Origin and evolution of auxin-mediated acid growth

Abstract

The classical acid growth theory suggests that auxin stimulates cell expansion by triggering apoplast acidification via plasma membrane (PM)-localized H⁺-ATPase. Here, we reconstructed the origin and evolutionary history of auxin-mediated acid growth. Comparative phylogenomic analysis showed that most core components of acid growth originated in Charophyta and then underwent subclass expansion and functional innovation during plant terrestrialization. In Charophyceae algae Chara braunii, we found that PM H⁺-ATPase has formed a core regulatory module with TMK and PP2C.D, which can be activated by photosynthesis-dependent phosphorylation through light rather than auxin. Despite the lack of canonical auxin receptor TRANSPORT INHIBITOR RESPONSE 1/AUXIN SIGNALING F-BOX (TIR1/AFB), auxin elicits significant internodal elongation and transcriptional reprogramming in C. braunii, implying the existence of an ancient auxin-mediated growth mechanism. We propose that the evolution of acid growth represents a neofunctional adaptation to terrestrial environments, in which PM H⁺-ATPase in carbon concentrating for photosynthesis was utilized to acidify apoplast for cell expansion, and the core components responsible for acid growth eventually established a regulatory network in land plants by connecting with the TIR1/AFB pathway.

Keywords: acid growth; auxin; cell expansion; evolution; plant terrestrialization.

Fig. 1.

Origin and evolution of core components responsible for auxin-mediated acid growth. (A) Identification of the orthologs of core components responsible for acid growth in 30 representative plant species. Circle size represents gene copy number. The ancient whole-genome duplication/triplication events were labeled on the different branches of the phylogenetic tree based on previous reports, and the named duplication events are shown alongside their Greek letter (38, 39). (B) Gradual coevolution model of auxin-mediated acid growth. The emergence of prominent features across various evolutionary stages is illustrated.

Fig. 2.

Functional evolution of PM H⁺-ATPases from aquatic to terrestrial. (A) Maximum likelihood tree of P-type H⁺-ATPases isolated and reconstructed from a monophyletic group of similar-sequence proteins. AtAHAs and CbHAs are highlighted. Additional representative P-type H⁺-ATPases from fungi were regarded as outgroups. Branches with bootstrap values greater than 50 are displayed. (B) The phosphorylation level of PM H⁺-ATPase of C. braunii in response to IAA and white light. The microsomal fractions were extracted from 12-hour-dark-adapted C. braunii thalli, which were pretreated in the liquid mSWC-2 medium containing 0.1% dimethyl sulfoxide (DMSO)/1 μM IAA/10 μM DCMU for 1 h, then were illuminated with white light at 1000 lx or kept in the dark for 6 h. The immunoblot assay was analyzed on an Mn²⁺-based phos-assay or a standard SDS-PAGE gel and probed with an anti-PM H⁺-ATPase antibody. The graph represents the phosphorylation level of PM H⁺-ATPase, quantified according to the signal intensity ratio of the phosphorylated H⁺-ATPase band to the sum of phosphorylated and unphosphorylated H⁺-ATPase bands. Values represent the mean and SEMs of three independent experiments. Paired t tests were used to calculate significant differences (P < 0.05). (C) PM H⁺-ATPase hydrolysis activities in C. braunii thalli under different treatments. Vanadate-sensitive ATP hydrolysis was measured by determining nicotinamide adenine dinucleotide (NADH) consumption. Values represent the means of three independent biological replications with SEMs. One-way ANOVA with Tukey’s test was used to calculate significant differences.

Fig. 3.

TMK and PP2C.D originated and formed a PM H⁺-ATPase-PP2C.D/TMK interacting module in Charophyta. (A) Maximum likelihood tree of TMKs and their domain architecture. Bootstrap values greater than 50 are indicated. (B) Maximum likelihood tree of the PP2C family and domain recombination events. Colors indicated by dots represent the ancestral species categories included in PP2C clades. PP2C.Ds are compared with PP2C.Cs. The table shows the distribution of PP2C.D-specific AMPC isoforms and their proportion in PP2C.Ds of each species. (C) Membrane-based yeast two-hybrid assays showing the interactions of CbTMKLC and CbPP2C.D with CbHA2. (D) Co-IP assays showing the association of the CbHA2 with CbTMKLC and CbPP2C.D in rice protoplasts transiently expressing indicated epitope-tagged fusion proteins.

Fig. 4.

Gene duplication–mediated neofunctionalization facilitates the establishment of acid growth in land plants. (A) Maximum likelihood tree of SAUR family. Bootstrap values are shown at key nodes. Colors indicated by dots represent the species categories included in SAUR family subgroups. (B) Nuclear-based yeast two-hybrid assays testing interactions between SAURs and PP2C.Ds. Plates were incubated for 24 h to visualize color differences. (C and D) Relative expression levels of CbSAUR and AtSAUR76-78 under treatment with 1 μM IAA or 5 ppm ethylene, as determined by RT-qPCR. RNA samples were extracted from 7-d-old Arabidopsis seedlings and vegetative stages of C. braunii thalli. Values are shown as means ± SEM; n = 3. Two-way ANOVA was used to calculate significant differences (P < 0.05) within each gene. (E) Maximum likelihood tree of the EXP superfamily in plants, bacteria, and fungi. The bootstrap values are shown for key nodes. Colors indicated by dots/branches represent the species categories included in each family. Species of Chlorophyta/Charophyta EXPs in the three clades are listed on the right.

Fig. 5.

The presence and absence of critical elements for auxin-binding and subcellular localization of ABP1/ABL. (A) Maximum likelihood tree of ABP1/ABL isolated and reconstructed from a monophyletic group of Cupin_2 domain-containing proteins. Red stars indicate an archaeal protein previously regarded as ABP1 (Sulfolobus acidocaldarius, YP_255873.1). Branches with bootstrap values greater than 45 are displayed. Solid circles on the right indicate the presence of a signal peptide, auxin-binding motif, and KDEL motif; hollow circles represent the absence of these elements. (B) Amino acid sequence alignment of auxin-binding motifs in representative ABP1/ABL sequences. Red asterisks represent conserved metal-core sites, and blue asterisks indicate conserved hydrophobic sites.

Fig. 6.

Auxin-induced internodal cell elongation and transcriptional reprogramming in C. braunii. (A) Elongation curves of the apical three internodal cells of C. braunii were examined under treatment with 0.1% DMSO (mock), 1 μM IAA, and 50 μM NPA. The algae were grown under a 10 h light: 14 h dark cycle. Each treatment was started after 1 h light period. Values are shown as means ± SEMs; n = 14. The positions of the three internodal cells are indicated on the Left. (Scale bars, 1 cm.) (B) Heatmap illustrating the expression levels of representative DEGs at 6 h. Normalized expression values (TPM) are shown in the cells. The cell colors correspond to the column value normalized as a relative value (Z score) indicated by the scale. (C) RT-qPCR verification of IAA-mediated CbRGLG2a gene upregulation in C. braunii thalli exposed or not exposed to 1 μM IAA and 10 mg/mL CHX for 6 h. Values are shown as means ± SEMs; n = 3. Unpaired t tests were used to calculate significant differences within CHX− and CHX+ groups.