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First published online March 10, 2006; 10.1104/pp.106.078592 Plant Physiology 141:121-134 (2006) © 2006 American Society of Plant Biologists Rice HYDROPEROXIDE LYASES with Unique Expression Patterns Generate Distinct Aldehyde Signatures in Arabidopsis1Section of Plant Biology, University of California, Davis, California 95616
HYDROPEROXIDE LYASE (HPL) genes encode enzymes that catalyze the cleavage of fatty acid hydroperoxides into aldehydes and oxoacids. There are three HPLs in rice (Oryza sativa), designated OsHPL1 through OsHPL3. To explore the possibility of differential functional activities among these genes, we have examined their expression patterns and biochemical properties of their encoded products. Transcript analysis indicates that these genes have distinct patterns and levels of expression. OsHPL1 is ubiquitously expressed, OsHPL2 is expressed in the leaves and leaf sheaths, whereas OsHPL3 is wound inducible and expressed exclusively in leaves. OsHPLs also differ in their substrate preference as determined by in vitro enzyme assays using 9-/13-hydroperoxy linolenic and 9-/13-hydroperoxy linoleic acids as substrates. OsHPL1 and OsHPL2 metabolize 9-/13-hydroperoxides, whereas OsHPL3 metabolizes 13-hydroperoxy linolenic acid exclusively. Sequence alignments of the HPL enzymes have identified signature residues potentially responsible for the substrate specificity/preference of these enzymes. All three OsHPLs are chloroplast localized as determined by chloroplast import assays and green fluorescent protein (GFP) fusion studies. Aldehyde measurements in transgenic Arabidopsis (Arabidopsis thaliana) plants overexpressing individual OsHPL-GFP fusions indicate that all rice HPLs are functional in a heterologous system, and each of them generates a distinct signature of the metabolites. Interestingly, these aldehydes were only detectable in leaves, but not in roots, despite similar levels of OsHPL-GFP proteins in both tissues. Similarly, there were undetectable levels of aldehydes in rice roots, in spite of the presence of OsHPL1 transcripts. Together, these data suggest that additional tissue-specific mechanism(s) beyond transcript and HPL enzyme abundance, regulate the levels of HPL-derived metabolites.
Plants have evolved multiple signal transduction pathways to ensure effective responses to biotic and abiotic challenges, as well as to developmental stimuli. One such pathway is the oxylipin pathway, which upon activation by environmental and developmental inputs, induces the de novo synthesis of biologically active compounds called oxylipins (Feussner and Wasternack, 2002  ; Howe and Schilmiller, 2002). The biosynthesis of these compounds is initiated by the coordinated action of lipases and lipoxygenases, leading to the oxygenation of free polyunsaturated fatty acids (FAs), mainly linoleic and  -linolenic acids, and the generation of 9- or 13-hydroperoxy FAs (9-/13-hydroperoxyoctadecatrienoic acid [9-/13-HPOT] and 9-/13-hydroperoxyoctadecadienoic acid [9-/13-HPOD]). These hydroperoxy FAs are metabolized by a group of cytochrome P450 enzymes present in different branch pathways, to generate the oxylipins (Feussner and Wasternack, 2002). Among the oxylipin branch pathways, the allene oxide synthase (AOS) and the hydroperoxide lyase (HPL) are considered to be the two major critical plant stress response pathways. These two branches compete for the same substrates, and are responsible for the production of lipid-based compounds involved in signaling as well as antimicrobial and antifungal activities. Much of the research to date on plant oxylipins is focused on the jasmonate family of molecules generated by the AOS branch pathway (Howe and Schilmiller, 2002). By contrast, the HPL branch pathway responsible for production of aldehydes and corresponding alcohols is not fully characterized.
The first gene encoding HPL was cloned and characterized from green bell pepper (Capsicum annuum; Matsui et al., 1996
The expression of HPL genes and the production of their metabolites in plants are induced by biotic and abiotic stimuli (Bate et al., 1998
Many plant species have more than one gene encoding HPL enzymes. For example, Medicago truncatula is reported to have two HPLs, whereas alfalfa (Medicago sativa) has three (Noordermeer et al., 2000
The HPL genes appear to be regulated by organ-specific and wound-induced transcriptional activation (Bate et al., 1998
Localization studies of HPL protein by tissue print immunoblot analysis of bell pepper showed that HPL is mainly located in the outer parenchymal cells of the pericarp (Shibata et al., 1995
Subcellular localization studies determined that HPLs are membrane proteins with diverse localization patterns. Of the many plant species examined, HPLs have been localized to the microsomes (Matsui et al., 1991
To examine the differential regulation and potential functional differences between HPL gene family members within the same species, we focused on rice (Oryza sativa), the model monocot, and cloned three HPL genes from this plant. The existence of two of these genes (OsHPL1 and 2) was recently reported (Kuroda et al., 2005
OsHPLs Belong to Two Distinct Phylogenetic Clusters Database searches enabled us to identify three HPL genes in rice: OsHPL1 (AK105964), OsHPL2 (AK107161), and OsHPL3 (AY340220), none of which contains any introns. All three genes were isolated and cloned by PCR-based amplification from rice genomic DNA, and their identity was confirmed by sequence analysis. OsHPL1 (1,533 bp) is predicted to encode a polypeptide of 511 amino acids with a calculated molecular mass of 55 kD. OsHPL2 (1,503 bp) is predicted to encode a polypeptide of 501 amino acids with a calculated molecular mass of 54 kD. OsHPL3 (1,464 bp) is predicted to encode a polypeptide of 488 amino acids with a calculated molecular mass of 54 kD. Analysis of the OsHPL's deduced amino acid sequences predicted the presence of domain structures typical of cytochrome P450 enzymes, including I- and K-helices, and a heme-binding domain (Heme-BD; Fig. 1A ). Further sequence analyses show that OsHPL1 and OsHPL2 have 84% amino acid identity with each other, and 34% to 45% identity with all other HPLs reported so far. Interestingly however, these HPLs show 42% to 56% amino acid identity with all reported AOSs. On the other hand, OsHPL3 shares 34% to 40% and 34% to 72% amino acid identity with AOSs and HPLs, respectively. OsHPL3 shares 43% amino acid identity with OsHPL1 and OsHPL2.
Because of the higher percentage of amino acid sequence identity between OsHPL1 and 2, and the AOSs, phylogenetic analyses were performed on both branch pathway enzymes, using the Vector NTI 9.0 Align X program. For simplicity however, here we present the phylogenetic analyses performed on a selected number of characterized AOSs. Based on these findings, the HPL enzymes are divided into two clusters. Members within each cluster not only share sequence similarity, but also a common substrate preference (Fig. 1B). OsHPL3 belongs to cluster I where all the characterized members are classified as 13-HPLs (CYP74B; Bate et al., 1998 ; Howe et al., 2000). OsHPL1 and OsHPL2 belong to cluster II whose characterized members metabolize 9-/13-hydroperoxides (CYP74C; Matsui et al., 2000; Itoh et al., 2002).
The AOS enzymes within the CYP74A subfamily are also classified, based on their substrate preference, into 13- and 9-/13-AOSs (Laudert et al., 1996
The expression levels and patterns of OsHPLs in roots, leaf sheaths, and leaves of mechanically wounded and unwounded rice plants were analyzed by semiquantitative reverse transcription (RT)-PCR using gene-specific oligonucleotides (Fig. 2, A and B ). These data show that OsHPL1 is expressed in all tissues examined, OsHPL2 transcripts are present in the leaves and leaf sheaths but are absent in the roots, and the OsHPL3 transcripts are detected exclusively in the leaves (Fig. 2B). Furthermore, there is no significant wound induction in the transcript levels of OsHPL1 and OsHPL2, whereas wounding caused a notable increase in OsHPL3 transcript levels. Abundance of the control transcript, Actin, remains constitutive in all tissues. Reactions without the RT step failed to amplify the OsHPLs. These data clearly indicate that the amplified products are not the result of contamination caused by the genomic DNA.
The same analyses were also performed on partially submerged rice plants, simulating natural growing conditions. These data were identical to those from nonsubmerged plants (Fig. 2B; data not shown).
The HPL-derived metabolites were analyzed by gas chromatography/mass spectrometry (GC/MS) in different rice tissues obtained from the same batches of plant material previously used in the transcriptional profiling experiments. These data show that the main constituents of the HPL pathway metabolites are the C6 aldehydes (hexanal and hexenal), and that C9 aldehydes (nonenal and nonadienal) are below the levels of detection (Fig. 3, A and B ). Among these two C6 aldehydes, hexenal is the predominant metabolite with its level, under all the experimental conditions examined, being approximately 5-fold higher than the hexanal level. Furthermore, these data clearly indicate that stress induced by wounding leads to a 5- and 10-fold increase in the levels of hexenal and hexanal in either nonsubmerged or partially submerged plants, respectively, as compared to the corresponding basal levels in the leaves. Interestingly, no aldehydes are detected in the leaf sheaths or in the roots, despite the presence of OsHPL transcripts in these tissues (Figs. 2B and 3, A and B). Moreover, a lack of notable induction in the OsHPL1 and 2 transcripts, in response to stress, does not correlate with the increase in the levels of C6 aldehydes in the leaves. On the other hand, the stress-inducible increase in the abundance of OsHPL3 transcripts correlates well with the increase in the levels of these aldehydes. This suggests that stress-inducible induction of the levels of aldehydes in rice leaves is primarily due to the activity of the OsHPL3-encoded product (Fig. 3, A and B).
OsHPLs Are Chloroplast Localized Sequence analyses of OsHPLs using ChloroP 1.1 (http://www.cbs.dtu.dk) predicted the localization of OsHPL1, 2, and 3 to the chloroplasts. To experimentally validate these predictions, we examined the ability of the in vitro translated OsHPLs to bind to isolated pea (Pisum sativum) chloroplasts in the presence of ATP. In contrast to OsHPL3, the in vitro translated products of OsHPL1 and OsHPL2 failed to bind to the organelle (Fig. 4A ). Fractionation of the chloroplast import reaction showed that OsHPL3 is absent from the soluble stromal fraction, but is associated with the envelopes. The presence of a lighter band detected in the thylakoids membrane fraction is most likely due to the contamination by the bulk envelopes within which the OsHPL3 resides (Fig. 4A). Furthermore, the appearance of a lower Mr form of OsHPL3 upon import indicates the proteolytic removal of an N-terminal transit peptide.
To further examine the in vivo localizations of OsHPLs, a C-terminal green fluorescent protein (GFP) translational fusion construct for each OsHPL, expressed under the control of the cauliflower mosaic virus 35S promoter, was introduced into Arabidopsis plants. The transgenic plants containing the OsHPL-GFP fusion constructs were used to isolate protoplasts that were subsequently imaged with an epifluorescent microscope. As shown (Fig. 4B), the fluorescence signals of all OsHPLs-GFP fusions were found in chloroplasts, as identified by imaging the autofluorescence of their chlorophyll. Control panels show that GFP protein expressed from a construct lacking the HPL fusion does not localize to the same organelle. These results confirmed that all three OsHPLs are localized to the chloroplasts.
To examine substrate preference of the recombinant OsHPLs in an in vitro enzyme assay, we had to identify the cleavage site of the signature peptides of these chloroplast-localized enzymes. Since the chloroplast import assays of OsHPL1 and 2 were not successful, we could not confirm the presence or absence of a cleavable transit peptide for these enzymes. Based on previous reports, most outer envelope-localized proteins lack a cleavable signature sequence (Cline and Henry, 1996 To examine the substrate preferences of each of the recombinant OsHPLs, we performed the enzyme assays with crude lysates harboring the recombinant proteins in the presence of either 9-/13-HPOT or 9-/13-HPOD as substrates.
These data in agreement with those previously published (Kuroda et al., 2005
Recombinant OsHPLs Catalyze Cleavage of 13-HPOT into (E)-2-Hexenal
To identify the catalytic properties of OsHPL enzymes, previously identified as OsAOSs (Agrawal et al., 2004
OsHPLs Are Functional in a Heterologous System The in vivo functionality of the OsHPLs in a heterologous system was examined by generation of transgenic Arabidopsis plants overexpressing OsHPLs-GFP fusion proteins. The expression levels of OsHPL-GFP fusion in each line was determined by western-blot analysis using commercially available GFP monoclonal antibodies. Lines with similar levels of OsHPLs-GFP together with the wild type and transgenic Arabidopsis expressing GFP alone were employed in GC/MS analyses to determine the composition and the levels of the HPL-derived metabolites in leaves (Fig. 7, A and B ). These analyses indicated that the levels and profiles of the C6 aldehydes in wild-type leaves are similar to those from the GFP overexpressers (data not shown). For simplicity, we have only included the data obtained from the latter line. Transgenic Arabidopsis overexpressing the OsHPLs-GFP fusions accumulates considerably higher levels of C6 aldehydes, namely hexenal and hexanal in the leaves, as compared to those from the GFP-overexpressing lines (Fig. 7B). Specifically, the plants expressing OsHPL1-, OsHPL2-, and OsHPL3-GFP fusions generate 25-, 82-, and 4-fold more hexanal, and 18-, 60-, and 68-fold more hexenal, respectively, as compared to GFP expressing Arabidopsis. In all instances, the predominant C6 aldehyde is hexenal. The highest fold difference between hexenal and hexanal (168-fold) is observed in OsHPL3-GFP lines. Plants overexpressing OsHPL1- and OsHPL2-GFP fusions contain 7- and 8-fold more hexenal than hexanal, respectively. The profile of the OsHPL3-GFP-derived metabolites in the corresponding transgenic plants reflects the observed in vitro substrate specificity of the OsHPL3 recombinant enzyme (Figs. 5, C and D and 7B). Interestingly however, C9 aldehydes were not detected in Arabidopsis plants overexpressing either OsHPL1 or OsHPL2, even though both enzymes metabolized 9- and 13-hydroperoxides with equal efficacy in vitro. Furthermore, in spite of the presence of comparable levels of the OsHPL-GFP fusion proteins in the leaves, these lines differ in their total levels of aldehydes (Figs. 7, A and B). Plants overexpressing OsHPL2-GFP and OsHPL3-GFP have comparable levels of total aldehydes, and they both contain 3-fold higher levels than the OsHPL1-GFP line.
Leaves from the nontransgenic control were devoid of any immunoreactive bands (Fig. 7A).
The metabolic profiling of HPL-derived metabolites in different rice tissues clearly indicated that the aldehydes are below the levels of detection in roots, in spite of the presence of OsHPL1 transcripts in this tissue. Due to the lack of availability of an HPL antibody that could recognize HPLs in general and OsHPL1 in particular, we could not correlate the presence of the transcript to the translated product. The availability of the transgenic Arabidopsis line producing OsHPL1-GFP fusion, however, provided us with the enabling tool to examine the presence of aldehydes in the roots in correlation to the enzyme levels. The selection of OsHPL1-GFP transgenic plants for these studies was based on the expression pattern analysis, which established the presence of the OsHPL1 transcripts in rice roots (Fig. 2B). The expression of OsHPL1 in the transgenic roots was confirmed by image analysis using an epifluorescent microscope (Fig. 8A ). The levels of OsHPL1 expressed in roots versus that present in the leaves of the same transgenic plant were determined by western-blot analysis using the GFP monoclonal antibodies. Those data show that OsHPL1 protein is expressed at similar levels in the root and leaves of the transgenic plant (Fig. 8B). Interestingly however, similar to the rice tissue, the aldehydes were below the levels of detection in roots of the transgenic Arabidopsis. In contrast, the leaves collected from the same plant, contain considerable levels of hexenal and hexanal as indicated on the GC/MS chromatograph (Fig. 8C).
Roots and leaves from the nontransgenic control were devoid of any immunoreactive bands (Fig. 8B).
We have cloned three HPL genes, OsHPL1 through OsHPL3, from the model monocot rice. These genes are differentially regulated as determined by their respective spatial expression patterns as well as the basal- and wound-inducible steady-state transcript levels. The OsHPL1 transcript is present in leaves, roots, and leaf sheaths, whereas that of OsHPL2 is not detectable in the roots. OsHPL3 transcript is present exclusively in the leaves. Moreover, wound-inducible induction of the mRNA abundance is most notable for OsHPL3. These data prompted us to compare promoter sequences of the OsHPLs for the presence of regulatory elements that may unravel the molecular basis of the differential spatial expression patterns and response to wounding. The OsHPL1 promoter contains two identical, unique cis-elements (ACTTTA) present at 49 and 222 bp upstream of the transcriptional start site. This motif is reported to be critical for the expression of rolB oncogene in leaves, stems, and roots of tobacco (Nicotiana tabacum; Baumann et al., 1999 ). Thus, the presence of this motif may explain the molecular basis for the ubiquitous presence of OsHPL1 transcripts in all tissues examined. The unique cis-element in OsHPL2 is the (CAANNNNATC) motif present 400 bp upstream of the transcriptional start site. This regulatory element is conserved among the clock-regulated, light-harvesting complex protein genes that are not expressed in roots or Bright-Yellow 2 nongreen culture cells (Piechulla et al., 1998). Therefore, it is tempting to speculate that, similar to the OsAOS1 gene (Haga and Iino, 2004), OsHPL2 is regulated by light and thus is absent in nonphotosynthetic organs. The OsHPL3 promoter contains a unique cis-element (AACGTGT) present 95 bp upstream of its transcriptional start site. This element is known as ExtA cis-element present in the promoter region of a wound-inducible extensin gene in Brassica napus (Elliott and Shirsat, 1998). The cis-element shared by all OsHPLs is the W box (TTGAC), a motif necessary for the binding of WRKY DNA-binding proteins responsible for the activation of pathogen-induced genes (Yu et al., 2001).
Detailed amino acid sequence comparison in conjunction with the examination of the substrate specificity/preference of the enzymes led to the grouping of OsHPLs into two phylogenetic clusters. OsHPL1 and OsHPL2, though highly similar to each other, are more similar to AOSs than to all the other HPLs. In fact, due to these higher levels of amino acid sequence identity with AOSs, OsHPL1 and 2 were previously clustered within the AOS family of enzymes and were designated as OsAOS3 and 4 (Agrawal et al., 2004
Based on the detailed sequence analysis, OsHPL1 and 2 form their own clade nested between HPLs and AOSs. Subsequent enzyme activity assays showed that OsHPL1 and 2, similar to closely related enzymes in the AOS and HPL group cluster, metabolize 9-/13-HPOT/D and thus belong to the CYP74C subfamily enzymes. We therefore placed OsHPL1 and 2 into cluster II, where the enzymes, in addition to their respective amino acid identity, also share similar activity profiles on 9-/13-hydroperoxides. OsHPL3 has higher percent amino acid identity with other HPLs than with AOSs. This enzyme belongs to cluster I, where in addition to sequence identity, the enzymes exclusively metabolize 13-hydroperoxides. Thus, in contrast to OsHPL1 and 2, OsHPL3 belongs to CYP74B subfamily enzymes. In fact, OsHPL3 is the only enzyme known to be active exclusively on 13-HPOT, as all the other characterized 13-HPLs metabolize both 13-HPOT and 13-HPOD (Bate et al., 1998 Detailed examination of multiple alignments of the catalytic regions of these enzymes led to the identification of two potential signature residues within the consensus sequences that may play a critical structural or functional role in determining the substrate preference of the enzymes. One potential signature is within the (L/I)(F/C)G(Y/F)(Q/R)(P/K) consensus sequence, where the C residue in 13-hydroperoxide-preferring enzymes is replaced by the more bulky and hydrophobic F residue present in all 9-/13-hydroperoxide-preferring enzymes. The second potential signature is within the Heme-BD [(PS(E/P)G(N/D)K(Q/I)C(A/P)(G/A)K(D/N)], where the nonreactive A residue present in all the characterized 13-hydroperoxide-preferring enzymes is replaced by the P in the 9-/13-hydroperoxides enzymes. The potential function of these signature residues in determining the substrate profile of these cytochrome P450 enzymes is yet to be examined.
The OsHPLs are similar in their subcellular localizations. A combination of import assays and GFP-fusion studies confirmed that OsHPL3 is similar to LeHPL, a member of CYP74B subfamily that is localized to the chloroplasts (Froehlich et al., 2001 In parallel with the transcriptional profiling, we also examined the levels of HPL-derived metabolites in rice. These data clearly show the lack of detectable levels of aldehydes in rice leaf sheaths and roots, in spite of the presence of OsHPL1 transcripts at levels similar to those in leaves, and the presence of OsHPL2 mRNA in the leaf sheaths (Figs. 2B and 3). This suggests additional tissue-specific mechanism(s) regulating the levels of these metabolites at posttranscriptional levels and/or substrate availability. Furthermore, unlike OsHPL1 and OsHPL2, the enhancement of OsHPL3 transcript levels in response to wounding correlates well with the induction of these aldehydes in leaves, thus suggesting that the OsHPL3-encoded enzyme is primarily responsible for the production of HPL-derived metabolites in response to stress (Figs. 2B and 3).
The expression of OsHPLs-GFP fusion constructs in Arabidopsis further provided us with the necessary tools to examine the differential functionality of these enzymes in vivo by measuring the profile and levels of aldehydes produced in plants that express similar levels of OsHPL enzymes as determined by western-blot analysis using anti-GFP monoclonal antibodies (Fig. 7A). The profiles of the HPL-derived metabolites in the OsHPLs-GFP-overexpressing lines are distinct from each other. Specifically, plants overexpressing OsHPL3-GFP accumulate significantly higher levels of hexenal and lower levels of hexanal as compared to those overexpressing OsHPL1- and OsHPL2-GFP (Fig. 7B). Plants with OsHPL1- and OsHPL2-GFP transgenes generated high levels of hexenal as well as hexanal, which reflect their ability to metabolize both 13-HPOT and 13-HPOD substrates. Although the recombinant OsHPL1 and OsHPL2 enzymes metabolize 13-hydroperoxides equally well, their expression in the transgenic line resulted in the production of higher levels of hexenal, the products of 13-HPOT, than hexanal, the products of 13-HPOD. This suggests that the availability of substrate regulated by upstream components of the HPL pathway determines the in vivo profiles of the aldehydes. Similarly, the absence of detectable levels of nonenal and nonadienal, derived from 9-hydroperoxides, in these plants also reflects the unavailability of the 9-HPOT/D substrates. In accordance with these data, it is reported that overexpression of Cs9HPL in tomato (Lycopersicon esculentum) did not result in detectable levels of nonenal or nonadienal (Matsui et al., 2001 The composition and the levels of the HPL-derived metabolites were examined in plants expressing similar levels of enzymes (Fig. 7, A and B). In spite of similar protein levels, Arabidopsis plants overexpressing OsHPL1-GFP contained only one third of the aldehyde levels as compared to each of those overexpressing OsHPL2- and OsHPL3-GFP. This suggests that in a heterologous system, OsHPLs have different activities, with OsHPL1 being the least active enzyme.
As stated above, we have established that rice roots do not contain aldehydes in spite of the presence of OsHPL1 transcripts at levels similar to those in the leaves. To begin to dissect the tissue-specific mechanism(s) involved, we exploited the transgenic Arabidopsis lines, overexpressing OsHPL1-GFP under the control of 35S promoter. Our analysis showed that although the roots have similar levels of fusion proteins as compared to the leaves, aldehydes are below the levels of detection in this tissue (Fig. 8, B and C). This is in contrast to previous reports showing the presence of aldehydes in the root tissues of red clover (Trifolium pratense) and melon (Kamm and Buttery, 1984
Plant Growth and Wound Treatments Arabidopsis (Arabidopsis thaliana) plants were grown in a 16-h-light/8-h-dark cycle at 22°C. Rice (Oryza sativa) L. cv Nipponbare plants were grown at 30°C in a greenhouse, under either nonsubmerged or partially submerged conditions. Rice seeds were germinated in soil and grown for 3 weeks before the seedlings were either submerged in water up to the first leaf, or they were kept in dry conditions. Tissues were collected from 4-week-old rice and Arabidopsis plants for the assays described in this report. Mechanical wounding of leaves and leaf sheaths was performed by puncturing the tissue many times using an 18-gauge needle. Tissues were harvested after 2 h of wounding, immediately frozen in liquid nitrogen, and stored at –80°C until use.
Since none of the OsHPL genes contains introns, genomic DNA isolated from rice L. cv Nipponbare was used for PCR-based amplification of these genes using the following gene-specific oligonucleotides: OsHPL1 (Forward: 5'-ATAGATATCGCATGCATGGCGCCGCCGCGAGCCAACTCCG-3' and Reverse: 5'-ATATACGTACTGCAGCGCGCGCCGCCGCTTGACACTATTA-3'), OsHPL2 (Forward: 5'-ATAGATATCGCATGCATGGCGCCACCGCCAGTGAACTCCG-3' and Reverse: 5'ATATACGTACTGCAGGCACGTGACGTCGACGTGCGTGCTA-3'), and OsHPL3 (Forward: 5'-ATAGATATCGCATGCATGGTGCCGTCGTTCCCGCAGCCGG-3' and Reverse: 5'-ATATACGTACTGCAGGAGAGAATGGCGGCAGCAAAGCTTA-3'). For each amplification, 30 PCR cycles were carried out using a Gene Amp PCR system 9700 (Applied Biosystems) in a 25 µL reaction mix containing 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 4% dimethyl sulfoxide (DMSO), 100 µM of each dNTP, 500 nM of each forward and reverse primer, 0.625 units of Taq DNA polymerase (Invitrogen), and 50 ng of the genomic DNA. Amplification was conducted at 94°C for 1 min, 94°C for 30 s, 55°C for OsHPL1, 63°C for OsHPL2, and 55°C for OsHPL3 for 1 min, 72°C for 90 s, and a 10-min extension step at 72°C. The amplified products were resolved by electrophoresis on a 1% (w/v) agarose gel. The band corresponding to each full-length gene was cut, purified using QIAquick Gel extraction kit (Qiagen), and cloned in pCR 2.1-TOPO Vector (Invitrogen) according to the manufacturer's instructions. The identities of these clones were confirmed by DNA sequencing. All DNA as well as polypeptide sequence analyses were performed using Vector NTI advance program 9 (Invitrogen).
Plasmids containing full-length DNA sequence of the OsHPLs were linearized by restriction endonuclease digestion at sites 3' of the coding regions. Transcriptions from the linearized plasmids were performed using either T7 or SP6 (Promega) RNA polymerase. Precursor proteins were synthesized using [35S]Met (Perkin Elmer) in a rabbit reticulocyte lysate (Cline et al., 1985 For fractionation, chloroplasts were utilized in import assays described above, but in a 300 µL reaction and incubated in light for 40 min. After import, reactions were treated with 200 µL of 0.5 M Na2CO3 on ice and in the dark for 15 min. Chloroplasts were then reisolated in 1 mL of 30% Percoll (GE Healthcare Bio-Sciences) by centrifuging at 1,900g, and then washed with 1 mL of import buffer. Reactions were resuspended in 80 µL of Tris-EDTA + Suc (10 mM Tricine, pH 7.5, 2 mM EDTA, and 0.6 M Suc). A 20 µL sample was taken as the whole chloroplast fraction (import). The remaining 60 µL were incubated on ice, freeze thawed in a –20°C freezer, and then centrifuged at 200g for 5 min. The pellet was resuspended in 40 µL 400 mM Suc, 40 mM Na-MES pH 6.5, 10 mM NaCl, and 5 mM MgCl2 from which 20 µL were taken as the thylakoid fraction. The supernatant was spun in an ultracentrifuge at 100,000g for 30 min, and the pellet was resuspended in 40 µL of sample buffer as the envelope fraction. The resulting supernatant was TCA precipitated and resuspended in 40 µL of sample buffer plus 0.5 M Tris, pH 8.0, as the stroma fraction. All samples were run on 15% acrylamide tris-Gly gels, fixed by a 15 min glacial acetic acid wash, and dried on filter paper. Gels were exposed on phosphor screens and visualized on a Storm Scanner (GE Healthcare Bio-Sciences).
The vector PQE-30 (Qiagen) was used for cloning OsHPL1 and 2, while OsHPL3-FL and OsHPL3 were cloned in pDEST17. All constructs contain His6-tag fusions at their N termini. For the in-frame fusion of the full-length OsHPL1 and OsHPL2 into the expression vector, their respective original constructs generated in pCR 2.1-TOPO vector were digested with SphI and PstI and the resulting fragments were individually subcloned into the SphI/PstI sites of the PQE-30 vector. These plasmids were sequenced and their authenticity was confirmed, after which they were transformed into M15 (pREP4) strain Escherichia coli. OsHPL3-FL and OsHPL3 were amplified by PCR with oligonucleotides designed for Gateway cloning. Primers used for amplification of OsHPL3-FL were: Forward: 5'-CACCATGGTGCCGTCGTTCC-3' and Reverse: 5'-GCTGGGAGTGAGCTCC-3'. OsHPL3 was amplified using the following primers: Forward: 5'-CACCCGGCCAATACCGGGG-3' and Reverse: 5'-GCTGGGAGTGAGCTCC-3'. PCR amplification was conducted as described above with Tm = 52°C, used for OsHPL3-FL and OsHPL3. Amplified products were cloned into the pENTR/D-TOPO vector, and subcloned into the Gateway destination vector pDEST17 by a LR reaction (Invitrogen). These plasmids were sequenced and transformed into BL21 E. coli cells.
Bacteria transformed with each of the above described plasmids or with the vector alone as the control were grown to an OD600 of 0.7 and then induced with 0.5 mM isopropyl
GFP fusions for stable expression were constructed by cloning the PCR-amplified, TOPO-cloned, and EcoRI-/BamHI-digested fragments of the full length of all three genes into the EcoRI/BamHI site of pEZS-NLGFP. Primers were designed to eliminate stop codons and fuse the coding sequences to the 5' end of the GFP gene. For OsHPL1, the primers used were: Forward: 5'-ATA-GAATTCATGGCGCCGCCGCGAG-3' and Reverse: 5'-ATAGGATCCGCTA-CTCCGCGCCGCGCG-3'. For OsHPL2, the primers used were: Forward: ATAGAATTCATGGCGCCACCGCCAGT-3' and Reverse: 5'-ATAGGATCC-GCTCCCGACGACGCCCGT-3'. OsHPL3 was amplified using the following primers: Forward: 5'-ATAGAATTCATGGTGCCGTCGTTCCC-3' and Reverse: 5'-ATAGGATCCGCGCTGGGAGTGAGCTCCC-3'. PCR amplifications were conducted as described above with a Tm = 55°C used for all genes amplified. GFP fusions for Arabidopsis transformation were created by subcloning the OsHPL1, OsHPL2, and OsHPL3 open reading frames from pEZS-NLGFP into the binary vector pMLBart, kindly provided by John Bowman (University of California, Davis) using NotI restriction sites with the GFP gene at the C terminus of each gene. The constructs were verified by sequencing, introduced into Agrobacterium EHA101 strain, and used to transform Arabidopsis plants by using the floral-dip method (Clough and Bent, 1998
Protoplast isolation was performed as previously described (Sheen, 2002 Protoplasts were visualized with a Nikon Eclipse 400 epifluorescence microscope (Nikon). GFP was visualized with a 450 to 490 nm excitation filter, a 495 nm dichroic mirror, and a 500 to 550 nm emission filter. Chlorophyll autofluorescence was visualized with a 528 to 553 nm excitation filter, a 565 nm dichroic mirror, and a 600 to 660 nm emission filter.
The hydroperoxide-metabolizing activity of the recombinant HPLs was measured spectrophotometrically by monitoring the decrease in A234 resulting from the disruption of the conjugated diene bond of the substrate (Zimmerman and Vick, 1970
Total RNA was isolated from leaves, leaf sheaths, and roots of wounded and unwounded rice plants grown in nonsubmerged or partially submerged conditions. RNA extraction was carried out by mixing 150 mg of ground and frozen tissue in 1 mL of TRIzol reagent (Life Technologies). Upon addition of chloroform (250 µL) the extract was vortexed briefly and incubated at room temperature for approximately 15 min. The RNA was subsequently precipitated in the presence of 200 mM NaCl, 133 mM Na-citrate, and 17% (v/v) isopropanol, and pelleted by centrifugation (14,000g, 30 min) at 4°C. RNA pellets were washed with ice-cold 70% (v/v) ethanol, air dried, and resuspended in diethyl pyrocarbonate-treated water. Transcriptional profiling of HPLs was based on the semiquantitative RT-PCR assays, conducted using 100 ng of DNaseI-treated RNA. A 516-bp amplicon was obtained using the following OsHPL1-specific primers: Forward: 5'-GGCGAGATCACCATCTCC-3' and Reverse: 5'-TTATACTCCGCGCCGCG-3'. A 666-bp amplicon was obtained using the following OsHPL2-specific primers: Forward: 5'-ATAGAATTCATGGCGCCACCGCCAGT-3' and Reverse: 5'-CGCGAGCCACGCCATG-3'. A 976-bp amplicon was obtained using the following OsHPL3-specific primers: Forward: 5'-CCTCAACAGGGCCCCTG-3' and Reverse: 5'-TTAGCTGGGAGTGAGCTCC-3'. A 1,134-bp amplicon was obtained using the following OsActin-specific primers: Forward: 5'-ATGGCTGACGCCGAGGATATC-3' and Reverse: 5'-TTAGAAGCATTTCCTGTGCACAAT-3'. RT-PCR reactions were conducted in a 20 µL reaction mixture containing 50 mM Tris-HCl (pH 8.3), 75 mM KCl, 3 mM MgCl2, 100 mM dithiothreitol, 500 µM of each dNTP, 1 µM of each reverse and forward primer, 200 units of Superscript III RT (Invitrogen), 40 units of RNase Out (Invitrogen), 5% DMSO, and 50 ng RNA for the amplification of the desired cDNA. The RT reactions were carried out at 50°C for 45 min followed by heating the sample to 70°C for 15 min to denature the reverse transcriptase. To amplify the DNA fragments, 2.5 µL of the RT reaction was used in a 25 µL reaction mix containing 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 4% DMSO, 100 mM of each dNTP, 500 nM of each of the forward and reverse primer, and 0.625 units of Taq DNA polymerase (Invitrogen). A total of 27 amplification cycles were performed at 95°C for 1 min, 94°C for 15 s, 50°C for 30 s, 72°C for 45 s, and 10 min of extension step at 72°C. The amplified products were resolved by electrophoresis on 1.0% (w/v) agarose and visualized by ethidium bromide staining. These experiments were repeated three times, resulting in an almost identical data set for each independent study.
One hundred milligrams of ground and frozen Arabidopsis tissue overexpressing the OsHPL-GFP fusions were boiled for 5 min in 200 µL of 2x SDS sample buffer. The mixture was centrifuged (14,000g, 5 min) and the protein content was determined by DC protein assay (Bio-Rad) according to the manufacturer's instructions. Equal amounts of proteins (20 µg) were loaded onto 7% SDS-PAGE. The separated proteins were transferred onto a Poly (vinylidene fluoride) PVDF membrane and probed using anti-GFP monoclonal antibodies (CLONTECH). The blots were finally developed using 0.33 mg/mL nitro blue tetrazolium chloride and 0.165 mg/mL 5-bromo-4-chloro-3-indoyl phosphate in 10 mL of Development buffer (100 mM Tris-HCl [pH 9.5], 100 mM NaCl, and 5 mM MgCl2).
To measure the aldehyde levels in the plants, the tissue was collected and frozen in liquid nitrogen. The material was ground rapidly and thoroughly with a pestle, and a weighed amount of the sample was introduced into a 4-mL screw-top Supelco vial containing 500 µL of 1% NaCl. The vial was then rapidly capped with the screw top having a polytetrafluoroethylene/silicone septum, and incubated for 30 min in a water bath at 50°C under continuous stirring. A 60-µm polydimethylsiloxane (PDMS)-coated solid phase micro extraction (SPME; Supelco) was used to measure the aldehydes released from the plant tissue. Measurements were done in triplicates. The headspace was sampled for 30 min with the PDMS-SPME and analyzed by GC-MS. GC-MS analysis was performed using a Hewlett and Packard 6890 series gas chromatograph coupled to an Agilent Technologies 5973 network mass selective detector. An HP-5MS column (30 m x 0.25 mm, 0.25 µm film thickness) was used with He (37 kPa) as carrier gas. The GC oven temperature was programmed as follows: 5 min 40°C, ramp to 225°C at 15°C/min and no hold time. Mass spectra in the electron impact mode were generated at 70 eV. Injection was performed by thermal desorption of the SPME in the injector at 200°C using the splitless injection mode. The compounds were identified by comparing the GC retention times and mass spectra with those of authentic reference compounds. The headspace was analyzed as described above and peak areas (mass-to-charge ratios 82 and 98) were determined. The aldehydes were quantified subsequent to careful preparation of calibration curves with n-hexanal and (E)-2-hexenal as standards. It is important to note that based on our analysis using the PDMS fiber, n-hexanal and (E)-2-hexenal have similar distribution constants, indicating that the fiber adsorbs these compounds equally well within a 10% marginal error.
We performed HPL-specific assays by analysis of the products derived from the recombinant enzymes. These assays were carried out in GC vials at room temperature with E. coli lysate (2 mg for OsHPL1 and OsHPL2, and 1 mg for OsHPL3) adjusted to a final volume of 1 mL of 0.1 M Tris-HCl (pH 7.0). Upon addition of 13-HPOT (0.1 mM final concentration) to the sample, the vial was rapidly capped with the screw top having a polytetrafluoroethylene/silicone septum, and incubated for 10 min at 40°C. A 60-µm PDMS microextraction fiber probe was inserted into the headspace of the vial and subsequently the probe was injected into the GC/MS and analysis was performed as described above.
We thank Raymond Sheehy for growing and maintaining the rice plants in the New Technologies Research Growers greenhouses. Received February 2, 2006; returned for revision March 1, 2006; accepted March 2, 2006.
1 This work was supported by Biostar and National Science Foundation grants awarded to K.D. 
2 Present address: Pacific Northwest National Laboratories, P.O. Box 999, Richland, WA 99352. 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: K. Dehesh (kdehesh{at}ucdavis.edu). Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.106.078592. * Corresponding author; e-mail kdehesh{at}ucdavis.edu; fax 530–752–5410.
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ABSTRACT
RESULTS
-D-thiogalactopyranoside for 3 h. Cells were sedimented by centrifugation (14,000g, 10 min), resuspended in a buffer containing 50 mM sodium phosphate (pH 7.0) and mini protease inhibitor mix (Roche Molecular Biochemicals), and sonicated three times on ice for 15 s and 9 W. Debris was pelleted by centrifugation (17,000g, 20 min) and the supernatant was used for the enzyme assays.