Download GFP-tagging of cell components reveals the dynamics of subcellular

The Plant Journal (2003) 33, 775–792
GFP-tagging of cell components reveals the dynamics of
subcellular re-organization in response to infection of
Arabidopsis by oomycete pathogens
Daigo Takemoto, David A. Jones and Adrienne R. Hardham
Plant Cell Biology Group, Research School of Biological Sciences, Australian National University, PO Box 475 Canberra,
ACT 2601, Australia
Received 20 September 2002; revised 21 November 2002; accepted 2 December 2002.
For correspondence (fax þ61 2 6125 4331; e-mail [email protected]).
Summary
Cytoplasmic aggregation, the rapid translocation of cytoplasm and subcellular components to the site of
pathogen penetration, is one of the earliest reactions of plant cells against attack by microorganisms. We
have investigated cytoplasmic aggregation during Arabidopsis–oomycete interactions. Infection by nonpathogenic Phytophthora sojae was prevented in the plant epidermal cell layer, whereas Peronospora
parasitica isolates Cala2 (avirulent) and Noks1 (virulent) could both penetrate into the mesophyll cell layer.
Epidermal cell responses to penetration by these oomycetes were examined cytologically with a range of
transgenic Arabidopsis plants expressing Green Fluorescent Protein (GFP)-tagged cell components. These
included plants containing GFP-TUA6 for visualizing microtubules, GFP-hTalin for actin microfilaments,
GFP-tm-KKXX for endoplasmic reticulum (ER), and STtmd-GFP for the Golgi apparatus. In all interactions,
actin microfilaments were actively re-arranged and formed large bundles in cytoplasmic strands focused on
the penetration site. Aggregation of ER membrane and accumulation of Golgi bodies at the infection site
were observed, suggesting that production and secretion of plant materials were activated around the
penetration site. Microtubules did not become focused on the penetration site. No difference was evident
between the responses of epidermal cells in the non-host, incompatible and compatible interactions. This
result indicates that the induction of cytoplasmic aggregation in Arabidopsis epidermal cells was neither
suppressed by the virulent strain of Peronospora, nor effective in stopping infection.
Keywords: Arabidopsis thaliana, oomycetes, actin microfilaments, microtubules, Golgi apparatus, endoplasmic reticulum.
Introduction
Recent use of Green Fluorescent Protein (GFP), fused to
target proteins, has enabled the study of protein localization in living plant cells and the labeling of selected organelles or cell components. There have been many reports of
the successful visualization of a variety of cell structures,
including microtubules (Marc et al., 1998; Ueda et al., 1999),
microfilaments (Klahre et al., 2000; Kost et al., 1998), endoplasmic reticulum (ER) (Benghezal et al., 2000; Haseloff
et al., 1997), the Golgi apparatus (Boevink et al., 1998;
Nebenführ et al., 1999), mitochondria (Köhler et al., 1997;
Niwa et al., 1999), vacuoles (Di Sansebastiano et al., 1998;
Mitsuhashi et al., 2000), and peroxisomes (Cutler et al.,
2000; Mano et al., 1999) using GFP-tagging. This approach
ß 2003 Blackwell Publishing Ltd
is an extremely powerful tool to monitor cell dynamics over
time in living cells during particular processes such as cell
division, vesicle and organelle trafficking, and in response
to various kinds of stimuli including temperature, light,
wounding, and pathogen attack.
Plant cell responses involved in the early stages of attack
by microorganisms have been of interest to plant pathologists for many years. The existence of cytoplasmic aggregation was first reported by Tomiyama (1956) in the potato–
Phytophthora system. This was defined as the rapid translocation of cytoplasm and subcellular components to the
site of pathogen penetration of the cell. Similar plant
responses to pathogens were observed in cowpea–Uro775
776 Daigo Takemoto et al.
myces (Heath and Heath, 1971), barley–Erysiphe (Kunoh
et al., 1985), parsley–Phytophthora (Gross et al., 1993), flax–
Melampsora (Kobayashi et al., 1994), onion–Magnaporthe
(Xu et al., 1998), and onion–Botrytis (McLusky et al., 1999)
interactions. This indicates that cytoplasmic aggregation is
a common phenomenon in the early response of plant cells
to pathogens in both non-host and race-specific interactions.
Pharmacological studies in the early 1980s first suggested the possibility that cytoplasmic aggregation during
the defense response was dependent on cytoskeletal reorganization in plant cells (Hazen and Bushnell, 1983;
Tomiyama et al., 1982). Cytochalasins, inhibitors of the
polymerization of actin, inhibited or delayed resistance
reactions such as cell death (Hazen and Bushnell, 1983;
Škalamera and Heath, 1998; Takemoto et al., 1999;
Tomiyama et al., 1982), callose deposition (Kobayashi
et al., 1997a; Škalamera and Heath, 1996), and pathogenesis-related (PR) protein expression (Takemoto et al., 1999),
as well as cytoplasmic aggregation. Moreover, treatment
with cytochalasins and microtubule polymerization and depolymerization inhibitors permitted the non-pathogen Erysiphe pisi to penetrate barley coleoptile cells and form
haustoria and secondary hyphae (Kobayashi et al.,
1997a). These results indicated that the cytoskeleton may
play an important role in plant resistance; however, details
of the precise contribution of cytoskeletal elements have
not yet been elucidated. As is generally the case in inhibitor
studies, it is difficult to determine what effects of the treatments apply specifically to the defense response and what
may be due to interference in other fundamental functions
played by the cytoskeleton in plant cell growth and development (see reviews by Nick, 1999; Staiger, 2000).
Over the last decade, a large amount of information on
plant–pathogen interactions has been obtained from studies of the infection of Arabidopsis by pathogens such as
Peronospora parasitica and Pseudomonas syringae. More
than 10 resistance genes (R-genes) have been characterized
at a molecular level (see review by Holub, 2001). Other
components involved in the expression of resistance have
also been identified, such as PR proteins (e.g. PR-1 to -5),
enzymes required for phytoalexin biosynthesis (e.g. PAD3),
signal transduction factors (e.g. NPR1, EDS1, and PAD4),
and possible rate-limiting factors of defense (e.g. RAR1 and
STG1) (Austin et al., 2002; Muskett et al., 2002; Parker et al.,
2000; Ryals et al., 1996; Zhou et al., 1999). However, cytological studies of Arabidopsis–pathogen interactions are
relatively limited.
There are some reports of the aggregation of plant material at the site of pathogen penetration in Arabidopsis. Koch
and Slusarenko (1990) described the aggregation of plant
material around the penetration hyphae of P. parasitica in
both compatible and incompatible interactions. Parker et al.
(1993) observed more callose accumulation in the incom-
patible interaction between Arabidopsis and P. parasitica
than in the compatible interaction. Donofrio and Delaney
(2001) investigated callose deposition in the compatible
interaction in detail and found that extrahaustorial callose
was present on nearly half of the haustoria examined. In the
newly established Arabidopsis–Phytophthora porri system,
the deposition of plant material and callose was described
as a specific response in the incompatible interaction
(Roetschi et al., 2001). However, as yet, there are no reports
that describe cytoskeletal or endomembrane re-arrangement within Arabidopsis cells in response to pathogen
attack.
In the present study, we have investigated the response
of epidermal cells in cotyledons of A. thaliana to attack by
three oomycete pathogens. Cytoplasmic re-organization in
response to penetration by P. sojae (non-pathogenic) and
P. parasitica isolates Cala2 (avirulent) and Noks1 (virulent)
was examined in transgenic Arabidopsis plants in which
selected cell components were tagged with GFP. Details of
the re-arrangement of cytoskeletal elements and components of the endomembrane system were revealed. Our
results indicate that cytoplasmic aggregation in Arabidopsis epidermal cells is not sufficient to stop P. parasitica
ingress in the compatible or incompatible interaction. Possible roles of cytoplasmic aggregation and cytoskeletal rearrangement in plant–pathogen interactions are discussed.
Results
The interaction between Arabidopsis and
Phytophthora sojae
Phytophthora sojae Kaufmann et Gerdemann (syn.
P. megasperma f. sp. glycinea Kuan et Erwin) is the causal
agent of stem and root rot of soybean. Phytophthora sojae
has only a limited host range and it is known that A. thaliana is not a host for P. sojae (Kamoun, 2001). The interaction between Arabidopsis and P. sojae was examined using
light microscopy, after staining with lactophenol trypan
blue of inoculated cotyledons, sampled from 6 h to 7 days
after inoculation. There were few zoospores moving on the
cotyledon surface, 6 h after inoculation, and most had
already encysted and germinated. At this point of time, a
variety of infection stages of P. sojae were present, from the
initiation of cyst germination to the formation of an appressorium-like swelling. One day after inoculation, most cysts
had germinated and developed appressorium-like structures which formed preferentially over anticlinal walls between epidermal cells (Figure 1a). Often, germ tubes grew
across the surface of the cotyledon and occasionally formed
more than one appressorium-like swelling (Figure 1b).
In rare instances, P. sojae grew through stomatal openings
or penetrated epidermal cells directly through the outer
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Arabidopsis response to oomycete pathogens
epidermal wall (data not shown). Occasionally, finger-like
structures that we interpret as developing haustoria formed
under the appressorium-like swelling (Figure 1b). In most
cases, P. sojae hyphae were unable to reach the plant
mesophyll cell layer. Plant cells around the penetration site
were generally not stained with trypan blue (staining of
plant cells with trypan blue is indicative of cell death),
although a few plant cells attacked by multiple P. sojae
germlings did undergo cell death (data not shown). In no
case was P. sojae found to colonize the mesophyll layer
(Figure 1c).
Morphological comparisons of incompatible and
compatible interactions between Arabidopsis thaliana
and Peronospora parasitica
The extent of Peronospora development in incompatible
and compatible interactions with Arabidopsis was examined. To monitor pathogen growth and plant cell death,
whole cotyledons of A. thaliana ecotype Columbia (Col-0)
were stained with lactophenol trypan blue 6 h to 7 days
after inoculation with P. parasitica isolates. Col-0 possesses
the RPP2 resistance gene which confers resistance to
P. parasitica isolate Cala2 but not to P. parasitica isolate
Noks1 (Holub et al., 1994). Thus, the interaction of Col-0 and
Cala2 is incompatible and that between Col-0 and Noks1 is
compatible.
Six hours after inoculation, most of the Cala2 or Noks1
conidia had germinated and penetrated the underlying
epidermis, although it should be noted that ungerminated
conidia tended to be lost during processing. Both Cala2 and
Noks1 preferentially penetrated between the anticlinal
walls of adjacent epidermal cells, and at 6 h after inoculation, both had started forming haustoria in the epidermal
cells (Figure 1d,j). By 12 h, mature haustoria had formed
below the penetration site, and for both isolates, about 10%
of the haustoria in epidermal cells were encapsulated by
plant cellular material (Figure 1l).
A difference between the incompatible and compatible
interactions became evident 1 day after inoculation. In the
Col-0–Cala2 interaction, densely-stained cells containing
mature or developing haustoria were observed in the
mesophyll layer (Figure 1e,f). Cala2 mycelium growth
usually terminated within a cluster of dead mesophyll cells,
but hyphae were seen occasionally beyond such a cluster
(Figure 1g). Blue-stained epidermal cells were occasionally
found next to the penetration site (Figure 1i) but did not
prevent growth of hyphae. Four days after inoculation,
there were many clusters of two to five dead mesophyll
cells in the Col-0–Cala2 interaction and some with more
than 10 dead cells (Figure 1h). Inoculated seedlings continued to grow normally, and conidiophore formation and
macroscopically visible symptoms of infection were never
observed for the Col-0–Cala2 interaction.
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At 1 day after inoculation in the compatible interaction
between Col-0 and Noks1, a number of large haustoria had
formed within the mesophyll layer in most infection sites,
and further mycelial growth and subsequent formation of
additional developing haustoria were observed (Figure 1k).
Blue-stained epidermal cells occasionally occurred at the
penetration site as seen for the Col-0–Cala2 interaction.
Two to three days after inoculation, extensive growth of
intercellular mycelia and the formation of large numbers of
haustoria were evident (Figure 1m). Mesophyll cell death,
indicated by dense staining, was observed in only a few
instances where there was massive infection. Up to this
stage, there were no macroscopically visible symptoms on
the inoculated cotyledon. Four days after inoculation, oogonia and antheridia appeared (Figure 1n). Conidiophores
emerged 4 to 5 days after inoculation (Figure 1o) and a
lawn of conidiophores was macroscopically obvious by
6 days on most of the inoculated cotyledons. The seedlings
became etiolated and finally collapsed by about 10 days.
In experiments in which transgenic Col-0 plants expressing GFP-TUA6, STtmd-GFP, GFP-tm-KKXX, or GFP-hTalin
were inoculated with P. parasitica, no differences in the
response of wild-type and transgenic plants to Noks1 or
Cala2 were detected (data not shown).
Quantitative comparison of the incompatible and
compatible interactions between Arabidopsis and
Peronospora parasitica
After lactophenol trypan blue staining, the numbers of
haustoria formed in mesophyll and epidermal cells and
the number of dead plant cells per infection site were
scored for a large number of penetration sites from 6 to
48 h after inoculation, in order to compare the incompatible
(Col-0–Cala2) and the compatible (Col-0–Noks1) interactions. In both interactions, more than 99% of the conidia
that remained on the cotyledon after processing produced
hyphae that penetrated between the anticlinal walls of two
epidermal cells. Up to 24 h after inoculation, the numbers
of developing and mature haustoria per infection site were
almost the same for both interactions. During this period,
the number of haustoria per infection site did not increase
dramatically, although many matured (compare the data
for 6 h with that for 24 h after inoculation in Figure 2a).
However, from 24 to 48 h after inoculation, Noks1 showed
massive production of haustoria while Cala2 produced only
a small number of additional haustoria from a few mycelia
that were not encased by dead plant cells (Figure 2a). This
dramatic difference was due mainly to haustoria production by Noks1 in mesophyll cells because the numbers of
haustoria formed in epidermal cells were almost the same
for both interactions over the time examined (Figure 2b).
At 6 and 12 h after inoculation, almost no plant cell
death was observed in either interaction. Differences in
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Arabidopsis response to oomycete pathogens
779
the frequency of plant cell death between the two interactions became evident 24 h after inoculation (0.29 0.04
dead cells per infection site for the incompatible interaction,
0.04 0.02 for the compatible). This difference became
more pronounced at 48 h (0.95 0.08 for incompatible,
0.13 0.04 for compatible) (Figure 2c). However, as was
the case for haustorium formation, the frequency of cell
death in the epidermis was the same in both interactions
during the time period studied (0.06 0.02 for both incompatible and compatible) (Figure 2c).
Microtubule distribution in Arabidopsis epidermal cells
during infection by oomycete pathogens
Transgenic Arabidopsis plants expressing GFP-tagged
a-tubulin (GFP-TUA6) (Ueda et al., 1999) were used to
visualize microtubules in living cotyledon cells. In uninoculated plants, the microtubules formed cortical arrays of
gently curving, brightly fluorescent strands underlying the
outer epidermal cell wall (Figure 3a). Divergence of bright
strands into fine strands of lower fluorescence intensity
suggests that most of the fluorescent strands represent
bundles of microtubules. Occasionally, weakly fluorescent
Figure 1. Cytological characterization of three Arabidopsis–oomycete interactions.
Arabidopsis thaliana ecotype Columbia (Col-0) was inoculated with Phytophthora sojae (non-pathogen), Peronospora parasitica isolate Cala2
(avirulent), or isolate Noks1 (virulent). Inoculated cotyledons were stained
with lactophenol trypan blue.
cy, cyst; c, conidium; as, appressorium-like swelling; h, haustorium; ps,
penetration site; dh, developing haustoria; eh, encapsulated haustorium,
an, antheridium; o, oogonium; cp, conidiophore. Bar ¼ 25 mm (a, b, f, l) and
50 mm (c–e, g–k, m–o).
(a–c) Non-host interaction of Col-0 with P. sojae. (a) A germinated cyst with
appressorium-like swelling on the surface of cotyledon. The micrograph
was taken 6 h post-inoculation (hpi). (b) One day post-inoculation (dpi). The
appressorium-like swellings were preferentially formed over the anticlinal
walls of plant epidermal cells. A finger-like structure is indicated by the
arrowhead. (c) Germ tubes growing across the surface of the cotyledon.
Insert at the top right is the region indicated by the square in the main panel
but focused within the epidermal cell. Note that P. sojae does not penetrate
into the mesophyll cell layer 5 dpi.
(d–i) Incompatible interaction of Col-0 with P. parasitica isolate Cala2. (d)
Initial stage of the infection by Cala2. Cala2 preferentially penetrates
between the anticlinal walls of adjacent epidermal cells and forms some
haustoria 6 hpi. (e) Mesophyll cell death beneath the penetration site. Image
at the top right is the same penetration site as in the main panel but focused
on the mesophyll cell layer. Note that the epidermal cells in direct contact
with Cala2 (indicated by arrowheads) do not show cell death 1 dpi. (f)
Haustoria formation by Cala2 in epidermal cells. Arrowheads indicate dead
mesophyll cells 1 dpi. (g) Clusters of dead mesophyll cells. Arrowheads
indicate mycelia growing beyond the cluster of dead cells in the mesophyll
layer 3 dpi. (h) Large cluster of dead cells in the mesophyll cell layer 4 dpi. (i)
Epidermal cell death adjacent to the penetration site (indicated by black
arrowheads). White arrowhead indicates the dead mesophyll cell 1 dpi.
(j–o) Compatible interaction of Col-0 with P. parasitica isolate Noks1. (j)
Initial stage of the infection by Noks1. Note that Noks1 initially showed a
similar pattern of infection as Cala2 in (d) 6 hpi. (k) Haustorial formation and
further infection of Noks1 without plant cell death 1 dpi. (l) Fully developed
and encapsulated haustoria formed in epidermal cell 1 dpi. (m) Heavy
colonization by Noks1 with numerous haustoria 3 dpi. (n) Oogonia and
antheridia formation 4 dpi. (o) Conidiophore formation 5 dpi.
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Figure 2. Time course of haustoria formation and induction of plant cell
death during the infection of avirulent (Cala2) and virulent (Noks1) Peronospora parasitica.
(a) Haustoria formation.
(b) Haustoria formation in epidermal cells.
(c) Induction of plant cell death. Haustoria or plant cell death were scored by
observation of inoculated plants after clearing and staining in lactophenol
trypan blue. The numbers of infection sites assessed are indicated above
each column.
cytoplasmic strands were seen running beneath the cortical
microtubule array in these cells (Figure 3a). Radial arrays of
microtubules were seen in stomatal guard cells (Figure 3a).
As in wild-type plants, 6 h after inoculation with zoospores of P. sojae, many appressorium-like swellings had
780 Daigo Takemoto et al.
Figure 3. Distribution of microtubules in plant epidermal cells after penetration by oomycete pathogens.
Images were collected 6 h after inoculation. Penetration sites and cytoplasmic strands are indicated by asterisks and closed arrowheads, respectively. The
junctions of epidermal cells are outlined by dashed lines. Except for (c), images of Green Fluorescent Protein (GFP) fluorescence are projections of optical
sections taken at 1-mm intervals from the outer epidermal wall through to immediately above the cortical cytoplasm adjacent to the inner periclinal wall of the
epidermal cell. Bar ¼ 10 mm.
(a) Tubulin network visualized by GFP-TUA6 in an uninoculated plant.
(b,c) GFP-TUA6 plant inoculated with the non-pathogen Phytophthora sojae. (b) Diffuse fluorescence has accumulated around the penetration site. Weakly
fluorescent cytoplasmic strands running through the cell focus on the penetration site (b, c). Image in (c) shows a single optical section taken at the centre of the
epidermal cell.
(d–f) GFP-TUA6 plant inoculated with the avirulent pathogen Peronospora parasitica isolate Cala2. Microtubules do not focus on the penetration site but diffuse
fluorescence is seen around the penetration site and in cytoplasmic strands (closed arrowheads) directed toward this region. A tendency for the microtubules to
form a circumferential arrangement around the penetration site is evident to varying degrees (most clearly seen in (f)).
(g–i) GFP-TUA6 plant inoculated with the virulent pathogen P. parasitica isolate Noks1. The pattern of fluorescence of GFP-TUA6 is similar to that observed after
inoculation with Cala2. Diffuse fluorescence is seen around the penetration site and in cytoplasmic strands (closed arrowheads) directed toward this region. Microtubules are not focused on the penetration site but in some cases form an approximately circumferential arrangement in cells surrounding the penetration site (e.g. (h)).
formed on the cotyledons of the GFP-TUA6 plants, and
hyphae had attempted to grow between the anticlinal walls
of the epidermal cells. Observation of GFP-TUA6 fluorescence revealed enhanced but diffuse fluorescence around
the penetration site (Figure 3b). Similar images were seen
following inoculation of the GFP-TUA6 plants with avirulent
(Figure 3d–f) and virulent (Figure 3g–i) isolates of P. parasitica. Fluorescent microtubule bundles could be seen to
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Arabidopsis response to oomycete pathogens
run through the region of diffuse fluorescence. This type of
diffuse fluorescence was not observed in wild-type plants
and is not due to autofluorescence of material deposited
around the penetration site (data not shown). In many
cases, the density and arrangement of microtubule bundles
in the cortical arrays near the penetration site did not
appear to be noticeably different from that seen in uninoculated plants. There was no evidence of microtubule
bundles being specifically focused on the penetration site
(Figure 3b,e,f,h,i). However, in some cells, the microtubules
appeared to be aligned circumferentially around the penetration site (Figure 3d,f,h). This was a global arrangement
that encompassed a number of epidermal cells in the
vicinity of the penetration site and was seen in all three
interactions. In addition, in cells adjacent to the penetration
site, greater numbers of broad, diffusely fluorescent cytoplasmic strands were observed traversing the central
vacuole of the cells (Figure 3c), and these did show a
tendency to focus on the infection site (Figure 3b,e,f,h).
These patterns of GFP-TUA6 fluorescence persisted in all
three Arabidopsis–oomycete interactions throughout the
duration of observations (up to 36 h after inoculation),
except that GFP fluorescence was not seen in epidermal
cells that had died (data not shown).
Re-organization of microfilaments in Arabidopsis
epidermal cells during oomycete infection
The actin microfilament network was observed in living
epidermal cells of transgenic Arabidopsis plants expressing GFP conjugated to the actin-binding domain (McCann
and Craig, 1997) of human talin (GFP-hTalin). The brightness of the fluorescence of the GFP-hTalin varied in the 14
independent transgenic lines that were isolated. In some
lines, presumably with high levels of transgene expression,
the network of fine cortical microfilaments beneath the
surface of epidermal cells could be clearly seen in addition
to the cytoplasmic microfilament cables in the centre of the
cells (Figure 4a). However, these transgenic plants tended
to show growth inhibition (e.g. shorter plants and smaller
leaves) compared with wild-type plants, and the cotyledons
tended to be angled downward rather than being held
horizontally. Nevertheless, these plants were still able to
grow and set viable seeds (data not shown). In transgenic
plants with apparently intermediate levels of GFP-hTalin
expression, microfilaments in cytoplasmic strands were
still very prominent, but the fine microfilaments in the cell
cortex were more weakly fluorescent and difficult to see
than in the plants expressing the transgene at a high level
(Figure 4b,c). However, as no effects on growth or morphology were detected in these latter plants, one line was selected and used for inoculation with the oomycete pathogens.
After their penetration along the anticlinal walls of epidermal cells, similar patterns of actin microfilament distriß Blackwell Publishing Ltd, The Plant Journal, (2003), 33, 775–792
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bution were observed in the GFP-hTalin plants inoculated
with the non-pathogen P. sojae (Figure 4d–f) and the
incompatible (Figure 4g–i) and compatible (Figure 4j–l) isolates of P. parasitica. Many relatively fine microfilaments
and a number of strongly fluorescent bundles of microfilaments became focused on the penetration site. In many
cases, small bundles of microfilaments coalesced to form
larger bundles near the penetration site (Figure 4e,i,k,
closed arrowheads). Re-organization of the microfilament
network was very active around the penetration site (Figure 5). During the 2-min intervals between the images
shown in Figure 5, both the more weakly fluorescent fine
microfilaments as well as the strongly fluorescent microfilament cables change their position, coalesce, or diverge.
Changes in structure of the endoplasmic reticulum
network after inoculation with oomycete pathogens
The Arabidopsis plants used to determine the distribution
of ER after inoculation with the three oomycete pathogens
were transformed with a chimaeric gene encoding GFP
fused between the signal peptide from Arabidopsis basic
chitinase at the N-terminus and the transmembrane
domain and short cytosolic tail containing the KKXX motif
of the tomato Cf-9 resistance protein (Jones et al., 1994) at
the C-terminus (Benghezal et al., 2000). The C-terminal
KKXX signal (dilysine motif) confers ER localization to
membrane-bound GFP (Benghezal et al., 2000). In GFPtm-KKXX plants, typical reticulate ER was clearly observed
in uninoculated plants as previously reported (Benghezal
et al., 2000).
After inoculation with any of the three oomycete pathogens, intense fluorescence rapidly accumulated around the
penetration site (Figure 6). The dynamics of this increase in
fluorescence around the invading hypha are illustrated in
Figure 6(a) after inoculation with P. sojae. In this example,
aggregation of ER is seen to begin before the hypha has
penetrated between the epidermal cells. We interpret the
strong fluorescence in cells on either side of the penetration
site as constituting sheet-like lamellae of ER cisternae.
These lamellae were continuous with the tubular ER network that extended throughout the cortical cytoplasm
underlying the outer epidermal cell walls. In many
instances, the mesh size of the ER network was smaller
(about 1–2 mm), closer to the penetration site than it was
further away (Figure 6c, open arrowheads; Figure 6b,g,i).
The tubular cisternae surrounding the intensely fluorescent
ER area at the penetration site sometimes expanded into
small lamellar cisternae (e.g. Figure 6f,i). Cytoplasmic
strands were invariably brightly fluorescent and were also
generally directed to the penetration site (Figure 6b,e,g,i,
closed arrowheads). The fluorescence of the cytoplasmic
strands tended to be more diffuse than that of the tubular
ER, a feature that is likely to be due to movement of the ER
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Arabidopsis response to oomycete pathogens
within the rapidly streaming strands. In a number of
instances, the fluorescence of the cortical ER network in
cells adjacent to the penetration site was lower in regions
distant from the penetration site (Figure 6d, x) than it was
close to the penetration site or in cells not immediately
adjacent to the infection site (Figure 6d, y). Following
inoculation with either Cala2 or Noks1 isolates of P. parasitica, the ER was seen to surround the neck but not the
distal regions of haustoria in the epidermal cells (Figure 6j).
This lack of fluorescence at the distal end of the haustoria
was observed in all of more than 20 haustoria examined.
Localization of the Golgi apparatus after inoculation with
oomycete pathogens
Boevink et al. (1998) reported the use of the transmembrane domain and signal anchor sequences of rat 2,6-sialyl
transferase (STtmd) to label the Golgi apparatus in plant
cells. In the present study, transgenic Arabidopsis plants
expressing STtmd-GFP were used to investigate the distribution of the Golgi apparatus in cells penetrated by
oomycete pathogens. As shown in Figure 7, Golgi stacks
became preferentially localized around the penetration site
in all three Arabidopsis–oomycete interactions. In contrast
to the situation for the ER, in which the brightly fluorescent
lamellae remained around the penetration site, the Golgi
stacks continued to move around the cell and the degree of
accumulation at the penetration site changed over time
(Figure 7c). Most of the Golgi stacks were transported along
cytoplasmic strands running through the centre of the
epidermal cells and focusing on the penetration site. Golgi
stacks moved towards the penetration site on some strands
and away from this site on other strands (data not shown).
As for the ER, Golgi stacks surrounded the neck but not the
haustorial body of Peronospora haustoria that formed in
the epidermal cells (Figure 7f). This lack of Golgi bodies
around the distal end of the haustorium was observed in all
of more than 20 haustoria examined.
Discussion
Green Fluorescent Protein has become a valuable tool for
investigations of subcellular structures in living cells.
Although transgenic plants with various GFP-tagged cell
783
components have been available (see Introduction section),
no reports of the use of these plants in studies of cell reorganization in response to pathogenic attack have yet
been published. In the present study, we first characterized
the non-host, incompatible and compatible interactions of
Arabidopsis with oomycete pathogens, and then monitored the re-organization of GFP-tagged cell components
during these interactions.
Comparison of the Arabidopsis responses to non-host,
incompatible and compatible oomycete pathogens
All three oomycete pathogens preferentially penetrated the
Arabidopsis cotyledons along the anticlinal walls between
epidermal cells, as has been commonly observed for the
infection of plant roots and shoots by oomycetes (e.g.
Hardham, 2001; Koch and Slusarenko, 1990; Stössel et al.,
1980). The progress of infection by P. sojae was halted
close to the initial penetration site in the epidermis. This
inhibition of further growth of P. sojae occurred in the
absence of epidermal cell death, indicating that other
mechanisms such as the formation of a barrier at the cell
wall effectively stopped pathogen ingress. In the Arabidopsis–P. sojae interaction, epidermal cell death was observed
only when multiple hyphae were in contact with a cell,
although contact with a single hypha of P. infestans or
P. porri can cause epidermal cell death in Arabidopsis
(Roetschi et al., 2001; Vleeshouwers et al., 2000).
Epidermal cell death was rarely observed during the
interaction of Col-0 with the incompatible Cala2 and compatible Noks1 isolates of P. parasitica. For the P. parasitica
isolates, no difference in the infection process was detected
during the first 24 h after inoculation, while the pathogens
grew through the epidermal cell layer. Both only occasionally formed haustoria in the epidermal cells. Cala2 and
Noks1 grew through the epidermis and into the mesophyll
cell layer, and it is within the mesophyll that differences
became evident between the incompatible and compatible
interactions with Col-0. Growth of Cala2 was, in general,
stopped quickly within the mesophyll, with few haustoria
being formed and the hyphae becoming surrounded by
dead mesophyll cells. Occasionally, Cala2 hyphae were
seen beyond a cluster of dead mesophyll cells. This may
reflect the inability of Col-0 to completely restrict growth in
Figure 4. Distribution of microfilaments in Arabidopsis epidermal cells after penetration by oomycete pathogens.
Images were collected 6 h after inoculation. Penetration sites are indicated by asterisks. The junctions of epidermal cells are outlined by dashed lines. Closed
arrowheads indicate sites where microfilament bundles merge to form larger cables. Images of Green Fluorescent Protein (GFP) fluorescence are projections of
optical sections taken at 1-mm intervals from the outer epidermal wall through to immediately above the cortical cytoplasm adjacent to the inner periclinal wall of
the epidermal cell. In all inoculated plants, both large and small bundles of actin microfilaments have become focused on the pathogen penetration site.
Bar ¼ 10 mm.
(a–c) Uninoculated GFP-hTalin plants showing relatively high (a), intermediate (b), and low (c) levels of transgene expression. Cortical microfilaments are most
clearly visualized in (a).
(d–f) GFP-hTalin plants inoculated with the non-pathogen Phytophthora sojae.
(g–i) GFP-hTalin plants inoculated with the avirulent pathogen Peronospora parasitica isolate Cala2.
(j–l) GFP-hTalin plants inoculated with the virulent pathogen P. parasitica isolate Noks1.
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 33, 775–792
784 Daigo Takemoto et al.
this interaction, although, contrary to a previous report
(Holub et al., 1994), no Peronospora sporulation was
observed under our conditions. Even during the interaction
of Cala2 with Oystese-1 (Oy-1), an ecotype of Arabidopsis
which is highly resistant as it has resistance genes at two
different loci (RPP2 and RPP3) conferring resistance to
Cala2 (Holub et al., 1994), no epidermal cell death was
observed (D. Takemoto, unpublished observations). Our
results are consistent with other observations (Botella
et al., 1998; Parker et al., 1993), which indicate that while
cell death may occur in epidermal cells in some interactions
(Koch and Slusarenko, 1990), in general, R-gene resistance
of Arabidopsis to P. parasitica is associated with cell death
in the mesophyll cell layer rather than the epidermal layer
(Figure 8). Reduced image quality in the mesophyll layer
restricted observations of GFP fluorescence in the present
study to the epidermis.
Observations of the GFP-tagged Arabidopsis plants using
confocal microscopy clearly revealed cytoplasmic aggregation at the penetration site within the epidermal cells during
interactions with all three oomycete pathogens. In other
studies, more extensive cytoplasmic aggregation (e.g.
Kobayashi et al., 1992) has been observed in resistant than
in susceptible plants, but in our study there were no
obvious differences between the non-host, incompatible
and compatible interactions with respect to the aggregation
of GFP-tagged cell components. Whether or not cytoplasmic aggregation was responsible for effectively stopping
the infection by P. sojae, despite failing to prevent the
ingress of either P. parasitica isolate, remains to be determined.
Re-arrangement of the plant cytoskeleton during
pathogen infection
Figure 5. Sequential images of the microfilament network around a Phytophthora sojae penetration site.
Images were collected from 6 h after inoculation at 2-min intervals. Top left
panel shows the differential interference contrast (DIC) image for the same
area as the other images. The germ tube and appressorium-like swelling are
outlined by a dashed black line. The junctions of epidermal cells are outlined
by dashed white lines. Arrowheads indicate microfilament strands that have
formed or increased in brightness during the time since the previous image.
The microfilament network was re-arranged actively around the penetration
site. Bar ¼ 10 mm.
In all three Arabidopsis–oomycete interactions, actin microfilaments were re-arranged to form large bundles focused
on the penetration site. Microfilament bundles from different parts of the cell merged and supported active cytoplasmic streaming directed towards the infection site. This rearrangement of actin microfilaments and the associated
formation of cytoplasmic strands focused on the infection
site is a consistent feature of the plant response in almost all
pathosystems studied to date, including barley–Erysiphe
(Baluška et al., 1995; Kobayashi et al., 1992), flax–Melampsora (Kobayashi et al., 1994), cowpea–Uromyces (Škalamera et al., 1998), onion–Magnaporthe (Xu et al., 1998),
and onion–Botrytis (McLusky et al., 1999). In some cases,
microfilament accumulation was greater in non-host or
resistant plants than it was in susceptible plants (e.g.
Kobayashi et al., 1992, 1994), although in the cowpea–Uromyces system, microfilament cables were formed around
the infection site in the susceptible host more frequently
than in the resistant host (Škalamera et al., 1998). Drug
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 33, 775–792
Arabidopsis response to oomycete pathogens
studies have also shown that de-polymerization of the actin
cytoskeleton inhibits formation of the cytoplasmic aggregate and reduces the plant’s ability to inhibit pathogen
attack (see Introduction).
In contrast to the behavior of the actin microfilaments, the
network of cortical microtubules did not become focused
on the penetration site in any of the three pathogen interactions. In the GFP-TUA6 plants, the most obvious change
was the appearance of strong but diffuse fluorescence
around the penetration site. Similar fluorescence was not
observed in wild-type plants or in any of the other GFPtransformants, indicating that the fluorescence was not due
to autofluorescence of material deposited at the penetration site. We consider it likely that the diffuse fluorescence
in the GFP-TUA6 plants is due to monomeric or dimeric
GFP-tubulin in the cytoplasm. The accumulation of the GFPtubulin subunits at the penetration site could be due to
increased tubulin synthesis, or, more likely, to localized
microtubule de-polymerization as has been reported previously in the parsley–Phytophthora interaction (Gross
et al., 1993) or after elicitor treatment in tobacco cells (Binet
et al., 2001). Our observations of diffuse GFP-tubulin
fluorescence in the cytoplasm at the penetration site may
reflect the retention of cytoplasmic proteins in the living,
non-permeabilized cells in contrast to the loss of soluble
proteins in cells permeabilized for immunocytochemical
labeling.
In some Arabidopsis epidermal cells, there was no apparent change in microtubule arrangement, but in others,
microtubules seemed to be aligned in a roughly circumferential array in epidermal cells surrounding the penetration
site. This type of alignment has not been previously
reported as part of the plant response to pathogen attack,
although similar global re-arrangements have been
observed in response to wounding (Hush et al., 1990)
and during leaf formation (Hardham et al., 1980). In both
barley–Erysiphe and flax–Melampsora interactions, microtubules become radially arranged and focused on the site of
contact with the pathogen (Kobayashi et al., 1992, 1994).
We note that in uninoculated plants, many cortical microtubules abut the anticlinal wall junctions at sharp angles,
and it is these microtubules that are apparently missing in
the cells forming part of the circumferential array. Perhaps
the circumferential arrangement we have observed has no
significance in itself, but is merely the result of preferential
de-polymerization of microtubules abutting the wall adjacent to the penetrating hypha. Thus, while the re-organization of actin microfilaments at the site of pathogen attack is
consistent, the behavior of microtubule arrays is quite
variable in different plant–pathogen systems. This may
be because microtubules do not play a direct role in the
response, although a number of studies have shown that
microtubule de-polymerization reduces the plant’s resistance (Kobayashi et al., 1997a,b). Alternatively, differences
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 33, 775–792
785
in the behavior of microtubule arrays may be influenced by
such factors as (i) the species of plant and pathogen
involved in the interaction, (ii) whether the pathogen is a
necrotroph or biotroph, (iii) whether the pathogen is growing intercellularly or is attempting to penetrate through the
plant cell wall, or (iv) whether the system involves cultured
cells or intact plant tissues. A number of Arabidopsis-based
necrotrophic and biotrophic pathosystems have now been
developed, involving true fungi such as Alternaria brassicicola (e.g. Penninckx et al., 1996), Erysiphe spp. (e.g. Adam
et al., 1999), Fusarium oxysporum (Mauch-Mani and Slusarenko, 1994), and Botrytis cinerea (e.g. Thomma et al.,
1998), as well as the oomycete pathogens P. porri (Roetschi
et al., 2001), Albugo candida (Holub et al., 1995), and
P. parasitica (Koch and Slusarenko, 1990). Future investigations of the response of the GFP-tagged Arabidopsis plants
described in the present study to this range of pathogens
may help elucidate the consistent and important features of
the cytoskeletal re-organization in these interactions.
Re-organization of the endomembrane system during
pathogen infection
The GFP-tm-KKXX plants revealed dramatic re-organization of the ER at the penetration site after inoculation with
the three oomycete pathogens. Strong fluorescence around
the penetration site indicated the existence of a dense
network of lamellar ER. The lamellar network merged into
a condensed reticulum of tubular ER. This latter structure
resembled the transitional form of ER between ‘perforated
sheets’ and tubular reticulum observed in growing cells
(Ridge et al., 1999). The function of the perforated sheets of
ER in growing cells is not clear, but may be involved with
the synthesis of large quantities of proteins and lipids. A
similar function could be envisaged in plant cells responding to pathogen attack because the production of many
defense proteins is induced in these cells. A similar reorganization of ER was reported in the compatible interaction of pea and E. pisi, following labeling of the ER by antiHDEL antibodies (Leckie et al., 1995). In this latter study, it
was suggested that the accumulation of ER might be
involved in exocytosis to generate new membrane and
materials to form the haustorial complex. This seems unlikely to be the case in the Arabidopsis–oomycete interactions in the present study because the accumulation of ER
membrane at the penetration site occurred in cells with or
without haustoria, and in cells containing haustoria, the ER
did not encompass the haustorium body. Concentration of
ER around the neck of the haustorium has also been
observed in the compatible interaction between wheat
and Puccinia graminis (Harder et al., 1978).
In contrast to the large accumulation of ER around the
penetration site, fluorescence of the tubular ER cisternae
was weaker on the opposite side of the cell. Cytoplasmic
786 Daigo Takemoto et al.
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 33, 775–792
Arabidopsis response to oomycete pathogens
strands focused on the penetration site contained ER cisternae, and the accumulation of ER membrane occurred
quite rapidly. It is thus likely that much of the accumulated
ER membrane was recruited from the existing ER network
throughout the cell. Although the reticulate cortical ER
usually does not show large-scale translocations and is
distinctive in appearance from the ER tubules in streaming cytoplasm (Quader and Schnepf, 1986; Ridge et al.,
1999), the ER may re-distribute dramatically in particular
circumstances. An example of this is the re-distribution of
ER from the cortical cytoplasm into the phragmosome
during cell plate assembly (Hepler, 1982; Nebenführ et al.,
2000).
Inoculation of Col-0 with the three oomycete pathogens
led to a concentration of Golgi stacks around the penetration site. In contrast to the stable accumulation of ER,
observation of the same penetration site over a period of
time revealed that the distribution of Golgi stacks changed
continually. Although individual Golgi stacks moved
towards or away from the penetration site, the localized
accumulation of Golgi stacks indicated that they stopped at
least temporarily near the penetration site. Golgi stacks
have recently been shown to move along cytoplasmic
strands in tobacco leaf cells in an acto-myosin-dependent
manner (Boevink et al., 1998; Nebenführ et al., 1999, 2000).
The GFP-labeled Golgi stacks displayed stop-and-go movements, oscillating between directed movement and random ‘wiggling’. Stacks moving along the same track often
paused at the same position, indicating that there were
specific sites that inhibit the movement. From this observation, the authors proposed a ‘recruitment model’ in which
Golgi stacks are recruited to active ER export sites (or to the
sites for secretion) by localized stop signals that transiently
prevent the stacks from leaving the area (Nebenführ et al.,
1999). If this model is applied to the Arabidopsis–oomycete
interactions, it is possible that the accumulation of ER at the
penetration site might be associated with a large quantity of
these ‘stop signals’, leading to the preferential accumulation of Golgi stacks. This would favor effective ER-to-Golgi
transport and site-specific secretion in response to pathogen penetration.
787
Concluding remarks
The aggregation of cytoplasm at the site of pathogen attack
is generally believed to be associated with the deposition of
material that will generate or strengthen the physical and
chemical barriers to pathogen ingress within the plant cell
wall. Accumulation of ER and Golgi bodies in the cytoplasmic aggregation is consistent with this role. The function of
cytoplasmic aggregation and the cell components responsible for its formation have been investigated mainly in
pharmacological studies using inhibitors of cytoskeletal
elements. These studies have shown that de-polymerization of either actin microfilaments or microtubules may
prevent or delay both non-host and race-specific plant
resistance (see Introduction), and that de-polymerization
of actin prevents cytoplasmic aggregation. The cytoskeleton is thus thought to play an important part in the mobilization of the plant defense response.
Currently, little is known about the factors that induce
cytoplasmic aggregation at the infection site. Pathogen
attack may generate both physical and chemical signals
that can be detected by the underlying plant cell. It has, for
example, been shown that local mechanical pressure can
trigger the translocation of cytoplasm to the site of stimulation (Gus-Mayer et al., 1998; Kennard and Cleary, 1997),
indicating that cytoplasmic aggregation could be induced
by the pressure generated by the penetrating hypha. However, in the barley–Erysiphe (Kunoh et al., 1985) and flax–
Melampsora (Kobayashi et al., 1994) interactions, it has
been observed that cytoplasmic aggregation or cytoskeletal
re-organization in the plant cell was induced before pathogen penetration began. In addition, the mitogen-activated
protein kinase (mps1) mutant of M. grisea, which is unable
to form a penetration peg, still elicits cytoplasmic aggregation on non-host onion cells (Xu et al., 1998). All these data
support the idea that a chemical elicitor released by the
pathogen is sufficient to induce cytoplasmic aggregation.
Responses, such as cytoskeletal re-organization or localized wall modifications in plant cells adjacent to cells in
direct contact with the pathogen, could also be triggered by
such an elicitor.
Figure 6. Re-organization of endoplasmic reticulum (ER) network in Arabidopsis after inoculation by oomycete pathogens.
Images were collected 6 h after inoculation. Asterisks indicate the penetration sites. Cytoplasmic strands containing ER are indicated by the closed arrowheads.
The junctions of epidermal cells are outlined by dashed lines. Except for (j), images of Green Fluorescent Protein (GFP) fluorescence are projections of optical
sections taken at 1-mm intervals from the outer epidermal wall through to immediately above the cortical cytoplasm adjacent to the inner periclinal wall of the
epidermal cell. Strong fluorescence around the penetration site indicated the existence of a dense ER network in all inoculated plants. Bar ¼ 10 mm.
(a–d) GFP-tm-KKXX plants inoculated with the non-pathogen Phytophthora sojae. (a) Sequential images of the initiation of ER accumulation around the P. sojae
penetration site. The germ tube and appressorium-like swelling are outlined by a white dashed line. Images were taken at 5-min intervals. Open arrowheads in (c)
indicate regions of the ER network that has a smaller mesh size than in regions distant to the infection site or in uninfected cells. Epidermal cells with (x) or
without (y) direct P. sojae contact are marked in (d).
(e–g) GFP-tm-KKXX plants inoculated with the avirulent pathogen Peronospora parasitica isolate Cala2.
(h–j) GFP-tm-KKXX plants inoculated with the virulent pathogen P. parasitica isolate Noks1. (j) Accumulation of ER membrane around the haustorium. This
image is a single optical section focused in the mid-plane of the haustorium. Top left micrograph shows the differential interference contrast (DIC) image of the
infection site shown in the main panel. h, haustorium.
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 33, 775–792
788 Daigo Takemoto et al.
Figure 7. Preferential localization of Golgi stacks around the penetration site of oomycete pathogens.
Images were collected 6 h after inoculation. Asterisks indicate the pathogen penetration sites. The junctions of epidermal cells are outlined by dashed lines.
Except for (f), images of Green Fluorescent Protein (GFP) fluorescence are projections of optical sections taken at 1-mm intervals from the outer epidermal wall
through to immediately above the cortical cytoplasm adjacent to the inner periclinal wall of the epidermal cell. Bar ¼ 10 mm.
(a–c) STtmd-GFP plants inoculated with the non-pathogen, Phytophthora sojae. In some cases, the rapid movement of the Golgi stacks during image capture
appears as weak trailing signals (indicated by arrowheads). (c) Changes in the accumulation pattern of Golgi stacks at the same penetration site. Images were
taken at 15-min intervals.
(d–f) STtmd-GFP plants inoculated with the avirulent pathogen Peronospora parasitica isolate Cala2. (f) Accumulation of the Golgi stacks around the haustorium
of Cala2. The image is a single optical section focused in the mid-plane of the haustorium. Lower left insert shows the differential interference contrast (DIC)
image of the infection site shown in the main panel. h, haustorium.
(g–i) STtmd-GFP plants inoculated with the virulent pathogen P. parasitica isolate Noks1.
Cytoplasmic aggregation is induced in non-host, incompatible and compatible interactions and is one of the earliest responses to pathogen attack. In many cases,
cytoplasmic aggregation and the associated deposition
of wall material and toxins may be sufficient to stop infec-
tion. Isolation of mutants with enhanced disease susceptibility (e.g. eds, pad, nim1/npri) indicated that even
susceptible plants possess a level of basal resistance able
to reduce disease severity (Glazebrook et al., 1996). Donofrio and Delaney (2001) investigated callose apposition for
ß Blackwell Publishing Ltd, The Plant Journal, (2003), 33, 775–792
Arabidopsis response to oomycete pathogens
789
Figure 8. Schematic diagram summarizing the interactions between Arabidopsis and the three oomycete pathogens. The non-pathogen, Phytophthora sojae, is
unable to penetrate through the epidermal cell layer, suggesting that there is effective non-host resistance at this stage. Both avirulent and virulent Peronospora
parasitica isolates can form haustoria in the epidermal cells without induction of plant cell death and can reach the mesophyll cell layer. In the incompatible
interaction, some haustoria are formed in the mesophyll cells; however, plant cell death occurs in the mesophyll cell layer and further growth of the pathogen is
inhibited. In the compatible interaction, numerous haustoria are formed in mesophyll cells. No plant cell death is observed and the virulent pathogen is able to
successfully colonize the cotyledon. HR, hypersensitive response.
wild type and systemic-acquired resistance (SAR) defective
(NahG and nim-1) Arabidopsis following infection by
P. parasitica. The NahG and nim-1 plants showed enhanced
disease susceptibility and a lower degree of accumulated
callose around haustoria compared with wild-type plants.
This circumstantial evidence suggests that localized accumulation of plant material may also be involved in basal
resistance. Because of the central role of cytoplasmic
aggregation in plant defense at various levels, it will be
important to determine in greater detail the molecular basis
of both its induction and formation. Further studies of the
response of GFP-tagged cell components of Arabidopsis to
a range of pathogens are likely to provide much new and
valuable information in this area.
Experimental procedures
Plants and growth conditions
The transgenic A. thaliana line containing GFP-TUA6 (GFP-tagged
alpha-tubulin) (Ueda et al., 1999) was obtained from Dr T. Hashimoto (Nara Institute of Science and Technology, Nara, Japan).
Arabidopsis containing STtmd-GFP (sialyltransferase short cytoplasmic tail and single transmembrane domain conjugated with
GFP) (Boevink et al., 1998) was obtained from Dr C. Hawes (Oxford
Brookes University, Oxford, UK). Generation of Arabidopsis containing GFP-tm-KKXX (GFP with Cf-9 transmembrane domain and
cytosolic tail with C-terminal dilysine motif that confers ER localization) was described previously (Benghezal et al., 2000). All
transgenic Arabidopsis plants were derived from the Columbia
(Col-0) ecotype. Arabidopsis plants were grown at 218C with 16 h
of light (50 mmol of photons m2 sec1) per day before pathogen
inoculation. To maintain the P. parasitica inoculum, Arabidopsis
plants were grown on soil (1 : 1 : 1 mixture of composted soil : sand : peat). For the inoculation assay, seeds of Arabidopsis were
surface sterilized and grown on agar plates containing 1 Muraß Blackwell Publishing Ltd, The Plant Journal, (2003), 33, 775–792
shige and Skoog salts (Murashige and Skoog, 1962), 2% (w/v)
sucrose, 1 vitamin mix (1000 vitamin mix contained 0.3% (w/
v) thiamine hydrochloride, 0.5% (w/v) nicotinic acid, and 0.05% (w/
v) pyridoxine hydrochloride) and 0.4% (w/v) Agargel (Sigma).
Construction of GFP-hTalin fusions and Arabidopsis
transformation
The ClaI-EcoRI GFP cassette lacking the stop codon was obtained
by PCR using pCBJ4 (Benghezal et al., 2000) and two oligonucleotides 50 -ATCGATGTCTAGAGGAGAAGAACTT-30 and 50 -GAATTCTTTGTATAGTTCATCCAT-30 . The EcoRI-BamHI talin actin-binding
domain (McCann and Craig, 1997) cassette with 5 Gly-Ala spacer
at the N-terminus was amplified by PCR using KIAA1027, containing human talin cDNA (provided by Kazusa DNA Research
Institute) and two oligonucleotides, 50 -GAATTCGGCGCTGGGGCAGGTGCCGGAGCTGGAGCGAACTTTGAG-30 and 50 -GGATCCTTAGTGCTCATCTCGAAG-30 . The amplified fragments were
cloned into the EcoRV site of Bluescript SK- (Stratagene) using
the TA cloning technique. The PCR products were verified by
sequencing with ABI Prism BigDye Terminator Cycle Sequencing
kits (Perkin-Elmer) according to the manufacturer’s instructions.
Both ClaI-EcoRI GFP and EcoRI-BamHI talin actin-binding domain
cassettes were ligated into a pUC19 vector containing the cauliflower mosaic virus (CaMV) 35S promoter and the tobacco mosaic
virus (TMV) omega leader sequence at the PstI-SalI site to generate
35S-GFP-hTalin/pUC19. The 35S-GFP-hTalin cassette was excised
from 35S-GFP-hTalin/pUC19 with HindIII and BamHI. The CaMV
35S terminator sequence was excised from CaMV-GR (Schena
et al., 1991) using BamHI and SacI. The 35S-GFP-hTalin cassette
and 35S terminator were cloned into HindIII-SacI digested
pSLJ7292 (described in http://www.jic.bbsrc.ac.uk/sainsbury-lab/
jonathan-jones/plasmid-list/plasmid.htm) to generate pCBJ-GFPhTalin.
Plant transformation was performed by the floral dip method
(Clough and Bent, 1998) using Agrobacterium tumefaciens strain
AGL-1 (Lazo et al., 1991), containing pCBJ-GFP-hTalin. Transformed seedlings were selected on agar plates (see Plants and
growth conditions section) containing 50 mg ml1 kanamycin, and
GFP fluorescence monitored under a fluorescence stereo microscope (model MZ FLIII, Leica, Germany).
790 Daigo Takemoto et al.
Plant inoculations
Inoculation of Arabidopsis plants with the P. parasitica isolates
and P. sojae was performed on plants grown for 10–14 days on
agar plates. Peronospora parasitica isolates Cala2 and Noks1 were
obtained via Dr D. Cahill (Deakin University, Geelong, Australia)
from Dr E. Holub (Horticulture Research International, UK) and
maintained by weekly subculturing on susceptible recipient plants
as described previously (Dangl et al., 1992). Peronospora parasitica conidia (sporangia) were washed from heavily sporulating
Arabidopsis leaves with water, collected by centrifugation at 350 g
for 5 min and re-suspended in water. For the P. parasitica inoculations, 1–2 ml drops of a suspension of 0.5–1 105 conidia ml1 in
water were placed on each cotyledon. Inoculated leaves were kept
at 168C with continuous light (50 mmol of photons m2 sec1).
Phytophthora sojae (H1169) was maintained on nutrient agar
plates (10% (v/v) cleared V8 juice, 0.002% (w/v) b-sitosterol,
0.01% (w/v) CaCO3 and 1.7% (w/v) Bacto agar) at 258C and subcultured 3 days before use. For zoospore production, 3-day-old V8
agar plates with P. sojae mycelium were washed with water every
hour over 5 h and left in a final wash overnight at 188C. Released
zoospores were collected by centrifugation at 350 g for 5 min and
re-suspended in water. For the P. sojae inoculations, 1–2 ml drops
of a suspension of 1 104 spores ml1 in water were placed on
each cotyledon. Inoculated leaves were kept at 228C with continuous light (50 mmol of photons m2 sec1).
Lactophenol trypan blue staining and light microscopy
To monitor the progression of oomycete infection and plant cell
death, infected leaves were stained as described by Koch and
Slusarenko (1990) with minor modifications. Infected leaves were
cleared in methanol for more than 24 h and boiled for 3 min in
lactophenol trypan blue stain (10 ml H2O, 10 ml of lactic acid,
10 ml of glycerol, 10 g of phenol, and 10 mg of trypan blue). After
the leaves had cooled to room temperature for 1 h, the stain was
replaced with 1 g ml1 chloral hydrate. Stained leaves were decolorized overnight and viewed using an Axioplan universal microscope (Zeiss, Germany).
Confocal laser scanning microscopy
Confocal fluorescence and concurrent differential interference
contrast (DIC) images were recorded on a TCS SP2 confocal
system (Leica, Germany) with a 63 NA 1.2 water-immersion
lens. Whole cotyledons were removed from Arabidopsis seedlings
6–36 h after inoculation with one of the three oomycete pathogens
and mounted between glass coverslips, with the inoculated surface facing the objective lens. A krypton–argon laser was used as
excitation source at 488 nm, and GFP fluorescence was recorded
between 495 and 520 nm. Observation of GFP fluorescence was
restricted to the epidermal cells because image quality was greatly
reduced in the underlying mesophyll layer, due at least in part to
the increased depth in the tissue. Except where stated otherwise,
images of GFP fluorescence shown in Figures 3–7 are projections
of optical sections taken at 1-mm intervals from the outer epidermal
wall through to immediately above the cortical cytoplasm, adjacent to the inner periclinal wall of the epidermal cell. The images
were stored as TIF files and processed with Adobe Photoshop 4.0
software (Adobe Systems Inc., USA).
Acknowledgements
We thank Drs Takeshi Hashimoto (Nara Institute of Science and
Technology, Japan) for providing GFP-TUA6, Chris Hawes (Oxford
Brookes University, UK) for STtmd-GFP plants, and Eric Holub
(Horticulture Research International, UK) and David Cahill (Deakin
University, Australia) for P. parasitica isolates. We also thank
Kazusa DNA Research Institute for providing plasmid KIAA1027,
containing human talin cDNA, Professor Jonathan Jones at the
Sainsbury Laboratory, John Innes Centre (Norwich, UK) for
pSLJ7292, and Daryl Webb for technical assistance with confocal
laser microscopy.
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