Download Protophloem Differentiation in Early Arabidopsis

Plant Cell Physiol. 48(1): 97–109 (2007)
doi:10.1093/pcp/pcl045, available online at www.pcp.oxfordjournals.org
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Protophloem Differentiation in Early Arabidopsis thaliana Development
Hélène Bauby 1, Fanchon Divol 1, Elisabeth Truernit 1, Olivier Grandjean 2 and Jean-Christophe Palauqui 1, *
1
INRA, Centre de Versailles, Institut Jean-Pierre Bourgin, Laboratoire de Biologie Cellulaire, Route de St-Cyr, 78026 Versailles cedex,
France
2
INRA, Centre de Versailles, Institut Jean-Pierre Bourgin, Plateforme commune de cytologie, Route de St-Cyr, 78026 Versailles cedex,
France
Introduction
During Arabidopsis embryogenesis, procambial cells
undergo coordinated, asymmetric cell divisions, giving rise to
vascular precursor cells (protophloem and protoxylem precursors). After germination, these cells terminally differentiate
into specialized conducting cells, referred to as protophloem
and protoxylem cells. Few readily identifiable markers of the
onset of specification and differentiation are available,
hampering the molecular genetic analysis of protophloem
development. Confocal microscopy was used to investigate
the patterning and differentiation of phloem cells during early
plant development. Longitudinal divisions of phloem initials
allowed the identification of protophloem precursor cells and
adjacent metaphloem initials along the length of the plant.
During germination, protophloem differentiation was observed
at two independent locations, in the cotyledons and the
hypocotyl. In both locations, differentiation was concomitant
with cell elongation. We identified five gene-trap lines
(PD1–PD5) with marker gene expression in immature
protophloem elements. The spatio-temporal marker expression
pattern of the lines divides them into two groups. The early
specification markers PD4 and PD5 were expressed in
developing organs before procambium formation and then
became restricted to phloem initial cells. The protophloem
precursor markers PD1–PD3 were expressed in differentiating
protophloem cells at different stages of their development. All
markers were expressed transiently and iteratively during the
differentiation of protophloem in newly formed organs.
Flanking genes were identified for four out of five gene-trap
insertion lines. The possible function of these genes with respect
to phloem differentiation is discussed.
The development of a flexible and efficient vascular
system in higher plants was probably one of the most
significant improvements leading to the conquest of land by
terrestrial plants. The vascular system connects plant organs
separated by distances of up to several meters and provides
for the transport and allocation of water, nutrients and
signaling molecules. Vascular strands are composed of
specialized elongated cells: the xylem, phloem and meristematic vascular cells. These cell types form a continuous
network from roots to shoots (Esau 1969).
Clonal fate and positional information control plant
organization. During Arabidopsis embryogenesis, procambial cells are characteristically arranged in continuous
strands. They acquire their elongated shape through
coordinated cell divisions, oriented parallel to the axis of
the emerging strand (Berleth et al. 2000). At the end of
embryogenesis, asymmetric divisions occur at particular
locations in the procambium, generating several files of cells
that become precursors of phloem and xylem cells (Scheres
et al. 1995). In the mature embryo, the specialized
conducting cells are specified but not yet fully differentiated. In Arabidopsis ecotype Ler, terminal differentiation
of the first vascular elements occurs within 3 d of germination (Busse and Evert 1999).
The mechanisms by which certain procambial cells are
selected for phloem differentiation remain unclear.
However, the stability of the patterning process in most
species suggests that positional cues and growth conditions
are of critical importance. The molecular details of the
specification of phloem and xylem cells are particularly
intriguing, as the underlying mechanism must reconcile
the stringent requirement to keep all parts of the vascular
system interconnected with the constraints imposed by the
developmental pattern of the various organs.
Vascular tissue patterning models have been developed
from examining natural variability and mutant phenotypes
(Carlsbecker and Helariutta 2005). These mutants have
Keywords: Phloem — Differentiation — Development —
Vascular tissue — Arabidopsis.
Abbreviations: AB, aniline blue; DAG, days after
germination; GFP, green fluorescent protein; GPI, glycosylphosphatidylinositol; GUS, b-glucuronidase; HAG, hours after
germination; PD, phloem differentiation; PIP-4,5-kinase,
phosphatidylinositol phosphate-4,5-kinase; PP, protophloem
precursor.
*Corresponding author: E-mail, [email protected]; Fax, þ33-130-833099
97
98
Protophloem differentiation in early A. thaliana development
provided evidence for the involvement of sterols (Clouse
et al. 1996, Carland et al. 2002, Cano-Delgado et al. 2004),
small peptides (Casson et al. 2002), cytokinin (Mahonen
et al. 2000, Inoue et al. 2001) and auxin (Hardtke and
Berleth 1998, Deyholos et al. 2000, Hamann et al. 2002) in
vascular patterning. However, very few vascular pattern
mutants have so far been identified, possibly due to the
difficulties in visualizing vascular tissue.
Mutant descriptions and screens, and molecular genetic
analyses of cell patterning processes are dependent upon the
proper identification of cell types and differentiation stages.
The genetic dissection of cell identity acquisition in internal
tissues is possible if distinct cell states can be distinguished
by identifying cell type characteristics and/or genetic
markers of cell fate. If gene expression profiles associated
with specific differentiation states are identified, it should
be easy to mark individual cell states along vascular
differentiation pathways. However, most gene expression
profiles analyzed to date are associated with the terminal
differentiation stage of various vascular cell types, and
very few mark the early stages of phloem formation. Two
mutants specifically involved in phloem formation have
been identified. The wol mutant generates too few
procambial cells and is therefore affected in the specification
of phloem tissue in the axis (root/hypocotyl) (Mahonen
et al. 2000). In the apl mutant, protophloem determination
seems to be correct but sieve elements are modified. Instead,
apl mutants develop a hybrid cell type with a xylem-like
structure (Bonke et al. 2003).
To gain more insight into the pattern and the process
of phloem development, we have identified characteristic
cellular features of protophloem precursors that are
suitable for rapid and high-resolution confocal microscopic
analysis. We have assigned cytological and molecular
markers to specific stages of protophloem development
and followed their progression in the early stages of
Arabidopsis development. Protophloem cell files are organized in a longitudinal and radial pattern that shows organspecific features. Screening and identification of gene-trap
lines with marker expression in immature protophloem
elements have validated our results and provide new data
for the analysis of spatial and temporal specificities of
phloem formation.
Results
In order to characterize the organization and the
process of phloem development, we monitored early phloem
patterning and differentiation in Arabidopsis thaliana
(ecotype Ws). Studies of the vascular tissue in plants are
limited by difficulties obtaining access to internal plant
tissues by classical microscopic techniques. The use of an
aniline blue (AB) staining technique (Bougourd et al. 2000)
in combination with confocal laser scanning microscopy
enabled us to study the differentiation state of phloem cells
in mature embryos and young seedlings of A. thaliana.
Confocal microscopy allowed us to collect stacks of optical
longitudinal sections (Z-stacks). With this, phloem strands
could be followed directly along the longitudinal axis of the
plant without the need for transversal sectioning.
Radial and longitudinal pattern of protophloem elements in
the mature embryo
The mature embryo of A. thaliana is composed of two
cotyledons and a root/hypocotyl axis. The cotyledons show
a bilateral symmetry in contrast to the radial symmetry of
the axis. Thus, we defined two planes of organization in the
mature embryo: a frontal plane that splits the cotyledons
into two, and a perpendicular sagittal plane (Fig. 1A).
In the axis of 20 embryos tested, xylem and phloem were
composed of two poles organized in bilateral symmetry.
The radial organization of the poles could be inferred from
the position of the cotyledons. The future xylem poles were
located in the frontal plane and the phloem poles were
located on both sides of the xylem plate in the sagittal plane.
To study the differentiation state of phloem cells in
the mature embryo, high-resolution confocal imaging
(Bougourd et al. 2000) was utilized. Embryos were stained
with AB and cleared with chloral hydrate. Detailed
observation of one phloem pole in the median part of the
hypocotyl indicated that this pole consisted of at least two
different cell types (Fig. 1B, C; Supplementary Movie
SM 1). One cell type was elongated and bone-shaped, with
a thinner middle section and bulging apical and basal ends.
Based on cell shape, spatial relationship to other procambium and pericycle cells, and the known locations of
protophloem cells at later stages of development (Esau
1969, Busse and Evert 1999, Baum et al. 2002), we identified
cells of this type as immature protophloem cells or
protophloem precursors (PPs). The nucleus of these PPs
was brightly fluorescent in AB-stained samples, indicating
the undifferentiated state of these cells (Fig. 1C). A second
cell type was observed in close proximity to the PPs,
molding itself to the curves of these cells (Fig. 1C).
We identified these cells as metaphloem initials. These
cells continued to divide and gave rise to functional
metaphloem cells.
By studying stacks of longitudinal sections, we were
able to trace both cell types back along the root/hypocotyl
axis. An initial cell located above the quiescent center
undergoes a first longitudinal division, giving rise to the
phloem initial and to a more internal cell. This phloem
initial then undergoes a second longitudinal division,
3–5 cells above the quiescent center (Fig. 1D). A third
longitudinal division in the vicinity of the collar
then generates the PP and the metaphloem initial.
Protophloem differentiation in early A. thaliana development
A
99
B
Sagittal
plane
Frontal
plane
C
Xylem
Phloem
D
E
3
EN
P
C
EP
2
1
QC
F
G
Fig. 1 Organization of the vasculature in the mature Arabidopsis thaliana embryo (Ws). (A) Scheme of the symmetry planes (frontal
and sagittal) of a mature embryo of A. thaliana. (B–G) Longitudinal section of AB-stained mature embryos of wild-type A. thaliana,
showing phloem organization. (B) Hypocotyl. (C) Magnification of an immature protophloem cell (bone-shaped, white arrow) and its
adjacent metaphloem initial (white arrowhead). (D) Root and collar. The black arrowheads show the consecutive longitudinal
cell divisions that give rise to protophloem and metaphloem precursor cell files. (E) Upper hypocotyl and cotyledonary node
showing the branching of immature phloem cells at the transition zone. (F) Abaxial face of the cotyledon, showing a central immature
protophloem cell (white arrowhead) and two flanking derivative cells (black arrowheads). (G) Cell wall thickening of protophloem
precursors is observed in developing seedlings (black arrows). QC, quiescent center; P, pericycle; C, cortex; EN, endodermis;
EP, epidermis. Scale bars ¼ 10 mm.
100
Protophloem differentiation in early A. thaliana development
These longitudinal divisions are not strictly oriented in the
same plane. Therefore, cells adjacent to the PPs may be
either internal or external to the precursors (Fig. 3B, E).
Taken together, PPs initiate at the root/hypocotyl junction
and are not yet specified in the embryonic root.
The organization of the phloem and xylem changes
significantly in the cotyledonary node. The two strands of
vascular tissue join to form a collateral vascular bundle in
the cotyledons (Esau 1969). We visualized this structure by
following the file of PPs in the cotyledonary node in stacks
of longitudinal optical sections. Each protophloem strand
was seen to split into two strands, with one strand supplying
one cotyledon and the other strand supplying the other
cotyledon (Fig. 1E). Thus the two initial PPs strands in the
axis produce four strands, with two strands leading into
each cotyledon. Both strands of phloem were visible in
each cotyledon until the branching of the distal loop of
secondary veins. Beyond this branching point, only one
strand of protophloem cells was generally observed in the
midvein (Fig. 1G). This strand was located on the abaxial
side of the cotyledon and was accompanied by two
adjacent cells. The three cells were tightly linked, and
were probably generated by two longitudinal divisions
during late embryogenesis (Fig. 1F). The specification of
PPs was completed in the midvein and in the distal loops
forming the secondary veins. Usually, PPs in the proximal
loops were not yet completely specified.
Size distribution of PPs reflects the organ-dependent
differentiation stage
One of the characteristics of differentiating protophloem cells is their ability to elongate and to give rise to
a functional conducting unit. Thus, we hypothesized
that cell length would be a relevant parameter of the
differentiation state. Therefore, we determined the average
length of PPs/phloem initials in each organ of five mature
embryos (Fig. 2A). PP size repartition shows that the most
elongated cells are located in the cotyledons, with longer
cells in the midvein than in the distal loop. Shorter cells
were present in the hypocotyl and even shorter phloem
initials in the root. Along the root/hypocotyl axis, PP cell
sizes increased from the collar to the upper middle part
of the hypocotyl (Fig. 2B). Cells were shorter at the
cotyledonary node. The increase of phloem cell size
coincided with the acquisition of the characteristic bone
shape. No correlation was observed between size and
inverted position of protophloem cells. These results suggest
that cell length is a relevant parameter for the study of
protophloem differentiation.
Initiation and progression of vascular tissue differentiation
during early seedling development
As we showed that PPs are not in the same
differentiation state in the different organs of the mature
embryo, we determined the kinetics of their differentiation
in 20 synchronized seedlings at 24, 48 and 72 h after
germination (HAG). In particular, we focused on three
consecutive features of differentiation that we could identify
in AB staining: (i) cell elongation; (ii) cell wall thickening;
and (iii) loss of the nucleus coinciding with a bright
fluorescence of the cell content. In parallel, we followed
the progression of protoxylem differentiation (secondary
cell wall thickening of xylem cells). At 24 HAG, protophloem precursors in the cotyledons displayed cell wall
thickening all along the midvein and the distal loop.
No nucleus was visible in these cells. In the middle/upper
part of the hypocotyl, PPs were highly fluorescent (Fig. 3A,
B). The fluorescent cells also exhibited thickened cell walls
(Fig. 3A, C). In the lower part of the hypocotyl, PPs were
still nucleated, less elongated and did not exhibit fluorescence. No major modifications were detected in the
embryonic root. At the same time, metaphloem initials
were already elongated in the cotyledons. In the hypocotyl,
whilst protophloem cells elongated, metaphloem initials
divided transversally, rapidly doubling in number along the
axis (Fig. 3A). Annular cell wall thickening of tracheary
elements started at the proximal midvein at the adaxial
side of the cotyledon (Fig. 3D). No secondary cell wall
thickening of protoxylem was detected in the hypocotyl.
These observations were consistent with previous descriptions of differentiating protoxylem (Dharmawardhana
et al. 1992; Pyo et al. 2004) (see also Supplementary
Movie SM2).
At 48 HAG, protophloem cell contents in the
hypocotyl and the cotyledons were no longer fluorescent
(Fig. 3E). In the root, meristem activity increased and
procambial cells and phloem initials divided basipetally
(data not shown). Phloem initials gave rise to PPs that
started to differentiate. Thus, protophloem differentiation
now progressed basipetally from the root meristem.
In parallel, secondary cell wall thickening of protoxylem
was seen in the middle/upper part of the hypocotyl.
At 72 HAG, differentiated protophloem cells appeared
completely dark in most parts of the seedling (Fig. 3F).
In the root, protophloem still differentiated basipetally,
as demonstrated by progressive cell wall thickening and cell
content darkening. Xylem cells were differentiating distally
from the phloem cells (Fig. 3G).
Taken together, these results suggest that protovascular elements are specified and begin to differentiate earlier
in the cotyledons than in the axis. Protophloem and
protoxylem differentiation was first observed in the midvein
and then extended to the distal loops and the cotyledonary
node (Fig. 6A1, A2). A second independent locus of
differentiation was observed in the upper part of the
hypocotyl. From there, protophloem differentiation
progressed towards the root (Fig. 6A2–A4). Later, a third
Protophloem differentiation in early A. thaliana development
A
101
50,00
Average size of phloem cells (µm)
45,00
40,00
35,00
30,00
25,00
20,00
15,00
10,00
5,00
0,00
# phloem initial
# phloem root
cell
# phloem
hypocotyl cell
# phloem
midvein cell
#phloem distal
loop cell
Organ
Average size of protophloem precursors (µm)
B
40
35
30
25
20
15
10
5
0
Quiescent 1
center
2
3
4
5
6
7
8
9
10 11
Cell number
12
13 14 15
16 17
18 19 Upper
hypocotyl
Fig. 2 Characteristics of phloem cells in mature embryos of Arabidopsis thaliana (Ws). (A) Average length of phloem cells in various
organs. Phloem initial: cells from the quiescent center until the second longitudinal division (n ¼ 10). Phloem root cell: cells from the
quiescent center to the collar (n ¼ 10). Phloem hypocotyl cell: cells from the collar to the cotyledonary node (n ¼ 20). Phloem midvein cell:
cells in the midvein of the cotyledons (n ¼ 20). Phloem distal loop cells: phloem cells in the distal loop of the secondary veins (n ¼ 20).
(B) Average size of phloem cells in the root/hypocotyl axis of mature embryos, from the initial located above the quiescent center (no. 1) to
the 19th cell located near the cotyledonary node. Error bars show standard deviation of the means.
starting point of differentiation was observed coming from
the root meristem (Fig. 6A5).
Identification of phloem markers by gene-trap screening
In order to identify reporter gene expression
profiles suitable for the study of phloem development
and its genetic determinism, we screened the Versailles
collection of Arabidopsis gene-trap mutants for plant
lines expressing the uidA reporter gene in immature
vascular tissues (Bechtold et al. 1993). Lines in which
the gene-trap construct is located inside a transcriptional
unit can generate chimeric proteins consisting of a
partial protein product of the transcriptional unit fused
to the b-glucuronidase (GUS) protein. Of 10,000 independent transformants, 100 transgenic lines displayed
reporter gene expression in the vascular tissues during
vegetative development 7 d after germination (DAG).
102
Protophloem differentiation in early A. thaliana development
Fig. 3 Confocal sections of AB-stained seedlings of wild-type Arabidopsis thaliana (Ws). (A) Hypocotyl of a 24-h-old seedling. Boneshaped protophloem precursors can be clearly observed. Black arrowheads show the corresponding transversal division of metaphloem
initials. (B) Hypocotyl of a 24-h-old seedling. The white arrow shows the classic orientation of a protophloem precursor and the white
arrowhead shows a protophloem precursor in inverted orientation. (C) Longitudinal section through the hypocotyl of a 36-h-old seedling.
White arrowheads point towards cell wall thickening of protophloem cells. (D) Adaxial section through the cotyledon midvein of a
36-h-old seedling showing the cell wall thickening of the first differentiating protoxylem element. (E) Hypocotyl of a 48-h-old seedling.
White arrows indicate protophloem cells with thickened cell wall and dark cell content. The black arrowhead shows adjacent cells in
inverted orientation. (F, G) Two different Z-stacks through a root of a 72-h-old seedling, showing that protophloem cells are differentiated
earlier than protoxylem cells. (F) A row of differentiating protophloem cells shows dark staining (black arrowhead). (G) Section showing
differentiating xylem cells with highly fluorescent secondary cell wall thickenings (white arrowhead). Scale bars ¼ 10 mm.
Phloem-specific expression was confirmed by analyzing
the GUS staining pattern in cross-sections of
inflorescence stems (Supplementary Fig. S1). We then
screened these lines for GUS activity between 0 and
72 HAG. We isolated five marker lines that showed
GUS expression in phloem tissue during early
differentiation (annotated PD1–PD5, for phloem
differentiation).
Protophloem differentiation in early A. thaliana development
103
Fig. 4 Early GUS expression patterns of the PD markers. (A) Mature embryo PD2 marker expression in dark field showing the branching
of phloem cell files in the cotyledonary node and the absence of expression in the lower part of the hypocotyl (white arrows). (B) Confocal
image of the upper part of the hypocotyl of an AB- and GUS-stained mature embryo with PD2 marker expression showing protophloem
precursor specificity. (C) PD3 marker expression in a mature embryo. PD3 GUS expression is also present in the epidermis (black arrows).
(D) PD4 marker expression in a mature embryo (dark field). PD4 GUS expression is restricted to the hydathodes and cotyledon veins, and
to two poles at the cotyledonary node in the sagittal plane (white arrows). (E) Confocal section of an AB- and GUS-stained mature embryo
with PD4 marker expression pattern showing protophloem precursor specificity. (F, G) PD5 marker expression in mature embryos. (G)
Magnification of the phloem poles in the hypocotyl, showing the expression of the PD5 marker in protophloem precursors and
metaphloem initials. (H) Confocal section of an AB- and GUS-stained embryonic cotyledon with PD5 marker expression in protophloem
precursors and metaphloem initials. (I, J, K, L) PD1 marker, 24- (I, J) and 36-h- (K) old seedlings. (L) Section of an AB- and GUS-stained
36-h-old embryo, PD1 marker expression pattern showing the protophloem precursor specificity. The PD1 marker is expressed only 24–
36 h after germination. (M, N) Primary root expression pattern of a 5 DAG seedling of PD5 and PD2 marker lines. Note that PD2 is also
expressed in the columella. Scale bars ¼ 100 mm (A, C, D, F, J, K), 10 mm (B, H), 20 mm (E, G, I).
Phloem specificity and patterning of PD markers during early
development
We determined the expression of each PD marker
during early phloem formation in 100 embryos/seedlings.
All PD marker lines, with the exception of PD1, displayed
GUS expression in the mature embryo. In PD1–PD4, the
GUS-expressing cells were identified as PPs on the basis of
their characteristic bone shape (Fig. 4B, E, H, I, L). In PD5,
reporter gene expression was detected in both PPs and
metaphloem initials (Fig. 4F–H).
While PD5 marker expression was seen throughout
the protophloem of the mature embryo (Fig. 4F), PD2
and PD3 marker expression was restricted to the midveins
of the cotyledon, the cotyledonary node and the
upper part of the hypocotyl (Fig. 4A, C). PD4 marker
expression was also restricted to the upper/middle part
of the embryo. It was detected in the hydathodes and
in the distal protophloem cells of the cotyledons (Fig. 4D).
Its expression was also present independently in two
internal zones at both sides of the meristem and was
104
Protophloem differentiation in early A. thaliana development
precisely located in the sagittal plane of the cotyledonary
node (Fig. 4D).
PD1 GUS expression was first observed between
24 and 36 HAG, in the region close to the cotyledon
midvein and/or in the vascular strand of the upper part of
the hypocotyl (Fig. 4J). It was not possible to discriminate
statistically whether the first expression event was in the
hypocotyl or in the cotyledon (data not shown).
Between 36 and 48 HAG, PD1, PD2, PD3 and PD4
marker expression progressed basipetally from the hypocotyl to the hypocotyl–root junction as shown for PD1
(Fig. 4J, K), whereas PD5 showed continuous marker
expression in the axis. Further progression of PD marker
expression was observed from the hypocotyl to the root.
As the seedlings developed, GUS expression in mature
protophloem cells gradually disappeared in all marker lines.
At 4 DAG, PD markers showed similar expression patterns
in the primary root. GUS expression initiated above the
root division zone in two symmetrical strands that could be
identified as protophloem strands. GUS activity gradually
disappeared towards the mature root (Fig. 4M). PD2 and
PD3 markers were also expressed in the differentiating root
cap (Fig. 4N) and root epidermis (data not shown).
Our data show that all PD markers are expressed in the
protophloem, but their spatial and temporal expression
patterns make them different and reflect the complexity
of phloem formation. Most of the PD marker expression
patterns progressed from the hypocotyl to the root,
supporting our previous observation in AB-stained seedlings. Furthermore, PD1 and PD3 marker expression
patterns indicated the presence of two independent foci of
protophloem differentiation during early seedling development: one coming from the midvein of the cotyledons and
another one from the upper part of the hypocotyl.
Progression of PD marker expression during shoot
development
In order to follow the chronology of PD marker
expression during the formation of a new organ, we took
advantage of the accurate description of vascular patterning
during Arabidopsis leaf development (Scarpella et al. 2004).
We followed the initiation and the progression of PD
marker expression during leaf primordia development
from 3 to 7 DAG. At 3 DAG, no vascular precursors are
present in the primordia, whereas at 7 DAG, differentiating
protoxylem and protophloem cells are present in the
midvein and in the first distal loop.
We identified and cloned the promoters driving GUS
expression in the PD1, PD3, PD4 and PD5 lines upstream
of the GFPER marker gene (see Materials and Methods).
Each construct was used to transform A. thaliana (ecotype
Ws) by floral dipping. Independent transformants for
each construct were obtained and homozygous plants
containing the construct pPDx::GFP were selected for
further investigation (PDx for PD1, PD3, PD4 and PD5).
For technical reasons, we were not able to identify the PD2
locus. pPDx::GFP expression in the transformants reflected
marker expression in the PDx lines (not shown).
pPD1::GFP expression and pPD3::GFP expression
was detected in the proximal protophloem of leaf primordia
at 4 DAG (Fig. 5A, E). It proceeded to the distal
protophloem (Fig. 5B, F) at 5 DAG and into the distal
loops at 6 DAG (Fig. 5C, H). In addition, pPD3::GFP
expression was found in the epidermis of leaf primordia
(Fig. 5D, G).
At 3 DAG, pPD4::GFP expression was detected in the
tip of leaf primordia (Fig. 5I). It was also seen in the future
midvein and started to invade the distal loops (Fig. 5I).
At 3.5 DAG, it invaded the proximal loops (Fig. 5J) with
a high GFP expression in the hydathode of the leaf tip.
At 4 DAG, a strong GFP expression appeared along the
leaf margins and in the secondary distal loops (Fig. 5K).
At 7 DAG, GFP expression was strong in leaf lobes and
defined the pre-pattern of the vascular tissue in the leaf.
Interestingly, GFP expression in the differentiated midvein
was restricted to the protophloem (Fig. 5L, M), and in the
margin of the leaf it was restricted to large isodiametric
cells (Fig. 5N).
pPD5::GFP expression was detected in the future
midvein axis in large isodiametric cells at 2.5 DAG
(Fig. 5O). The progression of this expression pattern
during development was thus very similar to the
pPD4::GFP expression pattern. At 7 DAG, GFP expression
was strong in the distal part of the leaf margin. In lower
order veins, expression was seen in large isodiametric cells
that were not yet specified as procambial cells (Fig. 5P, Q).
In the midvein of the same leaf, pPD5::GFP expression
was also restricted to the protophloem (Fig. 5R, S).
These results indicate that PD1 and PD3 markers are
expressed sensu stricto during protophloem differentiation.
In contrast, PD4 and PD5 reporter gene expression was
detected in the location of midveins and higher order
veins before procambium differentiation, and thus defined
the pre-patterning of the future veins. At later stages
of differentiation, the expression of these markers was
restricted to the phloem.
Genetic and molecular analyses of the gene-trap lines
For four out of five gene-trap lines, the GUS reporter
gene and resistance to kanamycin were co-segregating as
a typical single Mendelian trait. No differences were found
in the reporter gene expression analyses in segregating
populations of hemizygous plants, suggesting that patterns
were stable even in mutant backgrounds. Genomic regions
flanking the gene-trap insertions were identified. Detailed
loci analyses showed that in most cases the GUS gene
Protophloem differentiation in early A. thaliana development
A
PD1
I
4 DAG
5 DAG
B
PD1
3 DAG
6 DAG
C
PD1
J
3,5 DAG
D 3 DAG E
PD3
4 DAG
K
4 DAG
PD3
L
t
F
PD3
7 DAG
M
5 DAG
105
G
5 DAG
PD3
7 DAG
H
6 DAG
PD3
N
7 DAG
h
PD4
O
PD5
PD4
2,5 DAG
PD4
P
PD5
7 DAG
PD4
Q
PD5
PD4
7 DAG
R
PD5
PD4
7 DAG
S
7 DAG
PD5
Fig. 5 PD marker promoter GFP expression during leaf procambium and protophloem development. Marker identity and primordia age
(DAG) are indicated in the pictures. Images are from the abaxial face of young primordia/leaves. In the vein, expression of pPD4::GFP and
pPD5::GFP precedes the expression of pPD1::GFP and pPD3::GFP. Note epidermal pPD4::GFP expression at the primordium tip (t) and in
hydathodes (h) (L detailed in M), pPD5::GFP hydathode expression (P detailed in Q) and epidermal pPD3::GFP expression (D, E, F, G, H).
Note that expression of pPD4::GFP and pPD5::GFP in the central vein is becoming gradually restricted to protophloem cells (L detailed in
M, R detailed in S). Scale bars ¼ 50 mm.
was inserted close to the 50 end of a gene in the correct
orientation with respect to the transcriptional unit
(Supplementary Fig. S2). Two genes were annotated as
coding for putative membrane-bound proteins: first, a small
glycosylphosphatidylinositol
(GPI)-anchored
protein
(PD1) that could act as a potential signaling protein;
and secondly, a putative phosphatidylinositol phosphate
(PIP)-4,5-kinase (PD3) involved in intracellular signaling.
Two genes (PD4 and PD5) were annotated as proteins of
unknown function that are specific to multicellular plants.
The PD5 protein is a glycine-rich protein and the PD5 line
displays a short-root phenotype. The PD4 protein has
already been identified as corresponding to the BREVIS
RADIX locus in the Uk-1 ecotype and the protein was
described as a transcriptional activator (Mouchel et al.
2004). In Uk-1, the primary root length is reduced and the
root system is more branched. This phenotype was not
observed in PD4 lines, which suggests that PD4 is either
still functional in this line or that the Ws ecotype
could compensate for the phenotype. Lines PD1–PD3
had a wild-type phenotype. These data are summarized in
Supplementary Fig. S3.
Discussion
In this study, we provide a framework for the analysis
of vascular development in A. thaliana. Utilizing a
combination of confocal microscopy and new genetic
markers, we were able to visualize phloem precursor cells
at a high resolution. This has allowed studies of the
behavior of these cells during development without the need
for physical sectioning of plant tissues (Bougourd et al.
2000). Furthermore, the method makes it possible to
analyze the structure and patterning of vascular tissue
early in plant development and should provide a good tool
for further characterization of vascular mutants.
We have used the term ‘specification’ to define the
stage at which a cell adopts a predictable phenotype.
For example, some procambial cells are destined to become
phloem precursor cells. The differentiation of a protophloem cell starts with the acquisition of its final cell shape.
This is achieved by modification of the cell wall, which
ultimately leads to the formation of sieve plates. Functional
phloem tissue is formed by a continuous, rapid and
transient process, and this process may also be influenced
106
Protophloem differentiation in early A. thaliana development
by pre-existing mature phloem structures. These characteristics may hamper studies of phloem differentiation. These
problems were overcome by studying phloem specification
and differentiation at the very beginning of phloem
development. This approach can be used to study vascular
patterning in detail and to identify the key steps of
protophloem differentiation.
A
Provascular cells
Mature embryo (1)
24 HAG (2)
Protophloem precursors
Mature protophloem
36 HAG (3)
48 HAG (4)
72 HAG (5)
B
Pattern of phloem development
We have described the expression patterns of five
new molecular markers specific to several steps of phloem
differentiation. Based on our analyses, we were able to
determine the chronological order of expression of these
genes (Fig. 6B, and Supplementary Fig. S3). GUS expression in the PD4 and PD5 lines marks early provascular
patterns and is later restricted to the future phloem strand.
PD1, PD2 and PD3 markers are specific to protophloem
cells throughout plant development (Fig. 6B).
Only a few early phloem markers have been described
so far: the APL gene is expressed from the torpedo stage
onwards at the site of PP specification (Bonke et al. 2003).
It was also found to be expressed in metaphloem precursors
and companion cells. The AtHB8 gene was reported to be
expressed in the pre-procambial stage (Scarpella et al. 2004)
as early as the PD4 and PD5 markers, but its expression
was restricted to the procambium (Baima et al. 1995).
Conversely, the GT5211 reporter line shows expression in
the procambium and is progressively restricted to xylem
initials or precursors (Scarpella et al. 2004). Furthermore,
expression of the auxin efflux facilitator gene PIN1
coincides with the probable sites at which pre-procambial
cells are being specified. In addition, DR5rev::GFP expression, that reflects sites of auxin accumulation, indicates
the sites of vein formation (Scarpella et al. 2006). PD4
and PD5 markers display expression domains similar to
DR5rev::GFP and PIN expression patterns, respectively.
Thus, it is tempting to speculate that expression of the
respective PD4 and PD5 genes might be regulated by auxin
concentrations, or that the genes play a role in polar auxin
transport and its effects on phloem formation.
To explain the mechanism that controls the radial
pattern of the primary vascular tissues in the root, Aloni
et al. (2006) proposed that the primary xylem and phloem
strands are induced by alternating streams of high vs. low
IAA concentrations. Consistent with this hypothesis, there
is increasing evidence that the auxin concentration could act
as the driving force for the specification of vascular poles:
(i) the DR5 reporter gene is expressed in root protoxylem
cells; (ii) high auxin concentrations induce xylem differentiation (Aloni et al. 2003); and (iii) low levels of auxin are
needed for phloem differentiation (Aloni 2001). We show in
this study that xylem and phloem initials are specified at
two opposite locations in the mature embryo of A. thaliana.
PD4
Provascular cells
PD5
Procambium cells
Specification of xylem cells
Patterning
and
specification
Specification of phloem cells
Phloem initials
Protophloem
differentiation
Protophloem precursors
Metaphloem initials
PD3
PD2
PD1
Metaphloem
specification
Mature protophloem
CC
MST
Metaphloem
differentiation
Fig. 6 A model for early phloem development in Arabidopsis
thaliana. (A) Schematic diagram of phloem formation in mature
embryos and in seedlings at 24, 36, 48 and 72 HAG. In the mature
embryo, protophloem is differentiating in cotyledons and in the
cotyledonary node (red). A basipetal gradient of differentiation is
set up and protophloem is mature around 48 HAG (light blue).
At 72 HAG, root meristem is fully functional and gives rise to
new protophloem precursors (red). (B) Stages of vascular tissue
development, showing the appearance of PD marker expression.
Xylem is localized in the frontal plane and phloem in the
sagittal plane (Fig. 1A). Where does this symmetry come
from? We suggest that the acquisition of bilateral symmetry
at the heart stage of embryo development and consequently
the strong accumulation of auxin in the frontal plane
determines the position of xylem and phloem poles.
Consistent with this hypothesis, in leaves, xylem can
differentiate in the absence of phloem at the freely ending
veinlets (Horner et al. 1994) and hydathodes (Aloni et al.
2003), sites of presumably high auxin concentrations.
Protophloem specification and differentiation occur gradually
from the cotyledon to the root
Our data show that during embryogenesis phloem
differentiation in the cotyledons occurs earlier than in the
rest of the plant: (i) bone-shaped protophloem precursors
are longer in cotyledons than in the other organs;
(ii) expression of some PD markers is initially restricted to
the upper part of the plant; and (iii) metaphloem precursors
are already specified in cotyledons but not in the hypocotyl.
This suggests that protophloem development in cotyledons
is already underway before germination.
Protophloem differentiation in early A. thaliana development
The existence of a gradual differentiation process
was further confirmed during germination. The observed
pattern of transient fluorescence in protophloem cells,
the cell wall thickening and the progression of reporter
gene expression for most PD markers are consistent with
a time gradient of differentiation from the cotyledons
down to the root.
According to the PD1 marker expression pattern,
protophloem is initiated at distinct loci after germination
and then progresses almost simultaneously along the
cotyledons, hypocotyl and root. In the root, new protophloem elements are added acropetally from the meristem
onto the previously differentiated elements in the same
cell file, and consequently a continuous, uninterrupted
protophloem is established in the root (Fig. 6A1–A5). The
progression of expression of the PD1 marker is reminiscent
of the previously described ZCP4 marker of xylem
differentiation (Pyo et al. 2004), suggesting that initiation
and progression of phloem and xylem differentiation are
tightly linked. Consistent with our results is the previous
description of polar differentiation of the first protophloem
and protoxylem elements in Arabidopsis (Busse and
Evert 1999). However, this previous anatomical description
suggests that protophloem is differentiating almost simultaneously in cotyledon and hypocotyl. In the present study,
we demonstrate that protophloem is initiated discontinuously and then differentiation progresses simultaneously.
This new finding might be due to the use of a suitable
molecular marker (PD1) and the ability to scan the whole
phloem using confocal microscopy, with which we were able
to easily follow the early stages of phloem differentiation
in detail.
Identification of potential genes involved in specification and
differentiation of protophloem
To date, few genes have been identified as being
involved in phloem specification and/or differentiation
(APL and WOL). In our study, we identified four new
genes that are expressed during phloem formation.
Molecular analysis of the marker genes revealed that they
are plant specific and belong to multigene families.
The PD1 marker gene expression is localized in PPs,
indicating a proclivity to protophloem differentiation. The
PD1 gene encodes a small putative GPI-anchored protein of
7 kDa that belongs to a family of plant-specific proteins.
Analogous small GPI-anchored proteins in mammals,
such as CD24, are located on the cell surface and are
signal-transducing molecules. They communicate external
signals to the cell and function to control cell proliferation
or differentiation (Kay et al. 1991).
PD3 encodes a putative PIP-4,5-kinase annotated as
AtPIPK7 that belongs to a family of 10 genes (MuellerRoeber and Pical 2002). This class of proteins has been
107
shown to be involved in numerous signal transduction
pathways in eukaryotes, and one of them has been localized
to the procambium and may play a role in cell proliferation
(Elge et al. 2001). In plants, this family of proteins has an
additional N-terminal domain known as the membrane
occupation and recognition nexus (MORN) motif. MORN
repeats are thought to play a role in the targeting of
proteins to the plasma membrane of junctophilins, a class
of proteins that is present in the junctional complexes
between the plasma membrane and the endoplasmic
reticulum (Takeshima et al. 2000).
PD4/BRX belongs to a family of five proteins. It was
recently shown that all members of this family contain a
BRX domain, which is likely to be involved in protein–
protein interactions (Briggs et al. 2006). BRX is nuclear
localized and might act as a transcriptional activator
(Briggs et al. 2006).
PD5 is an unknown protein that belongs to a family of
five higher plant-specific genes. Preliminary observations
suggest that primary root development is altered in the
PD5 line (unpublished results). Such a phenotype is
encountered in other phloem mutants, such as apl and
wol. It may be that a modification of phloem transport
interferes with nutrients and/or auxin transport.
Materials and Methods
Plant material and growth
Arabidopsis thaliana ecotype Wassilevskaja (Ws) was used
as the wild type. The markers were obtained from insertion lines
of the Versailles collection (Ws ecotype). The original names
of the lines in the Versailles collection were CUM12 (PD1), ERA4
(PD2), EHD280 (PD3), EHH20 (PD4) and DYG25 (PD5). Seeds
were surface-sterilized and incubated in the cold for 2 d. They
were then sown on Arabidopsis medium (Estelle and Sommerville
1987) in Petri dishes and allowed to germinate and grow in a
growth chamber (200 mE m2 s1, 16 h day/8 h night, 208C,
70% humidity).
Cytoenzymological localization of GUS activity
GUS activity was detected as follows. Whole seedlings were
permeabilized in ice-cold (208C) 90% (v/v) acetone for 1 h at
208C (Hemerly et al. 1993) and washed twice, for 5 min each,
in 100 mM sodium phosphate buffer pH 7.2. They were then
infiltrated with 100 mM sodium phosphate buffer pH 7.2, 10 mM
sodium EDTA, 0.1% Triton X-100 and 1 mg ml1 5-bromo-4chloro-3-indolyl-b-d-glucuronic acid (X-gluc, Duchefa), to which
2.5 mM potassium ferrocyanide and potassium ferricyanide
were added, and incubated in the dark at 378C. PD5 seedlings
were incubated for 2 h. Other PD marker plants were incubated
overnight. Samples were rinsed three times in water, treated with
70% ethanol for 24 h and cleared in chloral hydrate for 24 h.
They were then fixed in Hoyer solution (chloral hydrate, arabic
gum, glycerol and H2O) on a glass slide. Samples were viewed
under differential interference contrast (DIC) optics, with a Zeiss
Axioplan II microscope (Zeiss, France). Images were acquired with
a Jenoptik ProGress C10plus digital camera (Clara Vision, France).
108
Protophloem differentiation in early A. thaliana development
Histological analysis
Floral stalks for resin embedding were fixed in 2% (v/v)
glutaraldehyde and 1% (v/v) paraformaldehyde in 0.1 M phosphate buffer, pH 7.2. After incubation at 48C for 24 h, samples
were washed in phosphate buffer, dehydrated using a series of
graded ethanol solutions and embedded in Technovit 7100 resin
(Heraeus Kulzer, Wehrheim, Germany), according to the manufacturer’s instructions. Sections (5 mm) were made on a rotary Jung
RM 2055 microtome (Leica Microsystems, Heidelberg, Germany)
equipped with metallic blades (Heraeus Kulzer).
Aniline blue staining and confocal microscopy
Mature embryos and 24-, 48- and 72-h-old seedlings were
stained with AB as previously described (Bougourd et al. 2000) and
mounted in Hoyer solution.
AB staining was observed by confocal microscopy,
using an inverted Leica TCS-SP2-AOBS spectral confocal laser
scanning microscope (Leica Microsystems, Mannheim, Germany).
Samples were excited with a 488 nm argon laser with emission at
516–721 nm for AB detection.
AB staining was combined with GUS staining in some cases.
GUS staining of seedlings was performed first. The seedlings were
then rinsed in water and AB staining was performed. GUS staining
resulted in the formation of crystals that could be observed with
the reflection mode of the confocal microscope. Samples were
excited with a 488 nm argon laser and the reflection of the signal
was detected at wavelengths between 485 and 491 nm.
GFP detection
Confocal microscopy was performed on an inverted Leica
TCS-SP2-AOBS spectral confocal laser scanning microscope
(Leica Microsystems, Mannheim, Germany). Samples were excited
with a 488 nm argon laser with emission at 502–547 nm for green
fluorescent protein (GFP) detection.
Plasmid construction and plant transformation
For all promoter GFP constructs, a region upstream of the
ATG start codon (promoter and 50 -untranslated region) was
cloned in the entry vector pDONR207 with the gateway BP
enzyme clonase mix (Invitrogen). AttB1 and attB2 sites were
added, respectively, to the 50 and 30 ends of PD promoter sequences
via PCR for Ws genomic DNA using Taq DNA polymerase
(Invitrogen) according to the manufacturer’s instructions. These
regions were subsequently cloned in the pBi101-R1R2-GFP binary
vector using the gateway LR enzyme clonase mix (Invitrogen),
according to the manufacturer’s instructions. The R1R2 ccdB
recombination cassette was inserted upstream from the GFPer gene
in the pBi101-GFP binary vector, to obtain pBi101-R1R2-GFP.
This vector was then propagated in Escherichia coli DB3.1.
For PD1, a region of 1,329 bp (primers 50 -AAAAAAGCA
GGCTTCAGTAAAGTTTCACAAGCT-30 and 50 -AAGAAAGC
TGGGTCTCTTTGTTTTCTGTTTCTTG-30 ) was used. For PD3,
a 1,926 bp region (primers 50 -AAAAAACAGGCTTTCTAAA
ACCTATCACACAA-30 and 50 -AAGAAAGCTGGGTCCTGG
GATTAG-CTTTTGGTG-30 ) was used. For PD4, a region of
3,859 bp (primers 50 -ACAAGTTTGTACAAAAAAGCAGGC
TGTATGTAGTTTAATTGGTGGCCATA-30 and 50 -ACCACT
TTGTACAAGAAAGCTGGGTTTTTGGTCTCTTTTTTGAGT
TGTT-30 ) was used. For PD5, a 1,880 bp region (primers
50 -AAAAAAGCAGGCTGCGGTGTAATCATTATTTCG-30
and 50 -AAGAAAGCTGGGTCGACGGGAAATGGTGGTTA
AT-30 ) was used.
These constructs were used to transform E. coli DH10B
(Invitrogen) and were then used with Agrobacterium tumefaciens
C58pMP90, for the transformation of wild-type Arabidopsis plants
(Ws) by floral dipping.
Supplementary material
Supplementary material mentioned in the article is
available to online subscribers at the journal website
www.pcp.oxfordjournals.org.
Acknowledgments
We thank Yka Helariutta and François Roudier for helpful
discussion and comments on this manuscript. We are indebted to
Matthieu Simon and SGAP (station de génétique et amélioration
des plantes) for the access to Versailles collection mutants.
We thank Jean-Denis Faure and Patrick Laufs for useful
discussions during the course of this work. We thank Krystyna
Goffron and Bruno Letarnec for plant care. This work was
partially supported by a grant from ‘ACI développement et
physiologie intégrative’.
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(Received October 4, 2006; Accepted November 18, 2006)