Distinct heat-shock element arrangements that mediate the heatshock, but not the late-embryogenesis induction of smallheat-shock proteins, correlate with promoter activationin root-knot nematode feeding cells

Marta Barcala Æ Alejandra Garcıa Æ Pilar Cubas Æ Concepcion Almoguera ÆJuan Jordano Æ Carmen Fenoll Æ Carolina Escobar

Received: 19 July 2007 / Accepted: 2 November 2007 / Published online: 28 November 2007

� Springer Science+Business Media B.V. 2007

Abstract Genes coding small heat-shock proteins

(sHSPs) show distinct behaviours with respect to environ-

mental and developmental signals. Their transcriptional

regulation depends on particular combinations of heat

stress cis-elements (heat-shock elements; HSEs) but many

aspects regarding their regulation remain unclear. Cyst and

root-knot nematodes induce, in the roots of infected plants,

the differentiation of special feeding cells with high met-

abolic activity (syncytia and giant cells, respectively), a

process accompanied by extensive gene expression chan-

ges. The Hahsp17.7G4 (G4) promoter was active in giant

cells and its HSE arrangements were crucial for this acti-

vation. In the present work, we provide further basis to

associate giant cell expression with the heat-shock

response of this gene class, by analysing additional pro-

moters. The Hahsp17.6G1 (G1) promoter, not induced by

heat shock, was silent in giant cells, while Hahsp18.6G2

(G2), which responds to heat shock, was specifically

induced in giant cells. In addition, a mutated Hahsp17.7G4

promoter version (G4MutP) with a strong heat-shock

induction was also induced in giant cells. The responses of

the different promoters correlated with distinct HSE con-

figurations, which might have implications on differential

trans-activation. Furthermore, the shortest giant cell and

heat-shock-inducible sHSP promoter version analysed in

tobacco (-83pb Hahsp17.7G4) fully maintained its

expression profile in Arabidopsis. Cyst nematodes did not

induce the Hahsp17.7G4 promoter, revealing additional

specificity in the nematode response. These findings, toge-

ther with the fact that the class I sHSP products of

endogenous genes accumulated specifically in tobacco giant

cells, support the idea that these nematode-induced giant

cells represent a transcriptional state very similar to that

produced by heat shock regarding this class of genes. The

high metabolic rate of giant cells may result in unfolded

proteins requiring class I sHSPs as chaperones, which might,

somehow, mimic heat-shock and/or other stress responses.

Keywords Heat-shock cis-element � Heat-shock factor �Late embryogenesis � Nematode � Giant cell �Promoter regulation

Introduction

Heat-shock proteins (HSPs) are ubiquitous proteins with a

great abundance and diversity in plants. They have been

divided into six different classes on the basis of their

subcellular localisation, immunological cross-reactivity

and sequence similarity (Waters et al. 1996; Scharf et al.

2001). Their role during heat shock has been often studied

(Sun et al. 2002; Yu et al. 2005); small heat-shock proteins

(sHSPs) form aggregates with other sHSPs and different

protein substrates, through which they act as molecular

chaperones (Lee et al. 1997; Heckathorn et al. 1998; Low

et al. 2000). sHSPs are not only induced after heat shock but

M. Barcala � A. Garcıa � C. Fenoll � C. Escobar (&)

Facultad de Ciencias del Medio Ambiente, Universidad de

Castilla-La Mancha, Campus de la Real Fabrica de Armas,

45071 Toledo, Spain

e-mail: carolina.escobar@uclm.es

P. Cubas

Centro Nacional de Biotecnologıa, Departamento de Genetica

Molecular de Plantas, Campus Universidad Autonoma de

Madrid, 28049 Cantoblanco, Madrid, Spain

C. Almoguera � J. Jordano

Instituto de Recursos Naturales y Agrobiologıa, Consejo

Superior de Investigaciones Cientıficas, Apartado 1052,

41080 Seville, Spain

123

Plant Mol Biol (2008) 66:151–164

DOI 10.1007/s11103-007-9259-3

also by other abiotic stimuli, such as cold, water deficit,

heavy metals, ozone, UV radiation and c-irradiation

(Almoguera et al. 1993; Banzet et al. 1998; Hong and

Vierling 2001; Sun et al. 2002). Furthermore, sHSP accu-

mulation is developmentally regulated, notably during seed

and pollen development (Coca et al. 1994; Wehmeyer and

Vierling 2000; Volkov et al. 2005) and fruit maturation

(Fray et al. 1990; Dure 1993; Low et al. 2000). The

expression of sHSP genes in these situations is regulated

mainly by cis-elements, described as heat-shock elements

(HSEs) present in their promoters. The HSEs are arranged in

clusters of three or more inverted repeats of the sequence 50-nGAAn-30, to which, heat-shock factors (HSFs) bind to

control transcription (Carranco et al. 1997; Nover et al.

2001). The role of these elements in sHSP genes expression

in response to different stimuli is beginning to be understood

and different combinations and/or particular sequences in

the HSE cores are crucial for their regulation under different

circumstances, such as embryogenesis or heat shock (Coca

et al. 1996; Almoguera et al. 1998, 2002b). In addition,

plant HSFs belong to gene families with high complexity

and more than 21 different HSFs have been described so far

(Nover et al. 1996, 2001). Their functional specificity has

been demonstrated only in a few cases. These include par-

ticular HSFs involved in transcriptional activation during

embryogenesis (Almoguera et al. 2002b), in apoptotic

responses (Yamanouchi et al. 2002), in early oxidative

stress responses to H2O2 (Davletova et al. 2005) or in the

heat-stress response (Mishra et al. 2002).

We found that the promoter of Hahsp17.7G4 (G4 in this

work), a sunflower gene that codes an sHSP expressed during

seed development and in response to water stress or heat

shock (Coca et al. 1996; Almoguera et al. 1998), is also

active in giant cells (GCs) (Escobar et al. 2003). So far, this

is the only sHSP promoter reported to respond to biotic

interactions, in particular to nematode infection (Meloido-

gyne spp.). A short version of the G4 gene promoter

(-83G4) includes only one HSE array (HSE I), which is

crucial for the G4 activation after root-knot nematode

infection in tobacco (Escobar et al. 2003). This minimal

promoter version is also up-regulated after heat shock and, to

a lesser extent, during late embryogenesis (Coca et al. 1996).

Cyst and root-knot nematodes are obligate sedentary

endoparasites which need plants for their reproduction. Upon

infection, they induce the transformation of a selected set of

root cells into a feeding site that contains a few highly spe-

cialised cells (syncytia for cyst nematodes and GCs for root-

knot nematodes) on which they feed. Despite accomplishing

similar functions, the ontogeny of both nematode-induced

cell types is very different. GCs are the result of a re-

differentiation process, linked to dramatic changes in gene

expression (Gheysen and Fenoll 2002; Ramsay et al. 2004;

Bar-Or et al. 2005; Jammes et al. 2005; Wieczorek et al.

2006). GC differentiation transforms a normal root xylem-

associated cell into a large, multinucleate cell with enlarged

nucleoli, whose dense cytoplasm is rich in mitochondria,

plastids, ribosomes, rough and smooth endoplasmic reticulum

and crystalline, electron-dense inclusions (possibly proteina-

ceous) frequently associated to spiny vesicles (Paulson and

Webster 1970; Bleve-Zacheo and Melillo 1997). These

characteristics are typical of a cell with very active protein

synthesis, possibly needed to attend the nematode demand as

its only nutrient source during the weeks needed for the

nematode to complete its life cycle (Grundler and Bockenhoff

1997). In this cellular context, a transcriptional state that

favours chaperone (such as sHSP) accumulation might par-

tially mimic the response to heat shock.

To obtain a deeper insight into the HSE-mediated reg-

ulation of sHSPs promoters after nematode infection in

comparison to that of the heat-shock and embryogenesis

responses, we have studied the activity in GCs of addi-

tional sHSP gene promoters, Hahsp17.6G2 (G2) and

Hahsp18.6G1 (G1), closely related to G4, that differ in

their HSE array structures and in their regulation during

late embryogenesis and heat shock (Coca et al. 1996;

Carranco et al. 1997, 1999; Almoguera et al. 2002b).

Our results show that the G2 promoter was activated after

nematode infection in tobacco transgenic plants. In contrast,

G1 was not induced after nematode infection. We have also

found that a mutated version of the G4 promoter (G4MutP),

which shows a greatly reduced late embryogenesis response

(Almoguera et al. 2002b), was active in GCs. Taken together,

our results indicate that transcriptional activation in response

to root-knot nematode infection rely on mechanisms different

from those operating during late embryogenesis but is similar

to those of the heat-shock response and that distinct HSE

array structures may be related to this behaviour. We have

also found that the G4 promoter is specific for root-knot

nematodes, since it is not induced in syncytia formed by the

cyst nematode Globodera tabacum. We have consistently

shown that the class I sHSPs encoded by endogenous genes

accumulate differentially in tobacco GCs, parallelling the

nematode-induction of the heterologous class I sHSP pro-

moters. Therefore, we suggest for these proteins a putative

role during GC formation and/or maintenance, perhaps sim-

ilar to their role during the heat-shock response.

Materials and methods

Infections with nematodes

Meloidogyne incognita infections

Transgenic tobacco seeds from T0 plants containing

different sHSP promoter constructs (Hahsp17.7G4,

152 Plant Mol Biol (2008) 66:151–164

123

Hahsp17.6G1 and Hahsp18.6G2) were provided by

Dr. J. Jordano (Coca et al. 1996; Carranco et al. 1999;

Almoguera et al. 2002b). The seeds were surface-sterilised

and selected in MS medium (Murashige and Skoog 1962)

supplemented with 50 lg/ll of kanamycin. For nematode

infection, resistant tobacco and Arabidopsis homozygous

plants were transferred to Gamborg B5 plates (as in

Escobar et al. 2003) and grown at 24–25�C, 70% humidity

in long day photoperiods. We tested at least five indepen-

dent lines per construct for the analyses in tobacco and four

lines in Arabidopsis.

The M. incognita MIK population (Escobar et al. 2003)

was maintained in an in vitro culture on cucumber roots.

The plates were kept in darkness at 28�C and 60%

humidity. Mature egg masses were collected from the

cucumber roots and kept in sterile tap water in the previous

conditions. After 3–4 days, freshly hatched J2 were used to

inoculate either tobacco or Arabidopsis roots. An average

of 10–12 J2s were added per root tip. Gall formation was

carefully followed by monitoring the infection every 24 h

under a Leica MZ6 microscope. Each newly formed gall

was single-labelled. Thus, days post-infection (dpi) were

measured with an error of less than ±24 h. A minimum of

30–80 galls were scored for each independent line tested.

Globodera tabacum infections

Tobacco plants were selected in MS medium (Murashige

and Skoog 1962) supplemented with kanamycin (50 lg/ll)

for 1 week. After this period, the seedlings were transferred

either to 50-ml conical tubes filled with 16 ml of sand

under sterile conditions and kept in a growth chamber at

24�C in long day periods or to Petri dishes with Gamborg

B5 medium, as previously described (Escobar et al. 2003).

Plants growing in conical tubes were watered twice a week

alternately with either water or liquid MS medium sup-

plemented with vitamins. The tubes were covered with

Parafilm1 to limit water loss. The cysts were sterilised as

in Heungens et al. (1996) with minor modifications. Plants

in tubes were infected with 150 Globodera tabacum J2 and

those grown in Petri dishes with 250 J2 per plate. On

average, 35 cysts were tested in each line.

Arabidopsis transformation and characterisation

Arabidopsis thaliana Col-0 plants were Agrobacterium

tumefaciens-infiltrated with the construct -83G4::GUS, as

described in Coca et al. (1996) using the floral dip method

(Clough and Bent 1998). Transformants were selected by

consecutive cycles of kanamycin selection and self-prog-

eny production. Only the lines with T1 and T2 progeny

showing a classical segregation rate for one locus were

considered as homozygous lines carrying a single copy

insertion. Seeds from T3 plants were used for nematode

infection and plant characterisation. To achieve embryo-

genesis histochemical analysis, every newly opened flower

was labelled in order to set its age. For this analysis, the

plants were grown in soil at 23–24�C and 70% humidity

and with a long day photoperiod.

Histochemical GUS assay

Histochemical assays of GUS activity were performed

mainly as described by Escobar et al. (2003). To avoid

temperature variations that could promote sHSP induction,

histochemical analyses were performed under the same

growth conditions in darkness. For heat-shock experiments,

the plants were placed at 42�C for 3 h and were allowed to

recover at growth conditions for another 3 h before car-

rying out the histochemical analysis. For histological GUS

localisation, positive galls were fixed and embedded in

Araldyte (TAAB, Aldermaston, UK) as in Escobar et al.

(2003). Micrographs from semi-thin sections (6–7-lm

thick) were taken with either Nikon 90i or Leica DM IRB

stereomicroscopes.

Western analysis

Protein extracts were obtained from hand-dissected tobacco

galls or uninfected or heat-shocked roots. The total protein

was extracted in 19 SDS buffer containing 62.5 mM Tris

HCl pH 6.8, SDS 1% (w/v), 2-mercaptoethanol 2.5% (v/v)

and a protease inhibitor cocktail (according to the manu-

facturer’s instructions; Sigma-Aldrich, St. Louis, Missouri).

Protein extracts were concentrated using Microcon YM-10

centrifugal filters (Millipore, Bedford, Massachusetts) and

quantified by Bradford assay. Samples were separated by

sodium dodecyl sulfate polyacrylamide gel electrophoresis

(SDS-PAGE) in a 12.5% polyacrylamide gel and transferred

to a PVDF membrane (Hybond-P, Amersham Biosciences

Technologies, UK) with a semi-dry transfer cell (Bio-Rad,

Hercules, California). PVDF membrane background binding

was blocked with 5% dry milk in Tris-buffered saline with

Tween (TBST) and incubated with either anti-class I sHSP

antibodies (Almoguera et al. 1993) or the respective pre-

immune sera, diluted 1:1000 in 5% dry milk. Anti-rabbit

IgG-AP-conjugated antibodies (Sigma-Aldrich, St. Louis,

Missouri) diluted 1:5000 were used as secondary antibodies,

followed by colorimetric detection with a nitrotetrazolium

blue/5-bromo-4-chloro-3-indolyl phosphate solution (NBT/

BCIP; Sigma-Aldrich, St. Louis, Missouri and Roche

Diagnostics GmbH, Mannheim, Germany).

Plant Mol Biol (2008) 66:151–164 153

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Immunolocalisation assay

Tobacco galls and uninfected control roots were sectioned

and incubated in freshly made fixative (4% (w/v) formal-

dehyde, 0.1 M PIPES, 0.25% (v/v) glutaraldehyde, 1 mM

DTT, 0.1% (v/v) Tween-20, 0.1% (v/v) Triton X-100)

twice for 10 min under vacuum. The fixative was replaced

with fresh solution and the samples were kept at 4�C

overnight. Then, they were dehydrated in ethanol (10%,

30%, 60%, 70%, 85%, 90% and 100% in water) at 4�C. At

the 90% and 100% ethanol steps, 0.1% (w/v) eosin Y was

added to stain the tissues. A complete dehydration was

carried out by incubating the samples in a 3:1, 1:1 and 1:3

series of ethanol (Histoclear; National Diagnostics,

Atlanta, Georgia). Finally, the samples were embedded in

Paramat extra Pastillated Gurr (BDH Lab Supplies, Poole,

UK).

Paraffin blocks were sectioned with a Leica RM 2125

rotary microtome. The sections (10-lm thick) were

mounted on poly-lysine-coated slides (Polysine, Menzel-

Glaser, KG, Germany). After de-waxing and rehydration,

the sections were incubated in primary antibody (1:200, at

4�C overnight), followed by the secondary antibody pre-

viously described for Western blot (1:200 for 45 min at

room temperature), both diluted in TBST.

Statistical analysis of data

The value distributions of the percentages of GUS-stained

galls were compared between plants carrying different

promoter::GUS fusions by the non-parametric test of U

Mann-Whitney and using the Statistical Package for the

Social Sciences (SPSS Inc., Chicago, Illinois). Pairwise

comparisons between all possible groups were performed

to analyse the differences of the effect of the chimaeric

gene on GUS expression. The level of significance was set

to P \ 0.05.

Results

Three differentially regulated sHSP gene promoters

show differential nematode response in tobacco

To gain insight into the regulation of sHSP genes through

their HSEs in GCs, we compared the promoter structure

and response of G4 with those of two sHSP genes, G1 and

G2, that code for very similar proteins but differ in their

expression patterns. We also analysed a mutant version of

the G4 promoter called the ‘‘perfect HSE mutant’’ G4MutP,

as it has a higher number of ‘‘perfect’’ core HSE repeats

(exactly matching the consensus sequence (50-nGAAn-30).

These changes in promoter structure are due to three point

mutations at positions -86 (TCAA changed by agAA),

-108 (TTT changed by TTc) and -116 (GGAA changed

by aGAA; Fig. 1A), which improves HSF binding and

maintains the heat-shock activation but reduces the late-

embryogenesis response (Almoguera et al. 2002b).

A detailed analysis of the promoter structure reveals

that, similarly to the G4 promoter (Escobar et al. 2003), the

G2 promoter has two HSE arrays, one proximal and one

distal (HSE I and HSE II, respectively; Fig. 1A). In con-

trast, G1 carries only one HSE array and in a distal position

(Fig. 1A). In addition, the G4 promoter contains three HSE

head–tail motifs (nGAAn; nTTCn), which include two

active head modules, one inactive head module and three

active tail modules in HSE I (ThTHTH; Fig. 1A). G2

carries fewer HSE head–tail motif combinations (THTHt)

in HSE I (Fig. 1A). In contrast, G1 contains a higher

number of HSE motifs as compared to G2. However, G1

HSE motifs present a distinct configuration (THHHtH;

Fig. 1A), as three adjacent active head modules are local-

ised in between an active and an inactive tail module

(Fig. 1A). Moreover, in G1, the distance between the HSE

and the TATA box is much longer than that of HSE I in G2

or G4 (49 bp as compared to 12 bp in G2 and 21 bp in G4;

Fig. 1A). Additionally, several putative CAAT boxes were

identified immediately upstream and in between the HSE

arrays of G2 and G4, at positions -177 and -72 bp for G4

and -141, -127 and -84 bp for G2. In contrast, a single

CAAT box was identified in the G1 promoter downstream

of the HSE array at position -67 pb.

Transgenic tobacco plants carrying translational fusions

of G2 and G1 promoters to the b-glucuronidase (GUS)

reporter gene (G2::GUS and G1::GUS) were infected with

Meloidogyne incognita and analysed 8–15 dpi, a stage

when GCs have a very active metabolism and during which

a high activity of the G4 promoter was previously reported

(Escobar et al. 2003).

Plants carrying the G2 promoter showed a blue signal

centred in galls (Fig. 2A), which corresponded to a blue

precipitate strongly concentrated in GCs, as shown in semi-

thin sections (Fig. 2F). A faint patchy expression was

occasionally detected along the root and at the root tip

(data not shown) in all of the tested lines. In contrast, G1

nematode inducibility was undetectable, as shown in galls

and semi-thin sections of galls (Fig. 2B, G). The mutated

version of the G4 promoter, G4MutP, containing point

mutations in the HSE arrays that convert inactive to active

modules (Fig. 1A; Nover et al. 2001), was active in galls,

with the signal centred in GCs (Fig. 2C, H).

Analysis of the frequency of blue gall occurrence con-

firmed that the G2 promoter responds to nematode

infection similarly to G4 and that G1 is not responsive to

this stimulus (Fig. 1B). No significant differences were

154 Plant Mol Biol (2008) 66:151–164

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Fig. 1 Small heat-shock protein (sHSP) promoter structures and

frequency of blue galls. (A) Structure of the different sHSP promoters

analysed based on Scharf et al. (2001). Core heat-shock elements

(HSEs) are referred as active head (nGAAn, nGAnn or nGnAn) or tail

(nTTCn, nnTCn or nTnCn) modules and are represented as white

boxes ( , ). Inactive head or tail modules lack G or C residues

and/or the other two core nucleotides (AA or TT) and are indicated

with grey boxes. HSE arrays are named I and II and the TATA box is

represented with a striped box. Numbers under promoter diagrams

indicate the position of the initial and last nucleotide of either the HSEarrays or the TATA box to the transcriptional starting point. The

numbers between slashes represent the distance to adjacent promoter

elements. (B) The histograms represent the frequency of blue galls

(10 dpi) of the different tobacco transformants. The data are means of

at least five independent lines per construct. The data from four

independent experiments per line were pooled. The table shows

activation of the G4, G2, G1 and G4MutP promoters after nematode

infection, heat shock and during late embryogenesis

Plant Mol Biol (2008) 66:151–164 155

123

found between G2 and G4 (P = 0.245), while the non-

heat-inducible G1 promoter showed differences to G2, G4

and G4MutP (P = 0.014). In addition, the mutations

introduced in G4MutP did not affect G4 promoter response

in GCs (Fig. 1B). Consequently, no significant differences

were found among G4MutP as compared to either G2 or

G4 (P = 1 and P = 0.149, respectively).

The results confirm that nematode responsiveness is not

a general feature of all sHSP promoters but a specific

characteristic of particular sHSP genes, such as G4,

G4MutP and G2, that are induced during heat shock but not

during embryogenesis (Coca et al. 1996; Almoguera et al.

2002b). Interestingly, promoter structural characteristics

correlate with the heat-shock and nematode inducibility of

the promoters (see discussion; Fig. 1A).

The G4-based constructs described in this work and in

Escobar et al. (2003) were translational fusions containing

the first 84 nucleotides of the G4 protein-coding sequence

fused in-frame to GUS (Almoguera et al. 2002a). To

investigate putative effects of the G4 coding region in the

translational fusions, we tested a transcriptional fusion of

the G4 promoter where these sequences were eliminated

(Bgl II; Fig. 1A; Almoguera et al. 2002a). From the anal-

ysis of nine independent transformants, we inferred that the

Fig. 2 GUS activity of feeding

sites from tobacco transgenic

plants carrying different

promoter::GUS constructs after

the infection of Globoderatabacum and Meloidogyneincognita (15–20 dpi and

10 dpi, respectively). (A, F) G2.

(B, G) G1. (C, H) G4MutP.

(D, I) Bgl II. The panels show

GUS activity as a blue colour in

whole-mount galls and as a

fuchsia precipitate in their

corresponding micrograph

sections taken under a dark

field. (E) GUS assay carried out

on cysts from G4 tobacco plants

after infection with Globoderatabacum. The scale bars in

panels A, B, D represent

250 lm; C 500 lm; E 200 lm;

F–H 50 lm; and I 100 lm.

Giant cells (GCs) are indicated

with asterisks (*) and nematode

as N

156 Plant Mol Biol (2008) 66:151–164

123

28 extra amino acids in G4 as compared to Bgl II were not

affecting the GUS expression in GCs (Fig. 1B, 2D, I).

Hence, the translational fusions used were clear indicators

of the promoter activity.

The G4 promoter is activated by root-knot

but not by cyst nematodes

It has been previously reported that some plant genes and/or

promoters are differentially regulated in the feeding cells

induced by cyst and root-knot nematodes (Vercauteren et al.

1998; Gheysen and Fenoll 2002; Mitchum et al. 2004).

Therefore, we decided to analyse the Meloidogyne spp.-

inducible G4 promoter upon infection with the cyst nema-

tode Globodera tabacum. We selected those transgenic lines

that showed a higher induction upon M. incognita infection;

one line contained the G4::GUS construct and two lines

had the minimal GC-responsive HSE promoter elements

(-83G4; Escobar et al. 2003). Syncytia at 10–20 dpi did not

show any GUS activity, either in in vitro-grown plants or in

hydroponic sand cultures, for either promoter construct

(Fig. 2E). Therefore, the G4 promoter responds to root-knot

but not to cyst nematodes.

Class I low-molecular-weight heat-shock proteins

accumulate in giant cells

In order to investigate whether G4 and G2 promoter acti-

vation in the transgenic lines is parallelled by the tobacco

endogenous genes and leads to the accumulation of sHSPs

in GCs, Western-blot assays were carried out in protein

extracts from tobacco galls and uninfected roots. Using an

anti-sHSP antibody that specifically recognises class I

sHSPs (Almoguera et al. 1993), a very faint band of the

expected size (about 18 kDa) could be detected in gall

extracts (Fig. 3A) but not in control uninfected root tissue

(Fig. 3A). Protein extracts from tobacco roots subjected to

heat-shock treatment and sunflower embryos were also

loaded as positive controls (Fig. 3A). To further confirm

that sHSPs were localised in gall cells, we performed im-

munolocalisation assays in gall sections and compared

them to control root sections using the same detection

system. A clear purple precipitate could be observed,

which was highly restricted to GCs (Fig. 3B), while the

surrounding tissues (Fig. 3B) control uninfected root sec-

tions (not shown), as well as gall sections incubated with

preimmune antisera did not show any signal (Fig. 3C). This

result indicates that the endogenous sHSP genes are highly

expressed at the protein level in GCs, thus, opening the

possibility of a biologically meaningful role for sHSPs in

GCs development and/or physiology.

The -83bp G4 promoter is also induced in giant cells

and during heat shock in transgenic Arabidopsis

Helianthus annum is a recalcitrant plant for transformation

(Lewi et al. 2006). Therefore, in order to assess whether

the nematode activation of sHSPs is conserved in plant

species other than tobacco, we transformed Arabidopsis. A

comparable response in Arabidopsis could simplify further

analysis by exploiting molecular genetics tools available in

Fig. 3 Abundance of class I sHSP in galls. (A) Western blot carried

out with anti-sHSP class I antibodies. Bands of approximately 18 kDa

appear in lanes loaded with total protein from sunflower embryos

(lane E, 4 lg) and heat-stressed tobacco roots (lane HR, 135 lg) used

as positive controls. A faint band can be detected in tobacco gall

protein extracts (lane G, 135 lg), whereas no signal was detected in

uninfected control root protein extracts (lane CR, 135 lg). (B, C)

Immunolocalisation of class I sHSPs in tobacco galls 10 dpi. Sections

were incubated with either anti-class I sHSP antibody (B) or pre-

immune serum (C). A purple precipitate centred in the GCs (*) is

detected in B. The scale bars represent 50 lm

Plant Mol Biol (2008) 66:151–164 157

123

Arabidopsis and this might help in finding the putative role

of the class I sHSPs and HSFs in GCs. Arabidopsis plants

carrying the minimal nematode-inducible promoter version

-83G4 fused to GUS were characterised after nematode

infection and also during plant development and heat

shock, since no information on the behaviour of the

Helianthus annus G4 promoter in Arabidopsis was avail-

able (Fig. 1A).

Arabidopsis transgenic plants carrying the -83G4::GUS

construct were analysed 7 days after nematode infection,

revealing a clear GUS signal inside galls (Fig. 4A),

although the percentage of positive galls for all tested lines

was reduced (30–50%) as compared to tobacco (Escobar

et al. 2003). Therefore, the nematode inducibility of the

sunflower promoter in tobacco plants was maintained in

Arabidopsis. Consistent with the activity observed in

tobacco (Escobar et al. 2003), semi-thin sections of the

positive galls showed a GUS signal located in GCs

(Fig. 4B), with a faint and diffuse signal in the surrounding

tissues, which might be due to diffusion of the GUS pre-

cipitate. Occasionally, patchy GUS expression in scattered

single cells was observed within the root in both infected

and uninfected plants (Fig. 4C).

During germination, an intense GUS expression signal

was found in cotyledons of seedlings at growth stages 0.5–

0.7 (as defined by Boyes et al. 2001; Fig. 5A). This coty-

ledon pattern disappeared completely by the 1.02–1.04

growth stages and GUS expression became restricted to the

shoot apical meristem (SAM; Fig. 5B). GUS activity could

not be detected any longer during development in any

vegetative tissues (Fig. 5C), apart from some occasional

activity in lateral root initiation points (data not shown). No

signal was detected in developing floral organs (Fig. 5D),

except for a very restricted blue signal at the base of the

siliques, 3–5 days post-anthesis (dpa) that decreased during

silique maturation (Fig. 5D, E). A faint signal was also

detected at the funiculus 1–5 dpa (Fig. 5F). However, no

GUS activity was detected during seed maturation either in

developing embryos or in mature seeds (Fig. 5F–I). In

tobacco, a low level of GUS activity was reported

in -83G4 developing seeds, albeit activity was not

completely abolished as in Arabidopsis (Coca et al. 1996).

We cannot rule out a transcriptional activation in Arabid-

opsis embryos below the detection level of our

histochemical assays or a different post-transcriptional

regulation of the -83G4 gene product in Arabidopsis as

compared to tobacco seeds, as proposed for similar chi-

maeric genes (Prandl et al. 1995).

After heat shock, all tissues of seedlings during germi-

nation responded to the treatment (data not shown;

Fig. 5J). Similarly, adult plants showed a strong blue

staining in the roots, in the leaf vascular tissue and espe-

cially intensely in the stem (Fig. 5K, L). During fruit

maturation, green siliques subjected to heat shock were

completely blue stained (Fig. 5M). As silique maturation

progressed, GUS signal was restricted to the funiculus, the

septum and the base of the silique (Fig. 5N). However, no

GUS signal was observed after heat shock in mature seeds

(Fig. 5N). In accordance with our observations in Arabid-

opsis, adult -83G4 tobacco plants showed a similar

response to heat shock, which was especially intense in

stems (Coca et al. 1996), suggesting a remarkable conser-

vation of the promoter regulation in two heterologous

systems. In this respect, the promoter activity of G1, G2

and G4 during heat-shock in tobacco was in agreement

with their mRNA accumulation in sunflower. In contrast,

the accumulation of G4 mRNAs in sunflower after water

stress did not correspond to its promoter activity in tobacco

(Coca et al. 1996; Carranco et al. 1999; Almoguera et al.

2002a).

Discussion

Many aspects regarding the regulation of sHSPs through

their HSEs remain unknown (Scharf et al. 2001). One

example is the sunflower G4 promoter, expressed in

tobacco GCs induced by root-knot nematodes, whose

expression is mediated by a specific array of HSEs

(Escobar et al. 2003). GC expression of various G4 pro-

moter versions indicated a parallel with their heat-shock-

mediated expression and also with some aspects of their

Fig. 4 Histochemical GUS assay in Arabidopsis plants carrying a -

83G4::GUS fusion, 7 dpi. (A) Gall after M. incognita infection

showing GUS activity as a blue precipitate. (B) Semi-thin section of a

gall. GUS signal, as a fuchsia precipitate, is mainly concentrated in

the GCs. (C) Scattered blue signals in roots. GCs are indicated with

asterisks (*) and nematode as N

158 Plant Mol Biol (2008) 66:151–164

123

late-embryogenesis regulation (Escobar et al. 2003). This

raises the issue of what could be the signals inside GCs that

induce the sHSP promoters and whether they are related to

their regulation during heat-stress and/or late embryogen-

esis. Therefore, we examined in further detail the

transcriptional responses of several sHSP gene promoters

in response to nematode infection. These promoters were

selected because they showed differences in their promoter

structures and in their regulation under heat shock or late

embryogenesis, which may correlate to their responses

after nematode infection.

The regulation of three sHSP promoters in giant cells

differs from their regulation during late embryogenesis

and is associated to different structural features

In a previous work, HSEs were identified as necessary for

the transcriptional activation of G4 in GCs, probably

through HSFs (Escobar et al. 2003) and partial differences

in its regulation after heat shock, embryogenesis and

nematode infection were described.

In the present work, we have studied the nematode-

inducibility of two additional sHSP promoters and one new

version of the G4 promoter. Results were compared to

previous observations of the G4 promoter. The mutant

version of the G4 promoter (G4MutP) showed the same

activation pattern as G4 after nematode infection (Fig. 1, 2)

but a very low activation during late embryogenesis

(Almoguera et al. 2002b). This indicates a clear difference

between GCs and late embryogenesis regarding the G4

promoter activation and suggests that imperfect repeats at

positions -86 and -108 are not crucial for the binding of

putative HSFs and/or other transcriptional regulators that

participate in nematode induction, whereas they were

essential for late embryogenesis activation (Fig 1;

Almoguera et al. 2002b).

The promoters of two additional sHSP genes (G1 and

G2) were tested for nematode-inducibility. These promot-

ers and that of G4 differ in their HSE array structure

(Fig. 1). The G1 promoter was inactive in GCs, as well as

during heat-shock (Figs. 1 and 2; Carranco et al. 1999),

despite its efficient activation in late embryogenesis

(Carranco et al. 1999), probably via HaHSFA9 or its

Fig. 5 Histochemical GUS assay of Arabidopsis transgenic plants

carrying a -83G4::GUS fusion. (A) During germination, GUS

activity is restricted to the cotyledons from stage 0.5 to 0.7 (Boyes

et al. 2001). (B) Stage 1.02 to 1.04 seedlings with GUS expression in

SAM. (C) No GUS staining in adult plants. (D) Flowers and siliques

0–5 dpa. (E) Blue staining at the base of the silique, 7–10 dpa. (F)

Silique 4–5 dpa with faint GUS staining at the funiculus (arrow-heads). (G, H) Siliques 15 and 18 dpa, respectively. (I) Embryos

isolated at 11, 15 and 18 dpa with no GUS staining. After heat-shock

treatment, b-glucuronidase is induced in: (J) Vegetative tissues of

seedlings; (K, L) Adult plants with GUS staining in leaf vascular

tissue and strong signal in stems; (M, N) GUS activity diminishes

through development of heat-stressed siliques from 5 to 9 dpa,

respectively. Scales bars in panel I represents 100 lm; A, B, D–F, J,

L and N 0.5 mm; and C, G, H, K and M 1 mm

Plant Mol Biol (2008) 66:151–164 159

123

orthologous tobacco HSF (Almoguera et al. 2002b). This

demonstrates that the presence of functional HSEs in an

sHSP promoter is not sufficient to confer GCs expression.

In contrast, the G2 promoter responded to nematodes

(Figs. 1 and 2), as well as to heat shock, but its transcripts

did not accumulate in late embryogenesis (Coca et al.

1996; Carranco et al. 1997). These results strongly suggest

that the HSFs involved in the nematode response in

tobacco are able to activate transcription from perfect

(G4MutP) or imperfect (G2, G4) HSEs but only from

promoter-proximal HSE locations, which are shared by the

G2 and G4 promoters but not by G1 (Fig. 1A). Further-

more, HSE II in G4 is dispensable for nematode activation

in tobacco and Arabidopsis (Escobar et al. 2003 and this

work). The heat-shock induction of sunflower sHSP pro-

moters in tobacco also depends on the presence of proximal

HSE, whereas distal HSE contributes to the developmental

regulation (Carranco et al. 1999). These also support a

higher similarity of the sHSP regulation during heat shock

and in GCs as compared to late embryogenesis.

Additionally, the G1 HSE array does not present the

alternate head-tail-head pattern observed in HSE I of G2

and G4, which could constitute a poor combination for

HSF primer binding (see results and Fig. 1). This fact

could explain, at least in part, their different regulation in

GCs and suggests that HSFs and other transcription factors

that participate in G1 promoter activation during late

embryogenesis differ from those involved in G2 and G4

expression in GCs and heat shock. In this respect, the

combination of active and inactive modules, as well as

their position, seem to be important for the proper inter-

action of HSFs and other factors in sHSP promoters (Scharf

et al. 2001; Bharti 2003), as described for the synergistic

effect between tomato HSFA1 and B1, as well as their

cooperation with other activators, such as HAC1 and

ASF1/2 (Bharti et al. 2004). Moreover, the HSE array in

G1 showed a low binding affinity to certain HSFs, such as

hHSF-1 (Carranco et al. 1997). In addition, in G1 the

distance between the HSE and the TATA box is much

longer than that of HSE I in G2 or G4 (Fig. 1). This dif-

ference may be relevant for the activation of G2, G4 and

-83G4 in GCs and not G1, as some HSFs interact directly

with elements of the basal transcription machinery (Reindl

and Schoffl 1998). Additionally, several putative CAAT

boxes were identified immediately upstream and in

between the HSE arrays of G2 and G4, whereas a single

CAAT box was identified in the G1 promoter downstream

of the HSE array. CAAT sequences placed in between and/

or immediately upstream to the HSEs have been described

to act cooperatively with HSE to increase promoter activity

in Gmhsp17.3-B (Rieping and Schoffl 1992). Therefore,

these CAAT boxes might participate in the activation of G2

and G4 after nematode infection but not in G1. All of these

structural differences of G2 and G4 as compared to G1

might be crucial for their differential activation in GCs.

Although the HSE I present in the -83G4 is crucial for

nematode activation in tobacco (Escobar et al. 2003), the

lines tested were translational fusions and included, besides

the promoter region, an additional 163 bp sequence

downstream from the transcription initiation site. A search

for putative cis-elements in the 246 pb that constitute the

minimal nematode-inducible sequence using three different

databases identified several homologies to known regula-

tory sequences from other plant genes (Escobar et al.

2003). Therefore, we cannot exclude the participation of

some of these putative cis-elements in the activation of

-83G4 in Arabidopsis GCs. However, in this work, we

found that a transcriptional fusion of the G4 promoter

where these 163 bp were eliminated (Bgl II; Almoguera

et al. 2002a) maintains its inducibility in tobacco GCs.

Thus, the possibility that the putative cis-elements identi-

fied could play a role in sHSP regulation in GCs is low.

Additionally, no relevant putative cis-elements in the

sequences between HSE I and the TATA box could be

identified in -83G4 (Fig. 1).

The sunflower -83G4 minimal promoter maintains its

regulation in Arabidopsis during development, heat

shock and nematode infection

We also demonstrate that both the nematode and the heat-

shock responses of the minimal promoter version -83G4

are conserved between tobacco and Arabidopsis. This short

promoter version, which does not contain the distal HSE

array (HSE II; Fig. 1), showed induction after heat shock in

vegetative tissues and after nematode infection in tobacco

(Coca et al. 1996; Almoguera et al. 1998; Escobar et al.

2003) and in Arabidopsis (Figs. 4 and 5). With respect to

late embryogenesis expression, the -83G4 promoter was

silent in Arabidopsis (this work) and had a low activity in

tobacco (Escobar et al. 2003). Therefore, the putative HSFs

and, possibly, other transcriptional regulators that must

interact with the G4 HSE I (Fig. 1) in GCs and during heat

shock seem to be functionally conserved in different plant

species.

Particular HSFs might be involved in sHSP gene

expression in giant cells

The data presented in this paper strongly suggest that the

regulation of sHSP promoters through HSEs in GCs

depends on HSFs, combinations of HSFs (Scharf et al.

1998; Rojas et al. 2002; Bharti et al. 2004) or combina-

tions of HSFs with other interacting proteins (Bharti et al.

160 Plant Mol Biol (2008) 66:151–164

123

2004; Diaz-Martin et al. 2005), which are different

from those involved in their regulation during late-

embryogenesis.

Plant HSFs show a high complexity and functional

specificity has been demonstrated in only a few cases

(reviewed by Baniwal et al. 2004). It is possible that still

uncharacterised HSFs might be involved in the nematode

response. The intrinsic characteristics of these HSFs

regarding binding to HSEs would match the observed HSE/

promoter structural requirements for nematode response in

tobacco and Arabidopsis.

Confirmation of the involvement of particular HSFs in

G4 and G2 promoter regulation after nematode infection

via loss-of-function mutants might be precluded by func-

tional redundancy amongst some members of the gene

family and lines carrying loss-of-function alleles for sev-

eral HSF genes might be necessary (Morimoto 1998; Miller

and Mittler 2006), since a master regulator of the heat-

shock response as in tomato (Mishra et al. 2002) has not

yet been identified in Arabidopsis (Reindl et al. 1997;

Wunderlich et al. 2003; Lohmann et al. 2004). GCs do not

seem to show a typical heat-shock stress cellular response.

Thus, it is possible that HSFs similar to HSFA3 from

tomato that could represent a developmentally regulated

HSF with expression only in rapidly dividing cells (Nover

et al. 2001) may be the crucial activators of the sHSP

promoters in GCs, whose differentiation involves repeated

endomitosis cycles (revised in Gheysen and Fenoll 2002).

Additionally, HSFs identified as important for the stress

response in Arabidopsis, such as AtHSF1 and AtHSF3,

might also participate in the activation of sHSPs promoters

in GCs (Wunderlich et al. 2003; Lohmann et al. 2004),

particularly AtHSF1, which is also activated during the cell

cycle (Reindl et al. 1997). Future work based on differen-

tially expressed HSFs in GCs identified in a transcriptome

analysis of micro-dissected Arabidopsis GCs (Barcala et al.;

unpublished) will help to distinguish amongst these multiple

possibilities and to assign functional roles of particular HSFs

in GC-specific expressions of sHSP genes.

A function for class I sHSPs in giant cells?

Our results show that class I sHSPs are present in galls and

are particularly abundant in tobacco GCs as compared to

the surrounding root tissues (Fig. 3). This is in complete

accordance with GUS expression data from several heter-

ologous promoters whose genes code for class I sHSPs

(Escobar et al. 2003 and this paper). sHSPs belong to a

large family of abundant proteins that play different roles

throughout plant life. They are not only induced by envi-

ronmental cues, such as extreme heat, heavy metals, water

stress etc. (Vierling 1991; Almoguera et al. 1993) but are

also developmentally programmed to be expressed during

embryogenesis. In prokaryotic cells, some sHSPs are

associated with inclusion bodies formed during high-level

protein expression (Laskowska et al. 1996). In GCs, crys-

talline inclusions have been detected in their cytosol

(Paulson and Webster 1970), perhaps related to the high

protein turnover that rapid GC growth predicts. The high

metabolic rate of these cells inferred from their ultra-

structure (Bird 1961; Bleve-Zacheo and Melillo 1997;

Gheysen and Fenoll 2002) might need chaperones to pre-

vent the aggregation of abundantly synthesised proteins.

Some sHSPs are particularly effective chaperones in pre-

venting protein aggregation, acting by an ATP-independent

mechanism through conformational changes both depen-

dent on and independent of heat (Sun et al. 2002; Haslbeck

et al. 2005). In addition, sHSPs are important for the pro-

tection of multiple cellular proteins and a wide variety of

cellular activities, such as transcription, cell signalling and

the assembly/disassembly of cytoskeletal components

(Basha et al. 2004), all of which are processes described to

be altered in GCs (Gheysen and Fenoll 2002; de Almeida

Engler et al. 2004). Nematode feeding habits should also

be taken into account, as they feed through disposable

‘‘feeding tubes,’’ thought to be formed by secretions

extruded from their stylets. These tubes are poorly

described crystalline structures found in the cytoplasm of

GCs and syncytia (Hussey and Mims 1991), which have to

be assembled and attached to the nematode stylet in each

feeding cycle and that might recruit plant chaperons as

well. Finally, the parasites might pre-digest proteins by

injecting proteases inside the feeding cells (Vanholme

et al. 2004; Shingles et al. 2007), which might also signal

chaperones overproduction.

Our finding that sHSP accumulate in GCs, together with

the great similarity between GCs and heat-shock responses

regarding sHSP promoter regulation, suggests that the

metabolic state of the GCs might somehow mimic a heat-

shock response or even a cellular state with conditions

similar to those elicited by osmotic, water or heat stress

when abundant unfolded proteins present in the cell need

chaperone activities. Hence, sHSP production in GCs

might be aimed at protecting protein activity during self-

generated stress conditions. GCs are nutrient sinks in which

soluble assimilates are continuously ingested by the nem-

atode and replenished by the plant (Hussey and Mims

1991). It is possible that cyclic changes in low-molecular-

weight metabolites result in cyclic alterations of the

osmotic balance in the GC. In this respect, several genes

involved in drought tolerance, including genes from car-

bohydrate metabolism, have been described as targets of

HSFs in Arabidopsis (Busch et al. 2005) and a recent work

by Swindell et al. (2007) suggests that sHSPs could con-

tribute to multiple stress tolerance in different plant

Plant Mol Biol (2008) 66:151–164 161

123

species. Moreover, HSFs and HSPs represent an interaction

point at the cross-talk of several stress response pathways.

At the moment, the signals that activate HSFs and other

possible transcription factors in GCs that lead to sHSP

activation are completely unknown. Future research,

including gain- and loss-of-function experiments by

exploiting molecular genetics tools available in Arabid-

opsis, will help in finding both the putative role of the class

I sHSPs in GCs and the transcriptional circuits that lead to

their accumulation.

Nematode-specificity in the transcriptional activation

of the G4 promoter

Syncytia and GCs show differential regulation of a set of

genes that share a common regulation during late-

embryogenesis and/or under different stress conditions,

such as drought (de Meutter et al. 2005). We had described

the induction of the G4 promoter in GCs induced by root-

knot nematodes (Escobar et al. 2003). Our results in

tobacco plants infected with the cyst nematode Globodera

tabacum now establish that G4 is a GC-specific promoter,

at least in tobacco, as we could not detect any GUS activity

in the syncytia of both in vitro and sand-grown plants

(Fig. 2). Furthermore, the minimal promoter version that

was active in tobacco GCs, -83G4, showed also a null

response after cyst nematode infection.

Expression studies of genes with common regulation

during late-embryogenesis, drought stress and nematode

infection (Van der Eycken et al. 1996; Escobar et al. 1999,

2003; de Meutter et al. 2005 and this study) have revealed

their complex regulation in nematode-feeding sites. Some

genes are syncytia-specific (ABI3 and CDeT-27-45; de

Meutter et al. 2005), whereas others are specific for GCs

(Hahsp17.7G4; Escobar et al. 2003). Other genes, such as

LEMMI9, are induced by both nematodes, although the

response to cyst nematodes is weaker (Van der Eycken

et al. 1996, 1999; de Meutter et al. 2005). Our results

support the view that complex sets of transcription factors

are differentially active in syncytia and GCs. Detailed

analysis of each particular promoter and its interacting

transcription factors in different physiological situations

will help in understanding the profound re-programming of

gene expression that takes place in nematode-feeding sites

and which leads to the widely described root morphologi-

cal changes elicited by plant parasitic nematodes.

Acknowledgements We thank Dr. Mugniery for kindly providing

the Globodera tabacum population and Ana Belen Yuste for her

technical help in the analysis of the Arabidopsis transformants. This

work was supported by grants from the Fundacion Ramon Areces and

the Ministerio de Educacion (AGL-2004-08103-C02-02) to CE, from

a collaboration project from the National Science Foundation of the

USA to Elizabeth Vierling and from the European Commission

(QLK5-1999-01501) and the Junta de Comunidades de Castilla-La

Mancha (JCCM, GC-02-011) to CF. MB was a recipient of an FPU

fellowship from the Ministerio de Educacion and AG from the ‘‘Junta

de Comunidades de Castilla-La Mancha.’’

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