- © 2020 American Society of Plant Biologists. All Rights Reserved.
Abstract
The aerial epidermis of land plants is covered with a hydrophobic cuticle that protects the plant against environmental stresses. Although the mechanisms of cuticle biosynthesis have been extensively studied in model plants, particularly in seed plants, the origins and evolution of cuticle biosynthesis are not well understood. In this study, we performed a comparative genomic analysis of core components that mediate cuticle biosynthesis and characterized the chemical compositions and physiological parameters of cuticles from a broad set of embryophytes. Phylogenomic analysis revealed that the cuticle biosynthetic machinery originated in the last common ancestor of embryophytes. Coexpansion and coordinated expression are evident in core genes involved in the biosynthesis of two major cuticle components: the polymer cutin and cuticular waxes. Multispecies analyses of cuticle chemistry and physiology further revealed higher loads of both cutin and cuticular waxes in seed plants than in bryophytes as well as greater proportions of dihydroxy and trihydroxy acids, dicarboxylic acids, very-long-chain alkanes, and >C28 lipophilic compounds. This can be associated with land colonization and the formation of cuticles with enhanced hydrophobicity and moisture retention capacity. These findings provide insights into the evolution of plant cuticle biosynthetic mechanisms.
The first land plants originated around 500 million years ago from ancestral freshwater algae and underwent dramatic diversification to grow successfully in diverse terrestrial habitats (Morris et al., 2018; Puttick et al., 2018; Huang et al., 2020). During the conquest of the land, plants encountered various environmental challenges, such as desiccation, ultraviolet radiation, mechanical damages, and pathogen infections. To adapt to dehydrating habitats, plants acquired the capacity to synthesize a hydrophobic skin, the cuticle, to cover the aerial surfaces and protect their tissues against desiccation, ultraviolet radiation, and other environmental stresses (Renault et al., 2017; Lee et al., 2020). As the interface between aerial plant organs and their environments, the cuticle is generally composed of a cross-linked cutin scaffold filled and covered with a mixture of cuticular waxes (Yeats and Rose, 2013).
Cutin consists of a large amount of cross-linked polyesters of oxygenated long-chain (C16 or C18) fatty acids and their derivatives as well as phenolic compounds such as coumaric and caffeic acids (Fich et al., 2016; Philippe et al., 2020). The biosynthesis of cutin starts with the formation of cutin monomers, which has been summarized in prior reviews (Yeats and Rose, 2013; Fich et al., 2016; Philippe et al., 2020). The biosynthesis of the cutin lipidic precursors mainly occurs in the endoplasmic reticulum and involves the esterification of plastid-derived C16 and C18 fatty acids to CoA by long-chain acyl-coenzyme A synthetase (LACS) proteins, the oxidation of acyl-CoA, mediated by CYP86A and CYP77A family cytochrome P450 enzymes and the protein HOTHEAD, as well as the synthesis of cutin precursor monoacylglycerols from CoA-activated oxygenated fatty acids by glycerol-3-phosphate acyltransferase (GPAT) enzymes. In addition, a recent study in Arabidopsis (Arabidopsis thaliana) revealed that EPOXIDE HYDROLASE1 (AtEH1) encodes a cytosolic epoxide hydrolase involved in the synthesis of polyhydroxylated cutin monomers (Pineau et al., 2017). The resulting cutin monomers are then exported to the apoplast from the cell by plasma membrane (PM)-localized ATP-binding cassette (ABC) transporters and are deposited into the cell wall (Philippe et al., 2020). Cutin synthase (CUS) enzymes of the GDSL motif lipase/esterase superfamily then catalyze cutin polymerization (Yeats et al., 2012; Philippe et al., 2016; Hong et al., 2017). Furthermore, the involvement of the Arabidopsis α/β-hydrolase family protein BODYGUARD (BDG) and acyltransferase DEFECTIVE IN CUTICULAR RIDGES (DCR) in cutin formation has been suggested by their mutant phenotypes (Yeats and Rose, 2013; Fich et al., 2016; Philippe et al., 2020). In addition, CYP98A family cytochrome P450 enzymes were reported to be involved in the metabolism of phenolic compounds that are components of moss cuticles, and the conjugation of both phenolic (ferulate) and aliphatic (ω-hydroxy acid) compounds is catalyzed by the transferase enzyme DEFICIENT IN CUTIN FERULATE (DCF) in Arabidopsis (Renault et al., 2017; Philippe et al., 2020).
Cuticular wax mixtures are mainly composed of very-long-chain (VLC; >C20) lipophilic compounds including fatty acids, alcohols, aldehydes, alkanes, esters, and ketones (Yeats and Rose, 2013; Lee and Suh, 2015). The molecular and biochemical mechanisms of cuticular wax biosynthesis in Arabidopsis have been summarized in previous reviews (Yeats and Rose, 2013; Lee and Suh, 2015). Cuticular wax biosynthesis shares the steps of acyl-CoA synthesis with the cutin biosynthetic pathway, and the generated C16 and C18 acyl-CoAs are elongated to VLC acyl-CoAs (up to C34) by the fatty acid elongase complex, which consists of β-ketoacyl-CoA synthase (KCS), β-ketoacyl-CoA reductase (KCR), 3-hydroxyacyl-CoA dehydratase (HCD), and enoyl-CoA reductase (ECR). In addition, ECERIFERUM2 (CER2) and its close homologs function in concert with the fatty acid elongase complex to further elongate the VLC acyl-CoAs beyond C28 (Haslam et al., 2012, 2015). VLC fatty acid derivatives, such as alcohols, alkanes, aldehydes, ketones, and wax esters, are produced from these elongated VLC acyl-CoAs through either alkane-forming or alcohol-forming pathways (Lee and Suh, 2015). In the alkane-forming pathway, the CER1/CER3/CYTOCHROME B5 (CYTB5) complex mediates the biosynthesis of VLC alkanes, which are further oxidized to secondary alcohols and ketones by the CYP95A family cytochrome P450 enzyme MIDCHAIN ALKANE HYDROXYLASE1 (MAH1; Yeats and Rose, 2013; Lee and Suh, 2015; Pascal et al., 2019). In the alcohol-forming pathway, the acyl desaturase CER17 mediates the formation of n-6 monounsaturated fatty acids, which are further converted to primary alcohols by the fatty acyl-CoA reductase CER4 (Lee and Suh, 2015; Yang et al., 2017). Additionally, the acyl-CoAs and primary alcohols are used as precursors for the biosynthesis of wax esters, catalyzed by WAX SYNTHASE/ACYL-COA:DIACYLGLYCEROL ACYLTRANSFERASE1 (WSD1) in the alcohol-forming pathway (Yeats and Rose, 2013; Lee and Suh, 2015). The resulting wax components are then transported to the PM through the canonical Golgi- and trans-Golgi network-mediated trafficking system involving GNOM-LIKE1 (GNL1) and ECHIDNA (ECH). The wax components are finally exported out of the cell by PM-localized ABCG half transporters, such as ABCG11 and ABCG12, as well as the glycosylphosphatidylinositol-anchored lipid transfer proteins (LTPGs; Yeats and Rose, 2013; Lee and Suh, 2015).
Cuticle biosynthesis is tightly regulated at the transcriptional levels, and multiple transcriptional regulatory mechanisms have been identified (Lee and Suh, 2015; Fich et al., 2016; Chen et al., 2020; Philippe et al., 2020). For instance, the Arabidopsis APETALA2 (AP2)-ethylene responsive factor-type SHINE (SHN) transcription factors directly regulate the transcription of genes involved in cuticle biosynthesis (Shi et al., 2011). Similarly, the zinc-finger transcription factor NFX1-LIKE2 (NFXL2) and MYB family transcription factors, such as MYB16, MYB30, MYB41, MYB94, MYB96, and MYB106, as well as the class IV homeodomain-Leu zipper family transcription factors HOMEODOMAIN GLABROUS1 (HDG1) and its homologs, were reported to control many aspects of Arabidopsis cuticle formation (Seo et al., 2011; Oshima et al., 2013; Lee et al., 2016). In addition, posttranscriptional regulatory components, such as the exoribonuclease AtCER7, and posttranslational regulatory components, including the RING domain-containing E3 ubiquitin ligase AtCER9 and the Kelch repeat F-box protein SMALL AND GLOSSY LEAVES1 (AtSAGL1), have been implicated in the regulation of cuticle synthesis in Arabidopsis (Hooker et al., 2007; Lü et al., 2012; Kim et al., 2019).
Recent comparative genomic analyses suggested that components of the cuticle biosynthetic machinery evolved in charophyte algae (Lee et al., 2020; Philippe et al., 2020), which represent the immediate ancestors of land plants. However, many details in the evolution of the plant cuticle biosynthetic machinery remain obscure. In this study, we performed a comprehensive evolutionary analysis of genes involved in cuticle biosynthesis from sequenced genomes spanning all major plant lineages and characterized the chemical composition of cuticles from a broad set of embryophytes. In addition, physiological parameters of cuticles from representative land plant lineages, such as hydrophobicity and moisture retention capacity, were measured and compared.
RESULTS
Cuticle Biosynthetic Machinery Originated in the Last Common Ancestor of Embryophytes
To trace the origin of the cuticle biosynthetic machinery, we first selected 41 plant species covering all major plant lineages and identified orthologs of 32 components responsible for cuticle biosynthesis in their genomes (see “Materials and Methods”; Supplemental Table S1). Comparative genomic analysis suggested that genomes of all the tested algae, including streptophyte algae Penium margaritaceum, Mesotaenium endlicherianum, Spirogloea muscicola, Chara braunii, and Klebsormidium nitens, contain only partial sets of genes that comprise cuticle biosynthetic pathways, such as six families of genes related to cutin biosynthesis (LACS, HOTHEAD, CUS, BDG, CYP98A, and EH1), 11 associated with cuticular wax biosynthesis (LACS, KCS, KCR, HCD, ECR, CER1-like/CER3, CYTB5, MAH, CER4, CER17, and WSD), three transporters (GNL1, ECH, and ABCG), and four regulatory components (NFXL2, HDG1, CER7, and CER9; Fig. 1A; Supplemental Fig. S1). Notably, this phylogenomic analysis identified orthologs of all known components required for the biosynthesis of cutin and cuticular wax (LACS, CYP77A, HOTHEAD, GPAT, CUS, BDG, CYP98A, DCF, EH1, LACS, KCS, KCR, HCD, ECR, CER1-like/CER3, CYTB5, MAH, CER4, CER17, and WSD) as well as the components involved in vesicle trafficking for wax secretion (GNL1 and ECH), cuticle transport (ABCG and LTPG), and biosynthesis regulation (NFXL2, MYB, HDG1, CER7, CER9, and SAGL1) in the genomes of all the land plant species, or embryophytes, including bryophytes (Fig. 1A; Supplemental Fig. S1). This suggests that the cuticle biosynthetic machinery originated in the last common ancestor of embryophytes. In addition, the cutin biosynthesis gene CYP86A and the regulatory gene SHN were found in the genomes of the mosses Physcomitrella patens and Sphagnum fallax but not of the hornworts Anthoceros agrestis and Anthoceros angustus or the liverwort Marchantia polymorpha (Fig. 1A). Similarly, the cutin biosynthesis gene DCR and the wax biosynthesis gene CER2-like originated separately in lycophytes and seed plants, suggesting diversification events in the evolution of cuticle biosynthesis during land colonization (Fig. 1A).
Evolutionary analyses of plant cuticle biosynthetic genes. A, Ortholog survey of 32 cuticle biosynthetic genes in 41 genomic data sets from the Archaeplastida. Columns in orange, green, purple, and blue indicate the presence of orthologs of genes involved in cutin biosynthesis, cuticular wax formation, transport, and regulation, respectively. The ortholog numbers corresponding to the various plant species are labeled, and the phylogenetic trees of plants were based on Gao et al. (2018), Puttick et al. (2018), Han et al. (2019), and Jiao et al. (2020). B, Expansion and coevolution analysis of LACS, CUS, and EH1. Gene numbers of LACS, CUS, and EH1 in land plant and algal species are shown as numbers in rectangles and circles respectively, and the numbers of gene gains during evolution are labeled with red plus symbols.
As shown in Figure 1A, expansions were observed in gene families involved in cutin biosynthesis (LACS, CYP86A, CUS, and EH1) and cuticular wax biosynthesis (CER1-like/CER3, CER4, and WSD) in monilophytes and seed plants. To further investigate the gene expansion events, we counted the numbers of LACS, CYP86A, CUS, EH1, CER1-like/CER3, CER4, and WSD in 58 representative land plant species with a known phylogenetic relationship. As shown in Figure 1B and Supplemental Figure S2, the bryophytes had low copy numbers (zero or one) of LACS, CYP86A, CUS, EH1, CER1-like/CER3, CER4, and WSD, and there was evidence of gene expansion events in either monilophytes or seed plants, leading to at least three copies in most associated species. In addition, the copy numbers of LACS, CYP86A, CUS, EH1, CER1-like/CER3, and WSD increased after the divergence of eudicots and monocots (Fig. 1B; Supplemental Fig. S2). Notably, these gene duplication events occurred at overlap ancestral nodes, suggesting that the biosynthetic pathways of cutin and cuticular waxes underwent coexpansion and coevolution during land colonization (Fig. 1B; Supplemental Fig. S2).
Coordinated Expression of Cuticle Biosynthesis Core Genes Is Conserved in Land Plants
Genes involved in related biological processes tend to be coexpressed and closely connected in coexpression networks (Proost and Mutwil, 2018). As reported by studies in the model plant Arabidopsis, the expression of cuticle biosynthesis core genes, such as KCS, LACS, GPAT, KCR, CER1-like/CER3, and CER2-like, is regulated by a common group of SHN and MYB-type transcription factors (Lee and Suh, 2015). To learn more about the evolution of the cuticle biosynthetic machinery, we constructed comparative coexpression networks for cuticle biosynthesis genes in seven representative land plant species with annotated genomic and transcriptomic data. As shown in Figure 2 and Supplemental Table S2, cutin biosynthesis genes (LACS, CYP86A, CYP77A, CUS, BDG, and DCR), cuticular wax biosynthesis genes (KCS, KCR, HCD, ECR, CER2-like, CER1-like/CER3, CER4, and WSD), transporter genes (ABCG and LTPG), and regulatory genes (SHN and MYB) were closely connected in the comparative coexpression networks in the dicotyledonous species Arabidopsis and Solanum lycopersicum. Similar comparative coexpression networks composed of cutin and cuticular wax biosynthesis genes were generated for the monocotyledonous species Oryza sativa and Zea mays, suggesting that cuticle synthesis is mechanistically highly conserved among angiosperms (Fig. 2; Supplemental Table S2). In the gymnosperm Picea abies and the lycophyte Selaginella moellendorffii, the cutin biosynthesis gene CUS resided in coexpression networks together with genes involved in cutin biosynthesis (LACS, CYP86A, CYP77A, and DCF), cuticular wax biosynthesis (KCS, CER1-like/CER3, and WSD), transport (ABCG), and regulation (SHN). Additionally, a similar comparative coexpression network containing five cutin biosynthetic components (LACS, CYP86A, CYP77A, GPAT, and CUS), two cuticular wax biosynthetic components (KCS and CER1-like/CER3), and two transporters (ABCG and LTPG) was identified from the moss P. patens (Fig. 2; Supplemental Table S2).
Coexpression network analysis of cuticle biosynthetic modules in Arabidopsis, S. lycopersicum, O. sativa, Z. mays, P. abies, S. moellendorffii, and P. patens. Plant cuticle biosynthesis gene families are indicated by node colors listed on the left. The coexpression networks were reconstructed using the highest reciprocal rank (HRR) method, where genes with an HRR < 30 were connected with gray lines.
Changes in Cutin Loads and Composition, from Bryophytes to Seed Plants
To investigate the biological significance of the evolution of the cuticle biosynthetic machinery, we used gas chromatography-mass spectrometry to characterize the amount and composition of cutin in cuticles from 13 representative land plant species: three bryophytes (the hornwort A. agrestis, the liverwort M. polymorpha, and the moss P. patens), two lycophytes (S. moellendorffii and Huperzia selago), two monilophytes (Botrypus lanuginosus and Polypodium virginianum), two gymnosperms (P. abies and Ginkgo biloba), and four angiosperms (Z. mays, O. sativa, S. lycopersicum, and Arabidopsis). As shown in Figure 3A, the amounts of total cutin monomers, on a dry residue basis, were less in A. agrestis (1,224 μg g−1) and M. polymorpha (851 μg g−1) than in P. patens (1,582 μg g−1) or the lycophytes S. moellendorffii (2,027 μg g−1) and H. selago (2,955 μg g−1). In addition, the cutin loads in the monilophytes B. lanuginosus and P. virginianum were higher than in lycophytes and bryophytes but were lower than in seed plants (Fig. 3A).
Comparative analysis of cutin composition in 13 representative land plant species. A, Comparative analysis of amounts of total cutin monomers in A. agrestis, M. polymorpha, P. patens, S. moellendorffii, H. selago, B. lanuginosus, P. virginianum, P. abies, G. biloba, Z. mays, O. sativa, S. lycopersicum, and Arabidopsis. B, Heat map of the normalized amounts of cutin monomers (percentage of total cutin monomers) in 13 representative land plant species. C to E, Comparative analyses of amounts (percentage of total cutin monomers) of dihydroxy and trihydroxy acids (C), dicarboxylic acids (D), and phenolic compounds (E) in 13 representative land plant species. The phylogenetic trees were based on Gao et al. (2018), Puttick et al. (2018), Han et al. (2019), and Jiao et al. (2020). For A and C to E, three independent biological replicates were analyzed, and results are shown as means ± se. Asterisks indicate significant difference by Student’s t test (**P < 0.01).
Chemical composition analysis revealed that cutins of A. agrestis and M. polymorpha were composed of phenolic compounds (mainly hydroxybenzoic acid and coumaric acid), C16 fatty acid, and hydroxyhexadecanoic acid (Fig. 3B; Supplemental Table S3). Notably, they contained 15-hydroxyhexadecanoic acid and 10,15-dihydroxyhexadecanoic acid, which were not identified from representatives of other plant lineages, such as the moss P. patens (Fig. 3B). P. patens cutin was found to contain large proportions of C16 fatty acid and 10,16-dihydroxyhexadecanoic acid, which was also the case for the lycophytes S. moellendorffii and H. selago (Fig. 3B). Quantitative analysis showed that the amounts of cuticular C18 acids were much higher in seed plants, especially in angiosperms, than in any other seedless plant (Supplemental Fig. S3). Further quantitative analysis revealed that the proportions of C16 and C18 hydroxy acids, especially the dihydroxy and trihydroxy acids, among the total cutin monomers were much higher in seed plants than in seedless plants (Fig. 3C; Supplemental Fig. S4). Similarly, the proportion of dicarboxylic acids, including C16:0 diacid, C18:0 diacid, C18:1 diacid, and C18:2 diacid, among the total cutin monomers increased from less than 2% in the hornwort A. agrestis and the liverwort M. polymorpha to more than 15% in seed plants (Fig. 3D). Conversely, the proportion of phenolic compounds among the total cutin monomers decreased from more than 35% in bryophytes to less than 16% in seed plants (Fig. 3E).
Changes in Cuticular Wax Coverage and Composition, from Bryophytes to Seed Plants
We next analyzed the chemical composition of cuticular waxes from the same 13 representative land plant species by gas chromatography-mass spectrometry. As shown in Figure 4A, total cuticular wax loads, on a fresh weight basis, were far lower in A. agrestis and M. polymorpha (<0.1 μg g−1) than in P. patens (2.77 μg g−1). In addition, the amounts of cuticular waxes in the lycophytes S. moellendorffii (4.68 μg g−1) and H. selago (4.37 μg g−1) and the monilophytes B. lanuginosus (10.96 μg g−1) and P. virginianum (8.95 μg g−1) were less than those in gymnosperms and angiosperms (Fig. 4A).
Comparative analysis of cuticular wax composition in 13 representative land plant species. A, Comparative analysis of amounts of total cuticular wax in A. agrestis, M. polymorpha, P. patens, S. moellendorffii, H. selago, B. lanuginosus, P. virginianum, P. abies, G. biloba, Z. mays, O. sativa, S. lycopersicum, and Arabidopsis. B, Heat map of the normalized amounts of cuticular wax components (percentage of total cuticular waxes) from 13 representative land plant species. C to E, Comparative analyses of amounts (percentage of total cuticular waxes) of VLC alkanes (C), >C28 lipophilic compounds (D), and VLC fatty acids (E) in 13 representative land plant species. The phylogenetic trees were based on Gao et al. (2018), Puttick et al. (2018), Han et al. (2019), and Jiao et al. (2020). For A and C to E, three independent biological replicates were analyzed, and results are shown as means ± se. Asterisks indicate significant difference by Student’s t test (**P < 0.01).
An analysis of cuticular wax composition revealed that the cuticular waxes in A. agrestis and M. polymorpha were mainly composed of VLC fatty acids (Fig. 4B; Supplemental Table S4). Other VLC fatty acid derivatives, such as alkanes, alcohols, ketones, and wax esters, were not detected in extracts from A. agrestis and M. polymorpha (Fig. 4B). In contrast, the cuticular waxes of P. patens contained large amounts of VLC alcohols, fatty acids, and wax esters, which was also the case for the lycophytes S. moellendorffii and H. selago (Fig. 4B). Quantitative analysis revealed that the percentage of VLC alkanes increased from less than 0.2% in bryophytes to more than 33% in seed plants (Fig. 4C). In addition, >C28 lipophilic compounds, such as fatty acids, alcohols, aldehydes, alkanes, and esters, were present at higher levels in the cuticular waxes of seed plants, monilophytes, and lycophytes than in A. agrestis or M. polymorpha (Fig. 4D). In contrast, the proportion of VLC fatty acids was more than 69% in A. agrestis and M. polymorpha but less than 17% in seed plants (Fig. 4E). Taken together, this wax chemical composition analysis revealed that cuticular wax loads increased among land plant lineages from the early-diverging bryophytes to the later-diverging seed plants, which was accompanied by the growing proportions of VLC alkanes and >C28 lipophilic compounds as well as the declining proportion of VLC fatty acids.
Changes in Cuticle Hydrophobicity and Moisture Retention Capacity, from Bryophytes to Seed Plants
We next evaluated cuticle hydrophobicity by measuring the contact angle of water droplets on the phyllid or leaf surface of the 13 representative land plant species (Kurokawa et al., 2018). As shown in Figure 5A, the phyllid surfaces of A. agrestis and M. polymorpha exhibited contact angles of 65° and 52°, respectively, which were much less than the values for P. patens (92°), S. moellendorffii (109°), and H. selago (107°), indicating the lowest cuticle hydrophobicity of A. agrestis and M. polymorpha. The leaf surfaces of seed plants showed contact angles greater than 125° and had higher values than those of monilophytes, lycophytes, and bryophytes, indicating the highest cuticle hydrophobicity in seed plants (Fig. 5A).
Comparative analyses of cuticle hydrophobicity and moisture retention capacity in 13 representative land plant species. A, Hydrophobicity analysis in the phyllids or leaves of A. agrestis, M. polymorpha, P. patens, S. moellendorffii, H. selago, B. lanuginosus, P. virginianum, P. abies, G. biloba, Z. mays, O. sativa, S. lycopersicum, and Arabidopsis. Contact angles from at least 50 water droplets were separately measured, and five independent biological replicates were analyzed. Results are shown as means ± se. Asterisks indicate significant difference by Student’s t test (**P < 0.01). B, Comparative analysis of moisture retention capacity in the phyllids or leaves of A. agrestis, M. polymorpha, P. patens, S. moellendorffii, H. selago, B. lanuginosus, P. virginianum, P. abies, G. biloba, Z. mays, O. sativa, S. lycopersicum, and Arabidopsis. Numbers of hours in a dehydrating environment are shown at the bottom of the graphs. Three independent biological replicates were analyzed, and results are shown as means ± se. Asterisks indicate significant difference by Student’s t test (**P < 0.01). The phylogenetic trees were based on Gao et al. (2018), Puttick et al. (2018), Han et al. (2019), and Jiao et al. (2020).
Since a critical function of the hydrophobic cuticle is to limit transpirational water loss, we examined the moisture retention capacity. As shown in Figure 5B, A. agrestis and M. polymorpha maintained less than 23% phyllid moisture content after 5 h of exposure to a dehydrating environment but P. patens maintained greater than 29% phyllid moisture content, indicative of a higher capacity to restrict desiccation. In contrast, the seed plants maintained a higher leaf moisture content (51% to 76%) than S. moellendorffii (34%), H. selago (29%), B. lanuginosus (41%), and P. virginianum (44%), suggesting that cuticles of seed plants have the highest capacity to restrict water loss (Fig. 5B). These results indicated that cuticle hydrophobicity and moisture retention capacity increased from bryophytes to seed plants.
DISCUSSION
Origin of Basic Cuticle Biosynthetic Machinery in the Common Ancestor of Land Plants
Chemical analysis of plant fossils has previously led to the suggestion that plants evolved the capacity to deposit a cuticle approximately 450 million years ago and that this enabled land colonization (Niklas et al., 2017). Although some algae, such as K. nitens, have extracellular wax-like lipid deposits, true cuticles are thought to be unique to land plants (Kondo et al., 2016). In our phylogenomic analysis, orthologs of genes involved in cuticle biosynthesis, transport, and regulation were identified from the genomes of algae, consistent with the recent studies (Cheng et al., 2019; Jiao et al., 2020; Philippe et al., 2020). Notably, homologs of cuticle biosynthesis genes, such as LACS, HOTHEAD, KCR, HCD, ECR, CER1-like/CER3, CYTB5, MAH, CER4, CER17, GNL1, ECH, ABCG, NFXL2, CER7, and CER9, were identified in the genomes of most plant lineages, including chlorophytes, suggesting that they represent conserved biochemical functions involved in basic cellular processes that were later recruited for cuticle biogenesis (Fig. 1A). For instance, the posttranscriptional regulation genes CER7 and CER9 may have played a role, prior to the evolutionary appearance of the cuticle, in other important cellular processes, such as RNA processing and stress responses (Lü et al., 2012; Lee et al., 2020; Philippe et al., 2020). Similarly, LACS, CER17, and CYTB5 may have first functioned in generating fatty acid structures necessary for intracellular membranes (Yeats and Rose, 2013; Fich et al., 2016; Yang et al., 2017).
In this study, most of the cuticle biosynthetic genes were present in the genomes of three members of the Zygnematophyceae, the sister lineage to land plants, consistent with their evolutionary trajectory toward terrestrialization (Wickett et al., 2014; Puttick et al., 2018; Cheng et al., 2019; Jiao et al., 2020). However, no closely related homologs of the cuticle biosynthesis core genes CYP86A, CYP77A, GPAT, DCF, CER2-like, LTPG, SHN, and MYB were identified in P. margaritaceum, M. endlicherianum, and S. muscicola, suggesting that a canonical land plant-like cuticle originated in the last common ancestor of the Zygnematophyceae and land plants (Fig. 1A). Since a hydrophobic cuticle would likely act as an important barrier to the diffusion and absorption of dissolved atmospheric gases in an aquatic environment, the absence of a cuticle in algae may result from selection against cuticle biosynthesis (Niklas et al., 2017).
Our comparative genomic analysis revealed that homologs of most core genes known to be involved in cuticle biosynthesis in angiosperms are present in all bryophytes, including representatives of the hornwort, liverwort, and moss lineages, which were recently shown to have evolved from a common ancestor (Fig. 1A; Bowman et al., 2017; Li et al., 2020; Zhang et al., 2020). In addition, biochemical analysis showed that cuticles from A. agrestis and M. polymorpha have relatively high levels of fatty acids and phenolic compounds such as hydroxybenzoic acid and coumaric acid (Figs. 3 and 4). However, very few of them have been functionally evaluated or experimentally associated with cuticle formation. A previous study reported that the cuticles of two other species of liverwort, Astarella lindenbergiana and Conocephalum conicum, also mainly consist of fatty acids and phenolic compounds (Caldicott and Geoffrey, 1976). These phylogenomic and biochemical findings support the hypothesis that some, but not all, of the cuticle biosynthetic machinery originated in the last common ancestor of embryophytes (as summarized in Fig. 6).
A model for the origin and evolution of cuticle biosynthetic machinery during plant land colonization. Although several components mediating cuticle biosynthesis, transport, and even regulation emerged in algae, additional important cuticle biosynthetic machinery originated in the last common ancestor of embryophytes. Gene expansion and emergence occurred afterward, resulting in changes in cuticle chemical composition and contributions to plant physiology. During the evolutionary transition from bryophytes to seed plants, both cutin and cuticular wax loads increased, together with the proportions of dihydroxy and trihydroxy acids, dicarboxylic acids, VLC alkanes, and >C28 lipophilic compounds, while the relative amounts of phenolic compounds and VLC fatty acids decreased, leading to the formation of cuticles in seed plants with enhanced hydrophobicity and moisture retention properties. Ma, Million years ago.
Coexpansion and Coordinated Expression of Cuticle Biosynthesis Genes in Land Plants
Our phylogenetic analyses revealed that the cutin biosynthesis gene families CYP86A, LACS, CUS, and EH1 exhibited coexpansion with the cuticular wax biosynthesis genes CER1-like/CER3, CER4, and WSD in monilophytes and seed plants. We propose that the interconnected cutin and cuticular wax biosynthetic pathways were shaped by the common selection pressures (Fig. 1B; Supplemental Fig. S2). Moreover, gene coexpression analysis revealed that the core genes responsible for the biosynthesis of cutin and cuticular waxes are present in overlapping coexpression networks, consistent with regulation by a common set of transcription factors, such as SHN1/2/3 and MYB16/106 (Yeats and Rose, 2013; Lee and Suh, 2015). Notably, the coexpressed relationships of cuticle biosynthesis genes were evident lineages from bryophytes to seed plants, suggesting functional associations during plant evolution. These results also suggest that knowledge of function obtained from the study of model plants such as Arabidopsis may be used for functional inference in other plants, including crops and earlier diverging land plant lineages (Fig. 3; Ruprecht et al., 2017; Huang and Zhu, 2019; Lampugnani et al., 2019).
Comparative Analyses of Cuticle Chemical Composition and Physiological Parameters
In this study, a comparative analysis of cuticle chemical compositions revealed that both cutin and cuticular wax loads increased in proportion to the evolutionary time of divergence of the lineage from bryophyte taxa to seed plants (Figs. 3 and 4). Among the tested land plants, A. agrestis and M. polymorpha contained the lowest levels of cuticle components, which were mainly composed of fatty acids and phenolic compounds. Notably, phenolic compounds have a high ultraviolet radiation-absorbing capacity, which is consistent with their proposed protective role in bryophytes (Soriano et al., 2019) and more broadly in plants that inhabit terrestrial habitats. In addition, an enrichment in fatty acids would makes the cuticle of A. agrestis and M. polymorpha somewhat hydrophobic and might enhance tolerance to damage during rehydration and equilibration of water content with the surrounding environment. Recently, collapsed stomata were identified in hornworts, where they may function in concert with a cuticle to promote gas exchange and transpirational water loss, further contributing to the sporangial and spore maturation (Renzaglia et al., 2017). Notably, 15-hydroxyhexadecanoic acid and 10,15-dihydroxyhexadecanoic acid were identified in the cuticles of A. agrestis and M. polymorpha but not other plant lineages. A previous study reported that cuticles of two other species of liverwort, A. lindenbergiana and C. conicum, also contain these compounds, suggesting that they might be unique to cuticles of hornworts and liverworts (Caldicott and Geoffrey, 1976).
Consistent with previous studies, we found that the cuticle chemical composition of P. patens is similar to that of lycophytes, monilophytes, and seed plants (Buda et al., 2013; Renault et al., 2017). Further quantitative analysis showed that the proportions of dihydroxy and trihydroxy acids, dicarboxylic acids, VLC alkanes, and >C28 lipophilic compounds in cuticles increase in an evolutionary sequence from bryophytes to seed plants, consistent with previous studies (Fich et al., 2016). Fatty acids with two or three hydroxyl groups contribute to the branching of cutin chains, and dicarboxylic acids are involved in ester cross-linking between cutin chains (Fich et al., 2016). Therefore, elevated levels of these compounds may be associated with the formation of a more highly connected cutin matrix, which would likely have a major effect on biomechanical and physicochemical properties of the cuticle. Given that VLC alkanes and >C28 lipophilic compounds are more hydrophobic than other cuticle constituents, their enrichment in spermatophyte cuticles would likely contribute to the formation of cuticles with higher hydrophobicity and moisture retention capacity than those of bryophytes and lycophytes, which is consistent with our physiological data (Fig. 5).
To conclude, the results of this study are consistent with a model where a subset of the biosynthetic machinery required for cuticle formation evolved in aquatic algae but that canonical cuticles with important barrier properties originated in the last common ancestor of embryophytes. This evolutionary innovation was likely a critical adaptation to allow the colonization of different terrestrial environments, including arid habitats (as summarized in Fig. 6).
MATERIALS AND METHODS
Material Growth Conditions
Gametophytes of Anthoceros agrestis and Physcomitrella patens (‘Gransden’) were propagated and maintained on BCD medium. Gametophytes of Marchantia polymorpha (‘Takaragaike-1’) was maintained on one-half-strength Gamborg’s B5 plates. Selaginella moellendorffii plants were propagated on 0.5× Murashige and Skoog medium supplemented with 0.8% (w/v) agar at 24°C/22°C (day/night) under a 16/8-h light/dark photoperiod. Huperzia selago, Botrypus lanuginosus, Polypodium virginianum, Picea abies, Ginkgo biloba, Zea mays (‘B73’), Oryza sativa (‘Zhonghua11’), Solanum lycopersicum (‘Micro-Tom’), and Arabidopsis (Arabidopsis thaliana; Columbia-0) plants were grown in growth chambers set at 24°C/22°C (day/night) with a 16/8-h light/dark photoperiod.
Ortholog Identification
To trace the origin of cuticle biosynthesis, we first selected 41 plant species with annotated genomes as our model species and identified orthologs of components responsible for cuticle biosynthesis in these plant genomes (Supplemental Table S1). The selected model species represent all major plant lineages, including 13 angiosperms (Arabidopsis, Carica papaya, Glycine max, S. lycopersicum, Helianthus annuus, Nicotiana tabacum, Aquilegia coerulea, O. sativa, Brachypodium distachyon, Z. mays, Dendrobium catenatum, Spirodela polyrhiza, and Amborella trichopoda), four gymnosperms (Pinus taeda, P. abies, Gnetum montanum, and G. biloba), two monilophytes (Salvinia cucullata and Azolla filiculoides), one lycophyte (S. moellendorffii), five bryophytes (Sphagnum fallax, P. patens, M. polymorpha, A. agrestis, and Anthoceros angustus), five charophytes (Penium margaritaceum, Mesotaenium endlicherianum, Spirogloea muscicola, Chara braunii, and Klebsormidium nitens), six chlorophytes (Volvox carteri, Chlamydomonas reinhardtii, Dunaliella salina, Chlorella variabilis, Ostreococcus tauri, and Micromonas pusilla), four rhodophytes (Chondrus crispus, Porphyridium purpureum, Cyanidioschyzon merolae, and Galdieria surphurus), and one glaucophyte (Cyanophora paradox).
Based on the reported network controlling cuticle synthesis, a total of 32 cuticle biosynthetic components, including 11 cutin biosynthetic components (LACS, CYP86A, CYP77A, HOTHEAD, GPAT, CUS, BDG, DCR, CYP98A, DCF, and EH1), 12 cuticular wax biosynthetic components (LACS, KCS, KCR, HCD, ECR, CER2-like, CER1-like/CER3, CYTB5, MAH, CER4, CER17, and WSD), four transporting components (GNL1, ECH, ABCG, and LTPG), and seven regulatory components (SHN, NFXL2, MYB, HDG1, CER7, CER9, and SAGL1), were selected for the evolutionary analysis of cuticle biosynthesis.
For the ortholog identification in this study, protein sequences of AtLACS1/2/4, AtCYP86A2/4/8, AtCYP77A6, SlGPAT6/AtGPAT4/6/8, AtDCF, SlCUS1/AtCUS1/2, AtEH1, AtBDG1/3, AtHOTHEAD, AtDCR, AtKCR1, AtKCS1/2/6/9/20, AtECR, AtHCD, AtCER4, AtCER17, AtCER1/3, AtCYTB5B, AtWSD1/6/7, AtMAH1, AtCER2/26/26-like, AtGNL1, AtECH, AtLTPG1/2, AtABCG11/12/13/32, AtCER7, AtCER9, AtNFXL2, AtSAGL1, AtHDG1, AtMYB9/16/94/96/106, AtSHN1/2/3, and PpCYP98 were used as templates to perform BLASTP against all genes annotated in 41 representative genomes with an E value of 1e-5 (Supplemental Table S1). Domain annotation was performed with interproscan (http://www.ebi.ac.uk/interpro/), and peptide sequences assigned with the annotations AMP-dependent synthetase/ligase, Cytochrome P450, E-class, group I, Cytochrome P450, E-class, group I, HAD superfamily + Phospholipid/glycerol acyltransferase, Chloramphenicol acetyltransferase-like domain + Transferase, SGNH hydrolase superfamily + GDSL lipase/esterase, Alpha/Beta hydrolase fold, Glucose-methanol-choline oxidoreductase, Chloramphenicol acetyltransferase-like domain + Transferase, Short-chain dehydrogenase/reductase SDR, Very-long-chain 3-ketoacyl-CoA synthase, 3-oxo-5-alpha-steroid 4-dehydrogenase/very-long-chain enoyl-CoA reductase, Protein-tyrosine phosphatase-like, PTPLA, Fatty acyl-CoA reductase, Acyl-CoA desaturase + Fatty acid desaturase domain, Fatty acid hydroxylase + Uncharacterised domain Wax2, C-terminal, Cytochrome b5-like heme/steroid binding domain, O-acyltransferase, WSD1, Cytochrome P450, E-class, group I, Chloramphenicol acetyltransferase-like domain + Transferase, Guanine nucleotide exchange factor + Sec7 domain, Golgi apparatus membrane protein TVP23-like, Bifunctional inhibitor/plant lipid transfer protein/seed storage helical domain, P-loop containing nucleoside triphosphate hydrolase + ABC-2 type transporter, Exosome complex component RRP45, Zinc finger, RING-CH-type, Transcription factor NFX1 family, F-box domain + Kelch repeat type 1, Myb-like transcription factor, AP2/ERF domain + DNA binding domain, and Cytochrome P450, E-class, group I were individually filtered out as ortholog candidates of LACS, CYP86A, CYP77A, GPAT, DCF, CUS, BDG, HOTHEAD, DCR, EH1, KCR, KCS, ECR, HCD, CER4, CER17, CER1-like/CER3, CYTB5, WSD, MAH, CER2-like, GNL1, ECH, LTPG, ABCG, CER7, CER9, NFXL2, SAGL1, HDG1, MYB, SHN, and CYP98A and subjected to the protein alignment with MAFFT, and positions with above 50% gaps were removed using the Phyutility program v2.2.6 (Katoh and Standley, 2013). Substitution model analysis was performed using ProtTest (Darriba et al., 2011), and phylogenetic trees were reconstructed using IQTREE with 500 bootstraps (Nguyen et al., 2015).
Gene Number Estimation
To investigate the gene expansion events, 58 representative land plant species were selected, including 32 seed plants, eight monilophytes, five lycophytes, and 13 bryophytes. The numbers of CYP86A, CUS1, BDG, and CER1 were counted, and the numbers of genes in the ancestral nodes were estimated by the method described by Nam and Nei (2005).
Gene Coexpression Network Construction
The HRR-based coexpression network analysis for Arabidopsis, S. lycopersicum, O. sativa, Z. mays, P. abies, and S. moellendorffii was performed using CoNekT (Coexpression Network Toolkit) with the edge cutoff of 30 (Proost and Mutwil, 2018). For the P. patens gene coexpression analysis, a P. patens HRR-based coexpression network generated by Ruprecht et al. (2017) was downloaded from PlaNet (http://www.gene2function.de/down load.html) and analyzed with the edge cutoff of 30 (Mutwil et al., 2011).
Cutin and Cuticular Wax Composition Analysis
The cutin composition analysis was performed as described with some modifications (Renault et al., 2017; Lee et al., 2020). Briefly, about 2 g of air-dried phyllid or leaf samples from 6-week-old A. agrestis, M. polymorpha, P. patens, S. moellendorffii, H. selago, B. lanuginosus, P. virginianum, P. abies, G. biloba, Z. mays, O. sativa, S. lycopersicum, and Arabidopsis plants was delipidated by sequential washing with isopropanol containing 0.01% (v/v) butylated hydroxytoluene followed by a solvent mixture containing chloroform and methanol. The extracts were dried under N2 and the dry, delipidated tissue weights were recorded. The samples were then heated to 60°C overnight in a reaction mixture containing methanol, methyl acetate, sodium methoxide, pentadecalactone (internal standard), and methyl nonadecanoate (internal standard) to depolymerize the cutin. The methyl esters were extracted with dichloromethane, glacial acetic acid, and NaCl. After drying under a stream of nitrogen, the samples were derivatized with pyridine and bis-N,O-trimethylsilyl trifluoroacetamide and then subjected to analysis with a capillary gas chromatograph (5890 Series II; Agilent Technologies) and a flame ionization detector (6890 N; Agilent Technologies) with a mass spectrometer (MSD 5973; Agilent Technologies). The oven temperature program was set as initial temperature 80°C, increased to 200°C at 15°C min−1, and then increased to 280°C at 2°C min−1. Quantification was based on flame ionization detection peak areas relative to internal standard pentadecalactone and methyl nonadecanoate and the dry, delipidated tissue weights. The cutin compound was identified by employing the Agilent/HP ChemStation softwares (Agilent Technologies).
The cuticular wax composition analysis was performed as described by Seo et al. (2011) and Lee et al. (2020). Briefly, about 2 g of air-dried phyllid or leaf samples from 6-week-old A. agrestis, M. polymorpha, P. patens, S. moellendorffii, H. selago, B. lanuginosus, P. virginianum, P. abies, G. biloba, Z. mays, O. sativa, S. lycopersicum, and Arabidopsis plants was dipped into chloroform (Merck) for 30 s to extract the cuticular wax. The wax extracts dried under N2 were derivatized at 70°C for 30 min through reaction with bis-N,O-trimethylsilyl trifluoroacetamide and analyzed with the same gas chromatograph and column as used for cutin analysis. The oven temperature was programmed for 2 min at 50°C, 40°C min−1 to 200°C, 2 min at 200°C, 3°C min−1 to 320°C, and 30 min at 320°C. For the quantification of individual compounds, H2 was used as the carrier gas. The wax compound was identified based on the Agilent/HP ChemStation software (Agilent Technologies). Quantification was based on flame ionization detection peak areas relative to the internal standards heneicosanoic acid, n-octacosane, 1-tricosanol, and stearyl stearate.
Contact Angle Measurement
Phyllid or leaf samples were collected from 6-week-old A. agrestis, M. polymorpha, P. patens, S. moellendorffii, H. selago, B. lanuginosus, P. virginianum, P. abies, G. biloba, Z. mays, O. sativa, S. lycopersicum, and Arabidopsis plants, and the cuticle hydrophobicity was analyzed by measuring the contact angle of water droplets (Kurokawa et al., 2018). Briefly, a demineralized deionized water droplet of about 1 μL was placed on the indicated phyllid or leaf surface. Angles from at least 50 water droplets were separately measured using the contact angle system (SDP-300; Sindin), and five independent surface samples were statistically analyzed using Student’s t test.
Moisture Retention Capacity Measurement
Phyllid or leaf samples were harvested from 6-week-old A. agrestis, M. polymorpha, P. patens, S. moellendorffii, H. selago, B. lanuginosus, P. virginianum, P. abies, G. biloba, Z. mays, O. sativa, S. lycopersicum, and Arabidopsis plants, and the fresh sample weights were recorded immediately as FW0. Then, the phyllid or leaf samples were kept at room temperature (23°C ± 0.5°C), and relative humidity (50% ± 5%) and their fresh weights were recorded at 1, 3, and 5 h as FWn. At the end of the experiment, leaves were dried in a hot oven at 75°C for 20 h. The moisture retention capacity (MRC) was estimated using the following formula: MRC (%) = {(FWn − DW)/(FW0 − DW)} × 100%, where DW represents dry weight. Three independent biological replicates were statistically analyzed using Student’s t test.
Accession Numbers
Sequence data used for the ortholog identification in this study can be found in the GenBank database using the following accession numbers: AtLACS1/2/4 (At2g47240/At1g49430/At4g23850); AtCYP86A2/4/8 (At4g00360/At1g01600/At2g45970); AtCYP77A6 (At3g10570); SlGPAT6/AtGPAT4/6/8 (Solyc09g14350/At1g01610/At2g38110/At4g00400); AtDCF (At3g48720); SlCUS1/AtCUS1/2 (Solyc11g006250/At3g04290/At5g33370); AtEH1 (At3g05600); AtBDG1/3 (At1g64670/At4g24140); AtHOTHEAD (At1g72970); AtDCR (At5g23940); AtKCR1 (At1g67730); AtKCS1/2/6/9/20 (At1g01120/At1g04220/At1g68530/At2g16280/At5g43760); AtECR (At3g55360); AtHCD (At5g10480); AtCER4 (At4g33790); AtCER17 (At1g06350); AtCER1/3 (At1g02205/At5g57800); AtCYTB5B (At2g32720); AtWSD1/6/7 (At5g37300/At3g49210/At5g12420); AtMAH1 (At1g57750); AtCER2/26/26-like (At4g24510/At4g13840/At3g23840); AtGNL1 (At5g39500); AtECH (At1g09330); AtLTPG1/2 (At1g27950/At3g43720); AtABCG11/12/13/32 (At1g17840/At1g51500/At1g51460/At2g26910); AtCER7 (At3g60500); AtCER9 (At4g34100); AtNFXL2 (At5g05660); AtSAGL1 (At1g55270); AtHDG1 (At3g61150); AtMYB9/16/94/96/106 (At5g16770/At5g15310/At3g4600/At5g62470/At3g01140); and AtSHN1/2/3 (At1g15360/At5g11190/At5g25390; and PpCYP98, Phpat.022g067700).
Supplemental Data
The following supplemental materials are available.
Supplemental Figure S1. A model of the cuticle biosynthetic pathway in Arabidopsis and identification of orthologs of the core components.
Supplemental Figure S2. Copy number analysis of CYP86A, CER1-like/CER3, CER4, and WSD in the 58 representative land plant species with known phylogenetic relationships.
Supplemental Figure S3. Comparative analysis of amounts (percentage of total cutin monomers) of C18 acids in 13 representative land plant species.
Supplemental Figure S4. Comparative analysis of amounts (percentage of total cutin monomers) of C16 and C18 hydroxy acids in 13 representative land plant species.
Supplemental Table S1. Reference information for 41 representative genomes used in this study.
Supplemental Table S2. Gene lists associated with coexpression networks.
Supplemental Table S3. Absolute values of amounts of cutin constituents in 13 representative land plant lineages.
Supplemental Table S4. Absolute values of amounts of cuticular wax components in 13 representative land plant lineages.
Acknowledgments
We thank Dr. Hongzhi Kong (State Key Laboratory of Systematic and Evolutionary Botany, CAS Center for Excellence in Molecular Plant Sciences, Institute of Botany, Chinese Academy of Sciences) for valuable suggestions on this study and Dr. Bojian Zhong (College of Life Sciences, Nanjing Normal University) for help on reference genomes used in this study. We also thank the editor and three anonymous reviewers for their very helpful comments and advice on earlier versions of this article.
Footnotes
C.C. and L.K. conceived the project and designed research; L.K., Y.L., P.Z., and X.W. performed most of the experiments; B.X. and Z.G. provided technical and professional assistance; C.C. and L.K. analyzed the data and wrote the article with assistance from B.X. and Z.G.; C.C. supervised the project and completed the writing.
↵1 This work was supported by the National Natural Science Foundation of China (grant nos. 31701412 and 31701986), the Natural Science Foundation of Shandong Province (grant no. ZR2017BC109), the Qingdao Science and Technology Bureau Fund (grant nos. 17–1–1–50–jch and 18–2–2–51–jch), and the State Key Laboratory of Plant Physiology and Biochemistry (grant no. SKLPPBKF2002).
↵[OPEN] Articles can be viewed without a subscription.
- Received July 10, 2020.
- Accepted September 5, 2020.
- Published September 15, 2020.
REFERENCES
