Phylogenetic analysis, structural evolution and functional divergence of the 12-oxo-phytodienoate acid reductase gene family in plants
© Li et al; licensee BioMed Central Ltd. 2009
Received: 02 December 2008
Accepted: 05 May 2009
Published: 05 May 2009
The 12-oxo-phytodienoic acid reductases (OPRs) are enzymes that catalyze the reduction of double-bonds in α, β-unsaturated aldehydes or ketones and are part of the octadecanoid pathway that converts linolenic acid to jasmonic acid. In plants, OPRs belong to the old yellow enzyme family and form multigene families. Although discoveries about this family in Arabidopsis and other species have been reported in some studies, the evolution and function of multiple OPRs in plants are not clearly understood.
A comparative genomic analysis was performed to investigate the phylogenetic relationship, structural evolution and functional divergence among OPR paralogues in plants. In total, 74 OPR genes were identified from 11 species representing the 6 major green plant lineages: green algae, mosses, lycophytes, gymnosperms, monocots and dicots. Phylogenetic analysis showed that seven well-conserved subfamilies exist in plants. All OPR genes from green algae were clustered into a single subfamily, while those from land plants fell into six other subfamilies, suggesting that the events leading to the expansion of the OPR family occurred in land plants. Further analysis revealed that lineage-specific expansion, especially by tandem duplication, contributed to the current OPR subfamilies in land plants after divergence from aquatic plants. Interestingly, exon/intron structure analysis showed that the gene structures of OPR paralogues exhibits diversity in intron number and length, while the intron positions and phase were highly conserved across different lineage species. These observations together with the phylogenetic tree revealed that successive single intron loss, as well as indels within introns, occurred during the process of structural evolution of OPR paralogues. Functional divergence analysis revealed that altered functional constraints have occurred at specific amino acid positions after diversification of the paralogues. Most notably, significant functional divergence was also found in all pairs, except for the II/IV, II/V and V/VI pairs. Strikingly, analysis of the site-specific profiles established by posterior probability revealed that the positive-selection sites and/or critical amino acid residues for functional divergence are mainly distributed in α-helices and substrate binding loop (SBL), indicating the functional importance of these regions for this protein family.
This study highlights the molecular evolution of the OPR gene family in all plant lineages and indicates critical amino acid residues likely relevant for the distinct functional properties of the paralogues. Further experimental verification of these findings may provide valuable information on the OPRs' biochemical and physiological functions.
Plant responses to many biotic and abiotic stresses are orchestrated locally and systemically by signaling molecules known as jasmonates (JAs), which are derived from linolenic acid via the octadecanoid pathway [1, 2]. Jasmonic acid (JA) and other octadecanoids act as plant growth regulators in various developmental processes such as fruit ripening, pollen maturation, root growth and tendril coiling [3–5]. They are also potent modulators of defenses against insects and pathogens [2, 5–7]. Thus, the lipid-based octadecanoid pathway leading to JA has also been found to be an integral part of the signal transduction pathway. The 12-oxo-phytodienoic acid reductases (OPRs) are enzymes that catalyze the reduction of double-bonds adjacent to an oxo group in α, β-unsaturated aldehydes or ketones and are part of the octadecanoid pathway that converts linolenic acid to jasmonic acid [8, 9].
In plants, the OPR genes, which belong to the old yellow enzyme (OYE) family, are flavin mononucleotide (FMN)-dependent oxidoreductases and form multigene families. The first member of the OPR family in higher plants was identified from Arabidopsis thaliana, and named AtOPR1 . Subsequently, other OPR genes were identified in the tomato [11, 12], pea , rice [14–16] and maize  genomes. Earlier studies on the enzymatic activity of OPRs in Arabidopsis and tomato revealed that these enzymes have distinct substrate preferences and therefore have been classified into two groups, group I and II, depending on their substrate specificity [10, 12, 18, 19]. OPR group I enzymes preferentially catalyze the reduction of (9R,13R)-12-oxo-10, 15(Z)-octadecatrienoic acid (9R,13R-OPDA), while OPR group II enzymes preferentially catalyze (9S,13S)-12-oxo-10, 15(Z)-octadecatrienoic acid (9S,13S-OPDA), a natural biosynthetic intermediate precursor in JA biosynthesis. AtOPR3 and LeOPR3, belonging to group II, have been shown to efficiently reduce the natural isomer 9S,13S-OPDA to 3-oxo-2(2'(Z)-pentenyl)-cyclopentane-1-octanoic acid (OPC 8:0), the precursor of JA [12, 18, 19]. In contrast, AtOPR1/2 and LeOPR1/2, belonging to group I, were unable to catalyze this step [10, 18, 20].
The biological significance of plants having multiple OPRs is not clearly understood. To date, studies of the physiological role of OPRs have focused mainly on their expression in dicots and monocots. Among dicots, OPRs have been characterized in Arabidopsis and tomato. The expression levels of OPR mRNA and protein have also been analyzed in transgenic plants. OPRs in Arabidopsis have been shown to have tissue-specific expression patterns. AtOPR3 is transcribed more actively in flowers or anthers than in the roots and leaves . Conversely, AtOPR1/2 is transcribed more actively in roots and leaves . Furthermore, the transcription of OPRs can also be induced by wounds, pathogens, signaling molecules such as JA, methyl jasmonate (MeJA), salicylic acid (SA), abscisic acid (ABA) and ethylene, and other environmental stimuli [10, 20–22]. For example, AtOPR1/2 transcription is up-regulated transiently in response to wounding ; likewise, the expression of β-glucuronidase (GUS) under the control of the AtOPR1 and AtOPR2 promoters was up-regulated after stimulation by touch, wounding, and ultraviolet (UV) irradiation . In addition, mutants for AtOPR3, which encodes the enzymes belonging to group II, were shown to be deficient in the biosynthesis of JA and the males were sterile [21, 22]. Moreover, the AtOPR3 mutants accumulated 12-oxo-10, 15(Z)-octadecatrienoic acid (OPDA) following wounding and were resistant to fungal and insect attacks .
In monocots, more than 13 OPR genes have been found in the rice genome; OsOPR1 was the first OPR gene characterized at the biochemical and molecular level . This gene is rapidly and transiently up-regulated in response to a variety of environmental cues including JA, SA, ethylene and H2O2 . Similar results were found for the expression of OsOPR7 . Moreover, over-expression OsOPR7, clustered in the same group II, was able to compensate for the phenotype of AtOPR3 mutants, whereas OsOPR1, which is clustered in other groups, was unable to compensate for the same phenotype . In addition, an analysis of mRNA transcripts indicated that maize OPR genes exhibit organ-specific expression and can be rapidly and transiently up-regulated in response to a variety of biotic and abiotic stresses that include wounding, signaling molecules (JA, MeJA, SA, ABA and ethylene) and the presence of pathogens . In spite of ongoing studies, the function of OPRs in plants remains obscure.
Although biochemical and genetic studies in Arabidopsis and other species have led to important discoveries in understanding the function of OPRs, proven biological roles have been elucidated for only a few members of this family and in a limited number of species. Additionally, differences in family size among eukaryotes raise several questions regarding the evolution and functional divergence of the OPR gene family. Thus, a comprehensive comparative genome study is essential for understanding the evolution and function of the OPR gene family in plants. Here, we performed a comparative genomic analysis using a comprehensive bioinformatics/phylogenetic approach to elucidate the evolutionary history, structural evolution and putative functional divergence of the OPR gene family in plants. Firstly, we identified all OPR paralogues from eleven species (Chlamydomonas reinhardtii, Volvox carteri, Physcomitrella patens, Selaginella moellendorffii, Picea sitchensis, Oryza sativa, Sorghum bicolor, Zea mays, Arabidopsis thaliana, Populus trichocarpa and Medicago truncatula), representing the six major plant lineages with available genome sequences. Secondly, phylogenetic analysis was performed to trace back the evolutionary history of the OPR family in plants. Thirdly, exon/intron structure analysis was performed to gain insight into the possible mechanisms for structural evolution of the OPR gene family, because the exon/intron structural divergence within gene families is also a mechanism for the evolution of multiple gene families. Finally, functional divergence analysis suggests that changes in selective constraints and/or amino acid properties occurred after gene duplication, which led to subfamily-specific functional evolution after their diversification. This has also led us to predict the positive-selection sites or critical amino acid sites that may be of importance for the functional divergence of the OPR paralogues.
Identification of OPR genes and their homologues in plants
Number of OPR or OPR-like genes in 11 representative plants
Phylogenetic relationships and evolution of the OPR gene family in all plant lineages
Interestingly, phylogenetic analysis showed distinct differences between aquatic and land plants, not only in the number of OPR genes, but also in the number of subfamilies (Figure 1A). Therefore, based on the results obtained from phylogenetic analysis, we proposed a schematic pattern to account for the expansion and evolution of the OPR gene family in plants (Figure 2C). In this pattern, the OPR genes were plant-specific and originated before the divergence of green algae from land plants. The ancestral OPR gene evolved into the present sub. VII and the common ancestral OPR of land plants evolved after the divergence of aquatic plants (green algae) from land plants. Subsequently, the common ancestral OPR of land plants underwent one duplication and yielded two copies: one copy evolved into the present sub. II, while the other copy evolved into the present sub. VI in lower land plants (mosses and lycophytes) and sub. I in higher land plants (gymnosperms and angiosperms) (Figure 2C). The events leading to lineage-specific expansion, especially by tandem duplication, occurred in monocots after their divergence from dicots, and sub. III, IV and V were generated exclusively in monocots (Figure 2C).
Structural evolution of the OPR family genes
In addition, a total of 12 different introns have been found within all the genes of the OPR family across different lineage species, according to intron position (Figure 3). Further analysis the introns of OPR paralogues in sub. VII indicated that I5 and I7 exist only in VcOPR1 and CrOPR2/3, but not in CrOPR1 and VcOPR2 (Figures 1B, 3). However, CrOPR1 and VcOPR2 arose earlier than VcOPR1 and CrOPR2/3, according to the topology of the ML tree (Figure 1A). These observations suggest that I5 and I7 were most likely gained by the ancestor of VcOPR1 and CrOPR2/3 before the divergence between Chlamydomonas reinhardtii and Volvox carteri. The above-mentioned events of intron loss and I5 and I7 gain in OPR paralogues are consistent with previous findings by Lin et al. (2006)  and Roy and Penny (2007) [30, 31].
Additionally, the exon-intron structure of OPR genes showed that the length of introns within each individual OPR gene was distinct, with lengths varying from 47 to 2919 bp (data not shown). Further analysis of the introns of OPRs in Arabidopsis and rice indicated that the average intron length of Arabidopsis OPR genes was 165 bp, close to that of the entire Arabidopsis genome (168 bp) calculated from Arabidopsis genome TAIR 6.0 release. The average intron length of rice OPR genes was 559 bp, longer than that of the entire rice genome (393 bp) calculated from the TIGR rice genome release 5.0. Moreover, most of these introns are putative miniature inverted-repeat transposable elements (MITEs) or retrotransposons, which can be found in the TIGR Oryza Repeat Database. For example, short fragment insertions in the first intron (213 bp) of OsOPR01-1 and the second intron (600 bp) of OsOPR06-1 show high homology (> 90%) with the MITEs, while a long fragment insertion in the intron (~5.0 kb) of OsOPR06-4 shows high homology (> 90%) with the retrotransposons (Figure 2B). Similar cases were also found in maize and Sorghum OPR genes. These results suggest that the presence of indels within introns, caused by MITEs or retrotransposons, may have arisen during the structural evolution of the OPR gene family.
Variable selective pressures among amino acid sites under diversifying selection
Likelihood values and parameter estimates for the OPR genes in plants
Estimates of parametersc
Positively Selected Sitesd
ω = 1.047
p0 = 0.415, (p1 = 0.585) ω0 = 0.154, ω1 = 1
p0 = 0.271, p1 = 0.447, (p2 = 0.283) ω0 = 0.113, ω1 = 1, ω2 = 3.183
20, 22, 24, 35*, 39, 40*, 48, 51, 56*, 62*, 69*, 70*, 76, 83*, 102, 131, 132, 135, 151, 152, 157*, 158*, 163*, 170, 177, 205*, 207*, 227*, 228*, 233, 243*, 246, 248*, 249, 254*, 258*, 262, 267, 268, 301, 302, 303, 304, 305, 307*, 311, 314*, 319, 324*, 327*, 330*, 333, 334, 336, 345, 354, 357, 358, 359, 361
p0 = 0.274, p1 = 0.449, (p2 = 0.227) ω = 0.119, ω1 = 1.049, ω2 = 3.290
16, 20, 22, 24, 35*, 39, 40*, 41*, 48, 51, 56*, 62*, 66, 69*, 70*, 76, 83*, 98, 102, 131, 132, 135, 151, 152, 157*, 158*, 159, 163*,170, 177, 205*, 207*, 223, 224, 227*, 228*, 233, 243*, 245, 246, 248*, 249, 254*, 258*, 262, 267, 268, 301, 302, 303, 304, 305, 307*, 311, 314*, 319, 324*, 325, 327*, 330*, 333, 334, 336, 345, 354, 357, 358, 359, 361
p = 0.384, q = 0.264
p0 = 0.733, (p1 = 0.267) p = 0.378, q = 0.264, ω = 2.853
20, 22, 24, 35*, 39, 40*, 48, 51, 56*, 62*, 69*,70*, 76, 83*, 102, 131, 132, 135, 151, 152, 157*, 158*, 163*, 170, 177, 205*, 207*, 227*, 228*, 243*, 246, 248*, 249, 254*, 262, 267, 268, 301, 302, 303, 304, 305, 307*, 311, 314*, 319, 324*, 327*, 330*, 333, 334, 336, 345, 354, 357, 358, 359, 361
All three models that allow for the presence of positive-selection sites, i.e., M2a (positive selection), M3 (discrete) and M8 (beta &ω), do suggest the presence of such sites (Table 2). Allowing for the presence of positive-selection sites (with ω > 1) significantly improves the fit of the models. The comparison of models M1a and M2a should be stated as a test of the null hypothesis that all genes evolved under neutral conditions, versus the alternate hypothesis that some sites are under negative selection (ω < 1), some sites under neutral constraints (ω = 1) and some sites under positive selection (ω > 1). The neutral model (M1a) does not allow for sites with ω > 1, while the positive selection model (M2a) adds an additional site class, with the ω ratio estimated to be 3.183. The log-likelihood improvement was huge, as 2Δℓ = 807.50 should be compared with = 9.21 with df = 2 (Additional file 6). Comparison between M7 (beta) and M8 (beta and ω) produced similar results (Additional file 6). This could be explained by the fact that the majority of the protein was subjected to constant negative selection while a few sites underwent positive selection .
Analysis of functional divergence
Analysis of functional divergence between OPR subfamilies in plants
θ1 ± S.Ea
Critical amino acid sitesd
Sub. I vs II
0.220 ± 0.056
35*, 65, 70*, 110, 258*, 306, 324*
Sub. I vs III
0.280 ± 0.055
41*, 54, 69*, 80, 104, 243*, 258*, 312, 324*, 330*, 360
Sub. I vs IV
0.176 ± 0.070
Sub. I vs V
0.351 ± 0.083
62*, 98, 166, 212, 227*, 254*, 264
Sub. I vs VI
0.229 ± 0.063
47, 110, 157*, 212, 307*
Sub. I vs VII
0.377 ± 0.077
38, 43, 56*, 62*, 68, 85, 163*, 178, 207*, 227*, 245, 257,258*, 327*, 331, 332
Sub. II vs III
0.340 ± 0.064
41*, 65, 69*, 70*, 80, 85, 167, 169, 194, 201, 243*, 306, 308, 309
Sub. II vs IV
0.001 ± 0.022
Sub. II vs V
0.142 ± 0.133
Sub. II vs VI
0.269 ± 0.064
38, 65, 157*, 191, 223, 245, 248*, 307*, 308, 324*
Sub. II vs VII
0.114 ± 0.083
Sub. III vs IV
0.350 ± 0.086
21, 40*, 83*, 187, 254*, 307*, 308, 309
Sub. III vs V
0.437 ± 0.084
16 *62, 66, 70*, 82, 104, 158*, 176, 243*, 254*, 255, 258*, 264, 273, 275, 318, 330*
Sub. III vs VI
0.353 ± 0.068
21, 70*, 87, 158*, 243*, 248*, 255, 259, 309, 312, 314*, 318, 324*
Sub. III vs VII
0.378 ± 0.081
56*, 62*, 68, 69*, 85, 189, 205*, 228*, 233, 245, 324*, 330*
Sub. IV vs V
0.288 ± 0.155
Sub. IV vs VI
0.351 ± 0.083
47, 157*, 189, 223, 248*, 254*, 265, 307*, 308
Sub. IV vs VII
0.259 ± 0.153
Sub. V vs VI
0.159 ± 0.105
Sub. V vs VII
0.316 ± 0.162
Sub. VI vs VII
0.178 ± 0.072
Moreover, some critical amino acid residues responsible for the functional divergence were predicted based on site-specific profiles in combination with suitable cut-off values derived from the posterior probability of each comparison. In order to reduce false positives, Qk > 0.7 (Qk, posterior probability) was used as the cutoff to identify type-I functional divergence-related residues in all comparisons between the seven OPR subfamilies; the results are shown in Table 3. These results show distinct differences in the number and distribution of predicted sites for functional divergence within each pair (Table 3; Additional file 7). For example, 5 critical amino acid sites were predicted for the subfamilyI/VI pair and distributed in SSSU (Super Secondary Structure Unit) 1, 3, 4 and 6, while 16 critical amino acids sites were predicted for the subfamilyI/VII pair and distributed in SSSU 1, 2, 3, 4, 5 and 7 (Additional file 7). Further analysis revealed that 2 out of 5 sites in pair I/VI and 9 out of 15 sites in pair I/VII were distributed in α helices and only one site (site 62) was located in β2 in the subfamily I/VII pair (Additional file 7). Similar cases were found in other subgroup pairs (Additional file 7). In addition, 24 out of 69 positively selected sites detected under model M3 (discrete) implemented in the codeml program of the PAML v4.0 package were also found to be functionally divergent between OPR paralogues (marked by asterisks in Tables 2 and 3). The shifted evolutionary rate at specific amino acid sites throughout the Oxidored_FMN domain within each pair facilitated the functional divergence of OPR subfamilies during the long period of evolution.
Origin and evolution of the OPR gene family
The OPR gene family is present in all plant species and generally has multiple genes in each species. In this study, we comprehensively analyzed the phylogeny and evolution of the OPR gene family in all plant lineages, and the results showed that seven well-conserved OPR subfamilies exist in plants (Figure 1A). All five OPR genes identified from green algae fell into a separate clade (sub. VII) while all OPR genes from land plants were clustered into sub. I–VI (Figure 1A), suggesting that the OPR gene might have originated before the divergence of green algae and the ancestor of land plants. Moreover, the intron positions and the phases of adjacent exons in the Oxidored_FMN domain were conserved in the OPR genes in land plants (Figures 1B, 3; Additional file 3), suggesting that all land plant OPR genes might have originated from a common ancestor. Additionally, phylogenetic analysis suggested that lineage-specific expansion events occurred after the divergence between lower and higher land plants, leading to the generation of sub. VI in lower land plants and sub. I in higher land plants (Figure 2C). Similarly, lineage-specific expansion events also occurred in higher land plants (monocots), and sub. III, IV and V were generated after the divergence from dicots (Figure 2C).
Gene duplication, including tandem duplication, segmental duplication and genome duplication, continues to be a pervasive process and contributes to biological novelty in evolution [39, 40]. In this paper, the clustering distribution of OPR genes (Figure 2A) revealed that tandem duplication had an additional role in determining the current size of the OPR gene family. Moreover, the search for paralogues indicated that the SbOPR04-1/SbOPR06-3 pair was generated by segmental duplication (Figure 2B); the age estimation of OsOPR genes (data not shown) indicates that the divergence time within OPR gene pairs was between 20.5 and 36.9 million years ago (Mya), falling into the period of large-scale duplication events 30–40 Mya [41–43]. These observations suggest that large-scale duplication may also have been involved in the expansion of the OPR gene family in Sorghum and rice.
The above analysis reveals that the OPR gene family originated from a common ancestor of green plants, followed by lineage-specific expansion and divergence in each lineage and species during their evolution. Moreover, lineage-specific expansion, especially by tandem duplication, is likely to have contributed to the size of the OPR family and yielded multiple OPR subfamilies in land plants, especially in higher land plants such as monocots. Additionally, large-scale or segmental duplication may have been involved in the expansion of the OPR gene family in Sorghum and rice.
Successive intron loss for structural evolution
Gene duplication is a common phenomenon in plant genomes and continues to be a pervasive force in genome evolution . To date, several models for the evolution of genomes have been proposed based on comparative genome studies of model organisms [45–47], but little attention has been focused on the structural evolution of duplicated gene families. In fact, the structural diversity of gene family members is also a mechanism for the evolution of multiple gene families, and intron loss or gain can be an important step in generating structural diversity and complexity . In this study, we analyzed the structural diversity of OPR genes (Figure 1B) and found that single intron loss events occurred during the expansion and structural evolution of OPR paralogues. We found that most OPR family genes lost two or more introns, and the number and position of intron loss was distinctly different among OPR genes (Figure 4). Furthermore, the intron loss events occurred not only in different plant lineages from algae to angiosperms (Figure 4B), but also in each individual plant lineage, from the ancestral OPR of each individual plant lineage to the present individual OPR genes (Figure 4C). These results, in combination with the phylogenetic trees of the OPR gene family (Figure 1A), suggest that intron losses occurred successively rather than simultaneously. In addition to intron loss, intron gain may have occurred during the structural evolution of OPR paralogues in green algae (Figure 4).
Intron positions have been shown to be remarkably well-conserved over long evolutionary time intervals [49, 50], and mounting evidence suggests that lineage-specific intron loss may occur during the evolution of a gene family . In this paper, we observed that the intron positions and intron phases of the OPR family genes were well-conserved (Figures 1B, 3), and some introns (I1, I2, I4, I6) were conserved in all plant lineages (Figure 4B). This suggests that lineage-specific intron loss events might have occurred during the expansion and structural evolution of OPR genes and generated diversity of gene structure.
The most commonly used model for intron loss is mRNA-mediated intron loss [52, 53], but there are also other possibilities such as simple genomic deletion  and in-frame intron deletion . Recent studies [56–58] have indicated that introns closer to the 3' end of genes are preferentially lost, leaving the flanking exons to fuse and form a large exon at the 3' end. In this study, the results of exon-intron structure analysis (Figures 1B, 3) revealed that the introns of OPR genes in aquatic plants (green algae) were distributed relatively symmetrically among the coding sequences, while the introns of OPR genes in land plants were distributed asymmetrically, clustered at the 5' ends with the largest exon at the 3' end (Figures 1B, 3A, 4). Moreover, introns I8 to I12 (except for I9), located on the 3' end, were lost in land plants after the divergence from aquatic plants (Figure 4B). These findings suggest that multiple unique introns, i.e., I8 -I12, were lost during the evolution from aquatic plants to land plants and that mRNA-mediated intron loss was responsible for their deletion. In contrast, the presence of indels within introns would predict that other mechanisms for intron loss (e.g. simple genomic deletion and in-frame intron deletion) were involved.
Functional divergence in the OPR gene family
Functional innovations including pseudogene formation [59, 60], subfunctionalization , neofunctionalization  and subneofunctionalization  after gene duplication may result in altered functional constraints between the gene clusters of a gene family. In this study, the differences between exon/intron structures and the divergences in amino acid sequences among different subfamilies provided us with some hints that the OPR paralogues may have a variety of physiological functions. The results of the functional divergence analysis (Table 3) suggested that OPR genes should be significantly functionally divergent from each other, owing to the evolutionary rate differences at some amino acid sites. A reasonable explanation for these differences would be that due to amino acid mutations, the OPR family genes evolved some new subgroup-specific functions after divergence. Hence, functional divergence might reflect the existence of long-term selective pressure. Previous studies regarding the enzymatic activity of OPR in Arabidopsis and tomato showed that OPR enzymes could be classified into two groups (OPR I and II) based on their substrate specificity, which was determined by the substrate binding loop (SBL) [18, 19, 64, 65]. Site-specific profile analysis of OPR members showed that 24 out of the 69 positive-selection sites found under model M3 (discrete) were functionally divergent and only 2 out of 24 critical amino acid sites (E157 and A158) were located in the SBL (Figure 5; Tables 2, 3 and Additional file 7). Moreover, the majority of positive-selection sites or critical amino acid sites were distributed in α-helices, especially in α5, α6 and α7 (Figure 5; Additional file 7). These observations suggest that positive-selection pressure on the SBL, as well as α-helices, accelerated the functional divergence and formed multiple subfamilies in plants. Additionally, few positive-selection sites were distributed in β-strands (β1-β8), suggesting that the function of the 8 β-strands, clustered inside of the OPR protein, might be to define the conserved fold common to all OPRs and maintain the proteins' structural and/or conformational stability.
Studies on AtOPRs showed the AtOPR01-4 and AtOPR01-5, which belong to subfamily I, preferentially catalyze 9R,13R-OPDA and are predominantly expressed in roots , whereas AtOPR02-1, which belongs to subfamily II, catalyzes the reduction of 9S,13S-OPDA to form OPC 8:0 and is expressed in flowers and anthers [21, 22]. Studies of Atopr3 have shown that opr3 plants are deficient in the biosynthesis of jasmonic acid and male-sterile, whereas opr1 and opr2 plants are normal [19, 21, 22]. These studies, together with the results of the phylogenetic analysis of OPR paralogues in plants, suggest that OsOPR08-1, ZmOPR01-1 and ZmOPR04-1, which belong to subfamily II, are probable candidates for involvement in the JA biosynthesis pathway, while OsOPR04-1 and ZmOPR02-1, which belong to subfamily I, are likely part of a defense signaling pathway.
Additionally, the expression of OsOPR06-6 (OsOPR1), a member of subfamily III, in the leaves of two week-old seedlings is induced not only by hormones (JA, MeJA, SA) and environmental stress factors such as drought, salt, chilling, UV and O3, but also by protein phosphatase inhibitors such as cantharidin (CN), endothall (EN) and okadaic acid (OA) , while its expression in suspension-cultured rice cells is induced by JA and protein synthesis inhibitor cycloheximide (CHX) . The results of our phylogenetic analysis of the OPR gene family, together with the previous studies, suggest that subfamily III, existing exclusively in monocots, may have an important role in defense signaling pathways and the mitogen-activated protein kinase (MAPK) pathway. The other two subfamilies (IV and V) may represent pseudogene, subfunctionalization, or neofunctionalization families of genes. Further experiments need to be performed to elucidate the function of these genes in monocots.
This study provides a comparative genomic analysis addressing the phylogenetic relationships and evolution of the OPR gene family in eleven species representing six major lineages within the green plants. The results of the phylogenetic analysis revealed that seven well-conserved subfamilies exist in plants and that all OPR paralogues originated from a common ancestor of green plants. Lineage-specific expansion, primarily through tandem duplication, contributed to the size of the OPR gene family, and multiple subfamilies formed in land plants after divergence from aquatic plants. The exon/intron structure analysis showed that the gene structures were diverse, while the intron positions and intron phases were highly conserved across different lineages. These observations together with the results obtained from the phylogenetic analysis indicate that successive single intron losses, as well as indels within introns, were involved in the structural evolution of OPR paralogues. Finally, the functional divergence analysis between OPR paralogues suggested that significantly site-specific altered selective constraints acted on most OPR paralogues after gene duplication, leading to subgroup-specific functional evolution after their phylogenetic diversification. This study also demonstrates that amino acids critical for functional divergence are located in the regions including the substrate binding loop (SBL), as well as in α-helices (especially helices α5, α6 and α7), indicating the importance of these regions in OPR proteins. These data may provide valuable information for future studies of the function of this gene family, especially subfamilies III, IV and V in monocots.
Identification of OPR genes and their homologues in plants
To identify OPR genes and their homologues in plants, the BLASTP and TBLASTN programs were used to search the TIGR (The Institute for Genomic Research, http://www.tigr.org/), TAIR (The Arabidopsis Information Resource, http://www.arabidopsis.org/), MaizeGDB (Maize Genetics and Genomics Database, http://www.maizegdb.org/), PlantGDB (Plant Genome Database, http://www.plantgdb.org/), JGI (Joint Genome Institute, http://genome.jgi-psf.org/) and NCBI (The National Center for Biotechnology Information, http://blast.ncbi.nlm.nih.gov/Blast.cgi) non-redundant databases for protein sequences of the three previously reported OPRs in Arabidopsis [10, 20]. The Blast searches were performed with the following criteria: E value < 1 × e-05 and only OPR or OPR-like genes from plants were included. Moreover, proteins identified by the BLAST search algorithms were considered as potential homologues when amino acid identity was above 25% over a stretch of 200 amino acids. Then, the Pfam http://pfam.sanger.ac.uk/search and SMART http://smart.embl-heidelberg.de/ databases were employed to detect conserved domains with OPR or OPR-like protein candidates. Finally, based on the Pfam and SMART analysis, we refined the search results manually to further reduce hits with partially conserved functional domains and other false positives.
Sequence alignment and phylogenetic analysis
Multiple-sequence alignment is the first step in phylogenetic analysis and the alignment quality may have an enormous impact on the final phylogenetic tree [68–70]. Amino acid sequences of OPR genes and their homologues in plants were aligned using the EBI web tool Clustal W v2.0 program http://www.ebi.ac.uk/Tools/clustalw2/ with the default parameters. The GBlocks 0.91b program [70, 72] was then used to select the conserved blocks of the above alignment with the default parameters underlined. Thus, the poorly aligned positions, gap positions and divergent regions from the alignment were completely excluded from the phylogenetic analyses. The Akaike Information criterion (AIC) was implemented in ProtTest v1.4  to estimate the most appropriate model of amino acid substitution for tree-building analyses. ProtTest v1.4 is based on the PhyML program  for maximum likelihood (ML) optimizations, and the best-fit model considers the relative rates of amino acid replacement and the evolutionary constraints imposed by conservation of protein structure and function. Then, according to the best-fit model predicted by ProtTest v1.4, a rooted maximum likelihood tree was constructed from the Gblocks alignment using the PhyML v3.0 online program , and the reliability of interior branches was assessed with 1000 bootstrap resamplings. Considering the limitations of PhyML in tree-space searches, the Phylip v3.68 package  was used to reconstruct the ML tree under the Jones-Taylor-Thornton (JTT) model . Finally, the phylogenetic trees were displayed using MEGA v3.1 . In addition, another three phylogenetic trees were reconstructed with MEGA v3.1 from the Gblocks alignment, by employing the neighbor joining (NJ), minimal evolution (ME) and maximum parsimony (MP) methods, respectively.
Estimating the pattern of nucleotide substitution and positive-selection sites
The diversity of OPR genes was examined with molecular evolutionary analyses using ω, which is the ratio of nonsynonymous substitutions (dN) to synonymous substitutions (dS), and a simple and useful measurement of protein evolution . Considering that positive selection may act in very short episodes or on only a few sites during the evolution of duplicated genes, we calculated the ω ratio for various amino acid sites and detected the positive selection sites (ω > 1). First, accurate nucleotide sequences and related multiple protein sequence alignments were obtained with PAL2NAL , a program that constructs multiple codon alignments from matching protein sequences. Then, the resulting codon alignments and NJ tree were used in the program codeml from the PAML v4.0 package  to calculate the dN/dS (or ω) ratio for each site and to test different evolutionary models. The improved versions of site-specific models, recommended by Anisimova et. al.  and Wong et. al.,  were tested: Models M0 (one ratio), M1a (nearly neutral), M2a (positive selection), M3 (discrete), M7 (beta) and M8 (beta+ ω) were all used in this analysis. Model M0 assumed a constant ω ratio, while in models M1a and M2a ω is estimated from the date (0 <ω0 < 1) while ω1 = 1 is fixed. M7 and M8 assume a β-distribution for ω between 0 and 1. Models M2a, M3, and M8 allow for the occurrence of positively selected sites (ω > 1). Subsequent likelihood rate comparisons of M0 with M3, M1a with M2a, and M7 with M8, respectively, were performed to test which model fits the data better. The difference in log likelihood between the models, multiplied by two, was compared with a chi-square distribution with n degrees of freedom, n being the difference between the numbers of parameters of the two models. A significantly higher likelihood of the alterative model compared to the null model suggests positive selection. Finally, the Naive Empirical Bayes (NEB) and/or Bayes Empirical Bayes (BEB) approach were used to calculate the posterior probability that each site belongs to the site class of positive selection under each model.
Functional divergence and altered functional constraint analysis
To estimate the level of functional divergence and predict important amino acid residues for these functional differences among OPR subfamilies, the coefficients of type-I and type-II functional divergence (θI and θII) between any two clusters were calculated for each position in the alignment (Additional file 3), using the method suggested by Gu et. al. (1999, 2006) [38, 77], as implemented in the DIVERGE v2.0 package . This method is based on maximum likelihood procedures to estimate significant changes in the site-specific shift of evolutionary rate or site-specific shift of amino acid properties after the emergence of two paralogous sequences. The advantage of this method is that it uses amino acid sequences and, thereby, is not sensitive to saturation of synonymous sites. Type I designates amino acid configurations that are very conserved in gene 1 but highly variable in gene 2, or vice versa, implying that these residues have experienced altered functional constraints (i.e., different evolutionary rates) [77, 78]. Type II designates amino acid configurations that are very conserved in both genes but whose biochemical properties are very different (e.g., positive versus negative charge), implying that these residues may be responsible for functional specification [78, 79]. θI or θII values that are significantly greater than 0, suggest site-specific altered selective constraints or a radical shift of amino acid physiochemical properties after gene duplication. Moreover, a site-specific posterior analysis was used to predict amino acid residues that were crucial for functional divergence.
This research was supported by grants from the National Natural Science Foundation of China (No. 30571069 and No. 30800600), the Ph.D. Programs Foundation of Ministry of Education of China (No. 20060558093) and the Natural Science Foundation of Guangdong Province (No. 8151027501000016), P. R. China. We thank five anonymous reviewers for helpful comments.
- Blechert S, Brodschelm W, Holder S, Kammerer L, Kutchan TM, Mueller MJ, Xia ZQ, Zenk MH: The octadecanoic pathway: signal molecules for the regulation of secondary pathways. Proc Natl Acad Sci USA. 1995, 92 (10): 4099-4105.PubMed CentralView ArticlePubMedGoogle Scholar
- Weiler EW, Laudert D, Stelmach BA, Hennig P, Biesgen C, Kubigsteltig I: Octadecanoid and hexadecanoid signalling in plant defence. Novartis Found Symp. 1999, 223: 191-204.PubMedGoogle Scholar
- Creelman RA, Mullet JE: Jasmonic acid distribution and action in plants: regulation during development and response to biotic and abiotic stress. Proc Natl Acad Sci USA. 1995, 92 (10): 4114-4119.PubMed CentralView ArticlePubMedGoogle Scholar
- Liechti R, Farmer EE: The jasmonate pathway. Science. 2002, 296 (5573): 1649-1650.View ArticlePubMedGoogle Scholar
- Turner JG, Ellis C, Devoto A: The jasmonate signal pathway. Plant Cell. 2002, 14 (Suppl): S153-164.PubMed CentralPubMedGoogle Scholar
- Zavala JA, Baldwin IT: Jasmonic acid signalling and herbivore resistance traits constrain regrowth after herbivore attack in Nicotiana attenuata. Plant Cell Environ. 2006, 29 (9): 1751-1760.View ArticlePubMedGoogle Scholar
- Kniskern JM, Traw MB, Bergelson J: Salicylic Acid and Jasmonic Acid Signaling Defense Pathways Reduce Natural Bacterial Diversity on Arabidopsis thaliana. Mol Plant Microbe Interact. 2007, 20 (12): 1512-1522.View ArticlePubMedGoogle Scholar
- Schaller F: Enzymes of the biosynthesis of octadecanoid-derived signalling molecules. J Exp Bot. 2001, 52 (354): 11-23.View ArticlePubMedGoogle Scholar
- Liechti R, Farmer EE: Jasmonate biochemical pathway. Sci STKE. 2006, 2006 (322): cm3-PubMedGoogle Scholar
- Schaller F, Weiler EW: Molecular cloning and characterization of 12-oxophytodienoate reductase, an enzyme of the octadecanoid signaling pathway from Arabidopsis thaliana. Structural and functional relationship to yeast old yellow enzyme. J Biol Chem. 1997, 272 (44): 28066-28072.View ArticlePubMedGoogle Scholar
- Strassner J, Schaller F, Frick UB, Howe GA, Weiler EW, Amrhein N, Macheroux P, Schaller A: Characterization and cDNA-microarray expression analysis of 12-oxophytodienoate reductases reveals differential roles for octadecanoid biosynthesis in the local versus the systemic wound response. Plant J. 2002, 32 (4): 585-601.View ArticlePubMedGoogle Scholar
- Strassner J, Furholz A, Macheroux P, Amrhein N, Schaller A: A homolog of old yellow enzyme in tomato. Spectral properties and substrate specificity of the recombinant protein. J Biol Chem. 1999, 274 (49): 35067-35073.View ArticlePubMedGoogle Scholar
- Matsui H, Nakamura G, Ishiga Y, Toshima H, Inagaki Y, Toyoda K, Shiraishi T, Ichinose Y: Structure and expression of 12-oxophytodienoate reductase (subgroup I) genes in pea, and characterization of the oxidoreductase activities of their recombinant products. Mol Genet Genomics. 2004, 271 (1): 1-10.View ArticlePubMedGoogle Scholar
- Agrawal GK, Jwa NS, Shibato J, Han O, Iwahashi H, Rakwal R: Diverse environmental cues transiently regulate OsOPR1 of the "octadecanoid pathway" revealing its importance in rice defense/stress and development. Biochem Biophys Res Commun. 2003, 310 (4): 1073-1082.View ArticlePubMedGoogle Scholar
- Sobajima H, Takeda M, Sugimori M, Kobashi N, Kiribuchi K, Cho EM, Akimoto C, Yamaguchi T, Minami E, Shibuya N, et al: Cloning and characterization of a jasmonic acid-responsive gene encoding 12-oxophytodienoic acid reductase in suspension-cultured rice cells. Planta. 2003, 216 (4): 692-698.PubMedGoogle Scholar
- Agrawal GK, Tamogami S, Han O, Iwahashi H, Rakwal R: Rice octadecanoid pathway. Biochem Biophys Res Commun. 2004, 317 (1): 1-15.View ArticlePubMedGoogle Scholar
- Zhang J, Simmons C, Yalpani N, Crane V, Wilkinson H, Kolomiets M: Genomic analysis of the 12-oxo-phytodienoic acid reductase gene family of Zea mays. Plant Mol Biol. 2005, 59 (2): 323-343.View ArticlePubMedGoogle Scholar
- Schaller F, Hennig P, Weiler EW: 12-Oxophytodienoate-10,11-reductase: occurrence of two isoenzymes of different specificity against stereoisomers of 12-oxophytodienoic acid. Plant Physiol. 1998, 118 (4): 1345-1351.PubMed CentralView ArticlePubMedGoogle Scholar
- Schaller F, Biesgen C, Mussig C, Altmann T, Weiler EW: 12-Oxophytodienoate reductase 3 (OPR3) is the isoenzyme involved in jasmonate biosynthesis. Planta. 2000, 210 (6): 979-984.View ArticlePubMedGoogle Scholar
- Biesgen C, Weiler EW: Structure and regulation of OPR1 and OPR2, two closely related genes encoding 12-oxophytodienoic acid-10,11-reductases from Arabidopsis thaliana. Planta. 1999, 208 (2): 155-165.View ArticlePubMedGoogle Scholar
- Stintzi A, Browse J: The Arabidopsis male-sterile mutant, opr3, lacks the 12-oxophytodienoic acid reductase required for jasmonate synthesis. Proc Natl Acad Sci USA. 2000, 97 (19): 10625-10630.PubMed CentralView ArticlePubMedGoogle Scholar
- Sanders PM, Lee PY, Biesgen C, Boone JD, Beals TP, Weiler EW, Goldberg RB: The arabidopsis DELAYED DEHISCENCE1 gene encodes an enzyme in the jasmonic acid synthesis pathway. Plant Cell. 2000, 12 (7): 1041-1061.PubMed CentralView ArticlePubMedGoogle Scholar
- Tani T, Sobajima H, Okada K, Chujo T, Arimura S, Tsutsumi N, Nishimura M, Seto H, Nojiri H, Yamane H: Identification of the OsOPR7 gene encoding 12-oxophytodienoate reductase involved in the biosynthesis of jasmonic acid in rice. Planta. 2008, 227 (3): 517-526.View ArticlePubMedGoogle Scholar
- Guindon S, Lethiec F, Duroux P, Gascuel O: PHYML Online – a web server for fast maximum likelihood-based phylogenetic inference. Nucleic Acids Res. 2005, W557-559. 33 Web Server
- Abascal F, Zardoya R, Posada D: ProtTest: selection of best-fit models of protein evolution. Bioinformatics. 2005, 21 (9): 2104-2105.View ArticlePubMedGoogle Scholar
- Retief JD: Phylogenetic analysis using PHYLIP. Methods Mol Biol. 2000, 132: 243-258.PubMedGoogle Scholar
- Jones DT, Taylor WR, Thornton JM: The rapid generation of mutation data matrices from protein sequences. Comput Appl Biosci. 1992, 8 (3): 275-282.PubMedGoogle Scholar
- Guo AY, Zhu QH, Chen X, Luo JC: [GSDS: a gene structure display server]. Yi Chuan. 2007, 29 (8): 1023-1026.View ArticlePubMedGoogle Scholar
- Lin H, Zhu W, Silva JC, Gu X, Buell CR: Intron gain and loss in segmentally duplicated genes in rice. Genome Biol. 2006, 7 (5): R41-PubMed CentralView ArticlePubMedGoogle Scholar
- Roy SW, Penny D: On the incidence of intron loss and gain in paralogous gene families. Mol Biol Evol. 2007, 24 (8): 1579-1581.View ArticlePubMedGoogle Scholar
- Roy SW, Penny D: Patterns of intron loss and gain in plants: intron loss-dominated evolution and genome-wide comparison of O. sativa and A. thaliana. Mol Biol Evol. 2007, 24 (1): 171-181.View ArticlePubMedGoogle Scholar
- Nielsen R, Yang Z: Likelihood models for detecting positively selected amino acid sites and applications to the HIV-1 envelope gene. Genetics. 1998, 148 (3): 929-936.PubMed CentralPubMedGoogle Scholar
- Nekrutenko A, Makova KD, Li WH: The K(A)/K(S) ratio test for assessing the protein-coding potential of genomic regions: an empirical and simulation study. Genome Res. 2002, 12 (1): 198-202.PubMed CentralView ArticlePubMedGoogle Scholar
- Yang Z: PAML 4: phylogenetic analysis by maximum likelihood. Mol Biol Evol. 2007, 24 (8): 1586-1591.View ArticlePubMedGoogle Scholar
- Anisimova M, Bielawski JP, Yang Z: Accuracy and power of the likelihood ratio test in detecting adaptive molecular evolution. Mol Biol Evol. 2001, 18 (8): 1585-1592.View ArticlePubMedGoogle Scholar
- Gu X, Velden Vander K: DIVERGE: phylogeny-based analysis for functional-structural divergence of a protein family. Bioinformatics. 2002, 18 (3): 500-501.View ArticlePubMedGoogle Scholar
- Gu X: Functional divergence in protein (family) sequence evolution. Genetica. 2003, 118 (2–3): 133-141.View ArticlePubMedGoogle Scholar
- Gu X: A simple statistical method for estimating type-II (cluster-specific) functional divergence of protein sequences. Mol Biol Evol. 2006, 23 (10): 1937-1945.View ArticlePubMedGoogle Scholar
- Adams KL, Wendel JF: Polyploidy and genome evolution in plants. Curr Opin Plant Biol. 2005, 8 (2): 135-141.View ArticlePubMedGoogle Scholar
- Soltis DE, Soltis PS: Polyploidy: recurrent formation and genome evolution. Trends Ecol Evol. 1999, 14 (9): 348-352.View ArticlePubMedGoogle Scholar
- Goff SA, Ricke D, Lan TH, Presting G, Wang R, Dunn M, Glazebrook J, Sessions A, Oeller P, Varma H, et al: A draft sequence of the rice genome (Oryza sativa L. ssp. japonica). Science. 2002, 296 (5565): 92-100.View ArticlePubMedGoogle Scholar
- Vandepoele K, Simillion C, Peer Van de Y: Evidence that rice and other cereals are ancient aneuploids. Plant Cell. 2003, 15 (9): 2192-2202.PubMed CentralView ArticlePubMedGoogle Scholar
- Paterson AH, Bowers JE, Chapman BA: Ancient polyploidization predating divergence of the cereals, and its consequences for comparative genomics. Proc Natl Acad Sci USA. 2004, 101 (26): 9903-9908.PubMed CentralView ArticlePubMedGoogle Scholar
- Bowers JE, Chapman BA, Rong J, Paterson AH: Unravelling angiosperm genome evolution by phylogenetic analysis of chromosomal duplication events. Nature. 2003, 422 (6930): 433-438.View ArticlePubMedGoogle Scholar
- Hurley I, Hale ME, Prince VE: Duplication events and the evolution of segmental identity. Evol Dev. 2005, 7 (6): 556-567.View ArticlePubMedGoogle Scholar
- Wolfe KH, Shields DC: Molecular evidence for an ancient duplication of the entire yeast genome. Nature. 1997, 387 (6634): 708-713.View ArticlePubMedGoogle Scholar
- Kellis M, Birren BW, Lander ES: Proof and evolutionary analysis of ancient genome duplication in the yeast Saccharomyces cerevisiae. Nature. 2004, 428 (6983): 617-624.View ArticlePubMedGoogle Scholar
- Zhang Z, Kishino H: Genomic background predicts the fate of duplicated genes: evidence from the yeast genome. Genetics. 2004, 166 (4): 1995-1999.PubMed CentralView ArticlePubMedGoogle Scholar
- Lander ES, Linton LM, Birren B, Nusbaum C, Zody MC, Baldwin J, Devon K, Dewar K, Doyle M, FitzHugh W, et al: Initial sequencing and analysis of the human genome. Nature. 2001, 409 (6822): 860-921.View ArticlePubMedGoogle Scholar
- Roy SW, Fedorov A, Gilbert W: Large-scale comparison of intron positions in mammalian genes shows intron loss but no gain. Proc Natl Acad Sci USA. 2003, 100 (12): 7158-7162.PubMed CentralView ArticlePubMedGoogle Scholar
- Rogozin IB, Wolf YI, Sorokin AV, Mirkin BG, Koonin EV: Remarkable interkingdom conservation of intron positions and massive, lineage-specific intron loss and gain in eukaryotic evolution. Curr Biol. 2003, 13 (17): 1512-1517.View ArticlePubMedGoogle Scholar
- Niu DK, Hou WR, Li SW: mRNA-mediated intron losses: evidence from extraordinarily large exons. Mol Biol Evol. 2005, 22 (6): 1475-1481.View ArticlePubMedGoogle Scholar
- Roy SW, Gilbert W: The pattern of intron loss. Proc Natl Acad Sci USA. 2005, 102 (3): 713-718.PubMed CentralView ArticlePubMedGoogle Scholar
- Banyai L, Patthy L: Evidence that human genes of modular proteins have retained significantly more ancestral introns than their fly or worm orthologues. FEBS Lett. 2004, 565 (1–3): 127-132.View ArticlePubMedGoogle Scholar
- Robertson HM: Two large families of chemoreceptor genes in the nematodes Caenorhabditis elegans and Caenorhabditis briggsae reveal extensive gene duplication, diversification, movement, and intron loss. Genome Res. 1998, 8 (5): 449-463.PubMedGoogle Scholar
- Frugoli JA, McPeek MA, Thomas TL, McClung CR: Intron loss and gain during evolution of the catalase gene family in angiosperms. Genetics. 1998, 149 (1): 355-365.PubMed CentralPubMedGoogle Scholar
- Feiber AL, Rangarajan J, Vaughn JC: The evolution of single-copy Drosophila nuclear 4f-rnp genes: spliceosomal intron losses create polymorphic alleles. J Mol Evol. 2002, 55 (4): 401-413.View ArticlePubMedGoogle Scholar
- Krzywinski J, Besansky NJ: Frequent intron loss in the white gene: a cautionary tale for phylogeneticists. Mol Biol Evol. 2002, 19 (3): 362-366.View ArticlePubMedGoogle Scholar
- Wagner A: The fate of duplicated genes: loss or new function?. Bioessays. 1998, 20 (10): 785-788.View ArticlePubMedGoogle Scholar
- Lynch M, Conery JS: The evolutionary fate and consequences of duplicate genes. Science. 2000, 290 (5494): 1151-1155.View ArticlePubMedGoogle Scholar
- Lynch M, Force A: The probability of duplicate gene preservation by subfunctionalization. Genetics. 2000, 154 (1): 459-473.PubMed CentralPubMedGoogle Scholar
- Lynch M, O'Hely M, Walsh B, Force A: The probability of preservation of a newly arisen gene duplicate. Genetics. 2001, 159 (4): 1789-1804.PubMed CentralPubMedGoogle Scholar
- He X, Zhang J: Rapid subfunctionalization accompanied by prolonged and substantial neofunctionalization in duplicate gene evolution. Genetics. 2005, 169 (2): 1157-1164.PubMed CentralView ArticlePubMedGoogle Scholar
- Fox BG, Malone TE, Johnson KA, Madson SE, Aceti D, Bingman CA, Blommel PG, Buchan B, Burns B, Cao J, et al: X-ray structure of Arabidopsis At1g7 12-oxophytodienoate reductase isoform 1. Proteins. 7680, 61 (1): 206-208.View ArticleGoogle Scholar
- Malone TE, Madson SE, Wrobel RL, Jeon WB, Rosenberg NS, Johnson KA, Bingman CA, Smith DW, Phillips GN, Markley JL, et al: X-ray structure of Arabidopsis At2g0 12-oxophytodienoate reductase isoform 3. Proteins. 6050, 58 (1): 243-245.View ArticleGoogle Scholar
- Breithaupt C, Kurzbauer R, Lilie H, Schaller A, Strassner J, Huber R, Macheroux P, Clausen T: Crystal structure of 12-oxophytodienoate reductase 3 from tomato: self-inhibition by dimerization. Proc Natl Acad Sci USA. 2006, 103 (39): 14337-14342.PubMed CentralView ArticlePubMedGoogle Scholar
- Guex N, Peitsch MC: SWISS-MODEL and the Swiss-PdbViewer: an environment for comparative protein modeling. Electrophoresis. 1997, 18 (15): 2714-2723.View ArticlePubMedGoogle Scholar
- Ogdenw TH, Rosenberg MS: Multiple sequence alignment accuracy and phylogenetic inference. Syst Biol. 2006, 55 (2): 314-328.View ArticleGoogle Scholar
- Smythe AB, Sanderson MJ, Nadler SA: Nematode small subunit phylogeny correlates with alignment parameters. Syst Biol. 2006, 55 (6): 972-992.View ArticlePubMedGoogle Scholar
- Talavera G, Castresana J: Improvement of phylogenies after removing divergent and ambiguously aligned blocks from protein sequence alignments. Syst Biol. 2007, 56 (4): 564-577.View ArticlePubMedGoogle Scholar
- Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, Valentin F, Wallace IM, Wilm A, Lopez R, et al: Clustal W and Clustal X version 2.0. Bioinformatics. 2007, 23 (21): 2947-2948.View ArticlePubMedGoogle Scholar
- Castresana J: Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Mol Biol Evol. 2000, 17 (4): 540-552.View ArticlePubMedGoogle Scholar
- Guindon S, Gascuel O: A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Syst Biol. 2003, 52 (5): 696-704.View ArticlePubMedGoogle Scholar
- Kumar S, Tamura K, Nei M: MEGA3: Integrated software for Molecular Evolutionary Genetics Analysis and sequence alignment. Brief Bioinform. 2004, 5 (2): 150-163.View ArticlePubMedGoogle Scholar
- Suyama M, Torrents D, Bork P: PAL2NAL: robust conversion of protein sequence alignments into the corresponding codon alignments. Nucleic Acids Res. 2006, W609-612. 34 Web Server
- Wong WS, Yang Z, Goldman N, Nielsen R: Accuracy and power of statistical methods for detecting adaptive evolution in protein coding sequences and for identifying positively selected sites. Genetics. 2004, 168 (2): 1041-1051.PubMed CentralView ArticlePubMedGoogle Scholar
- Gu X: Statistical methods for testing functional divergence after gene duplication. Mol Biol Evol. 1999, 16 (12): 1664-1674.View ArticlePubMedGoogle Scholar
- Gu X: Maximum-likelihood approach for gene family evolution under functional divergence. Mol Biol Evol. 2001, 18 (4): 453-464.View ArticlePubMedGoogle Scholar
- Lichtarge O, Bourne HR, Cohen FE: An evolutionary trace method defines binding surfaces common to protein families. J Mol Biol. 1996, 257 (2): 342-358.View ArticlePubMedGoogle Scholar