We isolated a dehydrin-like (DHN-like) gene fragment, PpDHNA, from the moss Physcomitrella patens by PCR amplification using degenerate primers directed against conserved amino acid segments of DHNs of higher plants. The full-length cDNA was found to encode a 59.2-kDa glycine-rich protein, DHNA, with typical characteristics of DHNs, including the presence of several Y repeats and one conserved K segment. DHNA had a high sequence similarity with a protein from Tortula ruralis, Tr288, which is thought to be involved in cellular dehydration tolerance/repair in this moss. Northern and Western analysis showed that PpDHNA is upregulated upon treatment of plants with abscisic acid, NaCl or mannitol, indicating a similar expression pattern to DHNs from higher plants. To analyze the contribution of DHNA to osmotic stress tolerance, we generated a knockout mutant (dhnA) by disruption of the gene using homologous recombination. Growth and stress response studies of the mutant showed that dhnA was severely impaired in its capacity to resume growth after salt and osmotic-stress treatments. We provide direct genetic evidence in any plant species for a DHN exerting a protective role during cellular dehydration allowing recovery when returned to optimal growth conditions.
In order to withstand and survive periods of low water accessibility, plants have evolved a number of protective cellular responses including stomatal closure, accumulation of osmolytes, oxidative stress protection, increased levels of the phytohormone abscisic acid (ABA) and accumulation of novel proteins (Bray, 1997; Chandler and Robertson, 1994; Ingram and Bartels, 1996; Shinozaki and Yamaguchi-Shinozaki, 1997). Considerable research in the last decade has focused on the stress-induced gene expression in order to elucidate the genetic and molecular basis underlying plant cellular dehydration tolerance. This has led to the identification of several plant genes that are upregulated in response to water stress (Bray, 1997; Oono et al., 2003; Palva, 1994; Shinozaki and Yamaguchi-Shinozaki, 1997; Zhu et al., 1997). The gene products characterized can be divided in two groups: regulatory and response proteins (Bray, 1997). The regulatory proteins include kinases, phosphatases, transcription factors, phospholipase C and 14-3-3 proteins, all forming part of signal transduction cascades leading to gene induction or direct cellular responses. The actual environmental stress response is thought to be generated by a second group of proteins that include aquaporins, enzymes for osmolyte biosynthesis, chaperones, desaturases, proteases, detoxification enzymes and late embryogenesis abundant (LEA) proteins. Many of these proteins accumulate to high levels in response to cellular dehydration and are therefore suggested to play a role in abiotic stress tolerance, although their functional analysis remains fragmentary.
Some of the most studied proteins that accumulate in response to water stress in higher plants are the group 2 LEA proteins, or dehydrins (DHNs; reviewed by Close, 1997; Svensson et al., 2002). DHNs have been divided into five subclasses based on their conserved amino acid sequences forming the Y, S and K segments (Close, 1997; Svensson et al., 2002). The Y segment (DEYGNP) is usually found in one to three copies in the N-terminal region of these proteins. Many DHNs contain a stretch of 5 to 7 serine residues followed by three acidic amino acids, termed the S segment. These residues can be phosphorylated, (Godoy et al., 1994; Plana et al., 1991) which may result in their nuclear targeting (Jensen et al., 1998). The K segment (EKKGIMEKIKEKLPGH) is the only segment that is found in all DHNs and is usually present in several copies. This motif has been proposed to form an amphipathic α helix (Dure, 1993), which may allow interaction with both membranes and partially denatured proteins (Close, 1996). DHNs have been found in all higher plant species studied so far, including angiosperms and gymnosperms (Close, 1997). There are also a few reports showing the existence of similar proteins in lower land plants (Velten and Oliver, 2001).
During stress, DHNs accumulate in most tissues and cells and are mainly localized in the cytoplasm and nucleus of the plant cell, although some DHNs have been found to interact with the chloroplast (Mueller et al., 2003), mitochondria (Borovskii et al., 2002) and tonoplast (Heyen et al., 2002). Some DHNs have been shown to accumulate constitutively in the root tip, open stomata and cells surrounding the vascular tissue, suggesting that these proteins might play a role in specific cells or tissues under non-stress conditions (Nylander et al., 2001).
DHNs have been proposed to have a protective function during abiotic stress via a number of different mechanisms. These include improving or protecting enzyme activity under cold or dehydration conditions (Lin and Thomashow, 1992; Rinne et al., 1999; Sanchez-Ballesta et al., 2004); or acting as radical scavengers (Hara et al., 2003) or as membrane stabilizers (Close, 1996; Koag et al., 2003). Nevertheless, there is still a lack of compelling genetic evidence showing a direct role for DHNs in abiotic stress tolerance in higher plants. This has partly been hindered by the difficulties to study the response of a specific tissue or cell type during stress conditions and by the fact that DHNs are usually encoded by multiple genes with possible overlapping function. Thus, many studies with specific members of this protein family have failed to show a clear correlation between accumulation of individual DHNs and stress tolerance (Iturriaga et al., 1992; Kaye et al., 1998). However, recent studies have suggested a cold protective role for some members of this protein family (Bravo et al., 2003; Hara et al., 2003; Zhang et al., 2000). Moreover, experiments where multiple DHN genes were overexpressed simultaneously in Arabidopsis resulted in an enhancement of freezing tolerance, suggesting a role for DHNs in stress tolerance (Puhakainen et al., 2004).
Strategies based on reverse genetics can be very powerful to establish the function of genes by the generation of knockout mutants. Given the lack of efficient homologous recombination, disrupted individual genes in higher plants have been obtained by screening of random mutants in a few plant species. Recently, the moss Physcomitrella patens has become a model plant to study gene function by targeted gene disruption due to a high frequency of homologous recombination (Schaefer and Zrÿd, 1997). Moreover, the phenotypic characterization of mutants is facilitated both by the fact that the haploid gametophyte is the dominant stage of the plant life cycle and also by the relatively simple plant developmental pattern and organization. Recent studies demonstrated that P. patens exhibited a high tolerance against drought, salt and osmotic stress, making it an excellent system to study abiotic stress adaptation in plants (Frank et al., 2005).
In this study, we demonstrate the existence of a DHN gene in P. patens, PpDHNA. We show that this gene is upregulated by ABA, salt and osmotic stress and that a knockout mutant of PpDHNA shows severely impaired stress recovery. This demonstrates that DHNA plays an important protective role during stress, allowing for subsequent recovery of the plant.
Identification of a dehydrin gene from Physcomitrella patens
In order to identify DHN-like genes from P. patens we used degenerate primers designed to amplify the conserved Y and the K segments of known DHNs. A 900-bp PCR product was amplified from genomic DNA and cloned, using a primer combination described in Experimental procedures. Sequencing and subsequent analysis of the amplified product revealed that the partial gene fragment encoded several Y segments and one K segment found in DHNs of higher plants. The full-length cDNA corresponding to the identified partial sequence was isolated from mRNA purified from ABA-treated P. patens using 5′ and 3′ rapid amplification of cDNA ends (RACE-PCR). Based on the sequences of the resulting PCR products, we designed specific primers to amplify the full-length cDNA and the corresponding genomic sequence. The isolated sequence, termed PpDHNA, corresponded to a 1883-nt transcript consisting of a 111-nt 5′ untranslated region (UTR), a 1666-nt open reading frame (ORF) and a 106-nt 3′ UTR. The genomic organization of PpDHNA revealed the existence of three introns as shown in Figure 2(b).
The deduced amino acid sequence of PpDHNA encoded a putative highly hydrophilic protein of 554 amino acids with a molecular mass of 59.2 kDa, and an isoelectric point (pl) of 5.6. DHNA contained 11 imperfect Y segments and one conserved K segment. The protein was rich in glycine (19%), charged amino acids (24%), and lacked the amino acids cystein and tryptophan (Figure 1a). These characteristics were consistent with those of DHNs from higher plants (Close, 1996, 1997; Garay-Arroyo et al., 2000; Svensson et al., 2002). DHNA contained 11 repetitive blocks of 35 amino acids throughout the entire protein (Figure 1b). Within these repetitive blocks, a high content of conserved glycine residues was found and we therefore designated these segments as G repeats. Each G repeat starts with a Y-like segment. In the C-terminal part of the protein, a conserved K segment was found that showed 60% identity to the conserved 15 amino acids found in all DHNs (Svensson et al., 2002). A search in protein databases with the entire DHNA polypeptide showed a 78% similarity and a 66% sequence identity with the protein Tr288 (AAG00978) isolated from the desiccation tolerant moss Tortula ruralis (Scott and Oliver, 1994). The Tr288 protein contains 15 of the 35 amino acids repeats similar to those found in DHNA. The T. ruralis protein is called a rehydrin, which originates from the fact that the cDNA was isolated from polysomes of rehydrated plants. Tr 288 is thought to play a role in cellular dehydration repair (Scott and Oliver, 1994).
PpDHNA is present as a single gene in the Physcomitrella patens genome
All higher plants analyzed to date encode more than one member of the DHN protein family. Southern blot analysis was done to see whether other genes with nucleotide sequence homology to PpDHNA were present in the P. patens genome (Figure 2a). As the entire protein contains 11 repeats of 35 amino acids, two probes were used: Rev1-4, corresponding to 10 of the repeats described in Figure 1(b); and Prom1-8, containing the sequence that corresponds to the most C-terminal part of the protein including the K repeat (Figure 2b). Membranes containing digested genomic DNA from P. patens were washed under high- and low-stringency conditions as described in Experimental procedures. Our results indicated that the PpDHNA gene is probably a unique gene in the P. patens genome, as all bands hybridizing to the two probes originated from the genomic PpDHNA sequence. A weak hybridization to a 2.2- and a 3-kb EcoR1 fragment was observed with the probe Rev1-4, clearly in low-stringency but also in high-stringency conditions. We suggest that these signals are due to incomplete digestion as all other hybridization signals corresponded to expected genomic fragments of PpDHNA. The hybridization pattern obtained when using Prom1-8 as a probe did not suggest any additional homologous sequences in the P. patens genome.
Stress-induced expression of PpDHNA
To analyze whether PpDHNA is regulated in a similar way to DHNs from higher plants, transcript accumulation of this gene was monitored after treatment of plants with increasing concentrations of ABA, exposure to low temperature (4°C), salt (0.4 m NaCl) and osmotic stress conditions (0.6 m mannitol; Figure 3a). Expression of PpDHNA was also analyzed in plants transferred to standard growth medium after stress treatment. All ABA treatments resulted in a rapid and substantial accumulation of PpDHNA mRNA. Expression was detected after 0.5 h of treatment with 1 μm ABA. Higher transcript levels were detected when 10 and 100 μm of ABA were used, whereas no significant difference in gene induction was observed in response to these two higher concentrations. Transcript accumulation for this gene was also detected already after 0.5 h of treatment of plants with 0.4 m of NaCl or 0.6 m of mannitol. In both cases, when plants were returned to standard growth medium, transcript levels rapidly decreased and could not be detected after 2 to 4 h after transfer to optimal growth conditions.
To compare the stress-induced transcript levels of PpDHNA with the accumulation of the corresponding protein, we generated polyclonal antibodies raised against a purified recombinant protein obtained by expressing the full-length cDNA in Escherichia coli. Accumulation of DHNA in response to ABA, osmotic or salt stress treatments was studied by Western blot analysis (Figure 3b). Total protein was isolated at different time points from plants treated with 10 μm ABA, 0.6 m mannitol or 0.4 m NaCl and from plants returned to non-stress conditions after 3 days of salt or mannitol treatment. In all experiments, DHNA was detectable within 24 h after treatment. No DHNA protein could be detected after 1 to 4 h in stressed-treated plants returned to growth on standard medium, indicating that DHNA is rapidly degraded under non-stress conditions. DHNA accumulation corresponded well with the induction pattern of the gene, suggesting that the regulation of this gene was at the transcriptional level. In samples where high levels of DHNA were detected (such as in ABA treatments), lower molecular mass products could also be observed. We believe that these bands corresponded to degradation products of DHNA and not to other related proteins. This is supported by the fact that no bands were observed in Western blot analysis of the dhnA knockout mutant (Figure 4b).
Generation of a PpDHNA knockout mutant
To study the physiological function of DHNA, we generated a PpDHNA null mutant by homologous recombination using targeted gene disruption. The knockout construction contained the nptIIgene driven by the CaMV 35S promoter and the 3′ UTR of the ocs gene, flanked by 1034 and 941 bp of the 5′ and 3′ ends of the gene, respectively (Figure 4a). Twenty micrograms of linear DNA were used to transform 2 × 106 of protoplasts and, after kanamycin selection, 60 transformants were analyzed for accumulation of DHNA by immunodetection analysis and subsequent PCR amplification. Out of 60 kanamycin-resistant colonies tested, one (designated dhnA) showed no DHNA accumulation after 2 days of ABA-treatment as seen in Figure 4(b). To ensure that the lack of DHNA accumulation was due to disruption of the gene by homologous recombination, PCR analysis of genomic DNA of dhnA and wild-type (WT) colonies was performed. We used a forward primer located within the 5′ region of the gene together with a reverse primer outside the gene targeting construct in the 3′ end of PpDHNA (Figure 4a). A PCR product of 2049 bp was amplified from the WT, while dhnA yield a 3445-bp fragment, consistent with the expected size originating from a homologous recombination event in the gene PpDHNA (Figure 4c). These results were confirmed by Southern blot analysis using a probe covering the entire gene construct used for transformation (Figure 4d). As compared with that from a WT strain, dhnA had a different genomic organization in the PpDHNA locus as seen by the fragment pattern for both restriction enzymes used. Digestion with HindIII resulted in two bands of 2.8 and approximately 10 kb in dhnA and only one band of approximately 11 kb in the WT strain. EcoRV digestion yields three bands of 3, 2 and 1.2 kb in the mutant and one band of approximately 5 kb in the WT. These results were consistent with the restriction pattern expected for plants in which the disrupted gene construct integrated at a single location by homologous recombination at the WT DHNA locus, generating a dhnA knockout mutant.
dhnA is impaired in osmotic stress recovery
Phenotypic characterization was done under various growth conditions where the mutant dhnA was compared with the WT strain. No negative growth effects (phenotypic or dry weight) could be observed for dhnA when grown on standard medium as compared with the WT (controls in Figures 5–7). The ability to adapt to salt or osmotic stress treatments was monitored in dhnA and WT plants exposed for 14 days to concentrations ranging from 0.2 to 0.5 m NaCl or 0.6 to 0.9 m mannitol, respectively (Figure 5). We found that growth of both WT and dhnA plants was negatively affected at all salt or mannitol concentrations used in this study. Thus, both strains showed a similar level of stress response, manifested in the impaired growth phenotypes of the plants under stress conditions.
The degree of salt and osmotic stress tolerance was assessed in dhnA and in the WT by exposing plants to 0.5 m NaCl or 0.9 m mannitol for 14 days and later transferring them back to standard medium where plants were allowed to recover from the stress treatment for a period of 3 weeks. Photographs were taken at different time points during and after the stress (Figure 6), and quantitative data concerning stress survival was obtained by the determination of chlorophyll content and by fresh and dry weight measurements (Figure 7). Fourteen days after mannitol or salt treatment, WT plants had lost up to 75 or 85% of their fresh weight, respectively (Figure 7a). Dry weight values were found to be similar in non-stressed control plants compared with those observed after mannitol treatment, whereas up to a 52% reduction could be observed in the salt-treated plants. A rather similar result was observed for dnhA plants, where fresh weight loss after mannitol or salt treatments reached 80 or 88%, respectively, and dry weight values showed a reduction of 11 or 55% in mannitol- or salt-stressed plants when compared with non-stressed controls.
Chlorophyll content per fresh weight was not affected in either the WT or dhnA during stress conditions, compared to non-stressed controls. However, when plants were transferred back to standard growth medium after mannitol or salt treatment, both WT and mutant plants were completely bleached after 2 days of growth (Figures 6 and 7c). Nevertheless, a clear difference was observed between the WT and dhnA with respect to their ability to recover from stress. For plants treated with mannitol, 3 weeks after returning to optimal growth conditions, WT plants had reached up to 78% of the fresh weight values observed in non-stressed controls (Figure 7a). Similar results were obtained with the dry weigh (Figure 7b) and chlorophyll measurements (Figure 7c). In contrast fresh and dry weight values, as well as chlorophyll levels for dhnA did not increase after 3 weeks of exposure to normal growth conditions, indicating that dhnA plants were not able to resume growth after an osmotic stress generated after 14 days of mannitol treatment. Similar results were obtained with plants that had undergone NaCl treatments, although less dramatic differences between the WT and dhnA were observed. Three weeks after being transferred back to standard growth medium, WT plants had reached up to 94% of the fresh weight, 69% of the dry weight values and 77% of the levels of chlorophyll content compared with non-stressed control plants. In comparison, average fresh and dry weight and chlorophyll values for dhnA reached only 39%, 28% and 16%, respectively. These results indicated that the mutant showed delayed NaCl stress recovery compared with the WT. The recovery of salt-treated dhnA plants was first detected 15 days after the transfer to normal conditions, while the WT starts growing already within 10 days. Both treatments were repeated 3 times with different periods of stress before the transfer back to normal growth conditions (data not shown).
Taken together, these results show that DHNA plays an essential role during salt and osmotic stress in P. patens, which is important for the recovery and survival capability of the plant when resuming growth at more favorable conditions.
In this study, we report the isolation and characterization of a DHN-like gene from the moss P. patens. The predicted protein, DHNA, was a 59.2-kDa glycine-rich protein containing conserved amino acid segments found in DHNs from higher plants, including a C-terminal K segment found in all DHNs. DHNA migrated as a protein of 65–70 kDa, which is indicating a larger protein than the theoretically calculated molecular mass. This observation is true for both the bacterial produced recombinant protein as well as the plant-isolated protein. This is consistent with the observation that many, if not all, DHNs migrate at a much slower rate than expected from their deduced molecular mass in SDS-PAGE (Close, 1996). This phenomenon has been attributed to conformational and/or charge properties of this family of proteins.
DHNA contained several Y segments forming the N-terminal part of a 35-amino-acid-long repetitive segment. We have designated this repeat the G repeat due to its high content of conserved glycine residues (7/35). Apart from the short Y segment, this long repeat does not share any similarity to other DHNs or other proteins. However, repeated amino acid segments constituting nearly the entire protein can also be found in other DHNs (Houde et al., 1992; Neven et al., 1993; Welin et al., 1994). Other characteristics found in DHNs of other plants also shared by DHNA are the lack of the amino acids cysteine and tryptophane and a high content of glycine and charged amino acids (Garay-Arroyo et al., 2000; Svensson et al., 2002,). DHNA also remains soluble after boiling treatment like other DHNs of higher plants (data not shown).
The G repeat also showed high similarity to another moss protein from T. ruralis, Tr288, which contained 15 G repeats similar to those found in DHNA. It is possible that this block of conserved amino acids might be moss-specific as no similarity was found with any other proteins in the databases. The T. ruralis protein was termed a rehydrin, although it contains the K-segment characteristic of a DHN. The name rehydrin originates from the fact that it is a protein that accumulates during rehydration and its transcript is actively associated with polysomes of rehydrated, rapidly dried gametophytes (Scott and Oliver, 1994). Wood and Oliver (1999) demonstrated that, in T. ruralis, Tr288 transcripts accumulate during desiccation, when protein synthesis is inhibited, but are stored in messenger ribonucleoprotein particles for use upon rehydration. In the first hour following rehydration, Tr288 transcripts are actively associated with the protein synthesis apparatus indicating that the synthesis of Tr288 is important in the recovery of this moss from desiccation. These findings, together with our results, indicate that these related proteins play an important role in cellular dehydration protection/repair and that this function is evolutionary conserved in plants.
Tr288 and PpDHNA showed similarities in terms of regulation of gene expression, although there are interesting differences to be considered. PpDHNA transcript and protein accumulated in response to ABA treatments, which is characteristic for many genes encoding DHNs in higher plants. The salt- and mannitol-induced gene expression further supports an ABA-mediated gene regulation, as ABA is known to increase in response to these stresses in both higher and lower plants (Addicott, 1983). In contrast, no mRNA accumulation of Tr288 was observed in T. ruralis plants treated with ABA (Velten and Oliver, 2001). Consistent with this is the observation that T. ruralis seems to lack ABA accumulation in response to water stress (Bewley et al., 1993; Velten and Oliver, 2001). Both Tr288 and PpDHNA showed a similar gene expression pattern during a slow dehydration or salt stress. However, in contrast to PpDHNA, Tr288 is induced during early rehydration upon a rapid dehydration event. This observation and the fact that T. ruralis survives a very rapid dehydration made the authors postulate that tolerance to cellular dehydration is constitutive in some desiccation tolerant mosses like T. ruralis (Oliver et al., 2000). However, this tolerance was not sufficient to prevent cell damage, which is manifested during the rehydration process, and therefore processes of repair are needed during the initial stages of regrowth of the plant. In accordance with this idea, the authors suggested that Tr288 could either play a role in the repair process or in minimizing damage (Velten and Oliver, 2001). In contrast, our results in P. patens clearly show that PpDHNA is induced during the initial stages (2 h) of osmotic stress, followed by the accumulation of the protein. Upon returning plants to non-stressed growth conditions, no protein was detectable after 2 h, suggesting that DHNA plays its role during the stress, probably protecting from cell injury allowing plant recovery. It is likely that the rapid degradation of this protein upon resuming growth after a stress treatment is selective, as it is an important mechanism to regulate protein levels and activity in higher plants (Serino and Deng, 2003; Vierstra, 2003).
Southern blot analysis suggested that PpDHNA does not share any homologous sequences in the P. patens genome. This was supported by Western blot experiments, where no additional proteins were detected with the antibodies generated against DHNA. However, when searching the large expressed sequence tag (EST) collection for this plant, we found a clone encoding two conserved K segments of DHNs (BQ039653). When comparing the corresponding sequence of this clone with PpDHNA, sequence homology was only found in the K segment (37% identity). Similar results were obtained in T. ruralis, where no homolog to Tr288 was found in Southern blot experiments, although proteins with the K repeat were found in EST collections in this plant (Dr Melvin Oliver, USDA-ARS, Lubbock, TX, USA personal communication). The information available from the EST collections and the genome sequencing project (http://moss.nibb.ac.jp), together with our results, suggest that the genome of P. patens encode relatively few DHN-like proteins compared with seed plants (Choi et al., 1999; Svensson et al., 2002). The fewer proteins found in the moss could reflect the more simple structure of this plant compared with seed plants, where evolution and specialization of different DHNs could have accompanied the development of tissues and organs. The fact that DHNs have been found in both lower and higher land plants, but not in other organisms, suggests that these proteins could constitute part of a molecular mechanisms that evolved to prevent and protect from cellular dehydration when plants inhabited land.
The generation of the knockout mutant dhnA allowed us to perform functional studies concerning the physiological role played by this protein during salt and osmotic stress in P. patens. Extensive comparisons were made between WT and mutant plants, both growing as protonema (data not shown) and as colonies. We could not detect any aberrant growth in the mutant during non-stress conditions. Moreover, no significant phenotypical difference in the stress response was observed between WT or dhnA plants when growing at the salt or mannitol concentrations tested. However, when plants exposed to stress conditions were transferred back to optimal growth medium, a very clear difference in their ability to resume growth was observed. WT plants were able to survive 14 days of exposure to 0.5 m NaCl or 0.9 m mannitol. Both chlorophyll content and fresh and dry weight values started to increase after 11 days of stress recovery and reached levels close to those observed for the non-stressed control plants 21 days after being transferred to normal growth medium. In contrast, dhnA plants were not able to recover at all from our mannitol treatment and showed a very much delayed recovery from the NaCl treatment compared with WT plants. The fact that DHNA is rapidly degraded after the stress, suggests that the protein plays a protective role during stress, but this protection becomes evident when plants resume growth at non-stress conditions. We have provided direct genetic evidence showing the important role played by a DHN during abiotic stress tolerance in any plant species, although this has been postulated for a long time (Close et al., 1989; Dure et al., 1989; Mundy and Chua, 1988).
An observation supporting a protective role during stress recovery comes from the fact that DHNs accumulate to high levels in desiccated seeds of higher plants. It is logical to think that the protective function of these proteins is accomplished both during the initiation of desiccation as well as upon imbibition of the seeds. It is therefore conceivable that DHNs may play the same protective role during both the dehydration process and the initial recovery, as one does not preclude the other. When water is removed from the cell, the physiological and physical conditions change. These changes, which normally are manifested by smaller cell size and lack of metabolic activity, are not always harmful to the cell. According to Walters et al. (2002), these observations could simply be a consequence of water removal, which would be reverted upon rehydration. It is therefore possible that damages are not seen simply as differences between a hydrated or rehydrated state of the plant but are proportionally linked to the capacity of the plant to assume regrowth. This highlights the importance of studying the responses to stress not only during stress treatments but perhaps, even more importantly, upon recovery conditions.
To discuss the possible role of DHNA and DHNs in general, it is interesting to look at the mechanisms responsible for desiccation tolerance in anhydrobiosis. Several studies, especially in seeds and pollen, show the consequences of water removal at the cellular level. When water is removed, the cell volume diminishes, increasing the probability of molecular interactions causing denaturation of proteins and irreversible damage to membranes (Hoekstra et al., 2001). Among the molecules that accumulate in dehydration tolerant cells are osmoprotective compounds, sugars and amphipathic metabolites. Sugars are the only compounds that can substitute for water in severely dehydrated cells so as to preserve the structure and function of macromolecules (Hoekstra et al., 2001). Interestingly, it has been shown that amphipathic compounds are transferred from the cytoplasm to the membrane during the process of dehydration in seeds of Pisum sativum and Cucumis sativa (Buitnik et al., 2000; Golovina et al., 1998). It is therefore possible that sugars and amphipathic metabolites together could maintain the stability of membranes during cellular dehydration in a stress tolerant tissue.
When analyzing the amino acid composition of DHNA, we found 11 segments of 35 amino acids (G repeats) repeated throughout the protein. A more detailed analysis of the G repeats indicated that a stretch of 12 amino acids (EGIVDKAKDAVG) within the repeat theoretically can form a class A amphipathic α helix. The class A amphipathic α helix forms two surfaces of different characteristics, where one side is formed by negatively charged amino acids and the opposite by hydrophobic amino acids. The positively charged residues form the interphase between the two surfaces. The same characteristic has been proposed for the K segment of DHNs of higher plants where it has been postulated that this segment interacts with the plasma membrane or the hydrophobic part of denatured proteins (Close, 1996). We propose that DHNA may interact and preserve the membrane integrity by the theoretically amphipathic structures formed by this protein. As the same type of structure can be found in the K segment, it is possible that all DHNs interact with membranes for protection and preservation during periods of cellular dehydration. Most structural studies involving DHNs show no clear secondary structure for these proteins although in vitro studies indicated that the addition of SDS increased the α helicity of these proteins (Ismail et al., 1999). More evidence supporting formation of an α-helix structure, comes from in vitro studies of an LEA3 protein found in pollen of Typha latifolia. This protein showed no secondary structure under hydrated conditions, but upon both rapid and slow dehydration, the protein assumed an increased α-helix structure (Wolkers et al., 2001). This property was due to an 11-mer amino acid segment with the theoretical capacity to form an amphipathic α helix, in a similar way to the K segment of DHNs and the G repeat of P. patens. Recently, Koag et al. (2003) showed that maize DHN1 can bind to lipid vesicles containing acidic phospholipids. The authors presented evidence that vesicle binding results in an apparent increase of an α helicity of DHN1, suggesting that DHN1 may only exhibit amphipathic helical structure when associated with lipid bilayers. It is therefore conceivable that, as proposed for DHN1, DHNA can undergo such conformational changes under stress conditions, maybe leading to stabilization of membrane structures.
This study has shown the importance played by DHNA during osmotic stress recovery in P. patens. The conservation of amino acids between DHNA and DHNs of seed plants could imply a similar role for all DHNs. These findings suggest that DHNs may constitute part of a general molecular mechanism used by all land plants to protect them from injury during cellular dehydration. The exact protective function is still unclear but we postulate a membrane preservation mode of action involving the interaction of the G and K segments of DHNA with phospholipids or/and membrane proteins during stress conditions.
Plant material, growth conditions and stress treatments
Physcomitrella patens Gransden WT isolate (Schaefer et al., 1991) was used for all experiments described in this study. Plants were grown and maintained axenically on cellophane overlaid BCDAT medium (1.6 g l−1 Hoagland's 1 mm MgSO4, 1.8 mm KH2PO4 pH 6.5, 10 mm KNO3, 45 μm FeSO4, 1 mm CaCl2, 5 mm ammonium tartrate and 10 g l−1 agar) as described by Ashton and Cove (1977). To generate protonemal cultures plant material was macerated with a sterile mortle and pestle in 2 ml of sterile double distilled water. For micropropagation, moss colonies were cut with a scalpel and plant fragments were transferred to fresh medium with cellophane. All plants were grown at 22°C under a photoperiod of 16 h light, with a photon flux of 100 μmol m−2 sec−1. Three-week-old colonies were used for all experiments. ABA treatments were done by applying ABA to the media to a final concentration of 10 μm for Western blot studies and 1, 10 or 100 μm for Northern blot studies. Low temperature treatments were done at 4/2°C (day/night) with a photoperiod of 12 h. For the salt and osmotic stress treatments, P. patens colonies grown on BCDAT medium were transferred to medium supplemented with the indicated concentrations of NaCl or mannitol. For stress recovery assays, plants exposed to 0.5 m NaCl or 0.9 m mannitol for 2 weeks were returned to standard BCDAT medium and samples were harvested at different time points for the determination of fresh and dry weight and chlorophyll content. All experiments were repeated at least 3 times.
Isolation of PpDHNA partial genomic sequence
Degenerate primers were designed using the program codehop (Rose et al., 1998). The forward primer (primer Y: 5′-ccgatgcatcatactgacganwanggnaay-3′) was designed to cover the Y segment RHTDEYGNP and the reverse primer (primer K: 5′-tatgatgtccaggcagcttctcyttdat-3′) was designed using the conserved K segment. PCR amplification was done using Taq-polymerase hot start (Promega, Madison, WI, USA) with 50 ng of genomic DNA and 10 pmol of each primer. PCR conditions were: 5 min at 94°C; 35 cycles of 30 sec at 94°C, 30 sec at 50°C, 1 min at 72°C; and 7 min at 72°C. PCR reactions were analyzed on 1% agarose gels and amplified DNA was excised and purified using the Qiagen Gel Extraction kit (Qiagen GmbH, Hilden, Germany). Purified PCR products were cloned into the vector pCR 2.1-TOPO (Invitrogen, San Diego, CA, USA). Cloned fragments were sequenced using a cycle sequencing thermosequence kit (Amersham Pharmacia Biotech, Uppsala, Sweden) and an ABI 377 sequencer (Applied Biosystems, Foster City, CA, USA).
Amplification of the full-length cDNA from PpDHNA
Total RNA was extracted using the RNeasy Mini Kit (Qiagen GmbH) and, for the isolation of poly A+ RNA, the Oligotex mRNA kit (Qiagen GmbH) was used. The full-length cDNA sequence of PpDHNA was obtained by amplification of the 3′ and 5′ ends using the Marathon cDNA amplification Kit (Clontech, Palo Alto, CA, USA). 3′ end RACE-PCR reactions were done using the adaptor primer 1 (AP1) and the gene-specific primer Prom1 (5′-ccgacgcttacgtgcatggcaaccatc-3′). Touchdown PCR was performed as follows: 30 sec at 94°C ; five cycles of 5 sec at 94°C and 4 min at 72°C; five cycles of 5 sec at 94°C and 4 min at 70°C; and 25 cycles of 5 sec at 94°C and 4 min at 68°C. PCR products were separated in agarose gels and a 750-bp fragment was excised, purified using the Qiagen Gel Extraction kit and cloned into the vector pCR 4-TOPO (Invitrogen). Four clones were chosen for sequencing. The 5′ end was amplified using primers AP1 and Rev1 (5′-atggccaatcctgtacaccgctatc-3′) and the Expand Long Template PCR system (Boehringer Mannheim, Mannheim, Germany). PCR conditions were as follows: 2 min at 94°C; five cycles of 10 sec at 94°C and 4 min at 68°C; 25 cycles of 10 sec at 94°C, 30 sec at 65°C and 5 min at 68°C. Amplified products were cloned into the vector pCR 4-TOPO and sequenced. Gene-specific primers directed to the ends of the entire cDNA sequence were designed. A 1883-bp PCR fragment, corresponding to the full-length cDNA was amplified, cloned into the pT-Adv vector (Clontech) and sequenced. The sequence for PpDHNA cDNA was deposited in the GenBank database under accession number AY365466.
Southern blot analysis
Genomic DNA was extracted essentially as described by Dellaporta et al. (1983) with an additional RNase treatment and phenol extraction using fresh plant material. The genomic organization of the locus PpDHNA was analyzed by digesting 10 μg of genomic DNA with EcoRI, EcoRV, HindIII and performing double digestions with these enzymes. Restricted DNA was separated on 1% agarose gels and transferred to nylon filters (Hybond XL, Amersham Pharmacia Biotech) according to Sambrook et al. (1989). Membranes were prehybridized at 68°C in 5 × SSPE, 5 × Denhardt's solution, 0.2% SDS and 0.5 mg ml−1 denatured salmon sperm DNA. Hybridizations were performed at 68°C overnight. The DNA fragments Prom1-8 and Rev1-4, corresponding to nt 1298 to 1883 and 350 to 1324 of the cDNA, respectively, were used as probes and were labeled with α32P-dCTP using the Rediprime II random priming labeling system (Amersham Pharmacia Biotech). Filters were washed twice for 30 min at 65°C with 2 × SSC–0.5% SDS (low stringency) or twice for 30 min at 65°C with 1 × SSC–0.5% SDS (high stringency). The corresponding autoradiograms were thereafter analyzed on a PhosphoImager (Molecular Dynamics, Sunnyvale, CA, USA) or exposed in autoradiography films.
Total RNA was isolated from control or treated plant tissue corresponding to 12 to 30 colonies, using standard procedures based on phenol/chloroform extraction followed by LiCl precipitation. Five micrograms of total RNA separated in denaturing formaldehyde agarose gels were transferred to nylon membranes (Hybond XL), according to Sambrook et al. (1989). Prehybridization and hybridization conditions were as described for Southern blot analysis. A DNA fragment corresponding to the full-length cDNA of PpDHNA was used as a probe in all Northern blot experiments. All membranes were washed twice at 65°C with 2 × SSC, 0.5% SDS and twice with 1 × SSC, 0.5% SDS, for 20 min each. Hybridization membranes were exposed in autoradiography films. Ethidium bromide staining was used to ensure equal amounts of loading of RNA in the samples.
Polyclonal antiserum was raised against the DHNA protein produced in the E. coli expression vector pJTS1 (Svensson et al., 2000). The coding sequence of PpDHNA was fused to the tac promoter resulting in the plasmid pMVC4. Bacterial cells harboring the gene construct were grown at 37°C in Luria Bertoni (LB) medium supplemented with 50 μg ml−1 of ampicillin to an OD600 of 0.5–0.7. Protein synthesis was induced by the addition of isopropyl β-D-thiogalactopyranoside (IPTG) to a final concentration of 1 mm in the growth medium and cells were maintained at 37°C for 4 h. Cells were harvested by centrifugation at 4000 g for 45 min and thereafter resuspended in 25 ml of 20 mm Na2HPO4 pH 7.2 and 150 mm NaCl. Phenylmethylsulfonyl fluoride (PMSF) was added to a final concentration of 1 mm followed by a 30-min incubation with lysozyme (0.1 mg l−1) on ice. Lysates were thereafter sonicated 6 × 15 sec and centrifuged at 9000 g to pellet cell debris. The supernatant and the solubilized cell debris fraction were analyzed by SDS-PAGE. The recombinant DHNA was found only in the supernatant fraction as visualized by Coomassie-blue stained SDS-PAGE. The protein band corresponding to the recombinant protein was excised from the gel, solubilized using two connected syringes and polyacrylamide-protein solution was subsequently injected into rabbits. Serum was collected every 2 weeks after protein injections. The corresponding serum was tested after 3–4 protein injections and found to react specifically with the purified DHNA (data not shown).
Western blot analysis
Soluble plant proteins were extracted in 50 mm Tris-HCl, 250 mm sucrose, 5 mm EDTA, 10 mmβ-mercaptoethanol, pH 7.2 and 1 mm PMSF. Protein concentrations were measured with Bradford Bio-Rad protein assay (Bio-Rad Laboratories, CA, USA). For Western blot analysis 10 μg of soluble proteins were separated in 8% polyacrylamide gels and electroblotted onto nitrocellulose membranes (HybondTM-C Pure, Amersham Pharmacia Biotech). To ensure equal loading of protein samples, blots were stained with 0.5% ponceau red. Membranes were blocked in Tris-buffered saline (TBS; 20 mm Tris-HCl, 150 mm NaCl, pH 7.4) containing 5% (weight in volume, w/v) skimmed milk powder and 0.2% Tween 20 for 1 h at room temperature. The polyclonal antisera described above were used as primary antibodies against DHNA diluted 1/1500 in TBS containing 0.1% Tween. Membranes were incubated with the primary antibodies for 1 h at room temperature. Horseradish peroxidase-labeled goat anti-rabbit antibody (Bio-Rad Laboratories) was diluted 1/7500 in TBS-Tween 1%, and membranes were incubated for 45 min at room temperature. Washes were performed in TBS-Tween 1%, with three changes of washing solution. Protein reactions were visualized using the ECL detection system (Amersham Pharmacia Biotech).
Gene disruption construct
The transformation vector used, pUBW302, contained the nptII gene driven by the constitutive CaMV 35S promoter (Odell et al., 1985) and the 3′ UTR of the ocs gene, cloned into the pBluescript SK+ vector. The gene replacement construct, pMLS2, contained a 1034-bp genomic fragment from the 5′ region of the gene PpDHNA cloned upstream from the CaMV 35S promoter and a 941-bp genomic fragment corresponding to the 3′region of the gene inserted downstream of the ocs termination signal. The 5′ sequence of the genomic DNA (corresponding to nt 67 to 1107 of the genomic sequence) was PCR-amplified using gene-specific primers containing sequences for the restriction enzymes Xho1 (forward primer 5′-AAGAATCTCGAGACT TTTCGGTGTGTCGGAGAGA-3′) and HindIII (reverse primer 5′-GACTGCAAGCT TGGCTTTGTCCACAATGCC-3′) to facilitate the subsequent cloning of the fragment. The 3′ sequence of the gene (corresponding to nt 1679 to 2620 of the genomic sequence) was obtained using the cloned fragment obtained initially by PCR amplification of genomic DNA using degenerated primers. The fragment was cut out from the vector pUBW201 using the restriction enzymes XbaI and SacI and ligated into the same restriction sites of pUBW302 downstream of the selection cassette. The resulting plasmid (pMLS2) was sequenced to confirm the correct incorporation of the fragments.
Protoplast transformation of Physcomitrella patens
The generation of moss protoplasts and the subsequent transformation were done as described by Schaefer et al. (1991). Isolated protoplasts (7 × 105) were incubated with 20 μg of linearized plasmid DNA (pMLS2 cut with KpnI). After polyethylene glycol treatment, protoplasts were incubated for 5 days at 25°C on BCDAT medium supplemented with 10 mm CaCl2 and 0.44 m mannitol. Protoplasts were thereafter transferred to BCDAT medium supplemented with 5 g l−1 of glucose, 20 μg ml−1 of kanamycin and cultured for 15 days. Protoplasts were subsequently allowed to grow for 10 days on BCDAT medium without selection and finally returned to BCDAT medium containing 20 μg ml−1 of kanamycin. Plants showing growth after 2 weeks on selection medium were analyzed for the correct incorporation of the knockout construct.
Molecular characterization of transformants
To verify the lack of DHNA in the transformants due to targeted gene disruption, WT plants and transformants were treated with 50 μm ABA for 48 h and soluble proteins were isolated as described above. Immunodetection of proteins isolated from ABA-treated WT plants and putative mutants was performed using the polyclonal antiserum directed against DHNA. To confirm that the lack of protein accumulation was due to the correct incorporation of the gene targeting construct in the PpDHNA locus, PCR amplification of genomic DNA was performed from putative mutants using the following primers and reaction conditions: forward primer cDNA-P1 5′AATCAGTACGGAAG AGAACAGCAAGA corresponding to 125 to 151 of the cDNA of PpDHNA, and reverse primer pprev7 5′ATTTGGTTTCTTGCAGAAA CACCTGGTG corresponding to 1812 to 1840. Amplification was done with the DNA polymerase ThermoZymeTM (Invitrogen) and the PCR amplification conditions were as follow: 3 min at 94°C; 35 cycles of 1 min at 94°C, 1 min at 52°C, and 1.5 min at 72°C; followed by 7 min at 72°C. Transformants where PCR amplification gave an expected fragment size of 3445 bp were subjected to further genomic analysis using Southern blot hybridization. Ten micrograms of genomic DNA were cleaved with EcoRV and HindIII, separated in agarose gels, blotted and prehybridized as described above. A DNA fragment corresponding to the entire gene disruption cassette from the knockout construct, pMLS2, was labeled with α32P-dCTP using the Rediprime II random priming labeling system (Amersham Pharmacia Biotech) and used as a probe. Hybridizations were performed at 68°C overnight, membranes were washed twice for 30 min at 65°C with 5 × SSC-0.5% SDS and twice for 30 min at 65°C with 1 × SSC-0.5% SDS and exposed in autoradiography films.
Phenotypic analysis of PpDHNA disruptants
Chlorophyll content and fresh and dry weight of plants were determined in three independent experiments using 12 colonies per genotype per treatment and per time point. Dry weight was measured after incubation of individual plant colonies on cellophane discs for 16 h at 80°C. For the determination of chlorophyll content, each plant was ground up in a mortle containing 5 ml of 80% (volume in volume, v/v) acetone and the homogenized plant material was filtered to remove cell debris. Total chlorophyll was calculated as chlorophyll a + chlorophyll b (mg g−1 fresh weight) using the following formula: Chla mg g−1 = [(12.7 × Abs663) – (2.6 × Abs645)] × ml acetone mg−1 fresh tissue; Chlb mg g−1 = [(22.9 × Abs645) – (4.68 × Abs663)] × ml acetone mg−1 fresh tissue.
This work was supported by a research grant from the Swedish Research Council of Natural Sciences, Fondo Professor Clemente Estable, project 7088, Uruguay and Comisión Sectorial de Investigación Cientifica, Universidad de la República, Uruguay. We are grateful to Dr Paul Gill and Dr Melvin Oliver for valuable comments on the manuscript. We thank Marcel Bentancor, Ana Victoria García, Alejandra Bertone, Carlos Cerveñansky and Alejandro Olmos for their technical support.
Gene accession number: the complete genome sequence of PpDHNA has been deposited at the GenBank database and the accession number is AY365466.