TY - JOUR
AU - Zuo, Yi, Y
AB - Abstract The surface tension of xylem sap has been traditionally assumed to be close to that of the pure water because decreasing surface tension is thought to increase vulnerability to air seeding and embolism. However, xylem sap contains insoluble lipid-based surfactants, which also coat vessel and pit membrane surfaces, where gas bubbles can enter xylem under negative pressure in the process known as air seeding. Because of the insolubility of amphiphilic lipids, the surface tension influencing air seeding in pit pores is not the equilibrium surface tension of extracted bulk sap but the local surface tension at gas–liquid interfaces, which depends dynamically on the local concentration of lipids per surface area. To estimate the dynamic surface tension in lipid layers that line surfaces in the xylem apoplast, we studied the time-dependent and surface area-regulated surface tensions of apoplastic lipids extracted from xylem sap of four woody angiosperm plants using constrained drop surfactometry. Xylem lipids were found to demonstrate potent surface activity, with surface tensions reaching an equilibrium at ~25 mN m-1 and varying between a minimum of 19 mN m-1 and a maximum of 68 mN m-1 when changing the surface area between 50 and 160% around the equilibrium surface area. It is concluded that xylem lipid films in natural conditions most likely range from nonequilibrium metastable conditions of a supersaturated compression state to an undersaturated expansion state, depending on the local surface areas of gas–liquid interfaces. Together with findings that maximum pore constrictions in angiosperm pit membranes are much smaller than previously assumed, low dynamic surface tension in xylem turns out to be entirely compatible with the cohesion–tension and air-seeding theories, as well as with the existence of lipid-coated nanobubbles in xylem sap, and with the range of vulnerabilities to embolism observed in plants. Introduction Vascular plants are the only organisms that transport water by taking advantage of cohesion between water molecules to pull it through a hydraulic system rather than pushing it under positive pressure. Pulling stretches the distance between water molecules slightly and thereby creates negative pressure in the water. The pull is caused by evaporation of water from nanocapillaries in the cell walls of leaves (Askenasy 1895, Dixon and Joly 1895, Dixon 1914b), and the negative pressure is generated by the weight of the water column and the hydraulic resistance in the hydraulic system (Hacke et al. 2006, Lucas et al. 2013). Outside vascular plants, negative pressure in aqueous solutions also exists in other systems that possess nanocapillaries from which water evaporates, such as soils and rocks (Graham et al. 1997, Or and Tuller 2002), and such conditions have also been created in engineered devices that are often referred to as synthetic trees (Thut 1928, Wheeler and Stroock 2008, Duan et al. 2012, Vincent et al. 2012). All such systems encompass multiple phases and surfaces between them, including rigid solid walls that do not collapse under the negative pressure, a continuous aqueous phase, solid–liquid surfaces, gas bubbles and gas–liquid menisci in nanocapillaries. There has been surprisingly little research into surface phenomena in negative pressure systems, except for studies on heterogeneous bubble nucleation at solid surfaces that are in touch with the liquid under negative pressure (Wheeler and Stroock 2009, Chen et al. 2016, Menzl et al. 2016). Understanding the behavior of surfaces under negative pressure is complicated further by the presence of surface-active molecules (surfactants) in the system (Atchley 1989, Yount et al. 1977, Schenk et al. 2015). The hydraulic system of a vascular plant encompasses the entire plant, but the negative pressure is restricted to the parts outside living cells, including cell walls, intercellular spaces and the dead cells of the xylem, including vessels and tracheids. Together, these nonliving parts are referred to as the apoplast. Surfactants in xylem sap were first documented by Christensen-Dalsgaard et al. (2011) and were found to include amphiphilic lipids, including phospholipids, and proteins (Gonorazky et al. 2012, Schenk et al. 2017, 2018). A recent lipidomic analysis of lipids extracted from xylem sap found phospholipids (25–93%) and galactolipids (7–75%) (see Table 1), with total lipid concentrations ranging from 0.18 to 0.63 μmol−1 of xylem sap (Schenk et al. 2019). Table 1 Composition of xylem sap lipids (in % of molar composition) for the four studied species determined by direct-infusion electrospray ionization triple-quadrupole mass spectrometry, arranged by headgroups and expressed as mean ± standard error percentage of total lipids (Schenk et al. 2019). Acyl chains (not shown here) most commonly had 34 or 36 carbon atoms, and >98% of acyl chains were unsaturated, mostly featuring 1–3 double bonds for 34 carbon atom acyl chains and 3–5 double bonds for 36 carbon atom acyl chains (Schenk et al. 2019). Percent of all lipids . L. tulipifera . G. parviflora . D. buccinatoria . E. farinosa . Galactolipids  DGDG 7.8 ± 3.1 3.3 ± 0.7 9.1 ± 1.3 2.9 ± 0.9  MGDG 26.6 ± 12.6 7.1 ± 1.5 11.0 ± 2.1 4.0 ± 1.1 Phospholipids  PA 21.7 ± 9.3 48.9 ± 15.8 33.9 ± 11.5 49.6 ± 15.8  PC 21.2 ± 5.3 15.3 ± 3.9 19.3 ± 4.1 13.5 ± 5.8  PE 8.4 ± 4.2 11.1 ± 2.1 10.2 ± 2.3 8.7 ± 6.0  PI 4.8 ± 2.9 7.8 ± 3.5 11.3 ± 3.8 13.4 ± 8.8  PS 2.3 ± 1.1 3.4 ± 0.8 2.9 ± 1.1 2.8 ± 1.6  PG 1.2 ± 0.5 0.7 ± 0.4 0.6 ± 0.2 0.8 ± 0.5  LysoPG 4.4 ± 0.8 1.8 ± 0.8 1.2 ± 0.3 1.9 ± 0.3  LysoPC 1.5 ± 1.0 0.3 ± 0.1 0.2 ± 0.1 1.4 ± 0.4  LysoPE 0.3 ± 0.2 0.2 ± 0.1 0.2 ± 0.1 0.9 ± 0.5 Percent of all lipids . L. tulipifera . G. parviflora . D. buccinatoria . E. farinosa . Galactolipids  DGDG 7.8 ± 3.1 3.3 ± 0.7 9.1 ± 1.3 2.9 ± 0.9  MGDG 26.6 ± 12.6 7.1 ± 1.5 11.0 ± 2.1 4.0 ± 1.1 Phospholipids  PA 21.7 ± 9.3 48.9 ± 15.8 33.9 ± 11.5 49.6 ± 15.8  PC 21.2 ± 5.3 15.3 ± 3.9 19.3 ± 4.1 13.5 ± 5.8  PE 8.4 ± 4.2 11.1 ± 2.1 10.2 ± 2.3 8.7 ± 6.0  PI 4.8 ± 2.9 7.8 ± 3.5 11.3 ± 3.8 13.4 ± 8.8  PS 2.3 ± 1.1 3.4 ± 0.8 2.9 ± 1.1 2.8 ± 1.6  PG 1.2 ± 0.5 0.7 ± 0.4 0.6 ± 0.2 0.8 ± 0.5  LysoPG 4.4 ± 0.8 1.8 ± 0.8 1.2 ± 0.3 1.9 ± 0.3  LysoPC 1.5 ± 1.0 0.3 ± 0.1 0.2 ± 0.1 1.4 ± 0.4  LysoPE 0.3 ± 0.2 0.2 ± 0.1 0.2 ± 0.1 0.9 ± 0.5 Galactolipids: DGDG = digalactosyldiacylglycerol, MGDG = monogalactosyldiacylglycerol. Phospholipids: PA = phosphatidic acid, PC = phosphatidylcholine, PE = phosphatidylethanolamine, PI = phosphatidylinositol, PS = phosphatidylserine, PG = phosphatidylglycerol, LysoPG = lysophosphatidylglycerol, LysoPC = lysophosphatidylcholine, LysoPE = lysophosphatidylethanolamine. Open in new tab Table 1 Composition of xylem sap lipids (in % of molar composition) for the four studied species determined by direct-infusion electrospray ionization triple-quadrupole mass spectrometry, arranged by headgroups and expressed as mean ± standard error percentage of total lipids (Schenk et al. 2019). Acyl chains (not shown here) most commonly had 34 or 36 carbon atoms, and >98% of acyl chains were unsaturated, mostly featuring 1–3 double bonds for 34 carbon atom acyl chains and 3–5 double bonds for 36 carbon atom acyl chains (Schenk et al. 2019). Percent of all lipids . L. tulipifera . G. parviflora . D. buccinatoria . E. farinosa . Galactolipids  DGDG 7.8 ± 3.1 3.3 ± 0.7 9.1 ± 1.3 2.9 ± 0.9  MGDG 26.6 ± 12.6 7.1 ± 1.5 11.0 ± 2.1 4.0 ± 1.1 Phospholipids  PA 21.7 ± 9.3 48.9 ± 15.8 33.9 ± 11.5 49.6 ± 15.8  PC 21.2 ± 5.3 15.3 ± 3.9 19.3 ± 4.1 13.5 ± 5.8  PE 8.4 ± 4.2 11.1 ± 2.1 10.2 ± 2.3 8.7 ± 6.0  PI 4.8 ± 2.9 7.8 ± 3.5 11.3 ± 3.8 13.4 ± 8.8  PS 2.3 ± 1.1 3.4 ± 0.8 2.9 ± 1.1 2.8 ± 1.6  PG 1.2 ± 0.5 0.7 ± 0.4 0.6 ± 0.2 0.8 ± 0.5  LysoPG 4.4 ± 0.8 1.8 ± 0.8 1.2 ± 0.3 1.9 ± 0.3  LysoPC 1.5 ± 1.0 0.3 ± 0.1 0.2 ± 0.1 1.4 ± 0.4  LysoPE 0.3 ± 0.2 0.2 ± 0.1 0.2 ± 0.1 0.9 ± 0.5 Percent of all lipids . L. tulipifera . G. parviflora . D. buccinatoria . E. farinosa . Galactolipids  DGDG 7.8 ± 3.1 3.3 ± 0.7 9.1 ± 1.3 2.9 ± 0.9  MGDG 26.6 ± 12.6 7.1 ± 1.5 11.0 ± 2.1 4.0 ± 1.1 Phospholipids  PA 21.7 ± 9.3 48.9 ± 15.8 33.9 ± 11.5 49.6 ± 15.8  PC 21.2 ± 5.3 15.3 ± 3.9 19.3 ± 4.1 13.5 ± 5.8  PE 8.4 ± 4.2 11.1 ± 2.1 10.2 ± 2.3 8.7 ± 6.0  PI 4.8 ± 2.9 7.8 ± 3.5 11.3 ± 3.8 13.4 ± 8.8  PS 2.3 ± 1.1 3.4 ± 0.8 2.9 ± 1.1 2.8 ± 1.6  PG 1.2 ± 0.5 0.7 ± 0.4 0.6 ± 0.2 0.8 ± 0.5  LysoPG 4.4 ± 0.8 1.8 ± 0.8 1.2 ± 0.3 1.9 ± 0.3  LysoPC 1.5 ± 1.0 0.3 ± 0.1 0.2 ± 0.1 1.4 ± 0.4  LysoPE 0.3 ± 0.2 0.2 ± 0.1 0.2 ± 0.1 0.9 ± 0.5 Galactolipids: DGDG = digalactosyldiacylglycerol, MGDG = monogalactosyldiacylglycerol. Phospholipids: PA = phosphatidic acid, PC = phosphatidylcholine, PE = phosphatidylethanolamine, PI = phosphatidylinositol, PS = phosphatidylserine, PG = phosphatidylglycerol, LysoPG = lysophosphatidylglycerol, LysoPC = lysophosphatidylcholine, LysoPE = lysophosphatidylethanolamine. Open in new tab Xylem lipids are concentrated on the surfaces of xylem conduits and on the cellulosic and nanoporous pit membranes serving as connections between conduits (Figure 1). Since the 3D structure of pit membrane pores is characterized by various pore constrictions, which are typically below 20 nm in diameter (Choat et al. 2003, Zhang et al. 2017, 2019), these pit membranes prevent the entry of large gas bubbles into water-filled conduits that are under negative pressure (Zimmermann 1983, Schenk et al. 2015). However, surfactants associated with pit membranes would decrease surface tension and thereby increase the likelihood of gas entry through this porous medium (Cochard et al. 2009, Domec 2011, Schenk et al. 2015, 2017, 2018, 2019, Losso et al. 2017). To date, only limited data are available to indicate what the effects of natural xylem surfactants on surface tension in xylem may be (Christensen-Dalsgaard et al. 2011, Losso et al. 2017, Schenk et al. 2017). Figure 1. Open in new tabDownload slide Confocal microscopy images of transverse and tangential cell walls in secondary xylem after vacuum infiltration with FM®1-43 fluorophores (yellow color) to detect amphiphilic lipids, in vessel walls and bordered intervessel pits. Image (A) is a longitudinal view, while all other images (B-D) are transverse views. Because the secondary xylem area scanned was some distance below the injection point, the fluorophores do not occur in all conduits. Blue-colored walls show lignin autofluorescence. Lt, L. tulipifera; Gp, G. parviflora; Db, D. buccinatoria; Ef, E. farinosa; V, vessel; short arrows, intervessel walls with intervessel pits. All scale bars represent 20 μm. Reproduced and slightly modified from Schenk et al. (2018) with permission. Figure 1. Open in new tabDownload slide Confocal microscopy images of transverse and tangential cell walls in secondary xylem after vacuum infiltration with FM®1-43 fluorophores (yellow color) to detect amphiphilic lipids, in vessel walls and bordered intervessel pits. Image (A) is a longitudinal view, while all other images (B-D) are transverse views. Because the secondary xylem area scanned was some distance below the injection point, the fluorophores do not occur in all conduits. Blue-colored walls show lignin autofluorescence. Lt, L. tulipifera; Gp, G. parviflora; Db, D. buccinatoria; Ef, E. farinosa; V, vessel; short arrows, intervessel walls with intervessel pits. All scale bars represent 20 μm. Reproduced and slightly modified from Schenk et al. (2018) with permission. The behavior of lipid coatings in xylem under fluctuating negative pressure remains subject to speculation until more is known about their surface tension properties. Amphiphilic lipids aggregate on surfaces and are essentially insoluble in water, with very low critical micelle concentrations (CMCs; Marsh and King 1986). Thus, surface tension imparted by these lipids depends strongly on their local concentration at a surface, i.e., the surface density of lipid molecules, which may change over time, a phenomenon known as dynamic surface tension (Figure 2). Even at very low concentrations in xylem, amphiphilic lipids would minimize their free energy by quickly accumulating on gas–liquid surfaces (Gibbs 1906, Rice 1926) and imparting a surface tension that depends on concentration of lipids per interfacial area and not on the overall lipid concentration per water volume (Figure 2). This dynamic surface tension concept is very different from the surface tension imparted by soluble surfactants in water, such as Tween, Triton-X or alcohols, which below their CMCs depends on the concentration per volume of water and is essentially constant above CMC. Soluble surfactants have been used experimentally to test for effects of lowering surface tension in xylem (Sperry and Tyree 1988, Cochard et al. 2009, Hölttä et al. 2012, Losso et al. 2017). Figure 2. Open in new tabDownload slide Conceptual illustration of dynamic surface tension shown for the example of amphiphilic lipid films on gas bubbles in water, depending on both surface area and time. Sizes of lipid molecules and bubbles are roughly to scale. A = collapsed lipid film, showing both outward and inward folding of the lipid layer; B = lipid monolayer at equilibrium surface tension; C = bubble expansion leading to stretched monolayer with dense and stretched lipid domains and some gaps, which can potentially be filled in by the adjacent lipid micelles, if they are present and given enough time; D = bubble compression leading back to equilibrium surface tension. I = compression, II = expansion, III = compression. Note that bubbles A–D are each coated by the same number of lipid molecules. Figure 2. Open in new tabDownload slide Conceptual illustration of dynamic surface tension shown for the example of amphiphilic lipid films on gas bubbles in water, depending on both surface area and time. Sizes of lipid molecules and bubbles are roughly to scale. A = collapsed lipid film, showing both outward and inward folding of the lipid layer; B = lipid monolayer at equilibrium surface tension; C = bubble expansion leading to stretched monolayer with dense and stretched lipid domains and some gaps, which can potentially be filled in by the adjacent lipid micelles, if they are present and given enough time; D = bubble compression leading back to equilibrium surface tension. I = compression, II = expansion, III = compression. Note that bubbles A–D are each coated by the same number of lipid molecules. It is impractical to collect lipids directly from coatings on xylem surfaces, but fortunately they are also found in low concentrations in xylem sap. Here, we report research on the dynamic surface tension properties of lipids extracted from xylem sap of four woody angiosperm species measured using constrained drop surfactometry (CDS). The CDS is a novel droplet-based surface tensiometry technology developed in the Zuo laboratory (Figure 3) for studying the surface activity of pulmonary surfactants (Valle et al. 2015). Using CDS, we directly measured the time-dependent and surface area-regulated dynamic surface tensions of lipids extracted from xylem sap, which cannot be measured using standard methods such as the pendant drop technique. The technique was used previously for studying dynamic surface tension of freeze-dried xylem sap extract (Schenk et al. 2017), i.e., non-purified mixtures of proteins, amino acids, carbohydrates, lipids and salts. In addition, using a newly developed in situ Langmuir–Blodgett (LB) transfer technique (Yang et al. 2018), we revealed the ultrastructure of natural xylem lipid films using atomic force microscopy (AFM). Figure 3. Open in new tabDownload slide Schematic of the CDS for studying the surface activity of lipid films. A liquid droplet is ‘constrained’ on a 3- to 5-mm carefully machined pedestal with a knife-sharp edge. The lipid film can be periodically compressed and expanded by precisely regulating liquid flow into and out of the droplet with a motorized syringe. The surface tension and surface area of the lipid film are simultaneously determined from the shape of the droplet using axisymmetric drop shape analysis. ADSA, axisymmetric drop shape analysis. Figure 3. Open in new tabDownload slide Schematic of the CDS for studying the surface activity of lipid films. A liquid droplet is ‘constrained’ on a 3- to 5-mm carefully machined pedestal with a knife-sharp edge. The lipid film can be periodically compressed and expanded by precisely regulating liquid flow into and out of the droplet with a motorized syringe. The surface tension and surface area of the lipid film are simultaneously determined from the shape of the droplet using axisymmetric drop shape analysis. ADSA, axisymmetric drop shape analysis. We hypothesized that purified xylem sap lipid films would show similar dynamic surface tension to what is found in other natural lipid mixtures, such as pulmonary surfactants (Zuo et al. 2008, Parra and Pérez-Gil 2015), with nonconstant but surface area-regulated dynamic surface tension upon film compression and expansion (Schürch et al. 2010). Galactolipids in xylem lipid mixtures may affect the behavior of films especially at low surface tension, because their galactose headgroups strongly interact with each other, promoting lipid stacking and layering (Lee 2000, Bottier et al. 2007, Kanduč et al. 2017). Together with the xylem sap lipidomic analysis in a companion study (Schenk et al. 2019), our data provide a novel insight into the understanding of surface phenomena in xylem of vascular plants and their potential impacts on hydraulic function at the organ to whole-plant level. Materials and methods Extraction of xylem lipids Natural xylem lipids were extracted from xylem sap of four woody plant species, Distictis buccinatoria (DC.) A.H. Gentry (syn. Amphilophium buccinatorium (DC.) L.G. Lohmann, evergreen vine, Bignoniaceae), Encelia farinosa A. Gray ex Torr. (semi-deciduous desert shrub, Asteraceae), Geijera parviflora Lindl. (evergreen tree, Rutaceae) and Liriodendron tulipifera L. (deciduous tree, Magnoliaceae). These species represent different plant growth forms from four major clades distributed across the angiosperm phylogeny. Stems for xylem sap extraction were collected in August 2017. Because lipid concentrations in sap are very low, as lipids are mostly attached to xylem surfaces, samples (from two stems for Encelia and four stems for Distictis, Geijera and Liriodendron) were combined into one composite sample per species to ensure a sufficiently high concentration of lipids per sample for surface tension analysis and to create the same lipid composition as in other subsamples analyzed via mass spectrometry (Schenk et al. 2019). Analysis of composite samples does not allow statistical comparisons between species, but such comparisons were not the goal of this study. Most collections were done at dawn to ensure full hydration of the stems. The extraction protocol was described in detail in Schenk et al. (2019). Branches were cut from the plants at a length exceeding the longest vessel for each species, transported immediately to the lab and cut under water, with the final cuts made with a fresh razor blade. The bark was removed from the proximal end for ~4 cm length to expose the xylem cylinder and to avoid phloem contamination. The cut surface was thoroughly cleaned with deionized water using a high-pressure dental flosser (WP-100 Ultra Water Flosser, Water Pik Inc., Fort Collins, CO, USA) for 2 min to remove cell debris and cytoplasmic content from the surface. Xylem sap was extracted from the proximal end of the stem under vacuum into a glass test tube embedded in ice inside a 1-l Buchner flask. The flask was subjected to lab vacuum for ~30 s, after which the distal end of the branch was cut back by ~4 cm, followed by successive 2-cm cuts made to all the side branches until sap was observed dripping into the test tube. Further 1-cm cuts were made every minute to allow for slow, continuous extraction of xylem sap. Depending on stem size, 1–2 ml of sap were extracted from each stem, moved in a glass pipette into pre-weighed 1.5-ml LoBind Eppendorf microcentrifuge tubes, flash-frozen in liquid nitrogen and stored in the freezer until further processing. Samples were also taken to test for contamination of sap samples by lipids from living cell remnants (Schenk et al. 2017). This was done prior to xylem sap extraction by inserting the cleaned, cut ends of xylem into a glass vial containing 1 ml of nanopure water. Lipid concentrations in these controls for the four study species ranged from 1.1 to 6.7% of the lipids found in xylem sap (Schenk et al. 2019), which was too low to be included in surface tension measurements. Other tests have shown that xylem sap lipids are not contaminants that originate from repeated cuts of stems made during the vacuum extraction (Schenk et al. 2019), and the ordered nature of lipid layers observed on xylem surfaces (Figure 1) (Schenk et al. 2017, 2018) also rules out contamination as the source of xylem sap lipids. Xylem sap samples in LoBind Eppendorf microcentrifuge tubes were partially lyophilized in a freeze dryer (FreeZone 1 Liter Benchtop Freeze Dry System, Labconco, Kansas City, MO, USA) until only ~100 μl of sample remained in each tube. The remaining aqueous samples were combined into one sample for each species using a glass pipette and weighed. Methanol:chloroform 1:1 (both HPLC grades, Fisher Scientific, Waltham, MA, USA) was added to the combined aqueous sample to create an approximate 5:5:1 methanol:chloroform:water one-phase mixture. The mixture was then vortexed and centrifuged, and the supernatant was collected using a 1-ml glass syringe with PrecisionGlide needle. A new one-phase 5:5:1 mixture of methanol:chloroform:water was then added to the residue, the mixture was vortexed and centrifuged, and the supernatant again was collected and combined with the previously collected supernatant. Samples were dried in a desiccator outfitted with an in-line carbon filter capsule (model 6704-7500, Whatman, GE Healthcare Life Sciences, Marlborough, MA, USA) and sent to the Zuo lab at the University of Hawaii at Manoa for analysis. Other samples from these same collections extracted the same way were sent out for mass spectrometry analyses to determine the lipid compositions, which are reported in a separate paper (Schenk et al. 2019). Xylem lipids extracted via this protocol also include very low concentrations of hydrophobic proteins determined by Tris-Tricine SDS-PAGE to be between 5 and 18 kDa in size (data not shown). The very low concentrations have prevented the identification of these proteins to date. Constrained drop surfactometry Dynamic surface tension measurements and surfactant film imaging were carried out with CDS (Valle et al. 2015). Constrained drop surfactometry is a new generation of drop-based surfactometer. As shown in Figure 3, it uses a sessile drop (~3 mm in diameter) to generate the air–water surface, where the natural surfactant sample is spread and studied. The sessile drop is confined on a carefully machined pedestal that uses a knife-sharp edge to prevent the droplet from spreading at low surface tensions. The spread surfactant film can be compressed and expanded by withdrawing and injecting liquid into the droplet using a motorized syringe. Surface tension is determined photographically from the shape of the droplet using axisymmetric drop shape analysis (ADSA) (Yang et al. 2017). For CDS measurement, each freeze-dried xylem lipid sample was dissolved in 50 μl of chloroform. The samples were subjected to 5 min of ultrasonic treatment, followed by 1 min of vortex treatment to create a homogeneous solution, after which 1 μl of xylem lipid sample was spread at the surface of the pure water droplet (Milli-Q ultrapure water; Millipore, Billerica, MA, USA) of the CDS instrument, where it was left undisturbed for 1 min to allow film spreading and evaporation of the chloroform. Time-dependent dynamic surface tension was recorded with ADSA until an equilibrium surface tension (γe) was reached. Subsequently, the xylem surfactant film was compressed and expanded at a rate of 0.1 cm-2 min. The surface area-regulated surface tension was recorded during three compression–expansion cycles for each composite sample. All measurements were conducted at room temperature of 20 ± 1°C. Film imaging with AFM To determine if the lipid films were smooth monolayers at γe, the molecular organization and lateral structure of the spread xylem lipid films at γe were visualized via AFM. The films were LB-transferred from the air–water surface to a small piece of freshly peeled mica sheet by lifting the mica through the drop surface at a rate of 1 mm min as described in Yang et al. (2018). Topographic images of the xylem lipid films were obtained using an Innova AFM (Bruker, Santa Barbara, CA, USA). Samples were scanned in air in a contact mode using a silicon nitride cantilever with a spring constant of 0.12 N m-1 and a tip radius of 2 nm. Lateral structures were analyzed using Nanoscope Analysis (version 1.5). Results Time-dependent dynamic surface tension of xylem lipids Figure 4 shows the time-dependent dynamic surface tensions of xylem lipids extracted from D. buccinatoria, E. farinosa, G. parviflora and L. tulipifera. Reproducibility of these dynamic surface tension measurements within each composite sample can be found in Figure S1 available as Supplementary Data at Tree Physiology Online. After spreading at the air–water surface, the xylem lipids immediately reduced surface tension below 40 mN m-1. They further reduced the surface tension to an equilibrium value (γe) around 25 mN m-1 within 2–3 s. The γe for xylem lipids of all four species is very close and is consistent with that of the adsorbed film from fully hydrated phospholipid bilayers (Zuo et al. 2008). Figure 4. Open in new tabDownload slide Time-dependent dynamic surface tension of natural xylem lipids from four woody angiosperm species, measured via CDS and ADSA (Figure 3) starting with a pure water droplet at time zero (72 mN m-1) and after adding xylem lipids to the droplet at ~3 s, leading to an immediate drop in surface tension to around 37 mN m-1 and then reaching equilibrium surface tension, γe, around 25 mN m-1 within a few seconds. Figure 4. Open in new tabDownload slide Time-dependent dynamic surface tension of natural xylem lipids from four woody angiosperm species, measured via CDS and ADSA (Figure 3) starting with a pure water droplet at time zero (72 mN m-1) and after adding xylem lipids to the droplet at ~3 s, leading to an immediate drop in surface tension to around 37 mN m-1 and then reaching equilibrium surface tension, γe, around 25 mN m-1 within a few seconds. Surface area-regulated surface tension of xylem lipids Figure 5 shows the surface area-regulated dynamic surface tension of the xylem lipids for the four plant species. Reproducibility of these compression–expansion cycles within composite samples can be found in Figure S2 available as Supplementary Data at Tree Physiology Online. In these experiments, the equilibrium xylem surfactant films at γe were first compressed to ~60% of the original area and subsequently expanded to 160% of the original area. When an insoluble surfactant film at equilibrium is compressed, it enters a metastable compression regime. All four xylem lipid films were remarkably similar between species, despite the differences in lipid composition (Table 1). Lipid films quickly reach a minimum surface tension (γmin) around 19 mN m-1 when the films are compressed by <5% of the original area. This γmin is in line with the collapse surface tension reported for the two galactolipids in xylem lipid, monogalactosyldiacylglycerol (MGDG) and digalactosyldiacylglycerol (DGDG) (Gzyl-Malcher et al. 2008, Hoyo et al. 2016). The γmin does not change significantly with further area reduction by 40% of the lipid film. Upon expansion, all four xylem lipid films show an abrupt surface tension increase to γe, i.e., around 25 mN m-1. The surface tension is stabilized at γe until the film is expanded beyond its original surface area. With further expansion, the lipid film enters a metastable expansion regime, in which the surface tension increases continuously with area expansion to 160% of its original area. The film is then compressed again with decreasing surface tension to complete a cycle. Figure 5. Open in new tabDownload slide Surface area-regulated surface tension of xylem lipids from four woody angiosperm species. Film compression is indicated by solid symbols and film expansion is indicated by open symbols. Lipids from all four species show similar compression and expansion cycles. Cycles start at 100% relative surface area, followed by compression to 50–70% and expansion to 140–170% of surface area. Explanation of symbols: γmax = maximum surface tension on expanded surface area, γe = equilibrium surface tension at 100% relative surface area, γmin = minimum surface tension of collapsed lipid layers. Figure 5. Open in new tabDownload slide Surface area-regulated surface tension of xylem lipids from four woody angiosperm species. Film compression is indicated by solid symbols and film expansion is indicated by open symbols. Lipids from all four species show similar compression and expansion cycles. Cycles start at 100% relative surface area, followed by compression to 50–70% and expansion to 140–170% of surface area. Explanation of symbols: γmax = maximum surface tension on expanded surface area, γe = equilibrium surface tension at 100% relative surface area, γmin = minimum surface tension of collapsed lipid layers. Ultrastructure of xylem lipid films Figure 6 shows the AFM topographic images of the four xylem lipid films at the equilibrium surface tension (γe). Reproducibility of these AFM topographic images within composite samples can be found in Figure S3 available as Supplementary Data at Tree Physiology Online. At γe, the lipid film at the air–water surface reaches a dynamic equilibrium with lipid micelles and vesicles in the subphase. As shown in Figure 6, all xylem lipid films maintain a lateral structure of coexisting interfacial monolayer with randomly distributed protrusions. Although the thickness of the background layer is unknown, its smoothness shows it to be a monolayer, as lipids prone to multilayer formation would form a much bumpier surface. Height analysis indicates that the sizes of most protrusions are multiples of 4 nm, which is the thickness of a fully hydrated lipid bilayer (Nagle and Tristram-Nagle 2000). Hence, these protrusions most likely represent multilayers of lipids attached to the interfacial monolayer of the xylem surfactant under the equilibrium condition. The number and height of the protrusions are different for the xylem lipid from the four plant species. Lipids from Distictis and Liriodendron, which have the largest amounts of galactolipids (Table 1), have the fewest peaks, and most are around 8 nm in height. Although the number of 8-nm peaks is highest in Distictis and Liriodendron, a 12-nm peak was found for Encelia. These images support the conclusion that xylem lipids form monolayers at equilibrium surface tension but begin to create folds and stacks that are likely to increase upon further compression of the layer. Figure 6. Open in new tabDownload slide Atomic force microscopy topographic images of four natural xylem lipid films close to equilibrium surface tension (see Figure 5). The films were created by moving a clean mica sheet through a lipid film at equilibrium. (a) D. buccinatoria, (b) E. farinosa; (c) G. parviflora and (d) L. tulipifera. The AFM images in the first row have the same scanning area of 10 × 10 μm and the z-range of 20 nm. Images in the second row show the surface render of the zoom-in xylem surfactant film as indicated by the white boxes. White arrows indicate the heights of the multilayered structures. Figure 6. Open in new tabDownload slide Atomic force microscopy topographic images of four natural xylem lipid films close to equilibrium surface tension (see Figure 5). The films were created by moving a clean mica sheet through a lipid film at equilibrium. (a) D. buccinatoria, (b) E. farinosa; (c) G. parviflora and (d) L. tulipifera. The AFM images in the first row have the same scanning area of 10 × 10 μm and the z-range of 20 nm. Images in the second row show the surface render of the zoom-in xylem surfactant film as indicated by the white boxes. White arrows indicate the heights of the multilayered structures. Discussion An aqueous liquid under negative pressure is in a metastable state and is prevented from turning into vapor by the nucleation energy required to break hydrogen bonds between water molecules and form a void (Wheeler and Stroock 2009, Chen et al. 2016, Menzl et al. 2016). Pure water has a very high tensile strength, up to a theoretical maximum of −190 MPa or at least to an experimentally achievable maximum of −30 MPa (Caupin et al. 2012), which prevents homogeneous nucleation under negative pressure in plants (Oertli 1971, Stroock et al. 2014). Xylem sap, however, is not pure water, and the presence of salts, organic solutes and especially surface-active molecules would affect nucleation thresholds (Dixon 1914b, Hemmingsen 1978). Because of the perception that low surface tension would increase vulnerability to embolism, the surface tension of xylem sap has been traditionally assumed to be equal to that of the pure water (Oertli 1971, Tyree and Zimmermann 2002, Meyra et al. 2007). That assumption, however, has proven to be incorrect (Christensen-Dalsgaard et al. 2011, Domec 2011, Losso et al. 2017, Schenk et al. 2017). Moreover, the functional significance of surface-active insoluble lipids on xylem surfaces demands the consideration of not just the equilibrium surface tension of bulk xylem sap but also the concentration-dependent, local surface tension at phase interfaces in xylem. Here, we determined both time-dependent and surface area-regulated dynamic surface tensions of natural lipids extracted from xylem sap. It was found that when spread at the air–water surface, all xylem lipids quickly reach an equilibrium (γe) by reducing the surface tension of the air–water surface from 72 to ~25 mN m-1 within a few seconds (Figure 3), indicating a potent surface activity. When compressed below γe, the xylem lipid films maintain a supersaturated metastable state without collapse and reduce the surface tension to a minimum surface tension (γmin) around 19 mN m-1. Presumably further compression could further reduce the surface tension toward zero, as the lipid layers stack and fold. When expanded above γe, the xylem surfactant films maintain an undersaturated metastable state and increase the surface tension to a maximum (γmax) of 52 mN m-1 for E. farinosa and 68 mN m-1 for D. buccinatoria, i.e., closer to the surface tension of pure water (Figure 5). The shapes of surface tension curves as function of drop surface area are remarkably similar between species (Figure 5), despite considerable differences in lipid composition (Table 1). Our hypothesis that xylem lipids would show similar dynamic surface tension to other biological lipid mixtures, such as pulmonary surfactants, was supported by these findings. Dynamic changes in surface tension are crucial for the function of alveoli in the lungs (Parra and Pérez-Gil 2015), but it remains to be investigated what their functional importance is in the angiosperm xylem, where changes in pressure that affect gas–liquid interfaces typically show diurnal and seasonal fluctuations, not many fluctuations per minute, as in the lungs. Effects of compression and expansion rates on dynamic surface tension (see Figure S2 available as Supplementary Data at Tree Physiology Online) merit further investigation, as not all pressure changes in plants are slow. Embolism formation in a xylem conduit creates a sudden, local pressure increase (Hölttä et al. 2009), and it remains to be investigated how lipid layers on air–water interfaces would respond to such an extremely rapid change in surface conditions. The presence of substantial amounts of galactolipids in xylem sap (6.9% in Encelia to 34.4% in Liriodendron; Table 1) was very unexpected, as both are plastid lipids, and MGDG has never been found outside of plastids before (Dörmann and Benning 2002, Kalisch et al. 2016). Apoplastic galactolipids in xylem vessels most likely originate from the original cell content of the living vessels before programmed cell death, although this requires further research (Esau 1965, Esau et al. 1966, Schenk et al. 2019). The role of galactolipids in the dynamic surface tension patterns shown in Figures 4 and 5 requires further investigation. Galactolipids are typically highly unsaturated (Dörmann and Benning 2002, Kalisch et al. 2016), and unsaturated MGDG, which made up between 4.0 and 26.6% of all xylem lipids in this study, do not form bilayers by themselves, because their small headgroup and spreading acyl chains give them a cone shape that favors curved shapes of lipid layers (Lee 2000). However, strong headgroup interactions between galactolipids favor stacking, such as seen in the grana of thylakoids in chloroplasts (Lee 2000, Simidjiev et al. 2000, Bottier et al. 2007, Kanduč et al. 2017). Galactolipids may be especially prone to stacking and folding and could be responsible for the distinct inward folding of lipid coats seen in freeze-fracture electron micrographs of lipid-coated nanobubbles from xylem sap in Figure 8C in Schenk et al. (2017) and the multilayered structures seen under AFM, even at equilibrium surface tension, γe (Figure 6). Most xylem sap lipids are phospholipids, especially phosphatidic acid (PA) and phosphatidylcholine (PC) (Table 1), and the composition of phospholipid headgroups is likely to affect some of the differences in dynamic surface tension between species. For example, the headgroup of PA is small and negatively charged, with a tendency toward forming nanodomains and concave (inward) curvature of lipid layers (Kooijman et al. 2003). However, surface tension is more strongly affected by acyl chain characteristics, and these were very similar between species, with ~84% of acyl chain lengths of 17 and 18 carbon atoms, > 98% unsaturated, and with mostly 1–3 double bonds for chain lengths of 17 and 3–5 double bonds for chain lengths of 18. The similarity in chain lengths and degree of saturation could explain why minimum and equilibrium surface tension was remarkably similar between species. Further research will be required to determine the effects of lipid composition on the differences in surface tension between species, including their effects on folding, stacking and formation of nanodomains by phase separation, best studied via AFM at minimum and maximum surface tension. Comparison with previous surface tension measurements of xylem sap Both γe and γmin determined here are significantly lower than the traditionally estimated and measured surface tension of bulk xylem sap (Tyree and Zimmermann 2002, Christensen-Dalsgaard et al. 2011, Losso et al. 2017). A previous study of the surface tension of xylem sap of three angiosperm tree species using the pendant drop method (Christensen-Dalsgaard et al. 2011) found that given a long-enough time (i.e., 30 min), the surface tension of xylem sap can be reduced to the range of 52–58 mN m-1, most likely caused by a reduction of surface area due to evaporation from the drop, which was not in a vapor-saturated environment. In a later study, the surface tension of xylem sap extracted from two conifers was measured using the sessile drop method (Losso et al. 2017), finding that the equilibrium surface tension showed seasonal variations and fluctuated between 50 and 68 mN m-1 over the year. Minimum surface tensions reported by these researchers are closer to the maximum surface tension (γmax) determined in the present study. The differences between the present study and previous measurements can be explained by the following two facts. First, both previous measurements (Christensen-Dalsgaard et al. 2011, Losso et al. 2017) did not study the surface area-regulated, dynamic surface tension. Second, both previous measurements (Christensen-Dalsgaard et al. 2011, Losso et al. 2017) used extracted xylem sap without further purification and concentration. In contrast, the xylem lipids studied here are purified (by organic solvent extraction) and concentrated (by pooling multiple xylem sap samples). Consequently, the xylem lipids studied here show significantly more potent surface activity than the xylem sap studied before, as indicated by a much faster adsorption (reaching equilibrium within 5 s vs >30 min) and a much lower γe (25 vs > 50 mN m-1). A look at the xylem conduits shown in Figure 1 (also see Schenk et al. 2018) demonstrates why it makes sense to measure surface tension of lipid films rather than that of extracted xylem sap. Lipid layers on inner vessel wall surfaces and in bordered pits create high local lipid concentrations at locations where bubble nucleation is most likely to occur, while most of the surface-attached lipids are unlikely to appear in extracted and non-purified xylem sap. Therefore, the equilibrium surface tension measurements of extracted bulk xylem sap do not provide any information about the functional significance of surfactants in water-conducting cells of xylem tissue. On the other hand, while lipid extraction from sap is necessary to concentrate the lipids for realistic surface tension measurements, extraction removes other components found in sap, such as ions, proteins and carbohydrates, that may interact with lipids in xylem. Future research will have to compare the behavior of lipids in the presence and absence of these other xylem components. Our data therefore indicate that the xylem surfactant films studied previously were most likely at an undersaturated expansion state, as they were conducted with xylem sap with very low surfactant concentrations (Schenk et al. 2017, 2019), and using static drop-based methods that did not include drop compression (Christensen-Dalsgaard et al. 2011, Losso et al. 2017), except for potential evaporation from a pendant drop, potentially causing it to shrink and decreasing surface tension (Christensen-Dalsgaard et al. 2011). The droplet for surface tension measurements usually has a diameter of 3–4 mm, corresponding to a surface area of 0.15–0.4 cm2 (Figure S2 available as Supplementary Data at Tree Physiology Online). This drop size is required by the measurement principle of drop shape analysis. When the drop/bubble is smaller than the capillary length (e.g., 2.7 mm for pure water), it cannot be effectively deformed by gravity and thus maintains a nearly spherical shape that fails the surface tension measurements from shape analysis (Yang et al. 2017). The difference between our study and previous ones is not in the size of the droplet but in the changing concentration of surface-active molecules on the droplet’s surface, which is needed to understand the concentration-dependent local surface tension. Implications of dynamic surface tension of lipids for xylem function Dynamic surface tension commonly shows hysteresis between compression and expansion (Figure 5) (Ingenito et al. 1999, Zuo et al. 2008), due to processes such as the formation, dissolution and tearing of lipid domains (Kwan and Borden 2010), lipid layer folding during compression and unfolding during expansion and replenishment of lipids from micelles into stretched lipid films (Figure 2). Hysteresis may be important in the face of pressure fluctuations in xylem conduits. Xylem commonly experiences diurnal fluctuations in negative pressure that can exceed 2 MPa (Meinzer et al. 2016), with the most negative pressures occurring in the middle of the day and less negative ones often just before dawn. Large fluctuations in pressure would affect phase interfaces in xylem and cause area variations of the lipid-covered surface, thus actively regulating the surface tension. Hysteresis effects are most likely short-lived, as lipids can migrate quickly between micelles in bulk and phase interfaces, but hysteresis phenomena in lipid films could play an important role in response to sudden pressure fluctuations, such as a sudden local tension relief caused by the formation of an embolism (Hölttä et al. 2009). The most likely sources for emboli in xylem are pores in pit membranes between sap-filled and gas-filled conduits (Zimmermann 1983, Hacke and Sperry 2001, Meyra et al. 2007, Jansen and Schenk 2015). Estimated pore constrictions in fresh (fully hydrated, non-dried) pit membranes of angiosperms are much smaller (<20 nm in diameter) than previously assumed, while previous estimates of maximum pore sizes were based on a simplistic, 2D view of pores and affected by drying artifacts (Zhang et al. 2017, 2019). According to the Young–Laplace equation, the pressure difference required to force a bubble through a 20-nm pore, assuming a contact angle of zero (Meyra et al. 2007,Caupin et al. 2008), and a pore shape correction factor for non-cylindrical pores of 0.5 (Schenk et al. 2015), would be 7.2 MPa in pure water. Most angiosperms are vulnerable to air seeding far below that value (Choat et al. 2012). With lipids in pit membrane pores, if the surface tension of an air–water meniscus is reduced to 25 mN m-1, the equilibrium surface tension found in this study, a meniscus could pass through a 20-nm pore with a shape correction factor of 0.5 under a pressure difference of 2.5 MPa. This is a far more realistic value for air seeding to cause embolism in angiosperms than 7.2 MPa (Choat et al. 2012). The assumption that a rare pit membrane with a much larger pore will trigger embolism formation at a similar air-seeding pressure is frequently suggested in the literature (Wheeler et al. 2005, Plavcová et al. 2013). This hypothesis, however, does not consider the presence of xylem sap surfactants and the 3D structure of the pit membrane pore space, which consists of various pore constrictions with highly variable pore volumes, depending on the overall pit membrane thickness and the number of cellulose microfibril layers (Jansen et al. 2018, Kaack et al. 2019, Zhang et al. 2019). Low surface tension in xylem is therefore entirely compatible with the cohesion–tension and air-seeding theories and with the range of observed vulnerabilities to embolism in plants, while xylem sap with the high surface tension of pure water is incompatible with both the theories and observations. Xylem lipids not only coat solid surfaces but have also been found to coat nanobubbles in extracted xylem sap (Schenk et al. 2017). It is impossible to know for certain if lipid-coated nanobubbles exist in sap that is under negative pressure in intact xylem, but their existence in aqueous solution under fluctuating negative pressure is theoretically possible if their surface tensions were sufficiently low (Atchley 1989, Schenk et al. 2015, 2017). In fact, closely packed insoluble surfactant monolayers and the resultant regulation of surface tension due to surface area variations may be the only way to allow very small gas bubbles to exist in a liquid (Atchley 1989, Yount et al. 1977, Yount 1979, 1997). Otherwise, a small gas bubble in water would either dissolve instantaneously because of the excess Laplace pressure inside the bubble or expand into an embolism if the differences between the sap pressure and the internal gas pressure in the bubble exceeded the pressure originating from the surface tension (Atchley 1989, Schenk et al. 2017). Lipid-coated nanobubbles could originate from liquid–lipid–gas interfaces moving through highly variable pore volumes with multiple pore constrictions. The resulting nanobubbles need to be smaller than the critical size that causes bubble expansion depending on surface tension (Schenk et al. 2017). Bubbles larger than that critical size would expand and either continue to be stabilized by increasing area-dependent surface tension (Figures 2 and 5) or continue expanding to form a xylem embolism. Obviously, xylem in a living plant experiences constant changes in temperature, pressure and dissolved gas concentrations (Schenk et al. 2016), and simple equilibrium models cannot do justice to ever-changing conditions. The dynamic surface tension data presented in this study will allow the development of more realistic dynamic models to predict the formation and behavior of lipid-coated nanobubbles and of other phase interfaces in xylem, especially at pit membranes. Remarkably, the first one to suggest a role of colloids for enabling water transport under negative pressure in plants was no other than Henry Dixon, one of the originators of the cohesion–tension theory (Dixon 1914a, 1914b). The evidence presented in this study sheds a novel light on the long-standing question of how plants transport water under negative pressure, which remains incompletely understood, especially with respect to the mechanisms that prevent embolism formation and the temporal frequency of hydraulic failure. Conclusions Contrary to long-standing assumptions, the apoplast of angiosperm xylem contains amphiphilic lipids, which are insoluble and accumulate on surfaces, including at air–water interfaces. Using constrained drop surfactometry, we were able to study the time-dependent and surface area-regulated dynamic surface tensions of apoplastic lipids. These were extracted from xylem sap, because it is impractical to collect lipids directly from surfaces in the xylem. It is found that xylem lipids extracted from all four tree species demonstrate potent surface activity. Surface tensions of the xylem surfactants reach an equilibrium at 25 mN m-1 and vary between a minimum value of 19 mN m-1 and a maximum value of 68 mN m-1 when changing the surface area between 60 and 160% around the surface area at equilibrium. We therefore concluded that xylem lipid films in natural conditions vary most likely between nonequilibrium conditions of a supersaturated metastable compression state and an undersaturated metastable expansion state, depending on the local surface area available at phase interfaces, which could be affected by changes in liquid pressure. Open questions to be addressed in future research include the exact origin of xylem sap lipids, the behavior of surfactant-coated air–water–solid interfaces under negative pressure, effects of other types of molecules on the dynamic surface tension of lipid films in xylem and how exactly air seeding would work based on a dynamic surface tension concept. Our study provides a novel understanding of how xylem lipids both affect air seeding through the tiny pore constrictions of angiosperm pit membranes and can also stabilize resulting nanobubbles, thus collectively reducing the vulnerability to embolism at the organ and whole-plant level. Acknowledgments The authors thank Tatiana Melendres for assistance with xylem sap extractions, Greg Pongetti at the Fullerton Arboretum for allowing sampling of plants, and Aissa Do and Alison Miyamoto for conducting protein analyses of xylem lipid mixtures. Conflict of interest We declare that we have no conflict of interest. Funding The research was funded by the National Science Foundation IOS-1558108 and IOS-1754850 (H.J.S., S.J. and Y.Y.Z.) and CBET-1604119 (Y.Y.Z.). S.J. acknowledges funding from the German Research Foundation (Individual Research Grant No. 383393940). References Askenasy E ( 1895 ) Ueber das Saftsteigen. Verhandlungen des Naturhistorisch-medizinischen Vereins zu Heidelberg . 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TI - Dynamic surface tension of xylem sap lipids
JF - Tree Physiology
DO - 10.1093/treephys/tpaa006
DA - 2020-04-08
UR - https://www.deepdyve.com/lp/oxford-university-press/dynamic-surface-tension-of-xylem-sap-lipids-u9JR3OFMj3
SP - 433
VL - 40
IS - 4
DP - DeepDyve
ER -