Lipid-Protein Interplay: How the Plant Plasma Membrane H-ATPase Dynamically Interacts with Its Lipid Environment – A Review

Ram Kumar
Department of Plant Protection, Agritec Plant Research, Sumperk, Czech Republic
Correspondence to: Ram Kumar, kumar@agritec.cz

DOI: https://doi.org/10.70389/PJPB.100021

Additional information

• Ethical approval: N/a
• Consent: N/a
• Funding: No industry funding
• Conflicts of interest: N/a
• Author contribution: Ram Kumar – Conceptualization, Writing – original draft, review and editing
• Guarantor: Ram Kumar
• Provenance and peer-review:
  Unsolicited and externally peer-reviewed
• Data availability statement: Data sharing is not applicable to this article as no new data were created

Keywords: H⁺-atpase regulation, Lipid-protein interactions, Plant plasma membrane, Sterol influence, Sphingolipid role.

Peer Review
Received: 29 May 2025
Last revised: 24 July 2025
Accepted: 24 July 2025
Version accepted: 2
Published: 9 August 2025

Plain Language Summary Infographic

Introduction

The plasma membrane (PM) H⁺-ATPase is the primary electrogenic proton pump in plants. It actively uses ATP hydrolysis to extrude protons through the PM, generating a proton motive force (ΔμH⁺).¹ This electrochemical gradient is the fundamental energy currency that drives secondary active transport systems through proton-coupled symport and antiport mechanisms.¹ As the principal energizer of PM transport, PM H⁺-ATPase activity is physiologically indispensable, directly mediating critical processes including but not limited to: nutrient acquisition through proton-coupled transporters, cellular expansion via acid growth-dependent cell wall loosening, stomatal regulation for transpiration control, and stress-responsive signaling during pathogen defence.^2–4

The pleiotropic nature of PM H⁺-ATPase function in plant physiology^5,6 reflects its central role as the dominant source of free energy for PM transport systems. Current evidence suggests that approximately 70–80% of PM transporters depend on ΔμH⁺ as their primary energy source,⁷ necessitating precise regulation of PM H⁺-ATPase activity through multiple coordinated mechanisms. These include post-translational modifications such as phosphorylation/dephosphorylation cycles, modulation of the lipid microenvironment, and dynamic interactions with 14-3-3 regulatory proteins. Such comprehensive regulatory networks enable rapid metabolic adjustments in response to environmental fluctuations while maintaining essential cellular homeostasis under optimal and stress conditions.

Like all integral membrane transporters, the PM H⁺-ATPase interacts with its lipid environment through specific and general mechanisms. Specific interactions occur via direct lipid binding to the protein, while general interactions depend on the bulk physicochemical properties of the lipid bilayer.⁸ Structural studies of P-type ATPases have identified well-defined lipid-binding sites crucial in stabilizing the enzyme and regulating its catalytic activity.^9,10 Fundamental bilayer characteristics govern general lipid-protein interactions, including membrane thickness, fluidity, and dipole potential. Notably, electrostatic interactions between cytoplasmic domains of membrane proteins and anionic phospholipid head groups significantly modulate enzymatic activity.⁸ Furthermore, the catalytic cycle induces conformational changes in the protein’s hydrophobic regions, necessitating dynamic adaptation of the surrounding lipid matrix to prevent hydrophobic mismatch. Sterols emerge as important regulators in this context, constituting up to 30% of total PM lipids.¹¹ These molecules profoundly influence membrane properties by modulating thickness, fluidity, and permeability.^12,13 Sterols preferentially interact with saturated fatty acid chains of phospholipids and sphingolipids, and can affect membrane protein activity through both specific binding sites and indirect modulation of the local lipid environment at the protein-bilayer interface.^14,15

Notably, the original hypothesis of PM horizontal heterogeneity emerged from studies on detergent-resistant membrane fractions (DRMs), initially referred to as “lipid rafts.” Though contemporary high-resolution fluorescence microscopy techniques have demonstrated that none of the classical DRM markers, including enrichment in sterols and sphingolipids, presence of acylated integral membrane proteins, glycosylphosphatidylinositol-anchored proteins, and other characteristic components, are sufficient to establish protein segregation into specific membrane domains in vivo, definitively. While resistance to cold non-ionic detergent extraction remains a useful experimental tool for studying specific membrane properties.^16–20 It is now well-established that these biochemical fractions do not accurately reflect functional membrane domains in living cells.^21,22

Lipid-mediated regulation of H⁺-ATPases is increasingly recognized as a crucial mechanism linking crop stress adaptation (such as drought and salinity) with promising biotechnological applications. Multiple studies have shown that the composition and dynamics of membrane lipids, including anionic phospholipids and sterols, directly modulate the activity of plant PM H⁺-ATPases, promoting efficient proton pumping by forming specific contact sites within the transmembrane domain and enhancing pump activation in response to environmental stimuli.^23–26 Recent advances have revealed that the PATROL1 protein governs the translocation and activation of H⁺-ATPase in both roots and shoots—overexpression of PATROL1 leads to significant increases in root length, lateral root number, shoot dry weight, and shoot nitrogen content under drought conditions, identifying H⁺-ATPase regulation as a promising target for improving crop drought resilience and nutrient uptake.²⁷ Similarly, under salt stress, exogenous melatonin enhances H⁺-ATPase activity and synergistically operates with endogenous hydrogen sulfide to maintain K⁺/Na⁺ homeostasis, enabling better plant stress tolerance by supporting secondary active transport energized by proton gradients.²⁸

Genetic manipulation studies further show that specific transcription factors (e.g., MYB308) induce H⁺-ATPase gene expression, thereby facilitating root iron uptake in iron-limited soils.²⁹ Together, these findings highlight a mechanistic bridge from lipid environment modulation and lipid-microdomain engineering to advanced crop biotechnology. Proteomic assessment of detergent-resistant fractions of plant PMs showed the presence of P-type H⁺-ATPases.^18,21,30,31 Over the past few decades, research has been conducted to understand the molecular mechanisms underlying the regulation of plant cell PM H⁺-ATPase by phosphorylation. Some individual protein kinases/phosphatases involved in this process have also been identified.^32,33 Interestingly, many of these proteins belong to the family of receptor-like kinases, which have also been detected in detergent-resistant fractions of plant PM. Whether this indicates a tendency for H⁺– ATPases to colocalize with their regulatory kinases is a matter of debate.

This review aims to synthesize current knowledge on three critical aspects of plant PM H⁺-ATPase regulation: (i) the structural organization of the pump, (ii) the functional modulation by diverse membrane lipids, and (iii) the potential regulatory influence of PM lateral heterogeneity. While P-type ATPases are ubiquitously distributed across cellular membranes with distinct lipid compositions, a direct comparison of experimental data remains challenging due to system-specific variations in lipid-protein interactions affecting their hydrolytic and transport activities.^10,34 By critically evaluating structural, biochemical, and biophysical evidence, this work seeks to establish unifying principles of lipid-mediated regulation while highlighting plant-specific regulatory mechanisms of this essential proton pump.

Catalytic Cycle, Assembly, and Regulation of Plant PM H⁺-ATPase

Plant genomes encode multiple isoforms of PM H⁺-ATPase, forming a functionally diverse gene family. In Arabidopsis thaliana, the most abundant and well-characterized isoforms are AHA1 and AHA2 (Autoinhibited H⁺-ATPase 1 and 2), which have been extensively studied as model proton pumps in this plant.^35,36

As members of the P3A-type ATPase superfamily, plant PM H⁺-ATPases share a conserved structural architecture with other P-type ATPases.^37,38 The enzyme’s tertiary structure consists of a single polypeptide chain organized into 10 transmembrane α-helices (TM1-TM10) (Figure 1). Structural and functional studies reveal distinct roles for these domains. Catalytic Core (TM1-TM6 helices form the cytoplasmic region responsible for ATP hydrolysis and energy transduction). Structural Stabilization (TM7-TM10 helices maintain functional integrity, analogous to their role in sarco-/endoplasmic reticulum Ca²⁺-ATPase (SERCA)-type Ca²⁺-ATPases).³⁹ Conserved Mechanism critical aspartate residue undergoes phosphorylation during the catalytic cycle, inducing conformational changes that facilitate ion binding site accessibility and proton translocation across the membrane.^7,40 The cytosolic N- and C-terminal domains serve as regulatory modules, containing autoinhibitory regions that control pump activity through intramolecular interactions and post-translational modifications.

Structural insights into the proton transport mechanism of AHA2 in Arabidopsis thaliana. Crystallographic studies of AHA2, the PM H⁺-ATPase in A. thaliana, identified a highly conserved aspartic acid residue (Asp684) in the M6 helix that serves as the primary proton donor/acceptor.⁴¹ The adjacent Asn106 (located in the M2 domain) stabilizes Asp684 and is essential for directional proton transport against the electrochemical gradient.⁴² These residues are functionally conserved across plant and fungal H⁺-ATPases, underscoring their critical role in energy transduction.

The catalytic cycle of plant PM H⁺-ATPase operates through the well-established Albers-Post model, which involves the coordinated action of three cytoplasmic domains: the nucleotide-binding (N), phosphorylation (P), and actuator (A) domains.^41,43,44 Central to this mechanism is the reversible phosphorylation of a conserved aspartate residue (Asp329 in AHA2) within the P-domain’s DKTGT motif, which serves as the molecular switch driving conformational transitions between high-affinity (E1) and low-affinity (E2) states. The cycle initiates when the N-domain (containing the KGA motif) binds and hydrolyses ATP, transferring the γ-phosphate to Asp329 to form the phosphorylated E1P intermediate. This triggers a conformational shift to the E2P state, facilitating the release of protons to the apoplast. Subsequently, the A-domain (featuring the TGE motif) catalyzes aspartate dephosphorylation, returning the pump to the E2 state before spontaneous relaxation to the E1 ground state completes the cycle. The precise spatiotemporal coordination between these domains is critical, as both N- and A-domains must alternately access the same aspartate residue while maintaining strict coupling between ATP hydrolysis and proton transport. This intricate interplay ensures efficient proton translocation against electrochemical gradients while allowing rapid regulation of pump activity in response to cellular demands.

The C- and N-terminal domains are critical regulatory elements of plant PM H⁺-ATPases. The C-terminal domain, comprising approximately 100 amino acid residues, functions as an autoinhibitory domain suppressing pump activity in its non-phosphorylated state. This inhibition occurs through two distinct mechanisms: (i) steric blocking of proton access to transmembrane binding sites, and (ii) restriction of the actuator (A) domain’s mobility.^41,45 Phosphorylation of the penultimate threonine residue within this domain triggers 14-3-3 protein binding, displacing the autoinhibitory region and consequent enzyme activation.^46,47 Recent structural studies have revealed an additional layer of regulation through intermolecular interactions – the C-terminal domain of one AHA2 monomer can engage with amino acid residues in the A-domain of an adjacent monomer.⁴⁸ This finding suggests that PM H⁺-ATPases may function as homooligomeric complexes in native membranes, potentially enabling cooperative regulation of proton transport activity (Figure 2).

The N-terminal domain serves as a critical modulator of 14-3-3 protein binding to the C-terminal autoinhibitory domain. Structural studies have demonstrated that deletion of the first 10 N-terminal residues leads to enhanced phosphorylation of the penultimate threonine and consequent 14-3-3 protein association, suggesting the N-terminus normally exerts a restraining influence on this regulatory interaction.⁴² This finding reveals an intricate interdomain communication network within the H⁺-ATPase molecule. The functional architecture of P-type ATPases reflects a remarkable evolutionary optimization for active transport, featuring precise coupling between transmembrane helices and cytoplasmic domains. This coordinated arrangement enables thermodynamically unfavorable proton translocation against steep electrochemical gradients. Current understanding of PM H⁺-ATPase regulation primarily centers on reversible phosphorylation mechanisms, with extensive characterization of stimulatory phosphorylation at the C-terminal penultimate threonine, inhibitory phosphorylation at other regulatory sites, and coordinated actions of specific protein kinases and phosphatases.³⁷

In Arabidopsis thaliana, the predominant H⁺-ATPase isoforms AHA1 and AHA2 undergo critical phosphorylation at their penultimate threonine residues (Thr948 and Thr947, respectively) within the C-terminal autoinhibitory domain. This post-translational modification creates a binding platform for 14-3-3 regulatory proteins, inducing a conformational change that releases autoinhibition and activates proton transport. The resulting 14-3-3/H⁺-ATPase complex exhibits remarkable stability, becoming essentially irreversible upon binding of the fungal toxin fusicoccin.^46,49

This phosphorylation event serves as a central hub for auxin-mediated signaling. Recent mechanistic studies have demonstrated that the transmembrane kinase TMK1, activated by auxin, directly phosphorylates Thr947 in AHA2, triggering apoplastic acidification through pump activation.^50,51 Conversely, in the absence of auxin, protein phosphatases PP2C-D maintain pump inactivation by dephosphorylating this critical threonine residue. The auxin-responsive SAUR19 proteins counteract this inhibition by suppressing PP2C-D activity, thereby promoting extracellular acidification and subsequent root growth responses.⁵² Notably, auxin signaling also influences the spatial organization of these components within the PM. As demonstrated by Pan,⁵³ auxin induces sterol-dependent nanoclustering of TMK1 kinase, which is essential for establishing polarized membrane domains during auxin-mediated pavement cell morphogenesis. This finding reveals an additional layer of regulation, where membrane microdomain organization modulates the efficiency of H⁺-ATPase activation.

The PM H⁺-ATPase is activated by blue light through a signaling cascade initiated by the photoreceptors phototropic 1 and 2 (phot1 and phot2), which possess serine-threonine kinase activity.^54,55 In guard cells, blue light perception by phototropins triggers phosphorylation of the serine-threonine kinase BLUS1, which subsequently activates the Raf-like kinase MAP3K-BHP. This kinase cascade ultimately phosphorylates Thr881 and Thr947 in AHA2, leading to H⁺-ATPase activation.⁵⁵ The resulting proton extrusion induces membrane hyperpolarization, driving K⁺ uptake into the cytosol and promoting stomatal opening.⁵⁶ Notably, blue light also enhances the interaction between phot1 and AtRem1.3, a marker of sterol-enriched membrane nanodomains.⁵⁷ This suggests that phot1 recruitment to sterol-rich microdomains may facilitate efficient signal transduction during blue light responses.

In contrast to light-mediated activation, abscisic acid (ABA) signaling exerts complex regulation over H⁺-ATPase activity through the PYR/PYL/RCAR receptor-PP2C-SnRK2 signaling module. Under low ABA conditions, the protein phosphatase PP2C-A (ABI1) dephosphorylates Thr947 in AHA2 (and Thr948 in AHA1), maintaining the pump in an inactive state.⁵⁸ Upon ABA binding, PYR/PYL/RCAR receptors inhibit PP2C-A, allowing phosphorylation of the penultimate threonine, 14-3-3 protein binding, and subsequent H⁺-ATPase activation.⁵⁹ Additionally, ABA signaling may involve the SnRK2 kinase, though its direct phosphorylation of H⁺-ATPase in vivo remains unclear.⁶⁰ These regulatory mechanisms highlight the intricate balance between light- and ABA-mediated pathways in modulating H⁺-ATPase activity for stomatal movement and stress adaptation.

The activity of PM H⁺-ATPase is finely tuned through phosphorylation at distinct sites by various protein kinases. The receptor-like kinase PSY1R phosphorylates Thr881 in AHA2 in response to the peptide hormone PSY1, resulting in pump activation that is independent of 14-3-3 protein binding.^61,62 Conversely, phosphorylation at Thr924 or Ser931 by PKS5 inhibits H⁺-ATPase activity and prevents 14-3-3 association, demonstrating the opposing effects of site-specific phosphorylation.^61,62 The FERONIA kinase pathway provides another layer of regulation. Upon interaction with the RALF peptide, FERONIA phosphorylates Ser889 in AHA2, thereby suppressing proton transport and causing extracellular alkalization that inhibits cell growth.⁶³ Notably, FERONIA localizes to DRMs, and its abundance in these domains increases following flg22 treatment, coinciding with AHA2 recruitment.⁶⁴ While this suggests potential co-localization of regulator and target, the functional implications require further investigation.

Recently, QSK1 was identified as a nitrate-responsive kinase that phosphorylates AHA2 at Ser889 under low-nitrate conditions, through a complex with the nitrate transporter NRT1.1, linking nutrient sensing to proton transport regulation.⁶⁵ The diverse phosphorylation-mediated regulation of H⁺-ATPase involves kinases associated with DRMs and nanodomain organizers like flotillins and remorins.^53,57,66 This raises important questions about the lipid microenvironment preferences for H⁺-ATPase and its regulators. In yeast, altered membrane composition (sterols, sphingolipids, anionic phospholipids) affects protein clustering and PMA1 H⁺-ATPase activity.⁶⁷ Similarly, reduced anionic phospholipids and sterols in plants decrease H⁺-ATPase clustering, suggesting both electrostatic and hydrophobic interactions govern its lateral segregation. These findings highlight how membrane lipid composition may spatially organize H⁺-ATPase with its regulatory kinases to facilitate precise, stimulus-responsive control of proton transport.

Phospholipid Composition of Membrane and Its Regulation of P-type ATPase Activity

The functional activity of P-type ATPases is critically dependent on membrane phospholipid composition and the physicochemical properties of the lipid bilayer.¹⁰ Reconstitution studies with plant H⁺-ATPase from mung bean roots revealed that phosphatidylcholine (PC) and phosphatidylserine (PS) stimulate hydrolytic activity, while phosphatidylinositol (PI) exerts an inhibitory effect.⁶⁸ Similar lipid specificity has been observed for animal P-type ATPases, where the Na⁺/K⁺-ATPase shows optimal activity in membranes containing PS and PC with unsaturated fatty acids.¹⁰ Structural analyses have identified specific lipid-binding sites in several P-type ATPases, including SERCA and PM Ca²⁺-ATPase, suggesting a conserved regulatory mechanism across this enzyme family.^10,69

A key breakthrough in understanding lipid regulation came with the discovery that lysophosphatidylcholine (LPC), a signaling lipid produced by phospholipase A2 activity, directly activates plant H⁺-ATPase at micromolar concentrations.⁷⁰ This activation occurs through interactions with the N- and C-terminal regulatory domains and is independent of phosphorylation or 14-3-3 protein binding.⁷¹ The finding that LPC increases the pump’s affinity for ATP suggests it may act as a physiological regulator during stress responses, such as pathogen defence.⁷² The P4-ATPase subfamily, including yeast Drs2p and Arabidopsis ALAs, shares structural homology with other P-type ATPases but transports phospholipids rather than ions. These flippases are activated by specific anionic lipids; for example, Drs2p requires phosphatidylinositol-4-phosphate (PI4P) binding to a cavity formed by transmembrane domains M7, M8, and M10.⁷³ Cryo-EM structures reveal that electrostatic interactions between positively charged residues and PI4P’s phosphate group induce activating conformational changes.⁷⁴ This mechanism likely extends to plant flippases, where similar lipid-binding pockets may regulate activity.

The differential responses of P-type ATPases to membrane composition (e.g., unsaturated PC/PS activation vs. PI inhibition) suggest evolutionary adaptation to distinct membrane environments. This lipid sensitivity enables cells to fine-tune pump activity in response to membrane remodeling during stress or development. The conservation of lipid-binding sites across P-type ATPase families highlights the fundamental importance of lipid-protein interactions in regulating these essential membrane transporters. Biochemical studies of plant PM H⁺-ATPase have demonstrated distinct lipid dependencies in different species. In maize roots, partial solubilization with deoxycholate revealed that PS and phosphatidylglycerol (PG), but not PC, enhanced ATP hydrolysis rates without significantly altering Kₘ values.⁷⁵ Similar experiments with Vigna radiata root membranes showed PC, PS, and PG stimulated activity, while PI, phosphatidylethanolamine (PE), and phosphatidic acid (PA) were inhibitory.⁷⁶ These effects depend critically on lipid acyl chain properties, with unsaturated fatty acids generally promoting higher activity,⁷⁷ mirroring observations in animal Na⁺/K⁺-ATPase.

While reconstitution experiments using detergents, such as dodecyl maltoside, enable controlled studies of lipid-protein interactions,³⁴ they may introduce artifacts by disrupting native membrane organization. Recent work with AHA2-containing proteoliposomes demonstrated PS-specific activation independent of the C-terminal regulatory domain, suggesting the presence of transmembrane lipid-binding sites.⁷⁸ Comparative sequence analysis revealed conserved anionic lipid interaction motifs in P-type ATPases, supporting this hypothesis. Structural modulation of the N-terminus electrostatically interacts with PS-containing membranes, potentially influencing conformational transitions between E1 and E2 states.⁸ Molecular dynamics (MD) simulations confirm that the N-terminus reaches membrane-anionic lipids.⁷⁹ Membrane organization of the kinase FERONIA, which inhibits H⁺-ATPase via Ser899 phosphorylation, also regulates PS distribution and nanodomain formation,⁸⁰ suggesting potential coupling between lipid microenvironment and phosphoregulation. Future directions involve phospholipids creating a permissive environment that facilitates H⁺-ATPase conformational dynamics.^10,78 These findings highlight the need for native membrane studies complementing reconstitution approaches, to understand the lipid-mediated regulation of plant H⁺-ATPases.

Recent cryo-EM studies of AHA2 in proteoliposomes⁷¹ not only confirm PS-dependent stimulation but also reveal oligomeric interfaces stabilized by anionic lipids, mirroring the nanoscale clustering of H⁺-ATPases observed in plant PMs.⁸ These oligomers, likely facilitated by lipid-mediated electrostatic interactions (e.g., between the N-terminus and PS), suggest a dual regulatory mechanism: lipid binding primes individual pumps for activation while promoting higher-order assemblies that could enhance cooperative activity or spatial coordination in membrane microdomains. Notably, this aligns with in planta super-resolution data, which show that H⁺-ATPase nanoclusters are enriched in sterol/sphingolipid rafts,72 where a localized lipid composition may template oligomerization. The convergence of structural⁷¹ and cellular^8,72 evidence implies that lipid-dependent oligomerization is a conserved feature across P-type ATPases. However, its functional consequences (e.g., for signal amplification or lipid homeostasis) remain to be tested in living plants.

Glycerophospholipids Dependent Regulation of H⁺-ATPase Localization

Growing evidence suggests that sphingolipids play crucial roles in P-type ATPase biogenesis and membrane targeting. In yeast, PMA1 H⁺-ATPase requires ceramide association for proper endoplasmic reticulum (ER) exit and PM delivery, with sphingolipid depletion leading to vascular degradation.⁸¹ This dependence extends to functional oligomerization, as hexameric PMA1 complexes incorporate sphingolipids with very-long-chain (C26) fatty acids during Golgi processing before domain formation in the PM.^82,83 Similar mechanisms may operate in plants, where H⁺-ATPases and sphingolipids co-fractionate in detergent-resistant membranes.^17,84 Genetic and pharmacological studies reveal that sphingolipids critically influence H⁺-ATPase activity. Saccharomyces cerevisiae mutants with impaired sphingolipid biosynthesis show reduced proton transport and stress sensitivity. Candida albicans exhibits decreased CaPma1p activity and altered membrane domain association when sphingolipid synthesis is disrupted.⁸⁵Maize embryos treated with the ceramide synthase inhibitor fumonisin B1 exhibit a 20–30% reduction in H⁺-ATPase activity, which is reversible by exogenous ceramides.⁸⁶

These effects may involve both direct lipid-protein interactions and indirect consequences of altered membrane properties. Notably, fumonisin’s structural mimicry of long-chain bases suggests potential competitive binding at regulatory sites.⁸⁶ While yeast studies provide a compelling model for sphingolipid-dependent H⁺-ATPase regulation,⁸³ plant-specific mechanisms remain unclear. Key unknowns include. Whether plant H⁺-ATPases require sphingolipids for ER export and oligomerization. How 14-3-3 protein binding⁸⁷ interfaces with potential sphingolipid interactions. The relative contributions of glycosylceramides vs. free ceramides in modulating activity. Future directions for resolving these questions will require in planta tracking of H⁺-ATPase trafficking in sphingolipid mutants, high-resolution structural studies of lipid-protein interactions and characterization of plant-specific sphingolipid species in membrane microdomains. Current evidence suggests that sphingolipids likely influence plant H⁺-ATPases through both conserved (trafficking, membrane organization) and possibly unique (signaling metabolite) mechanisms compared to fungal systems.

H⁺-ATPase and the Role of Sterols in the PM

Over the past decades, numerous experimental studies have investigated the influence of membrane sterols on the transport and catalytic activities of P-type ATPases.^9,88–91 While these studies have established a correlation between sterol content and enzyme functionality, the findings remain inconsistent, and the underlying molecular mechanisms remain poorly understood.

The variability in experimental outcomes may stem from the differential effects of sterols on specific P-type ATPases. For instance, studies on animal Na+/K+-ATPase reconstituted into liposomes demonstrated that cholesterol depletion significantly reduced hydrolytic activity, with optimal function observed at 20 mol% cholesterol relative to total lipids.^9,92 This enhancement coincided with increased phosphointermediate formation, elevated total protein phosphorylation, and a decreased Kₘ value. Conversely, H⁺/K⁺-ATPase exhibited opposing behavior: elevated cholesterol levels partially inhibited ATP hydrolysis and completely abolished proton transport.⁹¹ Surprisingly, SERCA activity remained unaffected by sterol content in proteoliposomes.⁹³ While some studies suggest cholesterol indirectly modulates catalytic parameters via bilayer physicochemical properties,⁹⁴ SERCA’s insensitivity may reflect its localization in the ER, where sterol concentrations are substantially lower than in the PM.

Early studies on DRMs from yeast cells identified the presence of the yeast PM H⁺-ATPase, PMA1.⁹⁵ Initially, it was hypothesized that disruption of lipid rafts impaired PMA1 association with the PM. However, fluorescence microscopy later revealed that PMA1 forms its distinct clusters termed the membrane compartment of Pma1, which are ergosterol-independent.⁹⁶ Furthermore, Permyakov in 2012 demonstrated that ergosterol levels in Saccharomyces cerevisiae did not affect PMA1 biosynthesis, localization, or glucose-dependent activation.⁹⁷ Notably, yeast cells can adapt their membrane composition in response to sterol biosynthesis perturbations by modulating sphingolipid content. Guan in 2009 proposed that membrane proteins may recognize sterol-sphingolipid complexes, suggesting crosstalk between sterol and sphingolipid biosynthesis pathways.⁹⁸ Consequently, interpreting H+-ATPase activity in mutants with defective lipid biosynthesis remains a challenge.

Similarly, plant PM H⁺-ATPase activity in reconstituted liposomes depends on sterol composition.⁹⁰ Cholesterol and stigmasterol enhance proton transport and pump activity, particularly at low concentrations, whereas campesterol, sitosterol, 24-methylcholesterol, and ergosterol exhibit inhibitory effects across all concentrations.⁹⁹ Intriguingly, sterol content affects proton transport more significantly than ATP hydrolysis.⁹⁰ The authors speculated that the pump discriminates between sterol types, as sitosterol and stigmasterol differ only by a double bond at C-22. These findings imply that “direct, structurally specific lipid-protein interactions” may underlie these effects. The activation of PM H⁺-ATPase by cholesterol and stigmasterol, even at trace concentrations, further supports this hypothesis, as such minimal lipid levels are unlikely to alter bulk membrane physicochemical properties (e.g., fluidity) and thus must act via specific molecular interactions.

In a previous study, Lapshin et al.¹⁰⁰ demonstrated that sterol extraction from PM vesicles of Pisum sativum following MβCD treatment led to an increase in both the maximal rate of ATP hydrolysis (V_max) and active proton transport by H⁺-ATPase.¹⁰⁰ However, the stimulatory effect on H⁺ pumping was transient and did not correlate with the enzyme’s hydrolytic activity. The authors proposed that sterol depletion not only modulates H⁺-ATPase activity but also increases the passive proton permeability of the plasmalemma, possibly due to the differential sensitivity of secondary transporters to membrane sterol content.

Sterols are thought to regulate PM H⁺-ATPase through multiple mechanisms. The most straightforward involves their direct interaction with transmembrane domains of the protein. For instance, crystallographic studies of animal Na⁺/K⁺ ATPase and SERCA have identified specific lipid and sterol-binding sites that enhance the stability of these pumps.^9,10 Experimental evidence suggests that the association of cholesterol with PC at the cytoplasmic interface stabilizes transmembrane domains (TM8–10) and maintains the equilibrium between E1 and E2 conformational states during catalysis.^9,101 Additionally, certain membrane transporters and neurotransmitter receptors possess well-defined sterol-binding motifs (e.g., CRAC/CARC domains.^14,102 While cholesterol recognition sequences (CARC domains) have been identified in animal PM Ca²⁺-ATPase, they are absent in SERCA.¹⁰³ Given the structural conservation among P-type ATPases, plant PM H⁺-ATPase may similarly harbor sterol/lipid-binding sites that modulate its activity.

Sterols may also influence protein function indirectly by altering the physicochemical properties of the lipid bilayer, thereby affecting conformational dynamics.¹⁰⁴ For example, cholesterol-induced changes in bilayer properties can regulate protein partitioning between sterol-rich and sterol-poor membrane domains.¹⁰⁵ Notably, proteins with short transmembrane α-helices are energetically disfavored in cholesterol-rich regions. Furthermore, the impact of cholesterol on H⁺ permeability in artificial membranes varies with the phospholipid composition.¹⁰⁶ Cholesterol-mediated modulation of bilayer thickness has been implicated in Na⁺/K⁺-ATPase activity.¹⁰⁷ As highlighted by Clarke,⁸ membrane proteins dynamically adjust their lipid environment during catalytic cycles to mitigate hydrophobic mismatch, with sterols playing a key role in maintaining bilayer thickness.^8,12 Intriguingly, SERCA adopts a distinct mechanism: rather than inducing membrane deformation, its transmembrane helices tilt to accommodate the surrounding bilayer.³⁹

Future Direction and Conclusion

The influence of the lipid environment on the functional regulation of P-type ATPases is a complex and multifaceted process involving diverse molecular and cellular mechanisms. Although experimental studies on lipid-mediated modulation of ATPase activity began over three decades ago^68,88,93 and high-resolution structures of major P-type ATPases have since been elucidated,^43,108 key questions remain regarding the precise molecular mechanisms governing protein-lipid interactions within their native membrane environment. The plant PM exhibits a unique lipid composition, enriched in phytosterols and sphingolipids.^109,110 Significant advances have been made in understanding the role of sphingolipids in the yeast PMA1 proton pump, where they are critical for ER biosynthesis, oligomerization, and subsequent trafficking to the PM.^82,83 While the involvement of sphingolipids in the oligomerization and vesicular transport of plant H⁺-ATPases remains unexplored, their functional contribution to these processes is highly plausible.

Sterols appear to regulate PM H⁺-ATPase activity through both specific lipid-protein interactions and general membrane effects. However, whether PM H⁺-ATPases possess dedicated lipid-binding sites⁹ or conserved motifs such as CARC/CRAC domains¹⁴ remains unresolved. MD simulations could provide valuable insights into these mechanisms. A particularly intriguing avenue for future research is the potential colocalization of PM H⁺-ATPases with their regulatory proteins, as well as the role of membrane lipids in facilitating these interactions. The detection of PM H⁺-ATPases and their binding partners in DRMs enriched in sterols and sphingolipids has fueled speculation about their localization in membrane microdomains.

Lipid-mediated regulation of H⁺-ATPase activity forms a mechanistic bridge between membrane dynamics and plant adaptation to abiotic stress. These insights are fueling biotechnological innovations such as the design of engineered lipid microdomains to enhance pump performance, nutrient uptake, and crop sustainability under challenging environments.^27,111,112 However, detergent-based fractionation is now recognized as an outdated and unreliable method for assessing protein lateral organization in membranes. Instead, emerging evidence suggests that direct phosphorylation or dephosphorylation of PM H⁺-ATPases by partner proteins implies close physical proximity, possibly within functionally active membrane complexes. A critical unresolved question is the subcellular site of complex formation: (i) during pump biosynthesis in the ER, (ii) during transit through the trans-Golgi network, or (iii) at the PM following ligand-induced recruitment of receptor-like kinases/phosphatases. Addressing this will require advanced high-resolution imaging techniques, such as super-resolution fluorescence microscopy or cryo-electron tomography.

Acknowledgements

The author thanks Dr. Marek Seidenglanz (Agritec Plant Research S.R.O.) for consultations and support during the writing of this manuscript. Supported by the Ministry of Agriculture of the Czech Republic, institutional support MZE-RO1023.

References

1. Palmgren MG. Plant plasma membrane H⁺-ATPases: powerhouses for nutrient uptake. Annu Rev Plant Physiol Plant Mol Biol. 2001;52(1):817–45. https://doi.org/10.1146/annurev.arplant.52.1.817
2. Elmore JM, Coaker G. The role of the plasma membrane H⁺-ATPase in plant–microbe interactions. Mol Plant. 2011;4(3):416–27. https://doi.org/10.1093/mp/ssq083
3. Minami A, Takahashi K, Inoue S ichiro, Tada Y, Kinoshita T. Brassinosteroid induces phosphorylation of the plasma membrane H⁺-ATPase during hypocotyl elongation in Arabidopsis thaliana. Plant Cell Physiol. 2019;60(5):935–44. https://doi.org/10.1093/pcp/pcz005
4. Haruta M, Sussman MR. The effect of a genetically reduced plasma membrane protonmotive force on vegetative growth of Arabidopsis. Plant Physiol. 2012;158(3):1158–71. https://doi.org/10.1104/pp.111.189167
5. Morsomme P, Boutry M. The plant plasma membrane H⁺-ATPase: structure, function and regulation. Biochim Biophys Acta BBA Biomembr. 2000;1465(1–2):1–16. https://doi.org/10.1016/S0005-2736(00)00128-0
6. Sondergaard TE, Schulz A, Palmgren MG. Energization of transport processes in plants. Roles of the plasma membrane H⁺-ATPase. Plant Physiol. 2004;136(1):2475–82. https://doi.org/10.1104/pp.104.048231
7. Palmgren MG, Nissen P. P-type ATPases. Annu Rev Biophys. 2011;40(1):243–66. https://doi.org/10.1146/annurev.biophys.093008.131331
8. Clarke RJ. Electrostatic switch mechanisms of membrane protein trafficking and regulation. Biophys Rev. 2023;15(6):1967–85. https://doi.org/10.1007/s12551-023-01166-2
9. Cornelius F, Habeck M, Kanai R, Toyoshima C, Karlish SJD. General and specific lipid–protein interactions in Na, K-ATPase. Biochim Biophys Acta BBA Biomembr. 2015;1848(9):1729–43. https://doi.org/10.1016/j.bbamem.2015.03.012
10. Hossain KR, Clarke RJ. General and specific interactions of the phospholipid bilayer with P-type ATPases. Biophys Rev. 2019;11(3):353–64. https://doi.org/10.1007/s12551-019-00533-2
11. Cassim AM, Gouguet P, Gronnier J, Laurent N, Germain V, Grison M, et al. Plant lipids: key players of plasma membrane organization and function. Prog Lipid Res. 2019;73:1–27. https://doi.org/10.1016/j.plipres.2018.11.002
12. Dufourc EJ. Sterols and membrane dynamics. J Chem Biol. 2008;1(1–4):63–77. https://doi.org/10.1007/s12154-008-0010-6
13. Van Meer G, Voelker DR, Feigenson GW. Membrane lipids: where they are and how they behave. Nat Rev Mol Cell Biol. 2008;9(2):112–24. https://doi.org/10.1038/nrm2330
14. Fantini J, Di Scala C, Baier CJ, Barrantes FJ. Molecular mechanisms of protein-cholesterol interactions in plasma membranes: functional distinction between topological (tilted) and consensus (CARC/CRAC) domains. Chem Phys Lipids. 2016;199:52–60. https://doi.org/10.1016/j.chemphyslip.2016.02.009
15. Levitan I, Singh DK, Rosenhouse-Dantsker A. Cholesterol binding to ion channels. Front Physiol. 2014;5:65. https://doi.org/10.3389/fphys.2014.00065
16. Borner GH, Sherrier DJ, Weimar T, Michaelson LV, Hawkins ND, MacAskill A, et al. Analysis of detergent-resistant membranes in Arabidopsis. evidence for plasma membrane lipid rafts. Plant Physiol. 2005;137(1):104–16. https://doi.org/10.1104/pp.104.053041
17. Mongrand S, Morel J, Laroche J, Claverol S, Carde JP, Hartmann MA, et al. Lipid rafts in higher plant cells: purification and characterization of Triton X-100-insoluble microdomains from tobacco plasma membrane. J Biol Chem. 2004;279(35):36277–86. https://doi.org/10.1074/jbc.M403440200
18. Morel J, Claverol S, Mongrand S, Furt F, Fromentin J, Bessoule JJ, et al. Proteomics of plant detergent-resistant membranes. Mol Cell Proteomics. 2006;5(8):1396–411. https://doi.org/10.1074/mcp.M600044-MCP200
19. Peskan T, Westermann M, Oelmüller R. Identification of low-density Triton X-100-insoluble plasma membrane microdomains in higher plants. Eur J Biochem. 2000;267(24):6989–95. https://doi.org/10.1046/j.1432-1327.2000.01776.x
20. Takahashi D, Kawamura Y, Uemura M. Detergent-resistant plasma membrane proteome to elucidate microdomain functions in plant cells. Front Plant Sci. 2013;4:27. https://doi.org/10.3389/fpls.2013.00027
21. Jaillais Y, Bayer E, Bergmann DC, Botella MA, Boutté Y, Bozkurt TO, et al. Guidelines for naming and studying plasma membrane domains in plants. Nat Plants. 2024;10(8):1172–83. https://doi.org/10.1038/s41477-024-01742-8
22. Tanner W, Malinsky J, Opekarová M. In plant and animal cells, detergent-resistant membranes do not define functional membrane rafts. Plant Cell. 2011;23(4):1191–3. https://doi.org/10.1105/tpc.111.086249
23. Grandmougin-Ferjani A, Schuler-Muller I, Hartmann MA. Sterol modulation of the plasma membrane H⁺-ATPase activity from corn roots reconstituted into soybean lipids. Plant Physiol. 1997;113(1):163–74. https://doi.org/10.1104/pp.113.1.163
24. Anionic phospholipids stimulate the proton pumping activity of the plant plasma membrane P-type H⁺-ATPase; 2025. https://doi.org/10.3390/ijms241713106
25. Wielandt AG, Pedersen JT, Falhof J, Kemmer GC, Lund A, Ekberg K, et al. Specific activation of the plant P-type plasma membrane H⁺-ATPase by lysophospholipids depends on the autoinhibitory N- and C-terminal domains. J Biol Chem. 2015;290(26):16281–91. https://doi.org/10.1074/jbc.M114.617746
26. Kasamo K. Mechanism for the activation of plasma membrane H⁺-ATPase from rice (Oryza sativa L.) culture cells by molecular species of a phospholipid 1. Plant Physiol. 1990;93(3):1049–52. https://doi.org/10.1104/pp.93.3.1049
27. Katsuhama N, Sakoda K, Kimura H, Shimizu Y, Sakai Y, Nagata K, et al. Proton ATPase translocation control 1-mediated H⁺-ATPase translocation boosts plant growth under drought by optimizing root and leaf functions. PNAS Nexus. 2025;4(5):pgaf151. https://doi.org/10.1093/pnasnexus/pgaf151
28. Exogenous melatonin-mediated regulation of K⁺/Na⁺ transport, H⁺-ATPase activity and enzymatic antioxidative defence operate through endogenous hydrogen sulphide signalling in NaCl-stressed tomato seedling roots. Plant Biol. 2021;23(5):797–805. https://doi.org/10.1111/plb.13296
29. Fan Z, Wu Y, Zhao L, Fu L, Deng L, Deng J, et al. MYB308-mediated transcriptional activation of plasma membrane H⁺-ATPase 6 promotes iron uptake in citrus. Hortic Res. 2022;9:uhac088. https://doi.org/10.1093/hr/uhac088
30. Furt F, Lefebvre B, Cullimore J, Bessoule JJ, Mongrand S. Plant lipid rafts, fluctuat nec mergitur. Plant Signal Behav. 2007;2(6):508–11. https://doi.org/10.4161/psb.2.6.4636
31. Sandstrom RP, Cleland RE. Selective delipidation of the plasma membrane by surfactants: enrichment of sterols and activation of ATPase. Plant Physiol. 1989;90(4):1524–31. https://doi.org/10.1104/pp.90.4.1524
32. Falhof J, Pedersen JT, Fuglsang AT, Palmgren M. Plasma membrane H⁺-ATPase regulation in the center of plant physiology. Mol Plant. 2016;9(3):323–37. https://doi.org/10.1016/j.molp.2015.11.002
33. Haruta M, Gray WM, Sussman MR. Regulation of the plasma membrane proton pump (H⁺-ATPase) by phosphorylation. Curr Opin Plant Biol. 2015;28:68–75. https://doi.org/10.1016/j.pbi.2015.09.005
34. Morales-Cedillo F, González-Solís A, Gutiérrez-Angoa L, Cano-Ramírez DL, Gavilanes-Ruiz M. Plant lipid environment and membrane enzymes: the case of the plasma membrane H⁺-ATPase. Plant Cell Rep. 2015;34(4):617–29. https://doi.org/10.1007/s00299-014-1735-z
35. Axelsen KB, Palmgren MG. Inventory of the superfamily of P-type ion pumps in Arabidopsis. Plant Physiol. 2001;126(2):696–706. https://doi.org/10.1104/pp.126.2.696
36. Palmgren MG, Axelsen KB. Evolution of P-type ATPases. Biochim Biophys Acta BBA Bioenerg. 1998;1365(1–2):37–45. https://doi.org/10.1016/S0005-2728(98)00041-3
37. Palmgren M, Morsomme P. The plasma membrane H⁺-ATPase, a simple polypeptide with a long history. Yeast. 2019;36(4):201–10. https://doi.org/10.1002/yea.3365
38. Stéger A, Hayashi M, Lauritzen EW, Herburger K, Shabala L, Wang C, et al. The evolution of plant proton pump regulation via the R domain may have facilitated plant terrestrialization. Commun Biol. 2022;5(1):1312. https://doi.org/10.1038/s42003-022-04291-y
39. Norimatsu Y, Hasegawa K, Shimizu N, Toyoshima C. Protein–phospholipid interplay revealed with crystals of a calcium pump. Nature. 2017;545(7653):193–8. https://doi.org/10.1038/nature22357
40. Palmgren M. P-type ATPases: many more enigmas left to solve. J Biol Chem. 2023;299(11):105352. https://doi.org/10.1016/j.jbc.2023.105352
41. Pedersen BP, Buch-Pedersen MJ, Preben Morth J, Palmgren MG, Nissen P. Crystal structure of the plasma membrane proton pump. Nature. 2007;450(7172):1111–4. https://doi.org/10.1038/nature06417
42. Ekberg K, Wielandt AG, Buch-Pedersen MJ, Palmgren MG. A conserved asparagine in a P-type proton pump is required for efficient gating of protons. J Biol Chem. 2013;288(14):9610–8. https://doi.org/10.1074/jbc.M112.417345
43. Focht D, Croll TI, Pedersen BP, Nissen P. Improved model of proton pump crystal structure obtained by interactive molecular dynamics flexible fitting expands the mechanistic model for proton translocation in P-type ATPases. Front Physiol. 2017;8:202. https://doi.org/10.3389/fphys.2017.00202
44. Pardo JM, Serrano R. Structure of a plasma membrane H⁺-ATPase gene from the plant Arabidopsis thaliana. J Biol Chem. 1989;264(15):8557–62. https://doi.org/10.1016/S0021-9258(18)81827-0
45. Palmgren MG, Sommarin M, Serrano R, Larsson C. Identification of an autoinhibitory domain in the C-terminal region of the plant plasma membrane H (⁺)-ATPase. J Biol Chem. 1991;266(30):20470–5. https://doi.org/10.1016/S0021-9258(18)54948-6
46. Fuglsang AT, Borch J, Bych K, Jahn TP, Roepstorff P, Palmgren MG. The binding site for regulatory 14-3-3 protein in plant plasma membrane H⁺-ATPase: involvement of a region promoting phosphorylation-independent interaction in addition to the phosphorylation-dependent C-terminal end. J Biol Chem. 2003;278(43):42266–72. https://doi.org/10.1074/jbc.M306707200
47. Jahn T, Fuglsang AT, Olsson A, Brüntrup IM, Collinge DB, Volkmann D, et al. The 14-3-3 protein interacts directly with the C-terminal region of the plant plasma membrane H⁺-ATPase. Plant Cell. 1997;9(10):1805–14. https://doi.org/10.1105/tpc.9.10.1805
48. Nguyen TT, Sabat G, Sussman MR. In vivo cross-linking supports a head-to-tail mechanism for regulation of the plant plasma membrane P-type H⁺-ATPase. J Biol Chem. 2018;293(44):17095–106. https://doi.org/10.1074/jbc.RA118.003528
49. Olsson A, Svennelid F, Ek B, Sommarin M, Larsson C. A phosphothreonine residue at the C-terminal end of the plasma membrane H⁺-ATPase is protected by fusicoccin-induced 14-3-3 binding. Plant Physiol. 1998;118(2):551–5. https://doi.org/10.1104/pp.118.2.551
50. Li XW, Lu XX, Zhang ZJ, Huang J, Zhang JM, Wang LK, et al. Intercropping rosemary (Rosmarinus officinalis) with sweet pepper (Capsicum annum) reduces major pest population densities without impacting natural enemy populations. Insects. 2021;12(1):74. https://doi.org/10.3390/insects12010074
51. Lin W, Zhou X, Tang W, Takahashi K, Pan X, Dai J, et al. TMK-based cell-surface auxin signalling activates cell-wall acidification. Nature. 2021;599(7884):278–82. https://doi.org/10.1038/s41586-021-03976-4
52. Spartz AK, Ren H, Park MY, Grandt KN, Lee SH, Murphy AS, et al. SAUR inhibition of PP2C-D phosphatases activates plasma membrane H⁺-ATPases to promote cell expansion in Arabidopsis. Plant Cell. 2014;26(5):2129–42. https://doi.org/10.1105/tpc.114.126037
53. Pan X, Fang L, Liu J, Senay-Aras B, Lin W, Zheng S, et al. Auxin-induced signaling protein nanoclustering contributes to cell polarity formation. Nat Commun. 2020;11(1):3914. https://doi.org/10.1038/s41467-020-17602-w
54. Takemiya A, Yamauchi S, Yano T, Ariyoshi C, Shimazaki KI. Identification of a regulatory subunit of protein phosphatase 1 which mediates blue light signaling for stomatal opening. Plant Cell Physiol. 2013;54(1):24–35. https://doi.org/10.1093/pcp/pcs073
55. Hayashi M, Inoue SI, Ueno Y, Kinoshita T. A raf-like protein kinase BHP mediates blue light-dependent stomatal opening. Sci Rep. 2017;7(1):45586. https://doi.org/10.1038/srep45586
56. Fuji S, Yamauchi S, Sugiyama N, Kohchi T, Nishihama R, Shimazaki KI, et al. Light-induced stomatal opening requires phosphorylation of the C-terminal autoinhibitory domain of plasma membrane H⁺-ATPase. Nat Commun. 2024;15(1):1195. https://doi.org/10.1038/s41467-024-45236-9
57. Xue Y, Xing J, Wan Y, Lv X, Fan L, Zhang Y, et al. Arabidopsis blue light receptor phototropin 1 undergoes blue light-induced activation in membrane microdomains. Mol Plant. 2018;11(6):846–59. https://doi.org/10.1016/j.molp.2018.04.003
58. Miao R, Yuan W, Wang Y, Garcia-Maquilon I, Dang X, Li Y, et al. Low ABA concentration promotes root growth and hydrotropism through relief of ABA INSENSITIVE 1-mediated inhibition of plasma membrane H⁺-ATPase 2. Sci Adv. 2021;7(12):eabd4113. https://doi.org/10.1126/sciadv.abd4113
59. Miao R, Russinova E, Rodriguez PL. Tripartite hormonal regulation of plasma membrane H⁺-ATPase activity. Trends Plant Sci. 2022;27(6):588–600. https://doi.org/10.1016/j.tplants.2021.12.011
60. Planes MD, Niñoles R, Rubio L, Bissoli G, Bueso E, García-Sánchez MJ, et al. A mechanism of growth inhibition by abscisic acid in germinating seeds of Arabidopsis thaliana based on inhibition of plasma membrane H⁺-ATPase and decreased cytosolic pH, K⁺, and anions. J Exp Bot. 2015;66(3):813–25. https://doi.org/10.1093/jxb/eru442
61. Fuglsang AT, Guo Y, Cuin TA, Qiu Q, Song C, Kristiansen KA, et al. Arabidopsis protein kinase PKS5 inhibits the plasma membrane H⁺-ATPase by preventing interaction with 14-3-3 protein. Plant Cell. 2007;19(5):1617–34. https://doi.org/10.1105/tpc.105.035626
62. Fuglsang AT, Kristensen A, Cuin TA, Schulze WX, Persson J, Thuesen KH, et al. Receptor kinase-mediated control of primary active proton pumping at the plasma membrane. Plant J. 2014;80(6):951–64. https://doi.org/10.1111/tpj.12680
63. Haruta M, Sabat G, Stecker K, Minkoff BB, Sussman MR. A peptide hormone and its receptor protein kinase regulate plant cell expansion. Science. 2014;343(6169):408–11. https://doi.org/10.1126/science.1244454
64. Keinath NF, Kierszniowska S, Lorek J, Bourdais G, Kessler SA, Shimosato-Asano H, et al. PAMP (pathogen-associated molecular pattern)-induced changes in plasma membrane compartmentalization reveal novel components of plant immunity. J Biol Chem. 2010;285(50):39140–9. https://doi.org/10.1074/jbc.M110.160531
65. Zhu Z, Krall L, Li Z, Xi L, Luo H, Li S, et al. Transceptor NRT1. 1 and receptor-kinase QSK1 complex controls PM H⁺-ATPase activity under low nitrate. Curr Biol. 2024;34(7):1479–91. https://doi.org/10.1016/j.cub.2024.02.066
66. Wang Z, Zhao X, Wu P, He J, Chen X, Gao Y, et al. Radiation interception and utilization by wheat/maize strip intercropping systems. Agric For Meteorol. 2015;204:58–66. https://doi.org/10.1016/j.agrformet.2015.02.004
67. Spira F, Mueller NS, Beck G, Von Olshausen P, Beig J, Wedlich-Söldner R. Patchwork organization of the yeast plasma membrane into numerous coexisting domains. Nat Cell Biol. 2012;14(6):640–8. https://doi.org/10.1038/ncb2487
68. Kasamo K, Yamanishi H. Functional reconstitution of plasma membrane H⁺-ATPase from mung bean (Vigna radiata L.) hypocotyls in liposomes prepared with various molecular species of phospholipids. Plant Cell Physiol. 1991;32(8):1219–25. https://doi.org/10.1093/oxfordjournals.pcp.a078127
69. Meneghelli S, Fusca T, Luoni L, De Michelis MI. Dual mechanism of activation of plant plasma membrane Ca²⁺-ATPase by acidic phospholipids: evidence for a phospholipid binding site which overlaps the calmodulin-binding site. Mol Membr Biol. 2008;25(6–7):539–46. https://doi.org/10.1080/09687680802508747
70. Palmgren MG, Sommarin M. Lysophosphatidylcholine stimulates ATP dependent proton accumulation in isolated oat root plasma membrane vesicles. Plant Physiol. 1989;90(3):1009–14. https://doi.org/10.1104/pp.90.3.1009
71. Wielandt AG, Pedersen JT, Falhof J, Kemmer GC, Lund A, Ekberg K, et al. Specific activation of the plant P-type plasma membrane H⁺-ATPase by lysophospholipids depends on the autoinhibitory N-and C-terminal domains. J Biol Chem. 2015;290(26):16281–91. https://doi.org/10.1074/jbc.M114.617746
72. Wi SJ, yeon Seo S, Cho K, Nam MH, Park KY. Lysophosphatidylcholine enhances susceptibility in signaling pathway against pathogen infection through biphasic production of reactive oxygen species and ethylene in tobacco plants. Phytochemistry. 2014;104:48–59. https://doi.org/10.1016/j.phytochem.2014.04.009
73. Bai L, Kovach A, You Q, Hsu HC, Zhao G, Li H. Autoinhibition and activation mechanisms of the eukaryotic lipid flippase Drs2p-Cdc50p. Nat Commun. 2019;10(1):4142. https://doi.org/10.1038/s41467-019-12191-9
74. Villagrana R, López-Marqués RL. Plant P4-ATPase lipid flippases: how are they regulated? Biochim Biophys Acta BBA Mol Cell Res. 2024;1871(1):119599. https://doi.org/10.1016/j.bba mcr.2023.119599
75. Brauer D, Tu SI. Phospholipid requirement of the vanadate-sensitive ATPase from maize roots evaluated by two methods. Plant Physiol. 1989;89(3):867–74. https://doi.org/10.1104/pp.89.3.867
76. Kasamo K, Nouchi I. The role of phospholipids in plasma membrane ATPase activity in Vigna radiata L. (Mung Bean) roots and hypocotyls 1. Plant Physiol. 1987;83(2):323–8. https://doi.org/10.1104/pp.83.2.323
77. Kasamo K. Mechanism for the activation of plasma membrane H⁺-ATPase from rice (Oryza sativa L.) culture cells by molecular species of a phospholipid. Plant Physiol. 1990;93(3):1049–52. https://doi.org/10.1104/pp.93.3.1049
78. Paweletz LC, Holtbrügge SL, Löb M, De Vecchis D, Schäfer LV, Günther Pomorski T, et al. Anionic phospholipids stimulate the proton pumping activity of the plant plasma membrane P-type H⁺-ATPase. Int J Mol Sci. 2023;24(17):13106. https://doi.org/10.3390/ijms241713106
79. Polarity of the ATP binding site of the Na⁺, K⁺-ATPase, gastric H⁺, K⁺-ATPase and sarcoplasmic reticulum Ca2⁺-ATPase. Biochim Biophys Acta BBA Biomembr. 2020;1862(2):183138. https://doi.org/10.1016/j.bbamem.2019.183138
80. Smokvarska M, Bayle V, Maneta-Peyret L, Fouillen L, Poitout A, Dongois A, et al. The receptor kinase FERONIA regulates phosphatidylserine localization at the cell surface to modulate ROP signaling. Sci Adv. 2023;9(14):eadd4791. https://doi.org/10.1126/sciadv.add4791
81. Lee MC, Hamamoto S, Schekman R. Ceramide biosynthesis is required for the formation of the oligomeric H⁺-ATPase Pma1p in the yeast endoplasmic reticulum. J Biol Chem. 2002;277(25):22395–401. https://doi.org/10.1074/jbc.M200450200
82. Zhao P, Zhao C, Chen D, Yun C, Li H, Bai L. Structure and activation mechanism of the hexameric plasma membrane H⁺-ATPase. Nat Commun. 2021;12(1):6439. https://doi.org/10.1038/s41467-021-26782-y
83. Young MR, Heit S, Bublitz M. Structure, function and biogenesis of the fungal proton pump Pma1. Biochim Biophys Acta BBA Mol Cell Res. 2024;1871(1):119600. https://doi.org/10.1016/j.bbamcr.2023.119600
84. Minami A, Fujiwara M, Furuto A, Fukao Y, Yamashita T, Kamo M, et al. Alterations in detergent-resistant plasma membrane microdomains in Arabidopsis thaliana during cold acclimation. Plant Cell Physiol. 2009;50(2):341–59. https://doi.org/10.1093/pcp/pcn202
85. Urbanek AK, Muraszko J, Derkacz D, Łukaszewicz M, Bernat P, Krasowska A. The role of ergosterol and sphingolipids in the localization and activity of Candida albicans’ multidrug transporter Cdr1p and plasma membrane ATPase Pma1p. Int J Mol Sci. 2022;23(17):9975. https://doi.org/10.3390/ijms23179975
86. Gutiérrez-Nájera NA, Saucedo-García M, Noyola-Martínez L, Vázquez-Vázquez C, Palacios-Bahena S, Carmona-Salazar L, et al. Sphingolipid effects on the plasma membrane produced by addition of fumonisin B1 to maize embryos. Plants. 2020;9(2):150. https://doi.org/10.3390/plants9020150
87. Kanczewska J, Marco S, Vandermeeren C, Maudoux O, Rigaud JL, Boutry M. Activation of the plant plasma membrane H⁺-ATPase by phosphorylation and binding of 14-3-3 proteins converts a dimer into a hexamer. Proc Natl Acad Sci. 2005;102(33):11675–80. https://doi.org/10.1073/pnas.0504498102
88. Cornelius F. Cholesterol modulation of molecular activity of reconstituted shark Na⁺, K⁺-ATPase. Biochim Biophys Acta BBA Biomembr. 1995;1235(2):205–12. https://doi.org/10.1016/0005-2736(95)80006-2
89. Garcia A, Lev B, Hossain KR, Gorman A, Pham THN, Cornelius F, et al. Cholesterol depletion inhibits Na⁺, K⁺-ATPase activity in a near-native membrane environment. J Biol Chem. 2019;294(15):5956–69. https://doi.org/10.1074/jbc.RA118.006223
90. Grandmougin-Ferjani A, Schuler-Muller I, Hartmann MA. Sterol modulation of the plasma membrane H⁺-ATPase activity from corn roots reconstituted into soybean lipids. Plant Physiol. 1997;113(1):163–74. https://doi.org/10.1104/pp.113.1.163
91. Ray TK, Nandi J, Dannemann A, Gordon GB. Role of cholesterol in the structure and function of gastric microsomal vesicles. J Cell Biochem. 1983;21(2):141–50. https://doi.org/10.1002/jcb.240210205
92. Habeck M, Haviv H, Katz A, Kapri-Pardes E, Ayciriex S, Shevchenko A, et al. Stimulation, inhibition, or stabilization of Na, K-ATPase caused by specific lipid interactions at distinct sites. J Biol Chem. 2015;290(8):4829–42. https://doi.org/10.1074/jbc.M114.611384
93. Cheng KH, Lepock JR, Hui SW, Yeagle PL. The role of cholesterol in the activity of reconstituted Ca-ATPase vesicles containing unsaturated phosphatidylethanolamine. J Biol Chem. 1986;261(11):5081–7. https://doi.org/10.1016/S0021-9258(19)89217-7
94. Drachmann ND, Olesen C, Møller JV, Guo Z, Nissen P, Bublitz M. Comparing crystal structures of Ca²⁺ – ATP ase in the presence of different lipids. FEBS J. 2014;281(18):4249–62. https://doi.org/10.1111/febs.12957
95. Bagnat M, Chang A, Simons K. Plasma membrane proton ATPase Pma1p requires Raft association for surface delivery in yeast. Mol Biol Cell. 2001;12(12):4129–38. https://doi.org/10.1091/mbc.12.12.4129
96. Malinsky J, Opekarová M, Grossmann G, Tanner W. Membrane microdomains, rafts, and detergent-resistant membranes in plants and fungi. Annu Rev Plant Biol. 2013;64(1):501–29. https://doi.org/10.1146/annurev-arplant-050312-120103
97. Permyakov S, Suzina N, Valiakhmetov A. Activation of H⁺-ATPase of the plasma membrane of Saccharomyces cerevisiae by glucose: the role of sphingolipid and lateral enzyme mobility. PLoS One. 2012;7(2):e30966. https://doi.org/10.1371/journal.pone.0030966
98. Guan XL, Souza CM, Pichler H, Dewhurst G, Schaad O, Kajiwara K, et al. Functional interactions between sphingolipids and sterols in biological membranes regulating cell physiology. Mol Biol Cell. 2009;20(7):2083–95. https://doi.org/10.1091/mbc.e08-11-1126
99. Rossard S, Roblin G, Atanassova R. Ergosterol triggers characteristic elicitation steps in Beta vulgaris leaf tissues. J Exp Bot. 2010;61(6):1807–16. https://doi.org/10.1093/jxb/erq047
100. Lapshin NK, Piotrovskii MS, Trofimova MS. Sterol extraction from isolated plant plasma membrane vesicles affects H⁺-ATPase activity and H⁺-transport. Biomolecules. 2021;11(12):1891. https://doi.org/10.3390/biom11121891
101. Habeck M, Kapri-Pardes E, Sharon M, Karlish SJD. Specific phospholipid binding to Na, K-ATPase at two distinct sites. Proc Natl Acad Sci. 2017;114(11):2904–9. https://doi.org/10.1073/pnas.1620799114
102. Tan S, Zhang P, Xiao W, Feng B, Chen L, Li S, et al. TMD1 domain and CRAC motif determine the association and disassociation of MxIRT1 with detergent-resistant membranes. Traffic. 2018;19(2):122–37. https://doi.org/10.1111/tra.12540
103. Delgado-Coello B, Luna-Reyes I, Méndez-Acevedo KM, Bravo-Martínez J, Montalvan-Sorrosa D, Mas-Oliva J. Analysis of cholesterol-recognition motifs of the plasma membrane Ca²⁺-ATPase. J Bioenerg Biomembr. 2024;56(3):205–19. https://doi.org/10.1007/s10863-024-10010-5
104. Grosjean K, Mongrand S, Beney L, Simon-Plas F, Gerbeau-Pissot P. Differential effect of plant lipids on membrane organization: specificities of phytosphingolipids and phytosterols. J Biol Chem. 2015;290(9):5810–25. https://doi.org/10.1074/jbc.M114.598805
105. Lundbaek JA, Andersen OS, Werge T, Nielsen C. Cholesterol-induced protein sorting: an analysis of energetic feasibility. Biophys J. 2003;84(3):2080–9. https://doi.org/10.1016/S0006-3495(03)75015-2
106. Gensure RH, Zeidel ML, Hill WG. Lipid raft components cholesterol and sphingomyelin increase H⁺/OH− permeability of phosphatidylcholine membranes. Biochem J. 2006;398(3):485–95. https://doi.org/10.1042/BJ20051620
107. Cornelius F. Cholesterol-Dependent interaction of polyunsaturated phospholipids with Na, K-ATPase. Biochemistry. 2008;47(6):1652–8. https://doi.org/10.1021/bi702128x
108. Bublitz M, Poulsen H, Morth JP, Nissen P. In and out of the cation pumps: P-type ATPase structure revisited. Curr Opin Struct Biol. 2010;20(4):431–9. https://doi.org/10.1016/j.sbi.2010.06.007
109. Fougère L, Mongrand S, Boutté Y. The function of sphingolipids in membrane trafficking and cell signaling in plants, in comparison with yeast and animal cells. Biochim Biophys Acta BBA Mol Cell Biol Lipids. 2024;1869(3):159463. https://doi.org/10.1016/j.bbalip.2024.159463
110. Rogowska A, Szakiel A. The role of sterols in plant response to abiotic stress. Phytochem Rev. 2020;19(6):1525–38. https://doi.org/10.1007/s11101-020-09708-2
111. López-Marqués RL. Mini-review: Lipid flippases as putative targets for biotechnological crop improvement. Front Plant Sci. 2023;14:1107142. https://doi.org/10.3389/fpls.2023.1107142
112. McMillan HM. Lipid droplets: new roles as mediators of biotic and abiotic stress. Plant Physiol. 2024;197(1):kiae340. https://doi.org/10.1093/plphys/kiae340

Cite this article as:
Kumar R. Lipid-Protein Interplay: How the Plant Plasma Membrane H-ATPase Dynamically Interacts with Its Lipid Environment: A Review. Premier Journal of Plant Biology 2025;4:100021

Export to EndNote Export to Mendeley