Zippo M (1996) Xylem recovery 

from cavitation-induced embolism in young plants of Laurus nobilis: 

A possible mechanism. New Phytol. 132, 47–56. 

Salleo S, Trifilo P, Esposito S, Nardini A, LoGullo MA (2009) Starchto-sugar 

conversion in wood parenchyma of field-growing Laurus 

nobilis plants: A component of the signal pathway for embolism 

repair? Funct. Plant Biol. 36, 815–825. 

Salleo S, Trifilo P, LoGullo MA (2006) Phloem as a possible major 

determinant of rapid cavitation reversal in Laurus nobilis (laurel). 

Funct. Plant Biol. 33, 1063–1074. 

Salt DE, Prince RC, Baker AJM, Raskin I, Pickering IJ (1999) 

Zinc ligands in the metal hyperaccumulator Thlaspi caerulescens 

as determined using x-ray absorption spectroscopy. Environ. Sci. 

Technol. 33, 713–717. 


Sannigrahi P, Ragauskas AJ, Tuskan GA (2010) Poplar as a feedstock 

for biofuels: A review of compositional characteristics. Biofuels 

Bioprod. Bioref. 4, 209–226. 

Sasaki T, Chino M, Hayashi H, Fujiwara T (1998) Detection of several 

mRNA species in rice phloem sap. Plant Cell Physiol. 39, 895–897. 

Sawa S, Ito T, Shimura Y, Okada K (1999) Filamentous flower controls 

the formation and development of Arabidopsis inflorescences and 

floral meristems. Plant Cell 11, 69–86. 

Sawchuck T, Head P, Donner TJ, Scarpella E (2007) Time-lapse 

imaging of Arabidopsis leaf development shows dynamic patterns 

of procambium formation. New Phytol. 176, 560–571. 

Sawchuk MG, Donner TJ, Scarpella E (2008) Auxin transportdependent, 

stage-specific dynamics of leaf vein formation. Plant 

Signal. Behav. 3, 286–289. 

Scarpella E, Barkoulas M, Tsiantis M (2010) Control of leaf and 

vein development by auxin. Cold Spring Harb. Perspect. Biol. 2, 

a001511. 

Scarpella E, Francis P, Berleth T (2004) Stage-specific markers 

define early steps of procambium development in Arabidopsis 

leaves and correlate termination of vein formation with mesophyll 

differentiation. Development 131, 3445–3455. 

Scarpella E, Helariutta Y (2010) Vascular pattern formation in plants. 

Curr. Op. Dev. Biol. 91, 221–265. 

Scarpella E, Marcos D, Friml JÃ, Berleth T (2006) Control of leaf 

vascular patterning by polar auxin transport. Genes Dev. 20, 1015– 

1027. 

Scarpella E, Meijer AH (2004) Pattern formation in the vascular system 

of monocot and dicot plant species. New Phytol. 164, 209–242. 

Schaaf G, Schikora A, Haberle J, Vert G, Ludewig U, Briat J, Curie 

C, von Wirén N (2005) A putative function for the Arabidopsis 

Fe-phytosiderophore transporter homolog AtYSL2 in Fe and Zn 

homeostasis. Plant Cell Physiol. 46, 762–774. 

Schachtman DP, Goodger JQD (2008) Chemical root to shoot signaling 

under drought. Trends Plant Sci. 6, 281–287. 

Schaumlöffel D, Ouerdane L, Bouyssiere B, Łobiński R (2003) 

Speciation analysis of nickel in the latex of a hyperaccumulating 

tree Sebertia acuminata by HPLC and CZE with ICP MS and 

electrospray MS-MS detection. J. Anal. At. Spect. 18, 120–127. 

Schenk PM, Kazan K, Wilson I, Anderson JP, Richmond T, 

Somerville SC, Manners JM (2000) Coordinated plant defense 

responses in Arabidopsis revealed by microarray analysis. Proc. 

Natl. Acad. Sci. USA 97, 11655–11660. 

Scheres B (2010) Developmental biology: Roots respond to an inner 

calling. Nature 465, 299–300. 

Scheres B, Di Laurenzio L, Willemsen V, Hauser MT, Janmaat 

K, Weisbeek P, Benfey PN (1995) Mutations affecting the radial 

organisation of the Arabidopsis root display specific defects throughout 

the embryonic axis. Development 121, 53–62. 

Schlereth A, Moller B, Liu W, Kientz M, Flipse J, Rademacher EH, 

Schmid M, Jurgens G, Weijers D (2010) MONOPTEROS controls 

embryonic root initiation by regulating a mobile transcription factor. 

Nature 464, 913–916.


Schlesinger WH (1997) Biogeochemistry. Academic Press, San 

Diego. 

Schrader J, Baba K, May ST, Palme K, Bennett M, Bhalerao RP, 

Sandberg G (2003) Polar auxin transport in the wood-forming 

tissues of hybrid aspen is under simultaneous control of developmental 

and environmental signals. Proc. Natl. Acad. Sci. USA 100, 

10096–10101. 

Schreiber SG, Hacke UG, Hamann A, Thomas BR (2011) Genetic 

variation of hydraulic and wood anatomical traits in hybrid poplar 

and trembling aspen. New Phytol. 190, 150–160. 

Schuetz M, Berleth T, Mattsson J (2008) Multiple MONOPTEROSdependent 

pathways are involved in leaf initiation. Plant Physiol. 

148, 870–880. 

Schuler M, Lehmann M, Fink-Straube C, Bauer P (2010) The 

interaction of NAS genes and FRD3 in the long-distance transport of 

iron in Arabidopsis thaliana. 15th International Symposium on Iron 

Nutrition and Interactions in Plants. Budapest, Hungary. 

Schulz A (1992) Living sieve cells of conifers as visualized by confocal, 

laser-scanning fluorescence microscopy. Protoplasma 166, 153– 

164. 

Searle IR, Men AE, Laniya TS, Buzas DM, Iturbe-Ormaetxe I, Carroll 

BJ, Gresshoff PM (2003) Long-distance signaling in nodulation 

directed by a CLAVATA1-like receptor kinase. Science 299, 109– 

112. 

Secchi F, Zwieniecki MA (2010) Patterns of PIP gene expression in 

Populus trichocarpa during recovery from xylem embolism suggest 

a major role for the PIP1 aquaporin subfamily as moderators of the 

refilling process. Plant Cell Environ. 33, 1285–1297. 

Sevanto S, Holbrook NM, Ball MC (2012) Freeze/thaw-induced embolism: 

Probability of critical bubble formation depends on speed of 

ice formation. Plant Sci. 3, 1–12. 

Shah J, Tsui F, Klessig DF (1997) Characterization of a salicylic 

acid-insensitive mutant (sai1) ofArabidopsis thaliana identified in 

a selective screen utilizing the SA-inducible expression of the tms2 

gene. Mol. Plant-Microbe Interact. 10, 69–78. 

Shah J, Kachroo P, Nandi A, Klessig D (2001) A recessive mutation 

in the Arabidopsis ssi2 gene confers SA- and NPR1-independent 

expression of PR genes and resistance against bacterial and 

oomycete pathogens. Plant J. 25, 563–574. 

Shalit A, Rozman A, Goldshmidt A, Alvarez JP, Bowman JL, Eshed 

Y, Lifschitz E (2009) The flowering hormone florigen functions as 

a general systemic regulator of growth and termination. Proc. Natl. 

Acad. Sci. USA 106, 8392–8397. 

Shapiro AD, Zhang C (2001) The role of NDR1 in avirulence genedirected 

signaling and control of programmed cell death in Arabidopsis. 


Sharp RG, Davies WJ (2009) Variability among species in the apoplastic 

pH signalling response to drying soils. J. Exp. Bot. 60, 4363– 

4370. 

Shulaev V, Leon J, Raskin I (1995) Is salicylic-acid a translocated 

signal of systemic acquired-resistance in tobacco. Plant Cell 7, 

1691–1701. 

Shulaev V, Silverman P, Raskin I (1997) Airborne signaling by methyl 

salicylate in plant pathogen resistance. Nature 385, 718–721. 

Sieburth LE (1999) Auxin is required for leaf vein pattern in Arabidopsis. 


Sieburth LE, Lee DK (2010) BYPASS1: How a tiny mutant tells a big 

story about root-to-shoot signaling. J. Integr. Plant Biol. 52, 77–85. 

Sinclair SA, Sherson SM, Jarvis R, Camakaris J, Cobbett CS (2007) 

The use of the zinc-fluorophore, Zinpyr-1, in the study of zinc 

homeostasis in Arabidopsis roots. New Phytol. 174, 39–45. 

Slaymaker DH, Navarre DA, Clark D, Pozo OD, Martin GB, Klessig 

DF (2002) The tobacco salicylic acid-binding protein 3 (SABP3) 

is the chloroplast carbonic anhydrase, which exhibits antioxidant 

activity and plays a role in the hypersensitive defense response. 


Smith AM, Stitt M (2007) Coordination of carbon supply and plant 

growth. Plant Cell Environ. 30, 1126–1149. 

Smith JAC, Milburn JA (1980a) Osmoregulation, the control of 

phloem-sap composition in Ricinus communis. Planta 148, 28–34. 

Smith JAC, Milburn JA. (1980b) Phloem turgor, the regulation of 

sucrose loading in Ricinus communis. Planta 148, 42–48. 

Snow R (1935) Activation of cambial growth by pure hormones. New 

Phytol. 34, 347–360. 

Song JT, Koo YJ, Seo HK, Kim MC, CHoi YD, Kim JH (2008) 

Overexpression of AtSGT1, an Arabidopsis salicylic acid glucosyltransferase, 

leads to increased susceptibility to Pseudomonas 

syringae. Phytochemistry 69, 1128–1134. 

Song JT, Lu H, Greenberg JT (2004b) Divergent roles in Arabidopsis 

thaliana development and defense of two homologous genes, aberrant 

growth and death 2 and AGD2-LIKE DEFENSE RESPONSE 

PROTEIN 1, encoding novel aminotransferases. Plant Cell 16, 353– 

366. 

Song JT, Lu H, McDowell JM, Greenberg JT (2004a) A key role for 

ALD1 in activation of local and systemic defenses in Arabidopsis. 


Soyano T, Thitamadee S, Machida Y, Chua NH (2008) ASYMMETRIC 

LEAVES2-LIKE19/LATERAL ORGAN BOUNDARIES DOMAIN30 

and ASL20/LBD18 regulate tracheary element differentiation in 


Sperotto RA, Boff T, Duarte GL, Santos LS, Grusak MA, Fett 

JP (2010) Identification of putative target genes to manipulate Fe 

and Zn concentrations in rice grains. J. Plant Physiol. 167, 1500– 

1506. 

Sperotto RA, Ricachenevsky FK, Waldow VA, Fett JP (2012) Iron 

biofortification in rice: It’s a long way to the top. Plant Sci. 190, 

24–39. 

Sperry JS (1993) Winter xylem embolism and spring recovery in Betula 

cordifolia, Fagus grandifolia, Abies balsamea, and Picea rubens. 

In: Borghetti M, Grace J, Raschi A, eds. Water Transport in Plants 

under Climatic Stress. Cambridge University Press, Cambridge. pp. 

86–98. 

Sperry JS (2000) Hydraulic constraints on plant gas exchange. Ag. 

For. Meteorol. 2831, 1–11.

Sperry JS (2003) Evolution of water transport and xylem structure. Int. 

J. Plant Sci. 164, S115–S127. 

Sperry JS (2011) Hydraulics of vascular water transport In: P. W, ed. 

Mechanical integration of plant cells and plants. Springer-Verlag 

Berlin. pp. 303–327. 

Sperry JS, Christman MA, Smith DD (2012) Vulnerability curves by 

centrifugation: Is there an open vessel artifact, and are “r” shaped 

curves necessarily invalid? Plant Cell Environ. 35, 601–610. 

Sperry JS, Hacke UG, Oren R, Comstock JP (2002) Water deficits 

and hydraulic limits to leaf water supply. Plant Cell Environ. 25, 

251–263. 

Sperry JS, Holbrook NM, Zimmermann MH, Tyree MT (1987) Spring 

filling of xylem vessels in wild grapevine. Plant Physiol. 83, 414–417. 

Sperry JS, Tyree MT (1988) Mechanism of water stress-induced xylem 

embolism. Plant Physiol. 88, 581–587. 

Sperry JS, Tyree MT (1990) Water-stress-induced xylem embolism in 

three species of conifers. Plant Cell Environ. 13, 427–436. 

Spicer R, Groover A (2010) The evolution of development of the 

vascular cambium and secondary growth. New Phytol. 186, 577– 

592. 

Spoel SH, Dong X (2008) Making sense of hormone cross talk during 

plant immune response. Cell Host Microbe 3, 348–351. 

Spoel SH, Dong X (2012) How do plants achieve immunity? Defence 

without specialized immune cell. Nat. Rev. Immunol. 12, 

89–100. 

Spoel SH, Mou Z, Tada Y, Spivey NW, Genshik P, Dong X 

(2009) Proteasome-mediated turnover of the transcription coactivator 

NPR1 plays dual roles in regulating plant immunity. Cell 137, 

860–872. 

Stacey MG, Patel A, McClain WE, Mathieu M, Remley M, Rogers 

EE, Gassmann W, Blevins DG, Stacey G (2008) The Arabidopsis 

AtOPT3 protein functions in metal homeostasis and movement of 

iron to developing seeds. Plant Physiol. 146, 589–601. 

Stadler R, Wright KM, Lauterbach C, Amon G, Gahrtz M, Feuerstein 

A, Oparka KJ, Sauer N (2005) Expression of GFP-fusions in 

Arabidopsis companion cells reveals non-specific protein trafficking 

into sieve elements, identifies a novel post-phloem domain in roots. 


Staehelin C, Xie ZP, Illana A, Vierheilig H (2011) Long-distance 

transport of signals during symbiosis: Are nodule formation and 

mycorrhization autoregulated in a similar way? Plant Signal. Behav. 

6, 372–377. 

Stefanovic A, Ribot C, Rouached H, Wang Y, Chong J, Belbahri 

L, Delessert S, Poirier Y (2007) Members of the PHO1 gene 

family show limited functional redundancy in phosphate transfer 

to the shoot, and are regulated by phosphate deficiency via distinct 

pathways. Plant J. 50, 982–994. 

Stiller V, Lafitte HR, Sperry JS (2005) Embolized conduits of rice 

(Oryza sativa L.) refill despite negative xylem pressure. Am. J. Bot. 

92, 1970–1974. 

Stiller V, Sperry JS (2002) Cavitation fatigue and its reversal in intact 

sunflower plants. J. Exp. Bot. 53, 1155–1161. 


Strawn MA, Marr SK, Inoue K, Inada N, Zubieta C, Wildermuth MC 

(2007) Arabidopsis isochorismate synthase functional in pathogeninduced 

salicylate biosynthesis exhibits properties consistent with 

a role in diverse stress responses. J. Biol. Chem. 282, 5919– 

5933. 

Street N, Jansson S, Hvidsten T (2011) A systems biology model 

of the regulatory network in populus leaves reveals interacting 

regulators and conserved regulation. BMC Plant Biol. 11, 13. 

Suer S, Agusti J, Sanchez P, Schwarz M, Greb T (2011) WOX4 

imparts auxin responsiveness to cambium cells in Arabidopsis. 


Sussex IM (1954) Experiments on the cause of dorsiventrality in leaves. 

Nature 167, 651–652. 

Svistoonoff S, Creff A, Reymond M, Sigoillot-Claude C, Ricaud 

L, Blanchet A, Nussaume L, Desnos T (2007) Root tip contact 

with low-phosphate media reprograms plant root architecture. Nat. 

Genet. 39, 792–796. 

Sweetlove LJ, Kossmann J, Riesmeier JW, Riesmeier JW, 

Trethewey RN, Hill SA (1998) The control of source to sink carbon 

flux during tuber development in potato. Plant J. 15, 697–706. 

Tada Y, Spoel SH, Pajerowska-Muhktar K, Mou Z, Song J, Wang 

C, Zuo J, Dong, X (2008) Plant immunity requires conformational 

changes of NPR1 via S-nitrosylation and thoredoxins. Science 321, 

952–956. 

Takahashi H, Miller J, Nozaki Y, Takeda M, Shah J, Hase S, 

Ikegami M, Ehara Y, Dinesh-Kumar SP (2002) RCY1, anArabidopsis 

thaliana RPP8/HRT family resistance gene, conferring 

resistance to cucumber mosaic virus requires salicylic acid, ethylene 

and a novel signal transduction mechanism. Plant J. 32, 655– 

667. 

Takahashi M, Terada Y, Nakai I, Nakanishi H, Yoshimura E, Mori 

S, Nishizawa N (2003) Role of nicotianamine in the intracellular 

delivery of metals and plant reproductive development. Plant Cell 

15, 1263–1280. 

Takano J, Miwa K, Fujiwara T (2008) Boron transport mechanisms: 

Collaboration of channels and transporters. Trends Plant Sci. 13, 

451–457. 

Takano J, Noguchi K, Yasumori M, Kobayashi M, Gajdos Z, Miwa 

K, Hayashi H, Yoneyama T, Fujiwara T (2002) Arabidopsis boron 

transporter for xylem loading. Nature 420, 337–340. 

Takano J, Yamagami M, Noguchi K, Hayashi H, Fujiwara T (2001) 

Preferential translocation of boron to young leaves in Arabidopsis 

thaliana regulated by the BOR1 gene. Soil Sci. Plant Nutr. 47, 345– 

357. 

Takei K, Takahashi T, Sugiyama T, Yamaya T, Sakakibara H (2002) 

Multiple routes communicating nitrogen availability from roots to 

shoots: A signal transduction pathway mediated by cytokinin. 

J. Exp. Bot. 53, 971–977. 

Talke IN, Hanikenne M, Krämer U (2006) Zinc-dependent global 

transcriptional control, transcriptional deregulation, and higher gene 

copy number for genes in metal homeostasis of the hyperaccumulator 

Arabidopsis halleri. Plant Physiol. 142, 148–167.


Tamaki S, Matsuo S, Wong HL, Yokoi S, Shimamoto K (2007) 

Hd3a protein is a mobile flowering signal in rice. Science 316, 

1033–1036. 

Tan Q, Zhang L, Grant J, Cooper P, Tegeder M (2010) Altered 

phloem transport of S-methylmethionine affects plant metabolism, 

seed number in pea plants. Plant Physiol. 154, 1886–1896. 

Tanaka M, Wallace IS, Takano J, Roberts DM, Fujiwara T (2008) 

NIP6;1 is a boric acid channel for preferential transport of boron to 

growing shoot tissues in Arabidopsis. Plant Cell 20, 2860–2875. 

Taoka KI, Ham BK, Xoconostle-Cázares B, Rojas MR, Lucas WJ 

(2007) Reciprocal phosphorylation and glycosylation recognition 

motifs control NCAPPl interaction with pumpkin phloem proteins 

and their cell-to-cell movement. Plant Cell 19, 1866–1884. 

Teo G, Suzuki Y, Uratsu SL, Lamoinen B, Ormonde N, Hu WK, 

DeJong TM, Dandekar AM (2006) Silencing leaf sorbitol synthesis 

alters long-distance partitioning and apple fruit quality. Proc. Natl. 

Acad. Sci. USA 103, 18842–18847. 

Thibaud MC, Arrighi JF, Bayle V, Chiarenza S, Creff A, Bustos 

R, Paz-Ares J, Poirier Y, Nussaume L (2010) Dissection of local 

and systemic transcriptional responses to phosphate starvation in 


Thomas RJ (1972) Bordered pit aspiration in angiosperms. Wood Fiber 

3, 236–237. 

Thompson AJ, Mulholland BJ, Jackson AC, McKee JM, Hilton HW, 

Symonds RC, Sonneveld T, Burbidge A, Stevenson P, Taylor IB 

(2007) Regulation and manipulation of ABA biosynthesis in roots. 


Thompson MV (2006) Phloem: The long, the short of it. Trends Plant 

Sci. 11, 26–32. 

Thompson MV, Wolniak SM (2008) A plasma membrane-anchored 

fluorescent protein fusion illuminates sieve element plasma membranes 

in Arabidopsis, tobacco. Plant Physiol. 146, 1599–1610. 

To JP, Deruère J, Maxwell BB, Morris VF, Hutchison CE, Ferreira 

FJ, Schaller GE, Kieber JJ (2007) Cytokinin regulates type-A 

Arabidopsis Response Regulator activity and protein stability via 

two-component phosphorelay. Plant Cell 19, 3901–3914. 

To JP, Haberer G, Ferreira FJ, Deruère J, Mason MG, Schaller 

GE, Alonso JM, Ecker JR, Kieber JJ (2004) Type-A Arabidopsis 

response regulators are partially redundant negative regulators of 

cytokinin signaling. Plant Cell 16, 658–671. 

Tomatsu H, Takano J, Takahashi H, Watanabe-Takahashi A, Shibagaki 

N, Fujiwara T (2007) An Arabidopsis thaliana high-affinity 

molybdate transporter required for efficient uptake of molybdate 

from soil. Proc. Natl. Acad. Sci USA 104, 18807–18812. 

Trampczynska A, Küpper H, Meyer-Klaucke W, Schmidt H, 

Clemens S (2010) Nicotianamine forms complexes with Zn(ii) in 

vivo. Metallomics 2, 57–66. 

Truernit E, Bauby H, Dubreucq B, Grandjean O, Runions J, 

Barthélémy J, Palauqui JC (2008) High-resolution whole-mount 

imaging of three-dimensional tissue organization and gene expression 

enables the study of phloem development and structure in 


Truernit E, Bauby H, Belcram K, Barthélémy J, Palauqui JC (2012) 

OCTOPUS, a polarly localised membrane-associated protein, regulates 

phloem differentiation entry in Arabidopsis thaliana. Development 

139, 1306–1315. 

Truman W, Bennett MH, Turnbull CGN, Grant MR (2010) Arabidopsis 

auxin mutants are compromised in systemic acquired resistance 

and exhibit aberrant accumulation of various indolic compounds. 


Truman W, Bennett MH, Kubigsteltig I, Turnbull C, Grant M (2007) 

Arabidopsis systemic immunity uses conserved defense signaling 

pathways and is mediated by jasmonates. Proc. Natl. Acad. Sci. 

USA 104, 1075–1080. 

Tsukamoto T, Nakanishi H, Uchida H, Watanabe S, Matsuhashi 

S, Mori S, Nishizawa NK (2009) 52Fe translocation in barley as 

monitored by a positron-emitting tracer imaging system (PETIS): 

Evidence for the direct translocation of Fe from roots to young leaves 

via phloem. Plant Cell Physiol. 50, 48–57. 

Tudela D, Primo-Millo E (1992) 1-Aminocyclopropane-1-carboxylic 

acid transported from roots to shoots promotes leaf abscission in 

cleopatra mandarin (Citrus reshni Hort. ex Tan.) seedlings rehydrated 

after water stress. Plant Physiol. 100, 131–137. 

Tuominen H, Puech L, Fink S, Sundberg B (1997) A radial concentration 

gradient of indole-3-acetic acid is related to secondary xylem 

development in hybrid aspen. Plant Physiol. 115, 577–585. 

Turgeon R (2010a) The role of phloem loading reconsidered. Plant 

Physiol. 152, 1817–1823. 

Turgeon R (2010b) The puzzle of phloem pressure. Plant Physiol. 154, 

578–581. 

Turgeon R, Wolfe S (2009) Phloem transport: Cellular pathways, 

molecular trafficking. Annu. Rev. Plant Biol. 60, 207–221. 

Turnbull CGN, Booker JP, Leyser HMO (2002) Micrografting techniques 

for testing long-distance signalling in Arabidopsis. Plant J. 

32, 255–262. 

Turner S, Gallois P, Brown D (2007) Tracheary element differentiation. 

Annu. Rev. Plant Biol. 58, 407–433. 

Tuskan GA, Difazio S, Jansson S, Bohlmann J, Grigoriev I, Hellsten 

U, Putnam N, Ralph S, Rombauts S, Salamov A, Schein J, Sterck 

L, Aerts A, Bhalerao RR, Bhalerao RP, Blaudez D, Boerjan W, 

Brun A, Brunner A, Busov V, Campbell M, Carlson J, Chalot 

M, Chapman J, Chen GL, Cooper D, Coutinho PM, Couturier 

J, Covert S, Cronk Q, Cunningham R, Davis J, Degroeve 

S, Déjardin A, Depamphilis C, Detter J, Dirks B, Dubchak 

I, Duplessis S, Ehlting J, Ellis B, Gendler K, Goodstein D, 

Gribskov M, Grimwood J, Groover A, Gunter L, Hamberger B, 

Heinze B, Helariutta Y, Henrissat B, Holligan D, Holt R, Huang 

W, Islam-Faridi N, Jones S, Jones-Rhoades M, Jorgensen R, 

Joshi C, Kangasjärvi J, Karlsson J, Kelleher C, Kirkpatrick 

R, Kirst M, Kohler A, Kalluri U, Larimer F, Leebens-Mack J, 

Leplé JC, Locascio P, Lou Y, Lucas S, Martin F, Montanini B, 

Napoli C, Nelson DR, Nelson C, Nieminen K, Nilsson O, Pereda 

V, Peter G, Philippe R, Pilate G, Poliakov A, Razumovskaya 

J, Richardson P, Rinaldi C, Ritland K, Rouzé P, Ryaboy D,

Schmutz J, Schrader J, Segerman B, Shin H, Siddiqui A, Sterky 

F, Terry A, Tsai CJ, Uberbacher E, Unneberg P, Vahala J, Wall K, 

Wessler S, Yang G, Yin T, Douglas C, Marra M, Sandberg G, Van 

de Peer Y, Rokhsar D (2006) The genome of black cottonwood, 

Populus trichocarpa (Torr. & Gray). Science 313, 1596–1604. 

Twumasi P, Iakimova ET, Qian T, van Ieperen W, Schel JH, Emons 

AM, van Kooten O, Woltering EJ (2010) Caspase inhibitors 

affect the kinetics and dimensions of tracheary elements in xylogenic 

Zinnia (Zinnia elegans) cell cultures. BMC Plant Biol. 10, 

162. 

Tyree MT, Sperry JS (1988) Do woody plants operate near the point 

of catastrophic xylem dysfunction caused by dynamic water stress? 

Answers from a model. Plant Physiol. 88, 574–580. 

Tyree MT, Sperry JS (1989) Vulnerability of xylem to cavitation and 

embolism. Annu. Rev. Plant Physiol. Plant Mol. Biol. 40, 19–38. 

Tyree MT, Zimmermann MH (2002) Xylem Structure and the Ascent 

of Sap. Springer, Berlin. 

Uggla C, Moritz T, Sandberg G, Sundberg B (1996) Auxin as a 

positional signal in pattern formation in plants. Proc. Natl. Acad. 

Sci. USA 93, 9282–9286. 

Umehara M, Hanada A, Yoshida S, Akiyama K, Arite T, Takeda- 

Kamiya N, Magome H, Kamiya Y, Shirasu K, Yoneyama K, 

Kyozuka J, Yamaguchi S (2008) Inhibition of shoot branching by 

new terpenoid plant hormones. Nature 455, 195–200. 

Umehara M, Hanada A, Magome H, Takeda-Kamiya N, Yamaguchi 

S (2010) Contribution of strigolactones to the inhibition of tiller bud 

outgrowth under phosphate deficiency in rice. Plant Cell Physiol. 

51, 1118–1126. 

Vacchina V, Mari S, Czernic P, Marquès L, Pianelli K, Schaumlöffel 

D, Lebrun M, Łobiński R (2003) Speciation of nickel in a hyperaccumulating 

plant by high-performance liquid chromatographyinductively 

coupled plasma mass spectrometry and electrospray 

MS/MS assisted by cloning using yeast complementation. Anal. 

Chem. 75, 2740–2745. 

van Bel AJE (2003) The phloem, a miracle of ingenuity. Plant Cell 

Environ. 26, 125–149. 

van Bel AJE, Hafke JB (2005) Physicochemical determinants of 

phloem transport. In: Holbrook NM, Zwieniecki M, eds. Vascular 

Transport in Plants. Elsevier, Amsterdam. pp. 19–44. 

van de Mortel JE, Villanueva LA, Schat H, Kwekkeboom J, Coughlan 

S, Moerland PD, van Themaat EVL, Koornneef M, Aarts 

MGM (2006) Large expression differences in genes for iron and zinc 

homeostasis, stress response, and lignin biosynthesis distinguish 

roots of Arabidopsis thaliana and the related metal hyperaccumulator 

Thlaspi caerulescens. Plant Physiol. 142, 1127–1147. 

van der Biezen EA, Freddie CT, Kahn K, Parker JE, Jones 

JDG (2002) Arabidopsis RPP4 is a member of the RPP5 multigene 

family of TIR-NB-LRR genes and confers downy mildew 

resistance through multiple signalling components. Plant J. 29, 439– 

451. 

Van Doorn W, Beers E, Dangl J, Franklin-Tong V, Gallois P, Hara- 

Nishimura I, Jones A, Kawai-Yamada M, Lam E, Mundy J 


(2011a) Morphological classification of plant cell deaths. Cell Death 

Differ. 18, 1241–1246. 

Van Doorn WG, Hiemstra T, Fanourakis D (2011b) Hydrogel regulation 

of xylem flow: An alternative hypothesis. Plant Physiol. 157, 

1642–1649. 

Van Goor BJ, Wiersma D (1976) Chemical forms of manganese and 

zinc in phloem exudate. Physiol. Plant. 36, 213–216. 

Van Hoof NALM, Hassinen VH, Hakvoort HWJ, Ballintijn KF, Schat 

H, Verkleij JAC, Ernst WHO, Karenlampi SO, Tervahauta AI 

(2001) Enhanced copper tolerance in Silene vulgaris (Moench) 

Garcke populations from copper mines is associated with increased 

transcript levels of a 2b-type metallothionein gene. Plant Physiol. 

126, 1519–1526. 

Van Iepeeren W (2007) Ion-mediated changes of xylem hydraulic 

conductivity in planta: Fact or fiction? Trends Plant Sci. 12, 137– 

142. 

Van Iepeeren W, Van Gelder A (2006) Ion-mediated flow changes 

suppressed by minimal calcium presence in Chrysanthemum and 

Prunus laurocerasus. J. Exp. Bot. 57, 2743–2750. 

Van Norman JM, Frederick RL, Sieburth LE (2004) BYPASS1 negatively 

regulates a root-derived signal that controls plant architecture. 

Curr. Biol. 14, 1739–1746. 

Van Norman JM, Sieburth LE (2007) Dissecting the biosynthetic 

pathway for the bypass1 root-derived signal. Plant J. 49, 619–628. 

Van Wees SCM, De Swart EAM, Van Pelt JA, Van Loon LC, 

Pieterse CMJ (2000) Enhancement of induced disease resistance 

by simultaneous activation of salicylate- and jasmonate-dependent 

defense pathways in Arabidopsis thaliana. Proc. Natl. Acad. Sci. 

USA 97, 8711–8716. 

Vanneste S, Coppens F, Lee E, Donner TJ, Xie Z, Van Isterdael G, 

Dhondt S, De Winter F, De Rybel B, Vuylsteke M, De Veylder L, 

Friml J, Inze D, Grotewold E, Scarpella E, Sack F, Beemster 

GT, Beeckman T (2011) Developmental regulation of CYCA2s 

contributes to tissue-specific proliferation in Arabidopsis. EMBO J. 

30, 3430–3441. 

Vatén A, Dettmer J, Wu S, Stierhof YD, Miyashima S, Yadav SR, 

Roberts CJ, Campilho A, Bulone V, Lichtenberger R, Lehesranta 

S, Mähönen AP, Kim JY, Jokitalo E, Sauer N, Scheres B, 

Nakajima K, Carlsbecker A, Gallagher KL, Helariutta Y (2011) 

Callose biosynthesis regulates symplastic trafficking during root 

development. Dev. Cell 21, 1144–1155. 

Venugopal SC, Jeong RD, Zhu S, Chandra-Shekara AC, Navarre D, 

Kachroo A, Kachroo P (2009) ENHANCED DISEASE SUSCEP- 

TIBILITY 1 and salicylic acid act redundantly to regulate resistance 

gene expression and low oleate-induced defense signaling. PLoS 

Genetics 5, e1000545. 

Vera-Sirera F, Minguet EG, Singh SK, Ljung K, Tuominen HCL, 

Blázquez MA, Carbonell J (2010) Role of polyamines in plant 

vascular development. Plant Physiol. Biochem. 48, 534–539. 

Verbruggen N, Hermans C, Schat H (2009) Molecular mechanisms 

of metal hyperaccumulation in plants. New Phytol. 181, 759– 

776.


Vercammen D, van de Cotte B, De Jaeger G, Eeckhout D, Casteels 

P, Vandepoele K, Vandenberghe I, Van Beeumen J, InzéD,Van 

Breusegem F (2004) Type II metacaspases Atmc4 and Atmc9 of 

Arabidopsis thaliana cleave substrates after arginine and lysine. J. 

Biol. Chem. 279, 45329–45336. 

Verma DP, Hong Z (2001) Plant callose synthase complexes. Plant 

Mol. Biol. 47, 693–701. 

Vernooij B, Friedrich L, Morse A, Reist R, Kolditz-Jawhar R, Ward 

E, Uknes S, Kessmann H, Ryals J (1994) Salicylic acid is not 

the translocated signal responsible for inducing systemic acquired 

resistance but is required in signal transduction. Plant Cell 6, 959– 

965. 

Verret F, Gravot A, Auroy P, Leonhardt N, David P, Nussaume 

L, Vavasseur A, Richaud P (2004) Overexpression of AtHMA4 

enhances root-to-shoot translocation of zinc and cadmium and plant 

metal tolerance. FEBS Lett. 576, 306–312. 

Verret F, Gravot A, Auroy P, Preveral S, Forestier C, Vavasseur A, 

Richaud P (2005) Heavy metal transport by AtHMA4 involves the 

N-terminal degenerated metal binding domain and the C-terminal 

His11 stretch. FEBS Lett. 579, 1515–1522. 

Vierheilig H, Lerat S, PichéY(2003) Systemic inhibition of arbuscular 

mycorrhiza development by root exudates of cucumber plants 

colonized by Glomus mosseae. Mycorrhiza 13, 167–170. 

Vlot AC, Klessig DF, Park S-W (2008) Systemic acquired resistance: 

The elusive signal(s). Curr. Op. Plant Biol. 11, 436–442. 

Voesenek LACJ, Jackson MB, Toebes AHW, Huibers W, Vriezen 

WH, Colmer TD (2003) De-submergence-induced ethylene production 

in Rumex palustris: Regulation and ecophysiological significance. 


Voinnet O (2009) Origin, biogenesis, and activity of plant microRNAs. 

Cell 136, 669–687. 

Voitsekhovskaja OV, Koroleva OA, Batashev DR, Knop C, Tomos 

AD, Gamalei YV, Heldt HW, Lohaus G (2006) Phloem loading in 

two Scrophulariaceae species. What can drive symplastic flow via 

plasmodesmata? Plant Physiol. 140, 383–395. 

von Wirén N, Klair S, Bansal S, Briat J, Khodr H, Shioiri T, Leigh 

R, Hider R (1999) Nicotianamine chelates both Fe-III and Fe-II. 

Implications for metal transport in plants. Plant Physiol. 119, 1107– 

1114. 

Walley JW, Rowe HC, Xiao Y, Chehab EW, Kliebenstein DJ, Wagner 

D, Dehesh K (2008) The chromatin remodeler SPLAYED regulates 

specific stress signaling pathways. PLoS Pathol. 4, e1000237. 

Walter A, Silk WK, Schurr (2009) Environmental effects on spatial and 

temporal patterns of leaf and root growth. Annu. Rev. Plant Biol. 60, 

279–304. 

Walz C, Juenger M, Schad M, Kehr J (2002) Evidence for the presence 

and activity of a complete antioxidant defence system in mature 

sieve tubes. Plant J. 31, 189–197. 

Wang D, Amornsiripanitch N, Dong X (2006) A genomic approach to 

identify regulatory nodes in the transcriptional network of systemic 

acquired resistance in plants. PLoS Pathog. 2, e123. 

Wang S, Durrant WE, Song J, Spivey NW, Dong X (2010) Arabidopsis 

BRCA2 and RAD51 proteins are specifically involved in defense 

gene transcription during plant immune responses. Proc. Natl. Acad. 

Sci. USA 107, 22716–22721. 

Ward ER, Uknes SJ, Williams SC, Dincher SS, Wiederhold DL, 

Alexander DC, Ahl-Goy P, Metraux JP, Ryals JA (1991) Coordinate 

gene activity in response to agents that induce systemic 

acquired resistance. Plant Cell 3, 1085–1094. 

Wardlaw IF (1974) Temperature control of translocation. In: Bieleski 

RL, Fergupon R, Cresswell MM, eds. Mechanisms and Regulation 

of Plant Growth. Bull. Royal Soc., New Zealand. pp. 533–387. 

Wardlaw IF (1990) The control of carbon partitioning in plants. New 

Phytol. 116, 341–381. 

Wardlaw IF, Bagnall D (1981) Phloem transport and the regulation 

of growth of Sorghum bicolor (Moench) at low temperature. Plant 

Physiol. 68, 411–414. 

Wardlaw IF, Moncur L (1976) Source, sink and control of translocation 

in wheat. Planta 128, 93–100. 

Warmbrodt RD (1987) Solute concentrations in the phloem and apex 

of the root of Zea mays. Am. J. Bot. 74, 394–402. 

Waters BM, Chu H, DiDonato R, Roberts L, Eisley R, Lahner B, Salt 

D, Walker E (2006) Mutations in Arabidopsis yellow stripe-like1 and 

yellow stripe-like3 reveal their roles in metal ion homeostasis and 

loading of metal ions in seeds. Plant Physiol. 141, 1446–1458. 

Waters BM, Sankaran RP (2011) Moving micronutrients from the soil to 

the seeds: Genes and physiological processes from a biofortification 

perspective. Plant Sci. 180, 562–574. 

Waters BM, Uauy C, Dubcovsky J, Grusak MA (2009) Wheat 

(Triticum aestivum) NAM proteins regulate the translocation of iron, 

zinc, and nitrogen compounds from vegetative tissues to grain. 

J. Exp. Bot. 60, 4263–4274. 

Weichert N, Saalbach I, Weichert H, Kohl S, Erban A, Kopka J, 

Hause B, Varshney A, Sreenivasulu N, Strickert M, Kumlehn J, 

Weschke W, Weber H (2010) Increasing sucrose uptake capacity 

of wheat grains stimulates storage protein synthesis. Plant Physiol. 

152, 698–710. 

Weigel D, Jürgens G (2002) Stem cells that make stems. Nature 415, 

751–754. 

Weir IE, Maddumage R, Allan AC, Ferguson IB (2005) Flow cytometric 

analysis of tracheary element differentiation in Zinnia elegans 

cells. Cytometry A. 68, 81–91. 

Wenzel CL, Schuetz M, Yu Q, Mattsson J (2007) Dynamics of 

MONOPTEROS and PIN-FORMED1 expression during leaf vein 

pattern formation in Arabidopsis thaliana. Plant J. 49, 387–398. 

Werner T, Schmülling T (2009) Cytokinin action in plant development. 


West AG, Hultine KR, Sperry JS, Bush SE, Ehleringer JR (2008) 

Transpiration and hydraulic strategies in a pinyon-juniper woodland. 

Ecol. Appl. 18, 911–927. 

Wheeler JK, Sperry JS, Hacke UG, Hoang N (2005) Inter-vessel 

pitting and cavitation in woody Rosaceae and other vesselled plants: 

A basis for a safety vs. efficiency trade-off in xylem transport. Plant 

Cell Environ. 28, 800–812.

White MC, Decker AM, Chaney RL (1981) Metal complexation in 

xylem fluid: I. Chemical composition of tomato and soybean stem 

exudate. Plant Physiol. 67, 292–300. 

Wiersma D, Van Goor BJ (1979) Chemical forms of nickel and cobalt 

in phloem of Riccinus communis. Physiol. Plant. 45, 440–451. 

Wildermuth MC, Dewdney J, Wu G, Ausubel FM (2001) Isochorismate 

synthase is required to synthesize salicylic acid for plant 

defense. Nature 414, 562–571. 

Williams LE, Mills RF (2005) P1B-ATPases – An ancient family of 

transition metal pumps with diverse functions in plants. Trends Plant 

Sci. 10, 491–502. 

Williams LE, Pittman JK (2010) Dissecting pathways involved in 

manganese homeostasis and stress in higher plant cells. Plant Cell 

Monographs 17, 95–117. 

Windt CW, Vergeldt FJ, de Jager PA, van As H (2006) MRI of longdistance 

water transport: A comparison of the phloem and xylem 

flow characteristics and dynamics in poplar, castor bean, tomato 

and tobacco. Plant Cell Environ. 29, 1715–1729. 

Woffenden BJ, Freeman TB, Beers EP (1998) Proteasome inhibitors 

prevent tracheary element differentiation in zinnia mesophyll cell 

cultures. Plant Physiol. 118, 419–430. 

Wong CKE, Cobbett CS (2009) HMA P-type ATPases are the major 

mechanism for root-to-shoot Cd translocation in Arabidopsis 

thaliana. New Phytol. 181, 71–78. 

Wu Y, Zhang D, Chu JY, Boyle P, Wang Y, Brindle ID, De Luca 

V, Després C (2012) The Arabidopsis NPR1 protein is a receptor 

for the plant defense hormone salicylic acid. Cell Rep. 28, 639– 

647. 

Wycisk K, Kim EJ, Schroeder JI, Krämer U (2004) Enhancing the 

first enzymatic step in the histidine biosynthesis pathway increases 

the free histidine pool and nickel tolerance in Arabidopsis thaliana. 

FEBS Lett. 578, 128–134. 

Xia Y, Gao QM, Yu K, Lapchyk L, Navarre D, Hildebrand D, Kachroo 

A, Kachroo P (2009) An intact cuticle in distal tissues is essential 

for the induction of systemic acquired resistance in plants. Cell Host 

Microbe 5, 151–165. 

Xia Y, Yu K, Navarre D, Seebold K, Kachroo A, Kachroo, P (2010) 

The glabra1 mutation affects cuticle formation and plant responses 

to microbes. Plant Physiol. 154, 833–846. 

Xie B, Wang X, Zhu M, Zhang Z, Hong Z (2011) CalS7 encodes a 

callose synthase responsible for callose deposition in the phloem. 


Xoconostle-Cázares B, Xiang Y, Ruiz-Medrano R, Wang HL, 

Monzer J, Yoo BC, McFarland KC, Franceschi VR, Lucas WJ 

(1999) Plant paralog to viral movement protein that potentiates 

transport of mRNA into the phloem. Science 283, 94–98. 

Yaginuma H, Hirakawa Y, Kondo Y, Ohashi-Ito K, Fukuda H (2011) 

A novel function of TDIF-related peptides: Promotion of axillary bud 

formation. Plant Cell Physiol. 52, 1354–1364. 

Yamaguchi M, Kubo M, Fukuda H, Demura T (2008) Vascular-related 

NAC-DOMAIN7 is involved in the differentiation of all types of xylem 

vessels in Arabidopsis roots and shoots. Plant J. 55, 652–664. 


Yamaguchi M, Mitsuda N, Ohtani M, Ohme-Takagi M, Demura T 

(2011) VASCULAR-RELATED NAC-DOMAIN7 directly regulates 

the expression of a broad range of genes for xylem vessel formation. 


Yamaguchi M, Ohtani M, Mitsuda N, Kubo M, Ohme-Takagi M, 

Fukuda H, Demura T (2010) VND-INTERACTING2, a NAC domain 

transcription factor, negatively regulates xylem vessel formation in 


Yamamoto R, Demura T, Fukuda H (1997) Brassinosteroids induce 

entry into the final stage of tracheary element differentiation in 

cultured zinnia cells. Plant Cell Physiol. 38, 980–983. 

Yang SJ, Zhang YJ, Sun M, Goldstein G, Cao KF (2012) Recovery 

of diurnal depression of leaf hydraulic conductance in a subtropical 

woody bamboo species: Embolism refilling by nocturnal root 

pressure. Tree Physiol. 32, 414–422. 

Yoder JI, Gunathilake P, Wu B, Tomilova N, Tomilov AA (2009) 

Engineering host resistance against parasitic weeds with RNA 

interference. Pest Manag. Sci. 65, 460–466. 

Yokosho K, Yamaji N, Ma J (2010) Isolation and characterisation of 

two MATE genes in rye. Funct. Plant Biol. 37, 296–303. 

Yokosho K, Yamaji N, Ueno D, Mitani N, Ma J-F (2009) OSFRDL1 

is a citrate transporter required for efficient translocation of iron in 

rice. Plant Physiol. 149, 297–305. 

Yoo BC, Kragler F, Varkonyi-Gasic E, Haywood V, Archer-Evans 

S, Lee YM, Lough TJ, Lucas WJ (2004) A systemic small RNA 

signaling system in plants. Plant Cell 16, 1979–2000. 

Yoshida S, Iwamoto K, Demura T, Fukuda H (2009) Comprehensive 

analysis of the regulatory roles of auxin in early transdifferentiation 

into xylem cells. Plant Mol. Biol. 70, 457–469. 

Yoshimoto K, Noutoshi Y, Hayashi K, Shirasu K, Takahashi T, 

Motose H (2012) A chemical biology approach reveals an opposite 

action between thermospermine and auxin in xylem development in 

Arabidopsis thaliana. Plant Cell Physiol. 53, 635–645. 

Yu M, Hu CX, Wang YH (2002) Molybdenum efficiency in winter wheat 

cultivars as related to molybdenum uptake and distribution. Plant 

Soil 245, 287–293. 

Zeevaart JAD (1976) Physiology of flower formation. Annu. Rev. Plant 

Physiol. 27, 321–348. 

Zeevaart JAD (2006) Florigen coming of age after 70 years. Plant Cell 

18, 1783–1789. 

Zhang LY, Peng YB, Pelleschi-Travier S, Fan Y, Lu YF, Lu YM, Gao 

XP, Shen YY, Delrot S, Zhang DP (2004) Evidence for apoplasmic 

phloem unloading in developing apple fruit. Plant Physiol. 135, 1– 

13. 

Zhang WH, Zhou Y, Dibley KE, Tyerman SD, Furbank RT, Patrick 

JW (2007) Nutrient loading of developing seeds. Funct. Plant Biol. 

34, 314–331. 

Zhang XY, Wang XL, Wang XF, Xia GH, Pan QH, Fan RC, Wu 

FQ, Yu XC, Zhang DP (2006) A shift of phloem unloading from 

symplasmic to apoplasmic pathway is involved in developmental 

onset of ripening in grape berry. Plant Physiol. 142, 220– 

232.


Zhang Y, Cheng YT, Qu N, Zhao Q, Bi D, Li X (2006) Negative regulation 

of defense response in Arabidopsis by two NPR1 paralogs. 


ZhangY,FanW,KinkemaM,LiX,DongX(1999) Interaction of 

NPR1 with basic leucine zipper protein transcription factors that 

bind sequences required for salicylic acid induction of the PR-1 

gene. Proc. Natl. Acad. Sci. USA 96, 6523–6528. 

Zhang Y, Tessaro MJ, Lassner M, Li X (2003) Knockout analysis of 

Arabidopsis transcription factors TGA2, TGA5, and TGA6 reveals 

their redundant and essential roles in systemic acquired resistance. 


Zhao C, Johnson BJ, Kositsup B, Beers EP (2000) Exploiting 

secondary growth in Arabidopsis. Construction of xylem and bark 

cDNA libraries and cloning of three xylem endopeptidases. Plant 

Physiol. 123, 1185–1196. 

Zheng Q, Durben DJ, Wolf GH, Angell CA (1991) Liquids at large 

negative pressures: Water at the homogeneous nucleation limit. 

Science 254, 829–832. 

Zhong R, Demura T, Ye ZH (2006) SND1, a NAC domain transcription 

factor, is a key regulator of secondary wall synthesis in fibers of 


Zhong R, Lee C, Ye ZH (2010) Global analysis of direct targets of 

secondary wall NAC master switches in Arabidopsis. Mol. Plant 3, 

1087–1103. 

Zhong R, Lee C, Zhou J, McCarthy RL, Ye ZH (2008) A battery of 

transcription factors involved in the regulation of secondary cell wall 

biosynthesis in Arabidopsis. Plant Cell 20, 2763–2782. 

Zhong R, Richardson EA, Ye ZH (2007) The MYB46 transcription 

factor is a direct target of SND1 and regulates secondary wall 

biosynthesis in Arabidopsis. Plant Cell 19, 2776–2792. 

Zhong R, Ye ZH (2007) Regulation of cell wall biosynthesis. Curr. Opin. 


Zhong R, Ye ZH (1999) Ifl1, a gene regulating interfascicular fiber 

differentiation in Arabidopsis, encodes a homeodomain-leucine 

zipper protein. Plant Cell 11, 2139–2152. 

Zhong R, Ye ZH (2001) Alteration of auxin polar transport in the 

Arabidopsis ifl1 mutants. Plant Physiol. 126, 549–563. 

Zhou J, Lee C, Zhong R, Ye ZH (2009) MYB58 and MYB63 are 

transcriptional activators of the lignin biosynthetic pathway during 

secondary cell wall formation in Arabidopsis. Plant Cell 21, 248– 

266. 

Zhou JM, Trifa Y, Silva H, Pontier D, Lam E, Shah J, Klessig DF 

(2000) NPR1 differentially interacts with members of the TGA/OBF 

family of transcription factors that bind an element of the PR-1 gene 

required for induction by salicylic acid. Mol. Plant-Microbe Interact. 

13, 191–202. 

Zhou N, Tootle TL, Tsui F, Klessig DF, Glazebrook J (1998) PAD4 

functions upstream from salicylic acid to control defense responses 

in Arabidopsis. Plant Cell 10, 1021–1030. 

Zhou Y, Setz N, Niemietz C, Qu H, Offler CE, Tyerman SD, Patrick 

JW (2007) Aquaporins and unloading of phloem-imported water 

in coats of developing bean seeds. Plant Cell Environ. 30, 1566– 

1577. 

Zhu S, Jeong RD, Venugopal SC, Lapchyk L, Navarre D, Kachroo 

A, Kachroo P (2011) SAG101 forms a ternary complex with EDS1 

and is required for resistance signaling against turnip crinkle virus. 

PLoS Pathog. 7, e1002318. 

Zimmermann U, Schneider H, Wegner L, Haase A (2004) 

Water ascent in tall trees: Does evolution of land plants 

rely on a highly metastable state? New Phytol. 162, 575– 

615. 

Zwieniecki MA, Holbrook NM (2000) Bordered pit structure and vessel 

wall surface properties. Implications for embolism repair. Plant 

Physiol. 123, 1015–1020. 

Zwieniecki MA, Holbrook NM (2009) Confronting Maxwell’s demon: 

Biophysics of xylem embolism repair. Trends Plant Sci. 14, 530– 

534. 

Zwieniecki MA, Melcher PJ, Feild TS, Holbrook NM (2004) A potential 

role for xylem-phloem interactions in the hydraulic architecture 

of trees: Effects of phloem girdling on xylem hydraulic conductance. 

Tree Physiol. 24, 911–917. 

Zwieniecki MA, Melcher PJ, Holbrook NM (2001a) Hydraulic properties 

of individual xylem vessels of Fraxinus americana. J. Exp. Bot. 

52, 257–264. 

Zwieniecki MA, Melcher PJ, Holbrook NM (2001b) Hydrogel control 

of xylem hydraulic resistance in plants. Science 291, 1059– 

1062. 

(Co-Editor: Li-Jia Qu)

The Plant Vascular System: Evolution, Development and FunctionsF

You also want an ePaper? Increase the reach of your titles

Delete template?

Save as template?