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Plant Physiol. Apr 2006; 140(4): 1418–1436.
PMCID: PMC1435805

Comparative Analysis of the Heat Stable Proteome of Radicles of Medicago truncatula Seeds during Germination Identifies Late Embryogenesis Abundant Proteins Associated with Desiccation Tolerance1,[W]


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 reinduced, albeit that drought tolerance was increased. The abundance of 15 polypeptides was linked with DT, out of which 11 were identified as late embryogenesis abundant 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 three 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 late embryogenesis abundant proteins might exert different protective functions at high and low hydration levels.

Desiccation tolerance (DT) corresponds to the ability to survive nearly complete protoplasmic dehydration (approximately −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 nonreducing 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 five 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 (HS) 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 groups 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 (Arabidopsis thaliana) that are deficient in several HS 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, homolog 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 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 groups 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 overexpressed 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, and 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 (Zea mays; Niogret et al., 1996). When these genes are overexpressed in desiccation-sensitive systems such as yeast (Saccharomyces cerevisiae; Swire-Clark and Marcotte, 1999), leaves of rice (Oryza sativa; 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 (approximately −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 nonreducing 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 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 water/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 groups 1, 2, and 3 are submitted to posttranscriptional and posttranslational 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 focused on the HS proteome that resists 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 HS 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 that 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 HS proteins were compared between desiccation-tolerant radicles of nongerminated (NG) 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 reestablished. 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 reestablishment of DT were further analyzed in older emerged radicles, for which DT can no longer be reinduced by the osmotic treatment. This strategy allowed for the 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 Fourier transform infrared (FTIR) spectroscopy.


Changes in HS Protein Patterns in Relation to the Loss and Reestablishment of DT

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% relative humidity (RH) at 20°C (Table I). Previously, Buitink et al. (2003) showed that DT can be reestablished 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 postgerminative growth, corresponding to seeds with protruded radicles of 5 mm in length, DT can no longer be reestablished after the same PEG treatment (Table I).

Table I.
Characterization of the DT stages together with soluble protein and HS protein contents and average number of spots detected in M. truncatula radicles

To identify proteins involved in DT, we compared the HS proteome extracted from 2.8 mm long, desiccation-sensitive radicles with those from 16 h-imbibed NG 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 reestablished. In 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 three stages were analyzed by two-dimensional gel electrophoresis (2DE) using a nonlinear pI gradient (Fig. 1). For each stage, the spots from six to eight replicates were detected and compared to each other using the PDQuest software. To secure the quality of the data, spots of poor quality and very low raw volumes were discarded using criteria set 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 reestablishment 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 nine 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%).

Figure 1.
Two-dimensional electrophoresis of the HS proteome of NG radicles excised from M. truncatula seeds that were imbibed for 16 h using 24 cm nonlinear immobilized pH gradient strips (3–10). pI and molecular mass (in kilodaltons) are noted. Numbers ...
Table II.
Synopsis of expression profiles of HS proteins of M. truncatula radicles following the loss of DT during germination and the reestablishment of DT by a PEG treatment

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 subgroups: those that are only induced in the 2.8 mm long radicles after PEG treatment and are thus linked specifically to DT (subgroup A) and those that are also induced by the PEG incubation in 5 mm long radicles that remain desiccation sensitive (subgroup B). Out of the 23 spots, 11 were found to be associated specifically with the induction of DT (subgroup A) and seven 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 seven spots, five exhibited a significantly higher intensity in 2.8 mm PEG-treated radicles than in the 5 mm ones. They were therefore also linked to DT. The remaining five spots 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 reestablishment 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; Table I). Germinated seeds with an emerged radicle of 2.8 mm long were able to survive a desiccation treatment down to 1 g/g, but died at lower water content. Fifty percent 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 were thus considered desiccation tolerant (Fig. 2). In contrast, 5 mm long radicles were very sensitive to drying. Fifty percent 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 (Glycine max) 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 reinduced by the PEG incubation in 5 mm long radicles that remained desiccation sensitive (data not shown).

Figure 2.
The relation between DT and the water content of the radicles of M. truncatula at different intervals during fast drying. Germinated seeds exhibiting a protruded radicle of 2.8 and 5 mm were first incubated or not (control) in a PEG solution for 2 d then ...

Identification of HS Proteins Associated with DT

The 16 spots belonging to profile 9A and 9B and linked with DT were analyzed using matrix-assisted laser desorption ionization time of flight (MALDI-TOF) mass spectrometry and liquid chromatography-tandem mass spectrometry (LC-MS/MS). Out of these 16 spots, 11 were identified as six different LEA proteins, some of them being present as different isoforms (Table III). Another polypeptide was identified as a homolog of a pea (Pisum sativum) legumin precursor (Table III). Since the Mr 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.

Table III.
Identified HS polypeptides that are associated with DT in radicles from M. truncatula

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 Institute for Genomic Research (http://www.tigr.org/tdb/mtgi) and National Center for Biotechnology Information (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 six LEA proteins identified in this study in relation to DT belonged to four different groups according to Cuming's classification. One DT-linked spot (Table III; Fig. 1, spot 37) was identified as a homolog of Em6 of Arabidopsis, 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 members was detected in relation to DT and named PM25 due to the high similarity with GmPM25 from soybean (Table III; Fig. 1, spot 21). One isoform of DHN3 (spot 11) was linked to DT. According to Figure 3, it is a member of 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 a homogeneous clade. Three members of this group were found to be correlated to DT. Among them were two closely related proteins, homologs of PM18 and MP2 of soybean (Table III; Fig. 1, spots 16 and 19) and one unrelated protein, identified as a homolog of SBP65 of pea, a biotinylated protein (Table III, spots 2–6).

Figure 3.
Phylogenic tree of LEA proteins of M. truncatula. ClustalX was used to create an alignment of the following translated tentative consensus that was found in the Medicago database (http://www.tigr.org/tdb/tgi/mtgi). The alignment was bootstrapped (n = ...

Analysis of the changes in relative volume of Em6 and PM25 (Fig. 4, B and D) demonstrated that the PEG treatment significantly reinduced the expression of both proteins in the 2.8 mm long emerged radicles. Nonetheless, the reestablishment of DT did not lead to a similar reinduction of the protein abundance as was the case in the NG, 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 (Figs. 4 and and5).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 and 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; Supplemental Table I). Three spots corresponding to PM18 (spots 15, 16, and 17) showed a different expression profile (Fig. 5A). Spot 16 (Fig. 5, A and B) was found to be linked to DT whereas the most acidic polypeptide (15) increased upon osmotic treatment regardless of 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 four digested peptides, the masses of which matched some obtained by the theoretical digestion of the translated Medicago tentative consensus (TC) similar to PM18. The third member of group 3, SBP65, existed in six isoforms with different pIs (spots 1–5, Fig. 5, C and D) and Mr (spot 6, Fig. 1). Again, the different isoforms were classified in two profiles: 9A, reinduced only in 2.8 mm long PEG-treated radicles and 9B, also reinduced in 5 mm long PEG-treated radicles (Fig. 5D). Nonetheless, for all isoforms, the abundance was significantly higher in those tissues that were desiccation tolerant than in those that remained sensitive.

Figure 4.
Changes in three HS proteins identified as PM25 (A and B), Em6 (C and D), and MP2 (E and F) associated with DT in germinating radicles of M. truncatula. Representative 2D gels (A, C, and E) and relative spot quantities (B, D, and F) during germination ...
Figure 5.
Changes in spots identified as PM18 (A and B) and SBP65 (C and D) that are associated with DT in germinating radicles of M. truncatula. Representative 2D gels (A–C) and relative spot intensities (B and D) corresponding to PM18 (B, spots 15–18) ...

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 (group 2) that were identified in this profile were DHN3 and BudCar5, both being present in several isoforms (Fig. 6). DHN3 was present in three isoforms (spots 11–13, Fig. 6, A and 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 most acidic form (11) responded to the osmotic treatment by increasing approximately 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. 6, C and D). Dehydrins are known to be expressed under different types of stress and in different tissues, thus their induction upon PEG incubation was expected.

Figure 6.
Changes in several spots identified as DHN3 (A and B) and Budcar5 (C and D). Representative 2D gels (A) and relative spot intensities (B) during germination (white bars) and after a PEG treatment (black bars) in 2.8 and 5 mm long emerged radicles. Spot ...

To investigate whether the six DT-linked 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 HS 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 nine groups (Fig. 7; Supplemental Table II). Except for DHN3, the remaining five LEA proteins that were identified in relation to DT appeared to be 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 all the LEA proteins shown in the phylogenetic tree (Fig. 3) revealed that all LEA genes from groups 1 to 4 that were specifically expressed in seeds (Supplemental 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 two of the 23 known LEA genes (DIP, a dehydrin and CapLea1B, member of group 3) were not present in at least one of the seed libraries (Supplemental Table II).

Figure 7.
In silico expression analysis of LEA genes in M. truncatula identified in this study (see Table III). The number of ESTs present in various libraries that were found at http://www.tigr.org/tdb/tgi/mtgi and exhibited similar characteristics (organs, developing ...

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, Mr around 16,000) were also identified as homologs to the pea legumin precursor (theoretical Mr 65,000, TC85216). Four spots (22, 23, 24, and 25; Mr around 31,000; Fig. 1) were identified as homologs of the pea convicilin (TC100299) having a theoretical Mr of 78,300. These fragments of storage proteins belonged to profiles 2, 3, and 5 (Table II) and were not very abundant. Likewise, the HS proteome included proteins other than LEA proteins (for example, homologs of a Vicia faba transcription factor [TC94137], an ankyrin repeat protein 2 from Vitis aestivalis [TC100495], and a Gly-rich protein 2 of Nicotiana sylvestris [TC98399]; see Supplemental Table I).

Changes in MtPM25 and MtEm6 in Relation to DT 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 an 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 Escherichia coli. Rabbit polyclonal antibodies were raised against the purified 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 preimmune 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. 8, A and E). In 20 h imbibed radicles (approximately 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 reinduce the expression of both proteins in 2.8 mm long emerged radicles, albeit to lower levels than those found in NG radicles (Fig. 8, C and 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. 8, B and 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 d 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.

Figure 8.
Western-blot analysis of MtPM25 (A–D) and MtEm6 (E–H) in relation to DT: in radicles (A and E) and cotyledons (B and F) during seed imbibition; in emerged radicles (C and G) having a length of 2.8 and 5 mm, before and after the PEG treatment; ...

Secondary Structure Analysis of MtPM25 and MtEm6 Proteins

It has been established that LEA proteins of groups 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. FTIR 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 1,646 cm−1 with the amide-I band between 1,700 and 1,600 cm−1, D2O instead of water was used for the proteins in solution. Wolkers et al. (1998) demonstrated that intermolecular β-sheet formation can effectively be prevented by fast drying, probably because the time required for such nonintramolecular rearrangements is too short. For this reason, we studied the recombinant proteins after fast and slow drying in air of approximately 3% and 67% RH, respectively.

Figure 9.
FTIR 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. ...

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 (1,460 cm−1) that was lower than in the case of the fast-dried proteins (1,550 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 approximately 1,658 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 approximately 1,630 cm−1, which is less prominent in the fast-dried MtEm6. This is suggestive of a larger proportion of intermolecular β 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 Figure 10 with fast-dried MtEm6 protein. Peaks representing different secondary structures were selected on account of the second derivative spectrum (Fig. 10A). Coaddition of all the dashed peaks that were mathematically produced should result in a fit (crosses) that resembles the original absorption spectrum (Fig. 10B). 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 β 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% β sheet, and 53% unordered structures, whereas MtPM25 would consist of 33% α helix, 18% β sheet, and 49% unordered structures (Table IV). These figures concur fairly well with PELE predictions (San Diego Supercomputer Center Biology workbench: http://workbench.sdsc.edu). In water solution, MtEm6 is predicted to form 33% α helix, 3% β sheet, and 64% unordered structures, whereas 38% α helix, 14% β 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 1,540 cm−1 in D2O (Fig. 9). This band was considerably smaller than that upon fast drying and partly downshifted to 1,450 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).

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 ...
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 ...

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 β sheets upon slow drying, which appeared to be reversible upon rehydration. When indicated as percentages of α helix, β sheet, and unordered structures, MtEm6 consisted of 57%, 12%, and 31%, and 60%, 8%, and 32%, after fast and slow drying, respectively. Data for MtPM25 were 54%, 17%, and 29%, and 56%, 25%, and 19%, respectively.


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 reestablishment 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 reinduced by the same osmotic treatment. In total, 15 polypeptides were found, whose abundance was linked to DT. Among them 11 were identified, which represented six 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, ,5,5, and and8).8). Nonetheless, the causal relationship between the six 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 nonreducing 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 (for review, see Buitink and Leprince, 2004). In vitro, a mixture of LEA proteins and sugars forms a glass upon drying that exhibits physicochemical properties resembling those of cytoplasmic glasses, whereas a glass made of sugars 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 5 mm long radicles did not reestablish DT, it did lead nevertheless to an increased tolerance to drying, evident from the reduction in the threshold water content from 3.6 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 (Triticum aestivum) 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 homologs of MtEm6 were correlated with seed longevity of Brassica napus (Soeda et al., 2005) and the wheat homolog 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, overexpression of the wheat Em in yeast cells (Swire-Clark and Marcotte, 1999) and also of the Arabidopsis homolog 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 overexpressing the PM2 (Liu and Zheng, 2005). 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 nonreducing sugars, which act as osmolytes during hyperosmotic stress and 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, 2005; 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 absence of heat stability (Cuming, 1999; Ramanjulu and Bartels, 2002). Indeed, MtPM25 was the least hydrophilic from the six LEA proteins identified in this study. However, MtPM25 was HS and the contention that group 5 proteins are not HS 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 FTIR 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]; nematodes [Goyal et al., 2003]; and dehydrins [Soulages et al., 2003]), to 60% to 70% unordered (GmPM16, a soybean group 4 LEA protein [Shih et al., 2004] and Em proteins [Table IV; McCubbin et al., 1985]), and finally down to 50% unordered (MtPM25 [Table IV] and an Arabidopsis 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 groups 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 are 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 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 groups 3 and 4 LEA proteins mentioned above, this study shows that it is also the case for members of groups 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 cannot 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% to observe a change in the secondary structure (β-sheet formation). The corresponding hydration level is around 0.2 to 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 Suc 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 showing that dehydrins are submitted to posttranslational modifications such as phosphorylation 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 and6).6). This raises the question as to whether the regulation of posttranslational modifications might be also important for DT.

SBP65 exhibits a peculiar posttranscriptional 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 (Garay-Arroyo et al., 2000). Hydrophilins are defined as proteins having a Gly content >6% and a hydrophylicity index >1. The analysis of the HS 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 cannot 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 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 posttranslational modifications.


Plant Material and Treatments

Seeds of Medicago truncatula Gaertn. (cv Paraggio; Seedco 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 16 h-imbibed seeds prior to radicle emergence and seeds exhibiting a protruded radicle length of 2.8 mm (approximately 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 exhibited 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

Soluble proteins were extracted from 50 (western blots) and 100 to 300 (2DE) radicles at 4°C in 400 and 950 L of buffer, respectively (50 mm HEPES pH 8.0, 1 mm EDTA, and 14% [v/v] of the protease inhibitor cocktail complete Mini [Roche Diagnostics Molecular Biochemicals]), 43 units of Dnase I, and 5.3 units of Rnase A. After two consecutive centrifugations at 13,000g at 4°C, the resulting supernatant was heated for 10 min at 95°C, cooled for 15 min on ice, and centrifuged at 13,000g for 15 min at 4°C. The resulting supernatant corresponded to the HS fraction. Protein concentrations were assayed according to Bradford (1976). For the 2DE, the HS proteins were precipitated with 20% (w/v) trichloroacetic acid on ice then centrifuged at 13,000g 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], and 1% [v/v] biolytes from Bio-Rad).


Twenty-four centimeters of immobilized pH gradient (non linear from 3–10) strips (Bio-Rad) containing 500 μg of HS 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 isoelectric focusing 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, and 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) according to Gallardo et al. (2001) using a modified running buffer containing 15.6 mm Tris (pH 8.3), 120 mm Gly, and 0.1% (w/v) SDS. The experiments were set up in randomized block design where six gels corresponding to independent protein extractions from various desiccation-tolerant and -sensitive stages were run in parallel. Four to eight 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

Gels were stained with 0.08% (w/v) Brilliant 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 analyzed using the PDQuest 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% (maximum 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): the normalized spot quantities and the stage were, respectively, the variable and the factor, whereas the extraction and the 2DE 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 5 mm and 5 mm + 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 5 mm + 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 (Bio-Rad).

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 NH4HCO3, followed by dehydration with 100 μL of 50% (v/v) acetonitrile in 25 mm NH4HCO3. Proteins were reduced and alkylated by incubation for 1 h 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). 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). One microliter 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, and 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) coupled to a hybrid quadrupole orthogonal acceleration time-of-flight mass spectrometer (Q-TOF Global, Micromass-Waters). 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 s on the mass range 400 to 1,500 mass-to-charge ratio 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 optimized for various mass ranges and charge states of precursor ions. Mass data obtained by MALDI-TOF or LC-MS/MS were analyzed with the Protein Lynx Global Server software (Micromass-Waters). Protein identification was performed by comparing the data with the UniProt sequence databank or with The Institute for Genomic Research 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 Escherichia 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: GGGGACAAGTTTGTACAAAAAAGCAGGCTTAGATTACAAGGATGATGATGATAAGATGAGTCAAGAACAACCAAG and MtEm6_F: GGGGACAAGTTTGTACAAAAAAGCAGGCTCGATGGGGCATCATCATCATCTAAACAACAAAACCG) 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: GGGGACCACTTTGTACAAGAAAGCTGGGTCTTACTTAACATTTTCATTGAGCCTAGCAGCCGCA 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 an 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) Ara at 37°C. Bacterial proteins were extracted by sonication in 50 mm NaH2PO4 pH 8, 300 mm NaCl, 10 mm imidazol, and 1 mm phenylmethylsulfonyl fluoride. The 6xHis-tagged recombinant proteins were purified by nickel-nitrilotriacetic acid agarose (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, MtPM25 and MtEm6 recombinant proteins were separated from their tag by Ni-NTA affinity chromatography, desalted, and lyophilized.

Western-Blot Analysis

Twenty micrograms 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) for 1 h at 100 V in 25 mm Tris (pH 8.3), 192 mm Gly, and 20% (v/v) methanol using a mini-transblot system (Bio-Rad). The membrane was then blocked with Tris-buffered saline (TBS; 10 mm Tris-HCl pH 7.5 and 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) diluted 1:50,000 in TBST. After washing in TBST and TBS, immunodetection was performed by chemiluminescence according to Grelet et al. (2005).

FTIR 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 5 min and 1 h, respectively, but the samples were left overnight under these conditions before analysis. Protein samples in D2O were obtained by 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 water vapor in the case of samples in D2O. IR spectra were recorded at room temperature on a FTIR spectrometer (Perkin-Elmer, 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). The acquisition parameters were 4 cm−1 resolution, 32 co-added interferograms, 2 cm s−1 moving mirror speed, and 3,600 to 1,000 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 1,750 and 1,350 cm−1 was selected. This region contains the amide-I and 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) to fit Voigt line shapes to the original spectrum between 1,720 and 1,590 cm−1. Prior to curve fitting a straight base line passing through the ordinates at 1,720 and 1,590 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 1,698 and 1,618 cm−1. Assignment of structures was according to Byler and Susi (1986), Surewicz and Mantsch (1988), and Raussens et al. (1997).

Sequence data for MtPM25 and MtEm6 have been deposited with the GenBank data library under the respective accession numbers DQ206870 and DQ206712.

Supplementary Material

Supplemental Data:


We thank J. Brettner (SeedCo Australia) for the gift of the seeds, N. Sommerer (Institute National de la Recherche Agronomique Montpellier) for preliminary MALDI-TOF analyses, and B. Ly-Vu (Institut National d'Horticulture) for dissecting the thousands of radicles used in this study.


1This work was supported by grants from the Contrat de Plan Etat-Région-des Pays-de-la Loire 2000–2006, Institut National de la Recherche Agronomique, and Van Gogh Netherlands Organization for Scientific Research/EGIDE.

The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Olivier Leprince (rf.hni@ecnirpel.reivilo).

[W]The online version of this article contains Web-only data.

Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.105.074039.


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