Heat stable proteome associated with desiccation tolerance

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Running head:

Heat stable proteome associated with desiccation tolerance

Correspondence:

Dr O. LEPRINCE

UMR Physiologie Moléculaire des Semences

ARES

16 bd Lavoisier

F-49045 Angers

France

Tel : + 33 241 22 55 16

Fax : + 33 241 22 55 49

E-mail [email protected]

Research area:

Environmental Stress and Adaptation

Plant Physiology Preview. Published on February 3, 2006, as DOI:10.1104/pp.105.074039

Copyright 2006 by the American Society of Plant Biologists

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Comparative analysis of the heat stable proteome of radicles of

Medicago truncatula seeds during germination identifies late

embryogenesis abundant proteins associated with desiccation

tolerance

Julie Boudet, Julia Buitink, Folkert A. Hoekstra, Hélène Rogniaux, Colette

Larré, Pascale Satour, and Olivier Leprince

Unité Mixte de Recherche 1191 « Physiologie Moléculaire des Semences (Université

d’Angers, Institut National d’Horticulture, Institut National de la Recherche Agronomique) »,

Anjou Recherche Semences, 16 boulevard Lavoisier, 49045 Angers, France (J.B., P.S., O.L.);

Laboratory of Plant Physiology, Department of Plant Sciences, Wageningen University,

Arboretumlaan 4, 6703 BD Wageningen, The Netherlands (F.A.H.); Unité de Recherche

« Biopolymères, Interactions, Allergie » (Institut National de la Recherche Agronomique), rue

de la Géraudière, BP 71624, 44316 Nantes (H.R., C.L.)



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This work was supported in parts by grants from the Contrat de Plan Etat-Région-des Pays-

de-la Loire 2000-2006, INRA and Van Gogh NWO/EGIDE.

Corresponding author; e-mail [email protected]; fax 33-2-41-22-55-49.

The author responsible for distribution of materials integral to the findings presented in this

article in accordance with journal policy is: O. Leprince, [email protected]



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A proteomic analysis was performed on the heat stable protein fraction of imbibed

radicles of Medicago truncatula seeds to investigate whether proteins can be identified

that are specifically linked to desiccation tolerance (DT). Radicles were compared before

and after emergence (2.8-mm long) in association with the loss of DT, and after

reinduction of DT by an osmotic treatment. To separate proteins induced by the osmotic

treatment from those linked with DT, the comparison was extended to 5-mm long

emerged radicles for which DT could no longer be re-induced, albeit that drought

tolerance was increased. The abundance of 15 polypeptides was linked with DT, out of

which 11 were identified as LEA proteins from different groups: MtEm6 (group 1), one

isoform of DHN3 (dehydrins), MtPM25 (group 5) and three members of group 3: MP2,

an isoform of PM18 and all the isoforms of SBP65. In silico analysis revealed that their

expression is likely seed-specific, except for DHN3. Other isoforms of DNH3 and PM18

as well as 3 isoforms of the dehydrin Budcar5 were associated with drought tolerance.

Changes in the abundance of MtEm6 and MtPM25 in imbibed cotyledons during the

loss of DT and in developing embryos during the acquisition of DT confirmed the link of

these two proteins with DT. Fourier transform infrared spectroscopy revealed that the

recombinant MtPM25 and MtEm6 exhibited a certain degree of order in the hydrated

state, but that they became more structured by adopting α-helices and β sheets during

drying. A model is presented in which DT-linked LEA proteins might exert different

protective functions at high and low hydration levels.



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Desiccation tolerance corresponds to the ability to survive nearly complete protoplasmic

dehydration (ca. –300 MPa). This phenomenon is widespread across the plant kingdom,

including ferns, mosses, pollen and seeds as well as several whole angiosperms, the so-called

resurrection plants. In orthodox seeds, DT is acquired during maturation approximately

halfway through the seed filling phase. Upon seed imbibition, emerging radicles are the first

to lose their ability to tolerate air drying, followed by hypocotyls and cotyledons (Buitink et

al., 2003). At the seedling stage, tissues can no longer survive great losses of moisture. To

cope with the physical and biochemical challenges accompanying the desiccation process,

anhydrobiotes (i.e. desiccation-tolerant organisms) are endowed with an array of protective

mechanisms that act synergistically. They include the synthesis of protective molecules, the

ability to avoid free-radical induced injury during drying and the capacity to repress

metabolism in a coordinated fashion (Leprince et al., 2000; Walters et al., 2002; Avelange-

Macherel et al., 2005). The protective molecules identified and characterized so far are non-

reducing di- and oligosaccharides (Hoekstra et al., 2001; Buitink et al., 2002), small heat

shock proteins (Wehmeyer and Vierling, 2000) and late embryogenesis abundant (LEA)

proteins (Cuming, 1999).

LEA proteins are classified in at least 5 groups by virtue of similarity in their amino acid

sequences (Cuming, 1999; Wise, 2003). They are low complexity, highly hydrophilic and

mostly unordered proteins in the hydrated state and heat stable after boiling (Cuming, 1999;

Wise 2003). Generally, the presence of LEA proteins correlates well with DT. LEA proteins

accumulate to high levels in developing seeds during late maturation (Blackman et al., 1995;

Cuming, 1999; Buitink et al., 2002) and in dehydrating vegetative tissues of resurrection

plants (Ramanjulu and Bartels, 2002). Correlations between the disappearance of various

members of group 1, 2 and 3 LEA proteins and loss of DT during germination have also been

reported (Ried and Walker-Simmons, 1993, Whitsitt et al., 1997; Capron et al., 2000;

Gallardo et al., 2001). Seeds of the double mutant aba,abi3 of Arabidopsis that are deficient

in several heat stable polypeptides are also desiccation-sensitive, although they exhibit an

array of pleiotropic defects ranging from decreased accumulation of storage proteins to

vivipary (Meurs et al., 1992). A role of LEA proteins in DT has been demonstrated in the

bacterium Deinococcus radiodurans. Inactivation of a group 3 LEA protein, homologue of

the plant LEA76, leads to 75% reduction in viability of desiccated cultures (Battista et al.,

2001). In contrast, in seeds, direct in vivo evidence for a role of LEA proteins in tolerance to

complete water loss has not yet been secured. There exist several Arabidopsis mutants that

produce seeds devoid of one or two LEA proteins belonging to group 1, but remaining



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desiccation-tolerant (Carles et al., 2002), either arguing against a role for these proteins in DT

or revealing a possible functional redundancy between the different group members as

suggested by Ditzer et al. (2001). Conversely, dehydrins have been detected in seeds that

remain desiccation sensitive at shedding (Kermode, 1997). Nonetheless, in vitro experiments

do point to a protective role of LEA proteins against the deleterious effects of drying. For

instance, several LEA proteins from group 2, 3 and 4 were found to protect enzymes against

nearly complete loss of water brought about by rapid evaporation or vacuum drying (Goyal et

al., 2005; Grelet et al., 2005).

In addition to being present in anhydrobiotes, LEA proteins are also expressed in

desiccation-sensitive vegetative tissues as a response to stress involving changes in cellular

water potential (Cuming, 1999). Most of the experimental evidence shows that LEA proteins

that are over-expressed in vegetative tissues can improve tolerance to various degrees of

hyperosmotic stress (-1 to -6 MPa), induced by a partial loss of water, salt or freezing (Imai et

al., 1996; Swire-Clark and Marcotte, 1999; Cheng et al., 2002; Houde et al., 2004; Riera et

al., 2004). Whether LEA proteins play a similar role in seeds as they do in drought-tolerant

systems is unclear. LEA proteins are extremely diversified in terms of genotypic variability,

regulation, localization at the tissue and cellular level (Dure 1993; Cuming, 1999; Wise,

2003). Several LEA genes appear to be specifically expressed in seeds, such as the Em1 and

Em6 in Arabidopsis (group 1; Bies et al., 1998) and rab28 (group 5) in Arabidopsis (Arenas-

Mena et al., 1999) and maize (Niogret et al., 1996). When these genes are over-expressed in

desiccation-sensitive systems such as yeast (Swire-Clark and Marcotte, 1999), leaves of rice

(Cheng et al., 2002) and Arabidopsis seedlings (Borrell et al., 2002), an improved tolerance to

salt or drought is observed. During drying, seed tissues pass through hydration ranges that

also necessitate protection against drought. Thus, it can be argued that, in seeds, the role of

LEA proteins might be similar as in drought-tolerant vegetative tissues, their action being

confined to relatively high water contents (ca. -3.5 MPa; Hoekstra et al., 2001; Ramanjulu

and Bartels, 2002). In this case, no LEA proteins would be found to be specifically correlated

with tolerance to low water contents. Alternatively, they might exert several functions that

differ according to the hydration level reached by the seed tissues during drying, as is the case

for non-reducing sugars (Hoekstra et al., 2001; Ramanjulu and Bartels, 2002). Sugars are

thought to act as compatible solutes during the initial water loss and when the bulk water is

removed, they protect macromolecules by replacing water with OH groups and by forming a

glass, which stabilizes the macromolecular structures for long periods of time (Hoekstra et al.,

2001). Similarly, apart from their protective role in drought conditions, LEA proteins have



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been shown in vitro to prevent conformational changes of hydrophilic polypeptides when the

last hydration layer is removed (Wolkers et al., 2001) and to participate in the formation of a

glassy state, occurring at a water content below 0.10 g/g (g H2O/ g dry weight; Wolkers et al.,

2001; Shih et al., 2004). Therefore, this study investigates if there exist specific LEA proteins

whose abundance is associated with DT rather than drought tolerance in Medicago truncatula

seeds.

To comprehend the changes in LEA proteins simultaneously, comparative proteomic

analysis was carried out in desiccation-tolerant and -sensitive radicles during germination. In

addition, this approach allows to assess whether putative posttranslational modifications are

also associated with DT, considering that some LEA proteins from group 1, 2 and 3 are

submitted to post-transcriptional and post-translational modifications during seed maturation

and germination (Bies et al., 1998; Campalans et al., 2000). Furthermore, the phosphorylation

status of the acidic dehydrins was found to determine the protective activity (Riera et al.,

2004; Alsheikh et al., 2005). To facilitate the detection of LEA proteins, we focussed on the

heat stable proteome that resist coagulation upon heating at 95°C. By this method, the soluble

protein extract containing hydrophilic proteins should be enriched with LEA proteins and

devoid of storage proteins, which can represent up to 60% of the total proteome of M.

truncatula seeds (Gallardo et al., 2003; Watson et al., 2003). Apart from LEA proteins, other,

unidentified proteins with protective functions might be present in the heat stable fraction.

This is based on a recent argumentation that LEA proteins are members of a larger family of

osmotic stress proteins called “hydrophilins”, which are defined as proteins have a Gly

content > 6% and a hydrophylicity index > 1 (Garay-Arroyo et al., 2000).

Several transcriptomic and proteomic analyses both during seed development (Gallardo et

al., 2003; Hajduch et al., 2005) and germination (Gallardo et al., 2001; Soeda et al., 2005)

have given some insights into several seed-specific events such as synthesis of storage

reserves, desiccation, radicle protrusion and germination performance. Although several stress

proteins were identified, their abundance was not studied in relation to DT. In our work,

profiles of heat stable proteins were compared between desiccation-tolerant radicles of non-

germinated seeds and sensitive radicles after emergence out of the seed coat. To confirm the

link with DT, profiles were also studied in emerged radicles in which DT was re-established.

This can be brought about by exposing germinated seeds to an osmotic treatment for several

days (Leprince et al., 2000; Buitink et al., 2003). Those proteins that were expressed in treated

radicles upon re-establishment of DT were further analyzed in older emerged radicles, for

which DT can no longer be re-induced by the osmotic treatment. This strategy allowed for the



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discrimination of putative proteins linked to DT and osmotic tolerance. Eleven polypeptides

representing several forms of LEA proteins were found to be associated with DT. Among

them, two proteins belonging to group 1 and group 5 were further characterized during

maturation and germination by western blotting. To gain further insights into their function,

their secondary structure was compared in the hydrated and dry state after fast and slow

drying using FTIR (Fourier transform IR) spectroscopy.

RESULTS

Changes in Heat Stable Protein Patterns in Relation to the Loss and Re-establishment of

Desiccation Tolerance

In seeds of M. truncatula, DT of the radicle is maintained during the early phase of imbibition

and is lost when the radicle protrudes the seed coat (Table I). Germinated seeds with 2.8 mm

long protruded radicles are not able to survive a 3 d drying at 42% RH at 20°C (Table I).

Previously, Buitink et al. (2003) showed that DT can be re-established in these sensitive

radicles by incubating the germinated seeds in a solution of polyethylene glycol (PEG) having

a water potential of -1.7 MPa for 2 d. Table I shows that in these conditions, DT was restored

to 91% in 2.8 mm long emerged radicles. However, when germinated seeds are selected at a

later stage during post-germinative growth, corresponding to seeds with protruded radicles of

5 mm in length, DT can no longer be re-established after the same PEG treatment (Table I).

To identify proteins involved in DT, we compared the heat stable (HS) proteome extracted

from 2.8 mm long, desiccation-sensitive radicles with those from 16 h imbibed non-

germinated desiccation-tolerant radicles. To validate whether putative candidates were linked

to DT, the HS proteome was also analyzed from PEG-treated 2.8 mm radicles, in which DT

was re-established. In non-germinated (NG) radicles, the weight fraction of the HS proteome

corresponded to 28% of the total soluble proteins. During germination, the amount of HS

proteins decreased 2.3-fold (Table I). The osmotic treatment did not reverse this decrease; the

amount of HS proteins in PEG-treated radicles represented 15% of the total soluble proteins.

The HS fractions from the 3 stages were analyzed by two-dimensional gel electrophoresis

(2DE) using a non-linear pI gradient (Fig. 1). For each stage, the spots from 6 to 8 replicates

were detected and compared to each other using the PD Quest software. To secure the quality

of the data, spots of poor quality and very low raw volumes were discarded using criteria set



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by the software. Furthermore, to be included in the statistical analysis, each spot had to be

present in at least 50% of the gel replicates. In total, 391 spots satisfied these criteria and were

included in the reference gel. The number of detected spots differed significantly among the

stages (Table I). Concurrent with the decrease in the proportion of the HS fraction, the

number of spots decreased from 328 to 252 during the loss of DT (Table I). In contrast, the

PEG-induced re-establishment of DT led to a slight increase in the spot number (Table I). A

nested ANOVA and the Student-Newman-Keuls test (P < 0.05) classified 376 spots out of the

391 spots in 9 expression profiles (Table II). For the remaining 15 spots, the Student-

Newman-Keuls test did not reveal a significant difference in contrast to the nested ANOVA,

which gives a better estimate of the residual variance. Out of the 376 spots, only 54 remained

constant in the three stages (Table II). The profile with the highest number of spots (profile 2)

represented those spots that were more abundant in the desiccation-tolerant, NG stage

compared to the other two stages. Only 5.9% (23) of the total detected spots had an

expression profile associated with DT, that is, they were significantly more abundant in NG

and PEG-treated 2.8 mm radicles than in the untreated, 2.8 mm sensitive radicles (Table II,

profile 9). There were 32 spots that were associated with desiccation sensitivity, being more

abundant in the 2.8 mm radicles: they represented 8% of the total amount spots. Another

interesting group of spots are those found in profile 4, whose abundance increased

significantly upon the PEG treatment (44 spots, 11.2%).

Discrimination Between the Desiccation-Tolerant Proteome and the Osmotically

Induced Proteome

A total of 23 spots showed a higher abundance in both desiccation-tolerant stages compared

to the sensitive stage (profile 9, Table II). These spots were further analyzed using two

additional stages: 5 mm long, desiccation-sensitive emerged radicles before and after a PEG

treatment (Table I). As a result, the spots could be separated in two sub-groups: A) those that

are only induced in the 2.8 mm long radicles after PEG treatment and are thus linked

specifically to DT and B) those that are also induced by the PEG incubation in 5-mm long

radicles that remain desiccation-sensitive. Out of the 23 spots, 11 were found to be associated

specifically with the induction of DT (subgroup A) and 7 were induced both in 2.8 and 5-mm

long PEG-treated radicles (subgroup B), albeit not always to similar levels in both tissues.

Among these 7 spots, 5 exhibited a significant higher intensity in 2.8 mm PEG-treated

radicles than in the 5 mm ones. They were therefore also linked to DT. The remaining 5 spots



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could not be categorized in either of these two groups and are not further considered in this

study.

It is noteworthy that, although the PEG incubation of the 5 mm long radicles did not lead

to the re-establishment of DT, this treatment did result in an improvement of the tolerance to

drying. This is demonstrated by the assessment of the water content to which 50 % of the

population of germinated seeds can be dried and rehydrated without loss of viability of their

radicle (threshold water content, Fig. 2 and Table I). Germinated seeds with an emerged

radicle of 2.8 mm long were able to survive a desiccation treatment down to 1 g H2O/g DW

(g/g) but died at lower water content. Fifty % of survival was obtained at 0.3 g/g (Table I).

After the PEG treatment, 2.8 mm long emerged radicles were able to survive nearly complete

removal of water and thus considered desiccation-tolerant (Fig. 2). In contrast, 5 mm long

radicles were very sensitive to drying. Fifty % of death was obtained when the radicles were

dried to 3.6 g/g. However, after a 2 d incubation in the osmoticum, they had become more

tolerant to desiccation since the threshold water content decreased to 0.8 g/g (Table I).

Whitsitt et al. (1997) observed a similar effect for soybean seedlings: an incipient water

deficit decreased the sensitivity of seedlings to further dehydration. The division of profile 9

into the two subgroups A and B showed that certain spots could only be induced in those

tissues that become desiccation-tolerant, whereas others could also be re-induced by the PEG

incubation in 5 mm long radicles that remained desiccation-sensitive (data not shown).

Identification of Heat Stable Proteins Associated with Desiccation Tolerance

The 16 spots belonging to profile 9A and 9B and linked with DT were analyzed using matrix-

assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF) and

liquid chromatography-tandem mass spectrometry (LC-MS/MS). Out of these 16 spots, 11

were identified as 6 different LEA proteins, some of them being present as different isoforms

(Table III). Another polypeptide was identified as a homologue of a pea legumin precursor

(Table III). Since the Mw of this spot was much lower than expected, we suspected that the

onset of the digestion of storage proteins, which is known to occur during radicle growth

(Capron et al., 2000; Gallardo et al., 2001), yielded small, hydrophilic peptides. This legumin

fragment represented less than 0.05 % of the HS proteome. It was not therefore taken into

account for the remainder of this study. Finally, four polypeptides were not identified.

To find out to which groups the identified LEA proteins linked to DT belong to, a

phylogenetic tree was constructed with LEA proteins of M. truncatula after a search in the



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TIGR (http://www.tigr.org/tdb/mtgi) and NCBI (http://www.ncbi.nlm.nih.gov) databases

using a set of keywords and the PFAM domains characteristic of plant LEA proteins

(http://www.sanger.ac.uk/Software/Pfam/index.shtml). Twenty-five genes were obtained and

a phylogenetic tree with the protein sequences was generated using ClustalX (Thompson et

al., 1997) and TreeView (Page, 1996). In light of conflicting classifications of LEA proteins

(see Wise, 2003), those by Cuming (1999) and Dure (1993) as well as the PFam domains

(Bateman et al., 2004) are indicated in the tree (Fig. 3). The 6 LEA proteins identified in this

study in relation to DT belonged to 4 different groups according to Cuming’s classification

One DT-linked spot (Table III, Fig. 1 spot 37) was identified as a homologue of Em6 of A.

thaliana, and was the only representative of group 1 (G1). Figure 3 suggests that Cuming’s

group 5 is divided in two clusters of closely related genes (i.e. D34, PF04927 and D95,

PF03168) as previously suggested by Dure (1993). In the literature, the D34 family has been

classified successively in group 6 then 5 (Wise, 2003). In this work, one of its member was

detected in relation to DT and named PM25 due to the high similarity with GmPM25 from G.

max (Table III, Fig. 1 spot 21). One isoform of DHN3 (spot 11) was linked to DT. According

to Fig. 3, it is a member group 2 (G2) LEA proteins, also known as dehydrins (D11,

PF00257). Figure 3 shows that LEA proteins from Cuming’s group 3 (G3, D7, PF029877) do

not appear to form an homogeneous clade. Three members of this group were found to be

correlated to DT. Among them were two closely related proteins, homologues of PM18 and

MP2 of G. max (Table III, Fig. 1 spot 16 and 19) and one unrelated protein, identified as a

homologue of SBP65 of P. sativum, a biotinylated protein (Table III, spots 2-6).

Analysis of the changes in relative volume of Em6 and PM25 (Fig. 4B and D)

demonstrated that the PEG treatment significantly re-induced the expression of both proteins

in the 2.8 mm long emerged radicles. Nonetheless, the re-establishment of DT did not lead to

a similar re-induction of the protein abundance as was the case in the non-germinated,

desiccation-tolerant seeds. The three DT-linked LEA proteins that belong to group 3 (MP2,

PM18 and SBP65) were present as several isoforms, mainly differing in pI (Fig 4 and 5).

Based on the statistical analysis of their normalized intensities, they were categorized in

different profiles. MP2 was present in two forms (Fig. 4 E, F); the most abundant one was

linked to DT (spot 19) whereas a very faint spot (20) was classified in profile 4 (i.e. induced

by the PEG treatment only in 2.8 mm long PEG-treated radicles; data not shown,

supplementary table I). Three spots corresponding to PM18 (spot 15, 16 and 17), showed a

different expression profile (Fig. 5A). Spot 16 (Fig. 5A, B) was found to be linked to DT

whereas the most acidic polypeptide (15) increased upon osmotic treatment regardless of



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whether full DT is induced or not (profile 9B, Fig. 5B). The most basic spot (18) did not vary

significantly between NG and 2.8 mm radicles, but decreased in the 5-mm long protruded

ones. Furthermore, it did not respond to the PEG treatment (Fig. 5B). A similar pattern was

observed for spot 17. It was tentatively identified as a fourth isoform of PM18 because the

experimental trypsin digestion before the MALDI-TOF analysis produced 4 digested

peptides, the masses of which matched some obtained by the theoretical digestion of the

translated Medicago TC similar to PM18. The third member of group 3, SBP65, existed in 6

isoforms with different pIs (spots 1 to 5, Fig 5C, D) and Mw (spot 6, Fig. 1). Again, the

different isoforms were classified in two profiles: 9A, re-induced only in 2.8 mm long PEG-

treated radicles and 9B, also re-induced in 5 mm long PEG-treated radicles (Fig 5 D).

Nonetheless, for all isoforms, the abundance was significantly higher in those tissues that

were desiccation-tolerant than in those that remained sensitive.

Another group of proteins that show an interesting profile are those being induced upon

osmotic stress in protruded radicles of both stages, belonging to profile 4 (Table II). The two

dehydrins (G2) that were identified in this profile were DHN3 and BudCar5, both being

present in several isoforms (Fig. 6). DHN3 was present in 3 isoforms (spots 11-13; Fig 6 A,

B). The abundance of the two most basic forms (spots 12 and 13) increased as a response to

the PEG treatment only in the 2.8 mm long radicles. In contrast, the two most acidic forms

(10 and 11) responded to the osmotic treatment by increasing ca. 2-fold both in 2.8 and 5 mm

radicles. Spot 11 was classified in profile 9B (Table II) as mentioned earlier. The amount of

all Budcar5 isoforms (spots 31-33) increased sharply upon the PEG-incubation (Fig. 6C, D).

Dehydrins are known to be expressed under different types of stress and in different tissues,

thus their induction upon PEG-incubation was expected.

To investigate whether the DT-linked 6 LEA proteins were seed specific or expressed in

different tissues and/or under different stress conditions, their gene expression was analyzed

in silico. Also added to this analysis were two additional LEA proteins: CapLea1

(TC100264), a group 3 LEA protein representing the largest amount of the heat stable fraction

but whose levels remained constant in the radicles of different stages (spot 34, Fig. 1), and

BudCar5. In silico gene expression was expressed as the number of expressed sequence tags

(ESTs) corresponding to the LEA proteins based on the total number of ESTs present in the

particular cDNA library. Thirty-four libraries representing different organs submitted or not to

drought, nutrient and biotic stresses, developing seeds at different stages and germinating

seeds were selected and pooled into 9 groups (Fig. 7, supplementary table II). Except for

DHN3, the remaining 5 LEA proteins that were identified in relation to DT appeared to be



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seed specific (Fig. 7). DHN3 was expressed in several libraries, mainly in drought-stressed

leaves. The expression of BudCar5 was detected in all the libraries studied. The in silico

expression analysis of the all LEA proteins shown in the phylogenetic tree (Fig. 3) revealed

that all LEA genes from group 1 to 4 that were specifically expressed in seeds (supplementary

table II) were found to be linked to DT in this study. Only three other members of group 5 and

Lea5 (no classification) that were present in the seed libraries were not detected here.

Furthermore, only 2 of the 23 known LEA genes (DIP, a dehydrin and CapLea1B, member of

G3) were not present in at least one of the seed libraries (supplementary table II).

Whether the digestion of storage proteins yielded hydrophilic peptides during germination

and PEG incubation was further assessed by excising and identifying spots of low mass from

gels of radicles of NG seeds and 2.8 mm long emerged radicles of germinated seeds. Spots 35

and 40 (Fig. 1, Mw around 16 kDa) were also identified as homologues to the pea legumin

precursor (theoretical Mw 65 kDa, TC85216). Four spots (22, 23, 24, 25; Mw around 31kDa,

Fig. 1) were identified as homologues of the pea convicilin (TC100299) having a theoretical

Mw of 78.3 kDa. These fragments of storage proteins belonged to profile 2, 3 and 5 (Table II)

and were not very abundant. Likewise, the heat stable proteome included proteins other than

LEA proteins (for example homologues of a Vicia faba transcription factor (TC94137), an

ankyrin repeat protein 2 from Vitis aestivalis (TC100495) and a glycine-rich protein 2 of

Nicotiana sylvestris (TC98399); see supplementary table I).

Changes in MtPM25 and MtEm6 in Relation to Desiccation Tolerance during Seed

Maturation and Germination

The expression profiles of two of the six LEA proteins that were linked to DT were further

characterized to confirm the data obtained from the 2D proteomic analysis. The analysis was

extended to cotyledons during germination and embryos during seed development to ascertain

the abundance of these LEA proteins with DT. MtPM25 and MtEm6 were chosen because

they were represented by a single spot in the gels, thereby alleviating any complication with

the interpretation of western blots that were performed in one dimension. Full length cDNAs

corresponding to the MtPM25 (DQ206870) and MtEm6 (DQ206712) were obtained by

RACE. Sequences corresponding to a N-terminal poly-HIS tag and cleavage site for

enterokinase were added to the full length encoding sequence and the recombinant proteins

were expressed in E. coli. Rabbit polyclonal antibodies were raised against the purified



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recombinant MtPM25 and MtEm6. For each antibody, a signal at the expected molecular size

was detected both with protein extracts from radicles and the recombinant protein. The signals

were absent when the pre-immune serums were used (data not shown). During seed

imbibition, contents of both MtPM25 and MtEm6 in the radicles remained high for up to 15 h

(Fig. 8A, E). In 20 h imbibed radicles (ca 2.8 mm in length), MtPM25 was barely detectable,

whereas MtEm6 had already disappeared. In accordance with the proteomic analysis, the

osmotic treatment was found to re-induce the expression of both proteins in 2.8 mm long

emerged radicles, albeit to lower levels than those found in NG radicles (Fig. 8C, G). In 5 mm

long radicles, the PEG treatment only resulted in the appearance of a very faint signal. The

relationship between DT and the presence of both proteins was also confirmed for the

cotyledons during germination (Fig. 8B, F). In contrast to radicles, DT in cotyledons was

maintained for up to 24 h of imbibition and lost at 48 h. In parallel, MtEm6 and MtPM25

amounts decreased to barely detectable levels and disappeared. During seed development,

tolerance to rapid enforced drying was acquired between 14 and 22 days after pollination

(DAP, Fig. 8). Contents of MtPM25 increased at 14 DAP in parallel with the acquisition of

DT (Fig. 8D) whereas those of MtEm6 started to accumulate later at 18 DAP.

Secondary Structure Analysis of MtPM25 and MtEm6 Proteins

It has been established that LEA proteins of group 3 and 4 undergo an unordered to ordered

structure transition during the loss of water (Wolkers et al., 2001; Goyal et al., 2003; Shih et

al., 2004). Considering their divergence in the Kyte and Doolittle hydrophilicity profile

(http://ca.expasy.org/tools/protscale.html), possible differences in secondary structure of the

recombinant form of MtPM25 and MtEm6 were investigated. Whether changes in protein

conformation were induced upon drying was also assessed. FT-IR spectra of recombinant

proteins in the hydrated and dried state were recorded after removal of the His6x tag (Fig. 9).

To avoid interference of the H-O-H scissoring vibration of water around 1646 cm-1 with the

amide-I band between 1700 and 1600 cm-1, D2O instead of H2O was used for the proteins in

solution. Wolkers et al. (1998) demonstrated that intermolecular ß-sheet formation can

effectively be prevented by fast drying, probably because the time required for such non-

intramolecular rearrangements is too short. For this reason, we studied the recombinant

proteins after fast and slow drying in air of ca. 3 and 67% RH, respectively.

Superficial inspection of the IR-spectrum in the Amide I region (Fig. 9) revealed that in

D2O the proteins displayed a broadened band at a wavenumber position (1460 cm-1) that was



11

lower than in the case of the fast-dried proteins (1550 cm-1). This behavior in D2O may be

partly due to 2H exchange with protons in the protein backbone, which is particularly likely in

unordered structures (Raussens et al., 1997). On the other hand, the dominating band at ca.

1658 cm-1 in the fast-dried proteins may be the result of an increased α-helical content. In the

fast-dried MtPM25, there is evidence of a shoulder at ca. 1630 cm-1, which is less prominent

in the fast-dried MtEm6. This is suggestive of a larger proportion of intermolecular ß-sheet in

MtPM25 than in MtEm6 after fast drying. Finally, slow drying led to an increase in this

structure for MtPM25, which is not observed for MtEm6. An additional slow drying

experiment was performed at 85% RH over a KCl saturated solution. In this case, the

conformation remains similar to that after fast drying, indicating that the increase in β-sheet

structure occurs below 85% RH.

More detailed information on secondary structures in these proteins was obtained by a

curve fitting procedure on the original amide-I band according to Wolkers et al. (2001). An

example of the curve-fitting procedure is given in Fig. 10 with fast-dried MtEm6 protein.

Peaks representing different secondary structures were selected on account of the second

derivative spectrum (Fig. 10, A). Co-addition of all the dashed peaks that were

mathematically produced should result in a fit (crosses) that resembles the original absorption

spectrum (Fig. 10, B). Individual contributions by the various protein secondary structures

can thus be estimated. Table IV shows the curve-fitting results of the amide-I region of both

proteins in D2O and after fast or slow drying. In solution both proteins had between 30 and

40% α-helical structure, with MtPM25 having more extended ß-sheet than MtEm6. If both

random and turn structures were to be combined and considered as “unordered structures”,

even though a certain degree of order might exist for some of them, then MtEm6 would have

37% α-helix, 10% ß-sheet and 53% unordered structures, whereas MtPM25 would consist of

33% α-helix, 18% ß-sheet and 49% unordered structures (Table IV). These figures concur

fairly well with PELE predictions (SDSC Biology workbench: workbench.sdsc.edu). In water

solution, MtEm6 is predicted to form 33% α-helix, 3% ß-sheet and 64% unordered structures

whereas 38% α-helix, 14% ß-sheet and 48% unordered structures are expected for MtPM25.

Support for the largely unordered nature of both Lea proteins in water also comes from the

behavior of the amide-II band around 1540 cm-1 in D2O (Fig. 9). This band was considerably

smaller than that upon fast drying and partly downshifted to 1450 cm-1. The effect of D2O was

stronger for MtPM25 than for MtEm6. Apparently, the amide protons (N-H) were, to a

considerable extent, open for 2H-exchange from D2O, which is interpreted to mean that both

proteins have a fairly unordered structure in water (Haris et al., 1989).



12

Table IV further shows that either fast or slow drying led to a considerable increase in α-

helical structure in both proteins at the expense of the unordered structures. This

intramolecular rearrangement apparently was independent of the rate of drying. In contrast to

MtEm6, MtPM25 tended to form extended ß-sheets upon slow drying, which appeared to be

reversible upon rehydration. When indicated as percentages of α-helix, ß-sheet and unordered

structures, MtEm6 consisted of 57, 12, 31% and 60, 8, 32%, after fast and slow drying,

respectively. Data for MtPM25 were 54, 17, 29% and 56, 25, 19%, respectively.

DISCUSSION

To identify proteins involved in DT, a proteomic screening of the HS fraction of soluble

proteins from imbibed radicles of M. truncatula was combined with a physiological system

that enables the re-establishment of DT in 2.8 mm long emerged radicles by an osmotic

treatment. To separate the proteins induced by the osmoticum from those involved in DT, the

comparison was extended to emerged radicles of 5 mm long, for which DT could no longer be

re-induced by the same osmotic treatment. In total, 15 polypeptides were found, whose

abundance was linked to DT. Among them 11 were identified, which represented 6 LEA

proteins from different groups: MtEm6 (group 1), one isoform of DHN3 (dehydrins),

MtPM25 (group 5) and three members of group 3: MP2, the basic isoform of PM18 and all

the isoforms of SBP65 (Table III). Our in silico analysis revealed that the expression of all the

DT-linked LEA genes was apparently seed specific, except for one isoform of DHN3.

The abundance of all these proteins was associated with DT (Figs. 4, 5 and 8).

Nonetheless, the causal relationship between the 6 LEA proteins and DT remains difficult to

assess. It is possible to obtain dry and viable seeds from Arabidopsis and maize mutants with

very low or undetectable levels of Em transcripts (Williams and Tsang 1991; Carles et al.,

2002). It is not known whether in these mutants, other (LEA) proteins could compensate for

the absence of Em proteins. In addition LEA proteins might act synergistically with other

protective compounds in the dry state. For instance, the combination of LEA proteins and

non-reducing sugars offers better protection against protein aggregation after drying than each

component alone (Goyal et al., 2005). In vivo, cytoplasmic glasses are thought to be

composed of sugars and other compounds (reviewed in Buitink and Leprince, 2004). In vitro,

a mixture of LEA proteins and sugars forms a glass upon drying that exhibits physico-

chemical properties resembling those of cytoplasmic glasses, whereas a glass made of sugars



13

alone has different properties (Wolkers et al., 2001; Buitink and Leprince, 2004; Shih et al.,

2004).

This study also identified several LEA proteins that are linked to drought tolerance rather

than DT, such as several isoforms of DHN3 (spots 12 and 13) as well as isoforms of BudCar5

(Fig. 6). Indeed, the isoforms of these dehydrins were induced not only in the 2.8 mm long

radicles after PEG incubation, but also in the 5 mm long PEG-treated radicles. However,

although the treatment on the 5mm long radicles did not re-establish DT, it did lead

nevertheless to an increased tolerance to drying, evident from the reduction in the threshold

water content from 3.6 g/g to 0.8 g/g. In silico analysis shows that both dehydrin genes are

expressed in drought-stressed plants as well (Fig. 7). This observation concurs with those of

Black et al (1999) who showed that the induction of dehydrins in maturating wheat embryos

is not regulated by the same factors that induce DT. The presence of dehydrins in recalcitrant

seeds of temperate climate (Kermode, 1997), the absence of correlation between their

amounts and seed longevity (Wechsberg et al., 1994) together with the observation that

dehydrins protect enzyme activities only at water potentials above -3 MPa (Reyes et al., 2005)

all point to a protective function at high hydration levels. Thus, dehydrins might protect at

intermediate hydration levels (>0.8 g/g), whereas the DT-linked LEA proteins might play a

role below the hydration level corresponding to the threshold water content of 2.8 mm long

radicles (i.e. 0.3 g/g). In this respect, transcript levels of an homologues of MtEm6 were

correlated with seed longevity of Brassica napus (Soeda et al., 2005) and the wheat

homologue of MtEm6 was found to protect citrate synthase from aggregation due to

desiccation upon multiple freeze-drying cycles, supporting the hypothesis that Em6 can

protect macromolecules in the dry state (Goyal et al., 2005). Conversely, over-expression of

the wheat Em in yeast cells (Swire-Clark and Marcotte, 1999) and also of the Arabidopsis

homologue of MtPM25 in germinating seeds (Borrell et al., 2002) led to improved growth

under high NaCl, KCl, LiCl and sorbitol conditions. Recently, a similar observation was made

for E. coli over-expressing the PM2 (Liu and Zheng, 2002). The cellular water potential

resulting from incubation in these osmotic solutions (osmotic potential ranging from –2 to –6

MPa and equivalent to 96-98% RH) is much higher than those experienced by the dry seeds

(in this study: 42% RH equivalent to –180 MPa, Walters et al., 2002). These results argue for

a protective role of Em6, PM25 and MP2 during hyperosmotic conditions rather than at low

water contents. In the light of these observations, one could envisage that the DT-linked Em6,

MP2, MtPM25 and PM18 could exert more than one function upon water loss as

hypothesized for non reducing sugars, which act as osmolytes during hyperosmotic stress and



14

stabilizers of macromolecules in the dry state (Prestrelski et al., 1993; Allison et al., 1999;

Hoekstra et al., 2001). For example, at high moisture contents, DT-linked LEA proteins may

act as compatible solutes that preferentially exclude chaotropic agents (such as salts) from the

surface of macromolecules as suggested by the beneficial effects described above (Swire-

Clark and Marcotte, 1999; Borrell et al.; 2002; Liu and Zheng, 2002; Reyes et al., 2005).

Likewise, when the hydration shell is removed (i.e. water content less than 0.3 g/g), they

might exert their protective effects in the dry state, as was found for wheat Em, by replacing

water molecules by hydrogen bonding and/or forming a glass which stabilizes the system in

the dried state (Hoekstra et al., 2001; Wolkers et al., 2001; Buitink and Leprince, 2004).

Group 5 LEA proteins, to which MtPM25 belongs, have been reported to be a peculiar

group, with low hydrophilicity and no absence of heat stability (Cuming, 1999; Ramanjulu

and Bartels, 2002). Indeed, MtPM25 was the least hydrophilic from the 6 LEA proteins

identified in this study. However, MtPM25 was heat stable and the contention that group 5

proteins are not heat stable is questionable. To determine the structure of a member of group 5

LEA protein, FTIR analysis on MtPM25 was carried out and compared to that on Em6. The

data on the secondary structure of MtEm6 and MtPM25 in solution complement those

obtained for members of groups 3 and 4 using FT-IR spectroscopy as well as for dehydrins

and a member of the D95 family with other spectroscopy techniques. In the hydrated state,

LEA proteins exhibit a wide degree of disorder, ranging from unordered [group 1, Soulages et

al., 2002; group 3 LEA proteins from Typha latifolia pollen (Wolkers et al., 2001) and

nematodes (Goyal et al., 2003); dehydrins (Soulages et al., 2003)], to 60-70% unordered

[GmPM16, a soybean group 4 LEA protein (Shih et al., 2004) and Em proteins (Table IV,

McCubbin 1985)] and finally down to 50% unordered [MtPM25 (Table IV) and an

Arabidopis Lea14, a member of the D 95 family (PF03168; Singh et al., 2005)]. Likewise, the

nature and contents of ordered structures in solution varies greatly. For example, β−sheet

amounts range from 10% for MtEm6 to 40% for the Arabidopsis Lea14. Considering that the

ectopic expression of members of group 1 to 5 always improves the tolerance against salt or

water stress in bacteria, yeast and plants, albeit to various degrees, it is tempting to speculate

that the unordered domains of LEA proteins is responsible for alleviating the osmotic stress

endured by the tissues. It is noteworthy that the presence of β−sheets appears to be a common

feature of LEA proteins and apparently does not affect the heat solubility of the protein.

Considering that MtEm6 and MtPM25 are hypothesized to play a role in the dry state,

their conformation was also determined after drying. The removal of the water induced a

transition from a fairly disordered conformation to the formation of a considerable amount of



15

ordered structures (Table IV). Our study suggests that this behavior is yet another feature that

now appears to be common to all LEA proteins. Indeed, originally observed for the group 3

and 4 LEA proteins mentioned above, this study shows that it is also the case for members of

group 1 and 5. Both proteins show an increase in their α-helical and β-sheet contents (Table

IV). According to Wolkers et al. (1998), the β-sheet formation results from the replacement of

hydrogen bonding of water by intermolecular hydrogen bonds between peptide backbones.

When induced by drying, β-sheets were fully reversible and could be interconverted by

rehydration, in agreement with previous observations on the group 3 LEA protein from pollen

(Wolkers et al., 2001) and the group 4 GmPM16 of soybean (Shih et al., 2004). The PELE

program did predict the overall contributions of secondary structures of MtEm6 and MtPM25

in the hydrated state, but did not in the dry state. Also for the group 4 GmPM16, the structure

predictions can not be fulfilled in the dried state (Shih et al., 2004). This contrasts with the

findings of Goyal et al (2003) on a group 3 LEA from nematodes. Thus, caution must be

taken in extrapolating computer predictions to the dried state to understand the structure-

function relationship of LEA proteins at low water contents.

So at which hydration level do these proteins gain structure? Slow drying over saturated

salt solutions indicated that the proteins had to be dried below an equilibrium RH of 85% in

order to observe a change in the secondary structure (β-sheet formation). The corresponding

hydration level is around 0.2-0.3 g/g, or to the onset of the removal of the hydration shell

(Hoekstra et al., 2001; Walters et al., 2002). This is a significant finding considering the

hypothesis that DT-linked LEA proteins could play different protective functions depending

on the hydration level. Indeed, the gain of structure at low water contents occurs at a

hydration level that is below that experienced during hyperosmotic stress, but before the

protein is immobilized in the cytoplasmic glassy state. Furthermore, it was observed that the

rate of water loss influences the conformation adopted by MtEm6 and MtPM25 in the dry

state, which is in agreement with the data of Wolkers et al. (2001) on the group 3 LEA protein

from pollen. The presence of solutes such as sucrose also influences the change in

conformation during drying (Wolkers et al., 1998; 2001). We do not know whether this is also

the case for MtPM25 and MtEm6. However, in light of these observations, it is important to

test whether and how the nature and rate of drying influences the protective activity of LEA

proteins in vitro.

Both group 2 (DHN3 and BudCar5) and group 3 LEA proteins (MP2, PM18 and SBP65)

were detected as several isoforms. This observation extends previous experimental evidence



16

showing that dehydrins are submitted to post-translational modifications such as

phosphorlylation during seed development and germination (Campalans et al., 2000; Riera et

al., 2004). Phosphorylation of dehydrins has a functional importance during water and cold

stress (Riera et al., 2004; Alsheikh et al., 2005). Both PM18 and DHN3 have distinct isoforms

that are related to DT, whereas others are clearly not (Figs. 5 and 6). This raises the question

as to whether the regulation of post-translational modifications might be also important for

DT.

SBP65 exhibits a peculiar post-transcriptional modification. This DT-linked group 3 LEA

protein is known to be biotinylated in seeds of a wide range of species (Dehaye et al., 1997;

Capron et al., 2000). This protein has been shown to accumulate during the later stages of

seed development and to be degraded during germination (Dehaye et al., 1997; Capron et al.,

2000). It has been suggested that SBP65 constitutes a storage form of biotin that can be

released during germination and postgerminative growth (Dehaye et al., 1997). Whether its

role in DT is simply to store biotin or whether it has an additional, structural function in the

protection against drying remains to be ascertained.

Using a computational analysis, it has been argued that LEA proteins are members of a

larger family of stress proteins called “hydrophilins” that could be used as predictors of the

responsiveness to osmotic adaptation in prokaryotes and eukaryotes (Garray-Arroyo et al.,

2000). Hydrophilins are defined as proteins having a Gly content > 6% and a hydrophylicity

index > 1. The analysis of the heat stable proteome of M truncatula radicles does not support

this analysis on seeds. On the one hand, MtPM25 and SPB65, two polypeptides linked to DT

do not satisfy the hydrophilicity and Gly content criterium, respectively. On the other hand,

we identified several hydrophilic proteins, such as several fragments of legumin and vicilin

that matched those criteria and yet, they were not induced by the osmotic treatment.

Therefore, in seeds, the term "hydrophilins" can not adequately describe the LEA protein

family or water stress proteins.

Altogether, the data reported in this study suggest that the LEA proteins expressed in

seeds can be divided in two groups, those that are induced only in tissues that are desiccation-

tolerant, and those that are also induced in osmotically shocked radicles that remain

desiccation-sensitive but do increase their tolerance to drying. The first group contains LEA

proteins that seem to be seed specific, based on electronic northerns, whereas the second

group is represented by proteins that are also expressed in vegetative tissues. Possibly, the

proteins that are linked to DT might protect both at high hydration levels and at very low

water contents (<0.3 g/g). Further research should be focused on elucidating whether these



17

proteins play a role at water contents where the proteins have gained further order in their

secondary structures and whether their functions are regulated by post-translational

modifications.

MATERIALS AND METHODS

Plant Material and Treatments

Seeds of M. truncatula Gaertn. (cv Paraggio, Seedco Australia Co-Operative Ltd, Hilton,

Australia) were allowed to imbibe on filter paper in distilled water at 20°C in the dark for up

to 3 d. For the proteomic analysis, desiccation-tolerant stages were established as described in

Buitink et al. (2003). They correspond to 16h-imbibed seeds prior to radicle emergence and

seeds exhibiting a protruded radicle length of 2.8 mm (ca. 20-h imbibed) that were incubated

in a PEG 8000 solution having a water potential of -1.7 MPa for 2 d at 10°C. Desiccation-

sensitive stages corresponded to germinated seeds exhibiting a protruded radicle of 2.8 mm as

well as protruded radicles of 5 mm before and after the PEG treatment. PEG-treated seeds

were briefly rinsed in distilled water before further analysis. To determine the threshold water

content, control and PEG-treated seeds with a protruded radicle of 2.8 and 5 mm were dried

for up to 2 d at 20°C in circulating air at 42% RH. At different intervals during drying,

triplicates of 10 radicles were excised to determine their moisture content and the remainder

of the batch was allowed to imbibe as described above. Seeds that exhibiting a growing

radicle during rehydration were considered desiccation-tolerant. When planted, these seeds

developed normal seedlings similar to untreated seeds.

Plants were grown in a sterile mix of vermiculite and soil in a growth chamber at

24°C/21°C, 16 h photoperiod at 350 µM m-2 s-2. Flowers were tagged and developing seeds

were harvested as described in Gallardo et al. (2003). DT was determined by drying and

rehydrating isolated seeds as described above. Seeds that were able to germinate under the

above conditions were considered as desiccation-tolerant. Water content was determined

according to Buitink et al. (2003). For protein extraction, excised radicles and cotyledons of

germinating seeds and isolated embryos from developing seeds were immediately frozen in

liquid N2 and stored at –80°C before use.

Protein Extraction



18

Soluble proteins were extracted from 50 (western blots) and 100-300 (2-DE) radicles at 4°C

in 400 and 950 µL of buffer, respectively [(500 mM HEPES pH 8.0, 1 mM EDTA and 14%

(v/v) of the protease inhibitor cocktail complete Mini (Roche Diagnostics Molecular

Biochemicals, Meylan, France)], 43 U of Dnase I and 5.3 U of Rnase A. After two

consecutive centrifugations at 13,000 g at 4°C, the resulting supernatant was heated for 10

min at 95°C, cooled for 15 min on ice and centrifuged at 13,000 g for 15 min at 4°C. The

resulting supernatant corresponded to the HS fraction. Protein concentrations were assayed

according to Bradford (1976). For the 2D electrophoresis, the HS proteins were precipitated

with 20% (w/v) trichloroacetic acid on ice then centrifuged at 13,000 g for 10 min at 4°C. The

pellet was washed with 400 µl of cold acetone, air dried and resuspended in 500 µL of

rehydration buffer (6 M urea, 2 M thiourea, 4% (w/v) CHAPS, 20 mM dithiothreitol (DTT),

1% (v/v) biolytes from Bio-Rad (Hercules, CA, USA).

2D Electrophoresis

Twenty-four cm immobilized pH gradient (non linear from 3 to 10) strips (Bio-Rad)

containing 500 µg of heat stable proteins were rehydrated at 50 V for 12 h at 20°C.

Isoelectrofocusing ran at 20°C at 250 V for 5 h then at 8 kV until 60 kVh in a Bio-Rad

Protean IEF Cell. Thereafter, a two step equilibration was carried out by incubating each strip

at room temperature in 8 mL of solution: first step, 15 min in a buffer containing 8 M urea,

375 mM Tris pH 8.8, 20% (v/v) glycerol, 2% (w/v) SDS, 130 mM DTT; second step, 30 min

in the same buffer with 250 mM iodoacetamide instead of DTT. Size separation of proteins

was performed on vertical polyacrylamide gels (12% (w/v) acrylamide) in a Ettan Daltsix

Electrophoresis system (Amersham Biosciences, Orsay, France) according to Gallardo et al.

(2001) using a modified running buffer containing 15.6 mM Tris (pH 8.3), 120 mM Gly,

0.1% (w/v) SDS. The experiments were set up in randomized block design where 6 gels

corresponding to independent protein extractions from various desiccation-tolerant and -

sensitive stages were run in parallel. Four to 8 gels per stage were accumulated independently

until the coefficient of variance of the normalized intensity of 50 representative spots that

were present in all stages was below 25% (Asirvatham et al., 2002).

Gel Staining, Image and Statistical Analysis



19

Gels were stained with 0.08% (w/v) Brillant Blue G-Colloidal for 24 h, destained briefly in

5% (v/v) acetic acid and 25% (v/v) methanol then in 25% (v/v) methanol for 8 h. Stained gels

were scanned at 95.3*95.3 resolution with an optical density range of 0.05 to 3.13 using a GS

800 scanner (Bio-Rad). Digitalized gels were analysed using the PD-Quest 7.1 software (Bio-

Rad). Images were filtered (mode pepper outlier 7*7). To identify differentially expressed

protein spots, the gels corresponding to the NG, 2.8 mm and 2.8 mm+PEG stages were first

compared using a representative gel of the NG stage as the reference gel. After optimization

of the parameters for background subtraction and spot detection, the spots that were not

present in at least 50% of the gels and those exhibiting a quality below the set value of 20%

(max. value being 100%) were discarded. After spot matching, spot intensities were

normalized using the "total quantity in valid spot" method: the quantity of each spot in a gel is

divided by the total quantity of all the spots in the reference gel. The statistical method of

Schiltz et al. (2004) was used to compare the protein abundance among the stages. A nested

ANOVA (P< 0.05) was performed using the Statgraphics software (StatPoint Inc, Herndon,

VA): the normalized spot quantities and the stage were respectively the variable and the factor

whereas the extraction and the 2D electrophoresis were nested in the factor. The Bartlett test

(P<0.05) was used to confirm the applicability of the ANOVA. A multiple comparison of the

means using the Student-Newman-Keuls test (P<0.05) was then performed on the normalized

quantities that were found to be significantly different among the stages. To discriminate

proteins expressed during the PEG-induced DT from those expressed solely as the result of

the PEG treatment, digitalized 2DE gels both from the 5mm and 5mm+PEG stages were

included in the image analysis. The spots of interest that were revealed by the statistical

analysis described above were matched in the reference gel when they were detected in the

gels from the 5 mm and/or 5mm+PEG stages. Because a minority of spots were considered,

the spot quantity was normalized throughout the five stages using the "total density in gel

image" method: the spot quantity is divided by the total intensity value of the gel. Difference

in protein abundance was then submitted to statistical analysis as above. Experimental

molecular masses and pI were determined from digitalized gels using 2-D marker proteins

(Bio-Rad) and the calibration method of the PDQuest software (BioRad).

Mass Spectrometry and Protein Identification

Spots of interest were excised from the 2DE gels and subjected to in-gel tryptic digestion as

described in (Wilm et al., 1996). Briefly, gel slices were washed with 100 µL of 25 mM



20

NH4HCO3, followed by dehydration with 100 µL of 50% (v/v) acetonitrile in 25 mM

NH4HCO3. Proteins were reduced and alkylated by incubation for 1h at 57°C in the presence

of 10 mM DTT followed by 45 min at room temperature after adding 55 mM iodoacetamide.

Gel slices were washed with NH4HCO3 and dehydrated as described before. Gel slices were

vacuum dried and rehydrated with 10 µL of 12.5 ng µL-1 trypsin in 25 mM NH4HCO3

(Sequencing grade, Promega, Madison, WI). After an overnight incubation at 37°C, the

supernatant was collected and the tryptic fragments were measured by MALDI-TOF and/or

LC-MS/MS. MALDI-TOF mass spectrometry was performed on a M@LDI LR instrument

equipped with a conventional laser at 337 nm (Micromass-Waters, Manchester, UK). One µL

of the sample was mixed with 1 µL of the matrix preparation (2.5 g/L α-cyano-4-

hydroxycinnamic, 2.5 g/L 2,5-dihydroxy benzoïc acid, 70% (v/v) acetonitrile, 0.1% w/v

trifluoroacetic acid) and deposited onto the MALDI sample probe. LC-MS/MS analysis was

performed using a nanoscale HPLC (Famos-Switchos-Ultimate system, LC Packings, Dionex,

The Netherlands) coupled to a hybrid quadrupole orthogonal acceleration time-of-flight mass

spectrometer (Q-TOF Global, Micromass-Waters, UK). Chromatographic separations were

conducted on a reverse-phase capillary column (Pepmap C18, LC Packings) at a flow rate of

200 nL/min using a gradient from 2 to 50% of 0.08% (w/v) formic acid in acetonitrile. Mass

data were recorded for 1 sec on the mass range 400-1500 m/z using the Masslynx software

(Micromass-Waters), after which the two most intense ions were selected and fragmented in

the collision cell. The collision energy profiles were optimised for various mass ranges and

charge states of precursor ions. Mass data obtained by MALDI-TOF or LC-MS/MS were

analysed with the Protein Lynx Global Server software (Micromass-Waters). Protein

identification was performed by comparing the data with the UniProt sequence databank or

with TIGR Medicago EST databank (date of release: January 26, 2005).

Expression and purification of recombinant MtPM25 and MtEm6 proteins

Expression vectors for the overexpression of MtPM25 and MtEm6 proteins in E. coli were

constructed using the Gateway technology (Invitrogen). Full length cDNA was amplified by

PCR using a forward primer with an attB1 sequence (MtPM25_F:

GGGGACAAGTTTGTACAAAAAAGCAGGCTTAGATTACAAGGATGATGATGATAA

GATGAGTCAAGAACAACCAAG and MtEm6_F:

GGGGACAAGTTTGTACAAAAAAGCAGGCTCGATGGGGCATCATCATCATC)



21

followed by an additional sequence encoding the FLAG epitope and digestion site for

enterokinase (DYKDDDK) just prior to the start codon and a reverse primer with an AttB2

sequence flanking the stop codon (MtPM25_R:

GGGGACCACTTTGTACAAGAAAGCTGGGTCTTACTTAACATTTTCATTGAGCCTA

GCAGCCGCA and MtEm6_R:

GGGGACCACTTTGTACAAGAAAGCTGGGTTCACTTGTTCTGGCTCCTAC). PCRs

and in vitro BP and LR clonase recombination reactions were carried out according to the

manufacturer's instructions (Invitrogen) using pDON207 and as destination vector pDEST17

(Invitrogen). pDEST17 contains a N-terminal 6xHis-tag. pDEST17-PM25 and pDEST17-

Em6 were transferred into BL21-AI competent cells (Invitrogen) and recombinant protein

production was induced in the presence of 0.2% (w/v) arabinose at 37°C. Bacterial proteins

were extracted by sonication in 50 mM NaH2PO4, pH 8, 300 mM NaCl, 10 mM imidazol, 1

mM PMSF. The 6xHis-tagged recombinant proteins were purified by Ni-NTA affinity

chromatography (Ni-NTA Superflow, Qiagen) under native conditions according to the

manufacturer’s instructions. After digestion by enterokinase (EKMax, Invitrogen) according

to the manufacturer’s instructions, de-tagged MtPM25 and MtEm6 recombinant proteins were

separated from their tag Ni-NTA affinity chromatography, desalted and lyophilized.

Western blot analysis

Twenty µg of proteins per sample were separated by SDS-PAGE using 12% (w/v) acrylamide

separating gels. Following electrophoresis, the gels were transferred onto nitrocellulose

membranes (Schleicher and Schuell, Dassel, Germany) for 1 h at 100 V in 25 mM Tris (pH

8.3), 192 mM Glycine and 20% (v/v) methanol using a mini-transblot system (Bio-Rad,

Hercules, CA, USA). The membrane was then blocked with Tris buffered saline (TBS: 10

mM Tris-HCl pH 7.5, NaCl 150 mM) containing 1.5% Tween 20 for 45 min under constant

agitation and rinsed several times with TBS containing 0.05% (v/v) Tween 20 (TBST). The

membrane was incubated for 1 h at room temperature with a rabbit polyclonal antibody raised

against MtPM25 or MtEm6 (dilution 1: 10,000 in TBST). After washing in TBST, the

membrane was incubated for 1 h with an antirabbit IgG peroxidase conjugate (Biosource

International, Camarillo, CA, USA) diluted 1:50,000 in TBST. After washing in TBST and

TBS, immunodetection was performed by chemiluminescence according to Grelet et al.

(2005).



22

Fourier transform IR spectroscopy

Dry protein films were prepared by drying 5 µL droplets of a solution of lyophilized protein

in water (20 mg/mL) on circular (2*13 mm) CaF2 IR windows at 25°C. Fast drying was

carried out in a stream of dry air (3% RH) and slow drying above saturated NH4NO3 (67%

RH) in a ventilated box. The protein films lost most of their water within five min and 1 h,

respectively, but the samples were left overnight under these conditions before analysis.

Protein samples in D2O were obtained by the adding 0.5 µL D2O to the fast-dried specimens.

Each sample was hermetically sealed between IR-windows using a rubber O-ring and

mounted into a brass holder. These procedures were performed in a cabin continuously

purged with dry air (3% RH) to prevent rehydration of dry samples and exchange with H2O

vapour in the case of samples in D2O. IR spectra were recorded at room temperature on a

Fourier transform IR spectrometer (Perkin-Elmer, Beaconsfield, Buckinghamshire, UK,

model 1725) equipped with a liquid nitrogen-cooled mercury/cadmium/telluride detector and

a Perkin-Elmer microscope as described previously (Wolkers and Hoekstra, 1995). The

optical bench was purged with dry CO2-free air (Balston, Maidstone Kent, UK). The

acquisition parameters were 4 cm-1 resolution, 32 co-added interferograms, 2 cm s-1 moving

mirror speed, and 3600-1000 cm-1 wave number range. Spectral analysis and display were

carried out using Spectrum version 2.00 (Perkin Elmer). For protein studies the spectral

region between 1750 and 1350 cm-1 was selected. This region contains the amide-I and the

amide-II absorption bands of the protein backbones. Secondary structures were derived from

the shape of the amide I band, which is the most sensitive to the secondary structure of

proteins. Second derivative spectra were used to determine the number and the positions of

the secondary structure components as starting parameters for a curve-fitting procedure. To

quantify the contributions of these different components to the amide I band, a least square

iterative curve fitting was performed (Peakfit, Jandel Software, San Rafael, CA) to fit Voigt

line shapes to the original spectrum between 1720 and 1590 cm-1. Prior to curve fitting a

straight base line passing through the ordinates at 1720 and 1590 cm-1 was subtracted. The

proportion of each structure was calculated as the percentage of the sum of areas of all Voigt

bands having their maximum between 1698 and 1618 cm-1. Assignment of structures was

according to Byler and Susi (1986), Surewicz and Mantsch (1988) and Raussens et al. (1997).



23

Sequence data for MtPM25 and MtEm6 have been deposited with the Genbank data library

under the respective accession number DQ206870 and DQ206712.

ACKNOWLEDGMENTS

We thank J. Brettner (SeedCo Australia) for the gift of the seeds, N Sommerer (INRA

Montpellier) for preliminary MALDI-TOF analyses and B. Ly-Vu for dissecting the

thousands of radicles used in this study.

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30

FIGURE CAPTION AND LEGENDS

Figure 1. Two dimensional electrophoresis of the heat stable proteome of non germinated

radicles excised from Medicago truncatula seeds that were imbibed for 16 h using 24 cm non

linear immobilized pH gradient strips (3-10). pI and molecular mass (in kDa) are noted.

Numbers indicate the spots that were identified.

Figure 2. The relation between desiccation tolerance and the water content of the radicles of

Medicago truncatula at different intervals during fast drying. Germinated seeds exhibiting a

protuded radicle of 2.8 and 5 mm were first incubated or not (control) in a PEG solution for 2

d then dried at different intervals. The data from 2 independent experiments are pooled

together.

Figure 3. Phylogenic tree of LEA proteins of Medicago truncatula. ClustalX was used to

create an alignment of the following translated tentative consensus that were found in the

Medicago database (http://www.tigr.org/tdb/tgi/mtgi). The alignment was bootstrapped

(n=1000 replicates) to create the final tree with the bootstrap values indicated. Underlined

LEA proteins indicate those that have been found associated with desiccation tolerance (see

Table III). BudCAR5 (TC 100264 homologue of Q9M603 of M. sativa); CapLEA-1a

(TC94389 similar to 049816 of C. arietinum); CapLEA-1b (TC112317 similar to 049816 of

C. arietinum); DHN (dehydrin; TC101013 similar to 023957 of G. max); DHN3 (or dehydrin-

like, TC 100921, similar to Q945Q7 P. sativum); DHN-Cog (dehydrin-cognate, TC106659

similar to Q43430 of P. sativum); DIP (drought induced protein, TC95389 similar to Q941N0

of R. raetam); ECP31 (TC 96862 similar Q96245 of A. thaliana); Em6 (TC96799 homologue

to P93510 of R. pseudoacacia); Lea14 (TC 101891, similar to P46519 of G. max); LEA5

(TC105834 similar to O24422 of G max), LEA D34a (TC102411 weakly similar to P09444

LEA D34 of G. hirsutum); LEA D34b (combination of TC107159 and –158, both similar to

P09444 LEA D34 of G. hirsutum); LEA-likea (TC108292 weakly similar to Q6Z4J9 of O.

sativa); LEA-likeb (TC 95012 similar to Q8LCW6 of A. thaliana); PM32 (mitochondrial

LEA, TC101811 and TC78559 of Q5NJL5 of P. sativum); MP2 (maturation polypeptide;

TC95538 and TC87025 similar to Q39871 of G. max); MtPM25 (this study); PM1 (TC96465

similar to Q01417 of G. max); PM10 (TC100258, TC85220 similar to Q39801 of G. max);

PM18 (or 35 kDa seed maturation protein, TC96265 similar to Q9ZTY1 of G. max), PM22

(translated BQ124186 similar to Q9XER5 of G. max); PM24 (TC96521 weakly similar to



31

Q9SEL0 of G. max); RAB21 (TC 107383 similar to P12253 of O. sativa); SBP65 (seed

biotinylated protein, TC102224 similar to Q41060 of P. sativum); SMP LEA-4 (TC94509

similar to Q9FNW8 of G. tabacina). Whenever possible, the classification according to Dure

(1993) and Cuming (1999) together with the PFam assignments are indicated. The scale bar

(number of substitutions/site) corresponds to the relative branch length.

Figure 4. Changes in three heat stable proteins identified as PM25 (A, B), Em6 (C, D) and

MP2 (E, F) associated with desiccation tolerance in germinating radicles of Medicago

truncatula. Representative 2D gels (A, C, E) and relative spot quantities (B, D, F) during

germination (open bars) and after a PEG treatment (black bars) in 2.8 and 5 mm long emerged

radicles. Spot numbers are taken from figure 1. pI and molecular mass (in kDa) are indicated

on the left gel. For MP2, the histogram corresponds to the most preponderant isoform (spot

19). Different letters shown above the bars represent significant differences after multiple

comparison of the means (P<0.05).

Figure 5. Changes in spots identified as PM18 (A, B) and SBP65 (C, D) that are associated

with desiccation tolerance in germinating radicles of Medicago truncatula. Representative 2D

gels (A-C) and relative spot intensities (B, D) corresponding to PM18 (B, spots 15-18) and

SBP65 (D, spots 1-5) during germination (open bars) and after a PEG treatment (black bars)

in 2.8 and 5 mm long radicles. Spot number assignment is taken from figure 1. pI and

molecular mass (in kDa) are indicated on the left gel. Different letters shown above the bars

represent significant differences after multiple comparison of the means (P<0.05).

Figure 6. Changes in several spots identified as DHN3 (A, B) and Budcar5 (C, D)

Representative 2D gels (A) and relative spot intensities (B) during germination (open bars)

and after a PEG treatment (black bars) in 2.8 and 5 mm long emerged radicles. Spot number

assignment is taken from figure 1. pI and molecular mass (in kDa) are indicated on the left

gel. Different letters shown above the bars represent significant differences after multiple

comparison of the means (P<0.05).

Figure 7. In silico expression analysis of LEA genes in Medicago truncatula identified in this

study (see table III). The number of expressed sequence tags present in various libraries that

were found at http://www.tigr.org/tdb/tgi/mtgi and exhibited similar characteristics (organs,

developing seeds, stress, etc.) were averaged and expressed as % of detected EST x1000. The



32

absence of value means that the EST was absent in the libraries. The corresponding libraries

are: leaf (T10110, 4046, 4049), flower (#9D5, #ARC), nodulated root (T1617, T10109, 4047),

seed (#ARD, T10493, T10494, T11127), drought (5413), biotic stress (#A8V, T11031 T1581,

T10014), N deficit (#8GI, 5518, #8GF), P starvation (5415), plant microbe interaction

(T1748, 7263, T10173, T1815, T1510, T1707, T1682, #ARE, #CDE, 5520).

Figure 8. Western blot analysis of MtPM25 (A-D) and MtEm6 (E-H) in relation to

desiccation tolerance: in radicles (A, E) and cotyledons (B, F) during seed imbibition; in

emerged radicles (C, G) having a length of 2.8 and 5 mm, before and after the PEG treatment;

in embryos (D, H) during the acquisition of desiccation tolerance during seed maturation (10-

40 days after pollination, DAP). Twenty (embryos, radicles) and 50 µg (cotyledons) of

soluble proteins were hybridized with rabbit antibodies against recombinant MtPM25 and

MtEm6. Independent protein extractions and immunoblots were performed in triplicates and

yielded identical results. Percentage of desiccation tolerance (DT) and molecular marker mass

(kDa) are indicated. DS, dry seeds.

Figure 9. FT-IR absorption spectra in the amide region of detagged, recombinant MtPM25

and MtEm6. Conditions of the proteins were: (A) hydrated in D2O, (B) after fast drying (FD)

in an air stream of 3% RH, and (C) after slow drying (SD) in circulating air of 67% RH.

Figure 10. Curve-fitting procedure illustrated for the recombinant MtEm6 after fast-drying in

an air stream of 3% RH. The absorption maxima of the different protein secondary structures

in the amide-I band were selected on account of peak positions in the second-derivative

spectrum (A). With these different components a least square iterative curve fitting was

performed to fit Voigt line shapes to the original spectrum between 1720 and 1600 cm-1 (B).

Prior to curve fitting a straight base line passing through the ordinates at 1720 and 1600 cm-1

was subtracted. Co-addition of all of the dashed peaks that were mathematically produced

resulted in a fit (crosses) that resembled the original absorption spectrum (grey line).



33

Table I. Characterization of the desiccation tolerance stages together with soluble protein and heat stable protein contents and average number of spots detected in Medicago truncatula radicles. N.G. stage corresponds to 16 h-imbibed, non-germinated seeds; 2.8 and 5 mm stages correspond to germinated seeds exhibiting protruded radicles of 2.8 or 5 mm in length, before (untreated) and after a 2d osmotic treatment at –1.7 MPa using a polyethylene glycol solution (+ PEG). Percentages of desiccation tolerance represent the average (± SE) of 3 independent experiments using 50 seeds. The threshold water content (g H2O/g DW; g/g) is defined as the water content at which viability is reduced by 50% during drying and was derived from Fig. 2. Protein content values (average ± SE) were obtained from 15 replicates of 100 radicles. N.D. not determined.

Stages of germination and osmotic treatment 2.8 mm 5.8 mm

N.G. Untreated + PEG Untreated + PEG Desiccation tolerance (%) 100 0 91 ± 6 0 0 Threshold water content (g /g) <0.03 0.3 <0.03 3.6 0.8 Protein content (µg/radicle) 60 ± 4 36 ± 5 32 ± 4 29 ± 2 23 ± 2 Heat stable protein content (µg/radicle)

17 ± 1 7 ± 1 5 ± 1 5 ± 0 5 ±1

Number of detected spots 328 ± 6 252 ± 2 267 ± 2 N.D. N.D.



34

Table II. Synopsis of expression profiles of heat stable proteins of Medicago truncatula radicles following the loss of desiccation tolerance during germination and the re-establishment of desiccation tolerance by a PEG treatment. The stages analyzed were non-germinated (tolerant, 1), 2.8 mm long emerged radicles before (sensitive, 2) and after PEG treatment (tolerant, 3). The percentage of spots is relative to the total amount of spots detected in the analysis. As an aid to the profile description, the bar diagrams schematize changes in the spot abundance.

Profile type Characteristics Nb

spots % of total

spots

1 1 2 3

constant levels in the 3 stages 54 13.8

2 1 2 3

highest levels before germination, decreasing significantly in the 2.8mm stage and remaining low in 2.8mm+PEG stage

111 28.4

3 1 2 3

higher levels in 2.8 mm radicles than in both desiccation-tolerant stages

32 8.2

4 1 2 3

low levels before and after germination, increasing significantly in PEG-treated 2.8 mm long radicles

44 11.3

5 1 2 3

levels decreasing steadily in the 3 stages 7 1.8

6 1 2 3

levels increasing steadily in the 3 stages 37 9.5

7 1 2 3

high levels before and after germination, thereafter decreasing significantly upon the PEG treatment

19 4.9

8 1 2 3

low levels before germination, increasing significantly in the 2.8 mm stage and remaining constant in 2.8mm+PEG stage

50 12.8

9 1 2 3

increased levels in both desiccation tolerant stages 23 5.9



35

Table III. Identified heat stable polypeptides that are associated with desiccation tolerance in radicles from Medicago truncatula. Identifier of consensus sequences were retrieved from the Medicago database (MtGI version 8, at http://www.tigr.org/tdb/tgi/mtgi) except for spot 36 which was identified in the version 7. NA indicates no MALDI or LC MS/MS was used in the identification. The group designations of the LEA proteins according to Cuming (1999) are indicated in brackets.

Cate-gory

Spota Exp. Mr/pI

MALDIb Q-TOFc Homology

Best matching TC/ GB #

Theor. Mr/ pI

A 21 32.5/4.50 NA 8 (44) Seed maturation protein PM25 (G. max) (group 5) CA917414 24.9/4.80 A 37 15.2/5.71 7 12 (53) Em protein (Robinia pseudoacacia) (group 1) TC96799 10.7/5.93 A 2 76.7/5.80 NA 9 (45) Sbp65a protein (SBP65) (P.sativum) (group 3) TC102224 38.7/5.59 A 3 76.0/5.80 NA 10 (114) Sbp65a protein (SBP65) (P.sativum) (group 3) TC102224 38.7/5.59 A 6 65.8/6.43 NA 1 (18) Sbp65a protein (SBP65) (P.sativum) (group 3) TC102224 38.7/5.59 A 36 17.5/6.46 7 NA Legumin precursor (P. sativum) TC85216 64.9/5.40 A 19 37.0/6.10 9 NA Maturation polypeptide MP2 (G. max) (group 3) TC95538 38.4/6.54 A 16 41.1/5.55 NA 7 (45) 35 kDa seed maturation protein PM18 (G. max) (group 3) TC96265 36.3/5.52 B 1 77.7/5.78 NA 7 (67) Sbp65a protein (SBP65) (P.sativum) (group 3) TC102224 38.7/5.59 B 4 75.5/5.95 NA 11 (120) Sbp65a protein (SBP65) (P.sativum) (group 3) TC102224 38.7/5.59 B 5 77.5/6.01 5 NA Sbp65a protein (SBP65) (P.sativum) (group 3) TC102224 38.7/5.59 B 11 42.5/6.30 NA 2 (33) DHN3 (dehydrin-like protein, M. sativa) (group 2) TC100921 31.2/6.45

a) Spot identification number (see Fig. 1); b) numbers of peptide matched are given when a significant hit was found, c) number of sequenced peptides by nanoscale LC-MS/MS. The values in brackets indicate the total number of amino acids sequenced once.

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36

Table IV Band positions and individual contributions by the various secondary structures of the untagged recombinant forms of MtEm6 and MtPM25 in D2O (hydrated state) and after fast and slow drying, determined by curve-fitting of the composite amide-I band of the FTIR spectra (see Fig. 10 for details). Assignment of structures was according to Byler and Susi (1986), Surewicz and Mantsch (1988) and Raussens et al. (1997).

Em6 MtPM25 Conditions

Position ν (cm-1)

Area (%)

Position ν (cm-1)

Area (%)

Assignment

Hydrated (D2O)

1622 8 1625 18 Extended ß-sheet

1636 23 1638 26 Coil 1651 37 1653 33 α-helix 1667 20 1668 15 Turns 1680 10 1681 8 Turns 1692 2 1693 0 Extended ß-sheet Fast dried 1627 5 1627 11 Extended ß-sheet 1641 11 1639 14 Coil 1658 57 1658 54 α-helix 1678 20 1680 15 Turns 1692 7 1692 6 Extended ß-sheet Slowly dried 1623 5 1623 24 Extended ß-sheet 1641 15 1637 10 Coil 1655 60 1655 56 α-helix 1681 17 1680 9 Turns 1695 3 1690 1 Extended ß-sheet

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76

43

21.5

17

4.5 5.5 6.0 7.6

1 2 3 4 5

6

7 8 9

11 12 13

2425

181715 16

19 20

2322

26

21

14

29

30

27

28

37

31 3233

3536

39

38

34

41

40





0.1

CapLea-1a

CapLea-1b 1000

PM18

MP2 999

PM1

SMP-LEA4 1000

301

PM10

PM32 459

386

320

LEA5

LEA-D34a

PM24

LEA-D34b

PM25

ECP31 803

673

1000 1000

311

DHN-Cognate

Lea14

LEA-likea

LEA-likeb 1000

996

196

RAB21

BudCar5

DIP 1000

368

DHN

DHN3 569

111

144

172

SBP65

Em6

280

G1 (D19), PF00477

G5 (D34), PF04927

(D73) PF03242

G2 (D11) PF00257

G2 (D11) PF00257

G3 (D7) PF 02987

G5 (D95), PF03168

G3 (D7) PF 02987

G4 (D113) PF 03760

G3 (D7) PF 02987



4.5

37.5

31

15.2

18.7

5.7

NG

Radicle length2.8 mm

- PEG + PEG

A 5 mm

- PEG + PEG

G erm ination stagesNG 2.8 m m 5 m mN

orm

aliz

ed s

pot q

uant

ity (

X10

3 )

0 ,0

0,5

1,0

1,5

2,0 controlP EG treated

a

b

c

d d

21

Germ ination stagesNG 2.8 m m 5 m mN

orm

aliz

ed s

pot q

uant

ity (

x103 )

0

5

10

15

controlPEG treated

a

c

dd

b

37

B

C D

E

Germination stages

NG 2.8 mm 5 mmNor

ma

lize

d sp

ot q

uant

ity (

x 1

03 )

0

4

8

12

16

a

b

c

d d

19

F

42

37

6.1 6.5

2019

2.8 mm

- PEG + PEG

2.8 mm

- PEG + PEG

NG 5 mm

- PEG + PEG

5 mm

- PEG + PEGNG

21

37



Germination stagesNG 2.8 mm 5 mm

0

1

2

3

Germination stagesNG 2.8 mm 5 mmN

orm

aliz

ed s

pot q

uant

ity

(x10

3 )

0

1

2

3


0

2

4

6


4

7

11

1415 18

aa ab b

a a

c

aa a

aa a

b bb b

b

c

16 17

control PEG-treated

Germination stagesNG 2.8 mm 5 mmN

orm

aliz

ed s

pot q

uan

tity

( X

103 )

0

1

2

3

4


0

1

2

3

4


0

2

4

6

8

NG 2.8 mm 5 mm0

2

4

6

81 2 3 4 5


0

1

2

a

bc

bd

b

a

c

b b

a

b

c

ed

a

b

c

d b

a

bc

d d

Germination stages

control PEG-treated

5.3 5.8

NG

Radicle length

2.8 mm 5 mm+ PEG- PEG - PEG+ PEG

42

15 1515 15 1716 16 1617 17

18 1817

1816

18

C

B

18171615

75.5

61

5.8 6.0

A

D1 5

4

3

2

1 5

4

3

2

1 5

4

3

2

1 5

4

3

2

1 5

4

3

2

NG 2.8 mm 5 mm+ PEG- PEG - PEG+ PEG



A

8703

52

6.1 6.5

NG

Radicle length

+ PEG2.8 mm 5 mm

- PEG - PEG+ PEG

37

1112

13 11 11 1112 13 12 1312 13 131211

B

31 31 32 3332 33

31

Germination stages

NG 2.8 mm 5 mm

Nor

mal

ized

spo

t qua

ntity

( x

103 )

0

2

4

632

Germination stages

NG 2.8 mm 5 mm0

2

4

633

Germination stages

NG 2.8 mm 5 mm0

2

4

6

control PEG-treated

a

b

a aa a a aa a

b

b

cbb

D

5.8 5.9

25.2

21.6

C+ PEG

2.8 mm 5 mm- PEG - PEG+ PEG

NG

control PEG-treated


0

3

6

9


0

6

12

18


0

3

6

911 12 13

ab

ca

b

c c

cab

c

bd

a

a

Nor

mal

ized

spo

t qua

ntity

(X

103 )





D30262220181614

Maturation (DAP)

12 35 37 DS

25

37

% DT: 0 0 31 54 97 97 100 100

B E F

G

A

C

6 15 17 20 24Imbibition (h)

20 24 48 60Imbibition (h)

- - ++PEG:2.8 5

Radicle length (mm)

% DT: 0 96 0 0

0 0100% DT: % DT:

2.8 5Radicle length (mm)

- - ++PEG:

% DT: 0 96 0 0

100 98 59 0 0

3 6 15 20Imbibition (h)

4815 20Imbibition (h)

% DT: 100 0100100 100 98 0

H Maturation (DAP)

% DT:

30262220181614

0 31 54 97 97 10015

20

MtPM25 MtEm6

100







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