RESEARCH ARTICLE

INVITED SPECIAL ARTICLE For the Special Section: Wood: Biology of a Living Tissue

From the sap’s perspective: The nature of vessel surfaces in angiosperm xylem

H. Jochen Schenk1,3 , Susana Espino1, Sarah M. Rich-Cavazos1, and Steven Jansen2

Manuscript received 29 July 2017; revision accepted 14 December PREMISE OF THE STUDY: Xylem sap in angiosperms moves under negative pressure in conduits 2017. and cell wall pores that are nanometers to micrometers in diameter, so sap is always very 1 Department of Biological Science, California State University close to surfaces. Surfaces matter for water transport because hydrophobic ones favor Fullerton, Fullerton, CA 92831, USA nucleation of bubbles, and surface chemistry can have strong effects on flow. Vessel walls 2 Institute of Systematic Botany and Ecology, Ulm University, contain cellulose, hemicellulose, lignin, pectins, proteins, and possibly lipids, but what is the Albert-Einstein-Allee 11, D–89081, Ulm, Germany nature of the inner, lumen-­facing surface that is in contact with sap? 3 Author for correspondence (e-mail: [email protected]) Citation: Schenk, H. J., S. Espino, S. M. Rich-Cavazos, and S. Jansen. METHODS: Vessel lumen surfaces of five angiosperms from different lineages were examined 2018. From the sap’s perspective: The nature of vessel surfaces in via transmission electron microscopy and confocal and fluorescence microscopy, using angiosperm xylem. American Journal of Botany 105(2): 172–185. fluorophores and autofluorescence to detect cell wall components. Elemental composition doi:10.1002/ajb2.1034 was studied by energy-­dispersive X-­ray spectroscopy, and treatments with phospholipase C (PLC) were used to test for phospholipids.

KEY RESULTS: Vessel surfaces consisted mainly of lignin, with strong cellulose signals confined to pit membranes. Proteins were found mainly in inter-­vessel pits and pectins only on outer rims of pit membranes and in vessel-­parenchyma pits. Continuous layers of lipids were detected on most vessel surfaces and on most pit membranes and were shown by PLC treatment to consist at least partly of phospholipids.

CONCLUSIONS: Vessel surfaces appear to be wettable because lignin is not strongly hydrophobic and a coating with amphiphilic lipids would render any surface hydrophilic. New questions arise about these lipids and their possible origins from living xylem cells, especially about their effects on surface tension, surface bubble nucleation, and pit membrane function.

KEY WORDS amphiphilic lipids; cellulose; hydrophilic vs. hydrophobic; lignin; pectin; pit membrane; proteins; vessel-associated cells; xylem sap; xylem vessel wall.

Plant xylem is a unique evolutionary invention: It functions as a long distances. Somehow plants manage to move large volumes negative pressure hydraulic system that can move vast quantities of water in heterogeneous channels that range in diameter from of water and solutes over distances longer than 100 m. No other nanometers to a few hundred micrometers, that is, in nanoflu- kind of organism transports liquids under negative pressure, and idic and microfluidic systems where water moves very close to the most successful attempts to build an artificial negative pres- surfaces of other phases, both solid and gas (Eijkel and van den sure hydraulic system succeeded only to move a small amount of Berg, 2005; Squires and Quake, 2005). Plants move all this water water over a few centimeters in a single microchannel (Wheeler efficiently along these phase surfaces without constantly creating and Stroock, 2008). That experiment proved that it is possible gas bubbles in the system (Jansen and Schenk, 2015). What do we to move water under negative pressure, but it left the question actually know about these surfaces in xylem conduits (i.e., vessels unanswered how plants can do this so effectively and over such and tracheids)?

172 • American Journal of Botany 105(2): 172–185, 2018; http://www.wileyonlinelibrary.com/journal/AJB/ © 2018 Botanical Society of America February 2018, Volume 105 • Schenk et al.—The nature of vessel surfaces in angiosperm xylem • 173

It turns out that our knowledge of the surface characteristics of act as bubble nucleation sites (Creech et al., 2002; Wheeler and inner conduit walls is very limited. The structure and function of Stroock, 2008, 2009; Boatwright et al., 2015). Most attempts to cre- cell walls is very well known; after all, xylem forms wood, a major ate negative pressure in aqueous solution fail due to surface bubble building material. But what are the chemical and physical charac- nucleation below negative pressures of about -­1 MPa or even above teristics of the wall surfaces along which water travels on its way that (Smith, 1994). So how can vascular plants transport water in from roots to leaves? The lumen-facing­ surface layer of secondary hydraulic systems that are routinely below -­1 MPa negative pressure cell walls is usually called the S3 layer or tertiary wall (Liese, 1963; (Lambers et al., 2008; Choat et al., 2012)? Butterfield and Meylan, 1980; Panshin and de Zeeuw, 1980). It is Critics have questioned the existence of negative pressure in very thin in angiosperms, about 100–250 nm in vessels (Harada, plants ever since the cohesion–tension theory was first proposed 1965; Fromm et al., 2003; Xu et al., 2006b; Kim and Daniel, 2012) (Askenasy, 1895; Dixon and Joly, 1895), but the direct and indirect and is defined by the interwoven and more swollen nature of mi- evidence for negative pressure in plants is overwhelming (Jansen and crofibrils embedded in amorphous material, rather than the more Schenk, 2015). However, critics have pointed to seemingly contra- parallel orientation of thinner microfibrils observed in the underly- dictory evidence, such as the existence of hydrophobic compounds ing and much thicker S2 layer (Liese, 1963). Its chemical composi- in xylem (Schneider et al., 2003; Rösch et al., 2004; Zimmermann tion was found to be clearly different from the S2 layer, based on the et al., 2004; Westhoff et al., 2008). These valid observations must be easy destruction of the S2 layer by soft-­rot fungi, while the S3 layer accounted for to fully understand how plants transport water via can remain intact (e.g., Liese, 1955). The S3 layer was also suggested the cohesion–tension mechanism. Water in plants is normally satu- to extend from the cell lumen into pit borders (Kollmann and Côté, rated with dissolved gas or even supersaturated during temperature 1968). Layered on top of the S3 layer can be a warty layer, which is rise (Schenk et al., 2016), so it would seem that bubble formation formed by deposits of matrix substance, either lignin or lignin and should be almost unavoidable. Are the interior surfaces of the xy- hemicellulose (Liese, 1963; Kuo and Manwiller, 1986). Moreover, lem, the plants’ hydraulic system, free of any crevices or locations a thin coat of lipids remaining from live cell content during vessel with hydrophobic properties? One quick look at any scanning elec- maturation was suggested to be deposited on the lumen walls of tron micrograph of xylem lays to rest the idea that there are no sur- conduits (Scott et al., 1960; Esau, 1965; Esau et al., 1966b). A re- face irregularities (e.g., Butterfield and Meylan, 1980), so what do cent study of several angiosperm species confirmed the presence of plants do to prevent bubbles from nucleating there? such lipid layers on pit membranes of bordered pits and other vessel The other problem posed by xylem surfaces is their effect on lumen surfaces (Schenk et al., 2017). liquid flow. Water molecules stick firmly to hydrophilic surfaces, In recent years, a growing number of studies have found that xy- potentially forming boundary layers or so-called­ exclusion zones lem surfaces tend to be somewhere between hydrophilic and hydro- that can have an almost crystalline structure and allow almost no phobic, with observed aqueous contact angles ranging from about movement of water within these layers (Bonaccurso et al., 2002; 40° to a little more than 90° (Schneider et al., 2000; Zwieniecki and Zheng et al., 2006), while hydrophobic surfaces could allow water Holbrook, 2000; van Ieperen et al., 2001; Shen et al., 2008; Kohonen molecules to slip past (Bonaccurso et al., 2002; Sendner et al., 2009). and Helland, 2009; Brodersen et al., 2010; Kim and Lee, 2010; Lee However, flow in both hydrophobic and hydrophilic nanochannels et al., 2013; McCully et al., 2014; Knipfer et al., 2016), where a can be surprisingly high due to molecular interactions at the liquid– contact angle of 0° denotes a completely hydrophilic surface and solid boundary (Calabrò et al., 2013; Gruener et al., 2016). Again, contact angles above 90° a hydrophobic one. Other recent studies the nature of the solid surface matters greatly for the flow that can have shown that xylem surfaces, at least in angiosperms, feature a be achieved. complex spatial arrangement of surfaces with hydrophilic and hy- Here we follow up on a study by Schenk et al. (2017) to provide drophobic properties (McCully et al., 2014; Schenk et al., 2017), more specific information on the nature of xylem surfaces that are including coatings consisting at least in part of amphiphilic lipids in contact with xylem sap. Fluorescent tracers and autofluorescence (Schenk et al., 2017). The actual chemistry of xylem surfaces is less were used to examine exposed surfaces for lipids, lignin, cellulose, well known: Xylem cell walls contain cellulose, hemicelluloses, pec- proteins, and pectins using confocal laser scanning and fluorescent tins, lignin, proteins, and glycoproteins, but which of these occur light microscopy for five woody plant species from across the an- right at the surface? These questions matter for two main reasons: giosperm phylogeny. All these chemical compounds are known to (1) Hydrophobic surfaces are prone to forming surface bubbles exist in or on plant cell walls, but to date it has not been clear which through heterogeneous nucleation (Blander and Katz, 1975; Ryan of these may be found right at the xylem surface in contact with and Hemmingsen, 1998), and (2) surface chemistry and roughness water, i.e., on the innermost side of vessel lumina, in pit borders and have strong effects on flow in micro-­ and nanochannels (Gruener pit membranes. The goal of this study was to provide much needed et al., 2016). data for future studies and models of water movement in vascular Gas-­saturated aqueous solutions confined in systems that are plants. under subatmospheric or negative pressure are prone to forming gas bubbles through heterogeneous nucleation on surfaces, espe- cially on small surface irregularities, such as in crevices, or surface MATERIALS AND METHODS locations with hydrophobic properties (Blander and Katz, 1975; Creech et al., 2002; Wheeler and Stroock, 2009; Belova et al., 2011; Plant species Brennen, 2014). Most attempts to maintain or transport water or aqueous solutions in artificial systems under negative pressure with- The species studied included woody plants from five major an- out bubble formation only succeed under highly controlled condi- giosperm clades (APG III, 2009) and different growth forms: tions, when solutions are completely degassed, or when surfaces are Liriodendron tulipifera L. (winter-deciduous­ tree, Magnoliaceae, treated to be completely hydrophilic and free of crevices that could Magnoliales, magnoliid clade), Triadica sebifera Small (syn. Sapium 174 • American Journal of Botany

sebiferum, winter-­deciduous tree, Euphorbiaceae, Malpighiales, transverse plane from the center of the stem segment and the distal fabid clade), Geijera parviflora Lindl. (evergreen tree, Rutaceae, surface recut using a sledge microtome (Sledge Microtome GSL 1, Sapindales, malvid clade), Distictis buccinatoria (DC.) A.H.Gentry Schweingruber, Switzerland). The stem piece was then placed onto (evergreen liana, Bignoniaceae, Lamiales, lamiid clade), and a clean microscope cover glass on top of a drop of water and ob- Encelia farinosa Torr. & A.Gray (drought-deciduous­ desert shrub, served using the inverted microscope of a confocal laser scanning Asteraceae, Asterales, campanulid clade). All grow in the Fullerton microscope (model TCS SP2, Leica Microsystems). Arboretum or on the California State University Fullerton campus Imaging was done with a 63.0 × 1.20 objective in water, using in Fullerton, California, United States. All five species will be re- four scans per image at 1024 × 1024 resolution. For this publica- ferred to by their generic names in this paper. All imaging studies tion, most images were brightness-­enhanced for the entire image were done with a single plant from each species, with 2–3 samples to make the distribution of fluorescent signals more visible, as the per plant. For logistic reasons, Liriodendron was not available for all sole purpose of these experiments was to reveal the spatial distribu- studies and was included whenever possible. tion of surface features. Brightness in these images cannot be inter- preted as reflecting the concentrations of fluorophores and/or the Confocal microscopy using vacuum infiltration of fluorophores compounds that they attach to. One exception was the brightness in images of samples treated with FM 1-­43 vs. phospholipase C (PLC) To test for cell wall surface components and coatings, we used three + FM 1-­43, which were taken at the same confocal settings and not fluorescent dyes to infiltrate living xylem, then imaged the tissue edited in any way. Images for the two treatments were compared by with laser scanning confocal microscopy. Non-­infiltrated samples calculating the mean luminosity of the image layer for FM 1-43­ for were imaged as controls for each treatment and species. FM 1-­43 each image in Photoshop (version CC 2017, Adobe Systems, San (Molecular Probes, Life Technologies, Eugene, OR, USA) is an Jose, CA, USA). amphiphilic fluorophore that is virtually nonfluorescent in water and strongly fluorescent under cyan light excitation when bound Transmission electron microscopy (TEM) to lipids, including amphiphilic lipids, and biological membranes (Jelínková et al., 2010). It was used with 476 and 488 nm excita- Stems were wrapped in wet tissue to prevent them from drying tion and an emission window of 578 to 618 nm. Direct Red 23 out, which can lead to artifacts (Li et al., 2016), and were express-­ (No. 212490, Sigma Aldrich, St. Louis, MO, USA), also known as shipped to the Jansen lab in Ulm, Germany, for transmission elec- Pontamine Fast Scarlet 4B (Anderson et al., 2010; Thomas et al., tron microscopy. Small sectioning blocks of xylem (ca. 1 × 1 × 2 2013), fluoresces red under cyan to green light excitation when mm) were fixed with glutaraldehyde (2.5% v/v glutaraldehyde, 0.1 bound to cellulose and to a much lesser degree when bound to mol phosphate, 1% w/v saccharose, pH 7.3), washed three to four hemicellulose, both of which are hydrophilic molecules. Direct times in phosphate buffer for 5 to 10 min each, and postfixed with

Red 23 has an excitation maximum of 535 nm (green light) when a 2% w/v aqueous OsO4 solution for 2 h at room temperature. The bound to cellulose (Anderson et al., 2010), but cell walls in xylem samples were then dehydrated through a gradual propanol series show substantial autofluorescence under green light excitation, (30%, 50%, 70%, 90%), which was followed by en bloc treatment so we determined that excitation with 476, 488, and 496 nm pro- with 4.5% w/v uranyl acetate in ethanol. Unsaturated phospholip- vided a much clearer signal at an emission window of 560 to 660 ids can be rendered insoluble by treatment with OsO4, which binds nm without creating much cell wall autofluorescence. NanoOrange to double bonds in the unsaturated fatty acid chains, while reten- (Molecular Probes) was used for protein detection, because it is tion of incompletely esterified phospholipids can be improved by nonfluorescent in water and fluoresces strongly under excitation at en bloc treatment with uranyl acetate solution. Further dehydra- 488 nm when bound to hydrophobic regions of proteins (Grossart tion was achieved by three treatments of 100% propanol for 30 min et al., 2000), with an emission window of 543 to 603 nm. Lignin was each. The samples were then immersed twice for 5 to 10 min in detected via autofluorescence when excited at 405 nm (Donaldson 1,2-­propylenoxide and gradually embedded in Epon resin, which and Radotic, 2013) and an emission window of 430 to 490 nm. was polymerized at 60°C for 48 h. Transverse semithin sections Solutions of all fluorophores were prepared by dissolving them were cut with an ultramicrotome (Leica Ultracut UCT), stained in nanopure water (FM 1-43:­ 5 μg/mL; Direct Red 23: 0.01%; with 0.5% toluidine blue O in 0.1 M phosphate buffer and mounted NanoOrange: 1× working solution, following the manufacturer’s on microscope slides using Eukitt. Ultrathin sections (60–90 nm) instructions). Woody stems, approximately 5–8 mm in diameter were mounted on copper grids (Athena, Plano GmbH, Wetzlar, and about 10 cm long, were cut from the plants and submerged in Germany) and observed with a JEM-­1400 TEM (JEOL, Tokyo, deionized water. About 2.5 cm were cut under water from each end Japan) at 120 kV. Digital images were taken using a MegaView III of the stem to remove most bubbles in the xylem that may have camera (Soft Imaging System, Münster, Germany). been induced by the cutting. The cut ends were then recut with new razor blades to create clean and smooth surfaces. Bark was removed Testing for phospholipids on vessel lumen surfaces from about 1 cm length at the distal end to expose the xylem. The proximal end of each stem was connected to a vacuum flask using The FM 1-­43 dye used for confocal microscopy fluoresces when tight-­fitting latex tubing around the stem inside Tygon tubing. A bound to lipids, including phospholipids. It also fluoresces weakly short piece of Tygon tubing was fitted to the exposed xylem cyl- when bound to other hydrophobic surfaces. To test for the presence inder at the distal end, using latex tubing around the cylinder to of phospholipids on vessel lumen surfaces, we flushed an aque- create a tight fit. Then 1 mL of fluorophore solution was added into ous solution of phospholipase C (0.1% Purifine PLC, DSM Food the tubing and onto the exposed xylem surface. The lab vacuum Specialties, San Diego, CA) through the xylem under vacuum be- was turned on, then off once the solution was completely absorbed fore flushing with FM 1-43­ solution (5 μg/mL). Phospholipase C into the xylem. A 5-mm-­ ­long stem piece was then cut along its (PLC) cleaves the glycerol group and fatty acid chain(s) from the February 2018, Volume 105 • Schenk et al.—The nature of vessel surfaces in angiosperm xylem • 175

phosphate and head groups, creating water-­insoluble oil and water-­ Liriodendron could not be included in these EDX studies, because soluble products, such as phosphocholine (Dowd and Gilroy, 2010). the stem samples taken during the winter turned out to be dead. The conversion of amphiphilic phospholipids to oils was expected to result in a change in brightness and possibly different patterns of FM 1-­43 fluorescence. Conversely, if no phospholipids were present RESULTS on vessel lumen surfaces, then the addition of PLC should have no effect on FM 1-­43 fluorescence. Confocal microscopy Effects of PLC on the structure and location of lipid nanoparti- cles in xylem were also tested using transmission electron micros- All xylem cell walls showed strong lignin autofluorescence, seen as copy (TEM). An aqueous solution of phospholipase C (2 mL of blue in confocal images (Figs. 1 and 2), most strongly in primary 0.1% Purifine PLC) was flushed through the xylem, and stems were walls but also in all secondary walls. Lignin in secondary vessel wrapped in wet tissue and were express-­shipped to the Jansen lab in walls appears to occur throughout the secondary wall and up to Ulm, Germany, for transmission electron microscopy and prepared the wall surface in most species, while fibers in Distictis (Fig. 1M), as described above. Encelia (Fig. 1Q), and especially Liriodendron (Fig. 1A) appeared to have thin surface layers that were rich in cellulose and low in lignin. Testing for pectins The new infiltration method to characterize xylem vessel sur- face chemistry via confocal laser scanning microscopy produced To test for pectins, we used coriphosphine O, a fluorescent dye that remarkable results and made it possible to distinguish some sur- binds to esterified pectins but not to cellulose (Weis et al., 1988; face features from signals in cell walls and living cells that otherwise Sun et al., 2008; de Oliveira et al., 2014). Unbound coriphosphine might have obscured those signals. Since the vessels observed were O is also fluorescent, which is why it needs to be rinsed from sam- located at some distance (ca. 3–5 cm, depending on the species) ples and therefore cannot be used with our vacuum infiltration from the dye injection point, the fluorophore was mainly limited to approach for confocal microscopy. We therefore used it on pre- those that were cut open at the height of the injection. Control sam- pared semithin (ca. 500 nm thick) sections of xylem studied with ples imaged at the same wavelengths as the FM 1-43­ and Direct Red fluorescent microscopy. These sections were prepared in a similar 23 treatments showed very weak fluorescence in cell walls, most way as described above for TEM, except that they were glued onto likely because lignin shows some autofluorescence when excited microscope slides without embedding in Eukitt and no toluidine with 488 nm light (Xu et al., 2006a), but the images were almost blue staining was applied. The sections were stained for 2 min with black and are therefore not shown here. 0.03% w/v aqueous solution of coriphosphine O (Sigma Aldrich), The use of Direct Red 23 fluorescent dye (shown in red) revealed washed with distilled water, and imaged under excitation with blue that intervessel and vessel-parenchyma­ pit membranes were strongly light (450 to 490 nm) with a barrier filter at 520 nm. marked for cellulose in all species, both in tangential and transverse images (Fig. 1A, B, E, F, I, J, M, N, Q, P), except for Liriodendron Energy-­dispersive X-­ray spectroscopy (EDX) that showed a positive signal in intervessel pit membranes only in tangential (Fig. 1B), but not in transverse images (Fig. 1A). Other Small, fresh xylem samples (ca. 2 mm wide, 5 mm long, and 1 vessel lumen surfaces showed only weak signals for cellulose, such mm thick) were cut with a razor blade in a tangential direction as in Distictis, Encelia, and Liriodendron (Fig. 1A, M, Q). Lumen-­ and dried at room temperature. The samples were then vacuum-­ facing surfaces of fibers were much more clearly marked for cellu- coated with 10 nm of carbon using a MED010 apparatus (Balzers lose than those of vessels, especially in Liriodendron (Fig. 1A) and Union, Balzers, Lichtenstein). Intervessel pit membrane surfaces Encelia (Fig. 1Q, R). Any of these signals may have also been caused were examined at 20 keV using an FE-­scanning electron micro- in part by Direct Red 23 binding to hemicellulose, which is also scope (S-­5200 UHR FE-­SEM, Hitachi, Tokyo, Japan) connected to highly hydrophilic and always occurs with cellulose. an EDX-­unit (HIT S-65­ 136-­5 Detecting Unit, AMETEK GmbH, Proteins were detected via NanoOrange fluorescence (shown in Wiesbaden, Germany) using the software Genesis Spectrum (ver- light yellow) in intervessel pit membranes and pit chambers and, to sion 5.211, EDAX, Mahwah, NJ, USA). Fiber or vessel wall areas some degree, lining vessel lumen surfaces in four species (Fig. 1G, without bordered pits were scanned at a similar magnification and K, O, S), but not in Liriodendron, where only weak signals were de- used as the control sample. The spectra obtained provide relative tected, mostly inside cell walls and in living cells (Fig. 1C). Intervessel amounts of chemical elements. To be sure that peaks other than pit membranes were strongly marked for protein in the other four carbon and oxygen were clearly present, we ran the spectra until species (Fig. 1G, K, O, S), most clearly in Triadica (Fig. 1G). the total counts for carbon were between 15,000 and 30,000, which The lipid probe FM 1-43­ (shown in yellow; Fig. 1D, H, L, took about 10 to 15 min at count rates between 1000 and 2000·s−1. P, T) strongly marked pit chambers, both of intervessel and

FIGURE 1. Confocal micrographs of transverse and tangential cell walls in secondary xylem after vacuum infiltration with fluorophores to detect cellulose (Direct Red 23; red in A, B, E, F, I, J, M, N, Q, R), proteins (NanoOrange; light yellow to yellow in C, G, K, O, S), and amphiphilic lipids (FM 1-­43, yellow in D, H, L, P, T) in vessel walls and bordered intervessel pits. Images B, D, F, J, N, and R are longitudinal views; all other images are transverse views. Since the secondary xylem area scanned was some distance below the injection point, the fluorophores do not occur in all conduits. Blue walls indicate lignin autofluorescence. Species abbreviations and the cell wall compounds that are stained (cellulose, proteins, lipids) are shown in the up- per right corner of each image. Lt = Liriodendron tulipifera, Ts = Triadica sebifera, Gp = Geijera parviflora, Db = Distictis buccinatoria, Ef = Encelia farinosa, V = vessel, short arrows = intervessel walls with intervessel pits. Scale bars = 20 μm. 176 • American Journal of Botany February 2018, Volume 105 • Schenk et al.—The nature of vessel surfaces in angiosperm xylem • 177

vessel-­parenchyma pits, coatings on intervessel pit membranes Triadica: 1300 ± 90 nm (n = 13), Encelia: 1009 ± 67 nm (n = 7). (Fig. 2B), but not the membranes themselves, and most vessel No pit membranes could be observed in transverse sections of lumen surfaces. By coating the pit membranes, the fluorophores Liriodendron. showed dark pit membrane shapes very clearly (e.g., Fig. 2B, C), Treatment with phospholipase C (PLC) to break up phospho- despite the limited (ca. 200 nm) resolution of these images, re- lipids into their lipophilic and hydrophilic components followed by vealing distinct pillow shapes of these fully hydrated membranes FM 1-­43 caused most confocal images of the fluorescence emission that all appear thicker than measured for fresh material directly window for the fluorophore to become much brighter (≈2× greater fixed and observed under TEM (Fig. 2A). Based on the FM 1-­ luminosity) than when using FM 1-43­ alone while maintaining the 43 images, the following estimates were obtained for pit mem- same imaging settings during a session, indicating that PLC activity brane thickness at their thickest point (mean ± standard error): affected the intensity of fluorescence. Figure 2D–G shows examples Distictis: 1827 ± 146 nm (n = 16), Geijera: 791 ± 51 nm (n = 16), of this effect in Distictis and Geijera.

FIGURE 2. Transmission electron micrograph (A), confocal micrographs (B–G), and fluorescence micrographs (H–K) of secondary xylem. Intervessel pit membranes (arrows in A–C) appear much thicker when shown fully hydrated under confocal microscopy (B, after injection with FM 1-­43 to detect lipids; C, after injection with Direct Red 23 to detect cellulose) than TEM (A). Treatment with phospholipase C (marked with PLC in upper right corner) shows brighter signals for FM 1-43­ (E, G) than without phospholipase C (D, F). Images D–G were obtained during the same microscopy session with the same settings; brightness and contrast were not adjusted. Staining with coriphosphine O (H–K) indicates the presence of pectins in primary walls and cells (long arrows) surrounding vessels (V), but no pectins in intervessel pit membranes (short arrows). Species abbreviations and the cell wall compounds that are stained (lipids, cellulose, pectins) are shown in the upper right corner of each image. Db = Distictis buccinatoria, Gp = Geijera parviflora, Ts = Triadica sebifera, Ef = Encelia farinosa. 178 • American Journal of Botany

Fluorescence microscopy DISCUSSION Because coriphosphine O could not be used for confocal micros- What do these findings mean from the perspective of xylem sap? copy, it was used for regular fluorescence microscopy of perma- What types of surfaces are actually in contact with sap? And what nent semithin sections, i.e., sections that were not hydrated. The are the potential functional implications of these findings? To an- patterns (shown in red to orange) were consistent for all species swer these questions, it is helpful to distinguish between bordered studied in that pectins were not observed on any vessel lumen pits and other vessel lumen surfaces, with functional implications surfaces, except on the outer edges (the annulus) of intervessel discussed last. pit membranes and possibly in vessel-parenchyma­ pit membranes (Fig. 2H–K). It is not possible to determine from these images whether the fluorescent signals come from the vessel-­parenchyma Vessel lumen surface characteristics pit membranes directly, only from the protective layer between Overall, our findings agree with previous studies to show that the the pit membrane and the plasma membrane, or from both. lumen-­facing vessel surface consists of a mixture of cellulose, possi- Otherwise, evidence for pectins is clearly visible for primary cell bly hemicellulose, a substantial concentration of lignin, and lipids. walls and protective layers on the vessel-facing­ sides of vessel as- Cell wall surfaces of water-­conducting cells have sometimes been sociated parenchyma cells (Fig. 2H–K), but these locations are not assumed to be hydrophilic and consisting mainly of cellulose (Cary at vessel lumen surfaces. et al., 1968; Oertli, 1971). Our data do not support that assumption for vessels, because the cellulose-­specific Direct Red 23 did not pro- Transmission electron microscopy (TEM) vide strong signals for most vessel lumen surfaces other than pit membranes (Fig. 1). Cellulose, of course, is a major constituent of Observations of lumen-­facing vessel surfaces in the study spe- all plant cell walls, making up about half or more of the composition cies show abundant black nanoparticles lining inner vessel wall, even in secondary cell walls (Meier, 1985). Cellulose signals from pit membranes, and pit chamber walls, but also other vessel lu- Direct Red 23 may have been masked by embedded lignin (Thomas men surfaces, as shown by Schenk et al. (2017). The distribution et al., 2013). Direct Red 23 also binds to hemicelluloses (Anderson of these black nanoparticles is the same as that shown for lipids et al., 2010), which almost always occur with cellulose, crosslink- (Fig. 1D, 1H, 1L, 1P, 1T, 2A, 2B). High magnification reveals these ing microfibrils. We did not specifically look for hemicellulose in particles to be aggregates of different shapes, sometimes more this study, but other studies have found that xyloglucans and xy- or less circular and sometimes very irregular (Fig. 3C, F, H, J). lans are much more abundant than mannans in secondary walls They are not forming continuous layers but due to their location of vessels (Kaneda et al., 2010; Kim and Daniel, 2012), which was next to vessel lumen and pit membrane surfaces could be bro- confirmed for pit borders of poplar (Herbette et al., 2015), where ken-­up remnants of continuous layers. Pit membranes in vessels xylans and xyloglucans were detected at both the lumen-facing­ and that were in their final developmental stages could be observed pit-­chamber-­facing surfaces. in Distictis and showed a bright pit membrane with a somewhat Lignin autofluorescence appeared to extend all the way to the darker inner area (Fig. 3A). Remnants of the cytoplasm were seen inner cell wall surface in most vessels (Fig. 1). Previous studies have in pit borders and near inner vessel walls, which showed no dark found the S3 layer to have the same lignin concentration as the S2 lining compared to fully developed vessels (e.g., compare Fig. 3A layer in birch (Saka and Goring, 1985), be relatively low in lignin with 3B). in beech (Fromm et al., 2003), and appearing to have a different Intervessel pit membranes after PLC treatment become generally lignin composition than the S2 layer (Gierlinger, 2014), such as more electron dense and greyer (Fig. 2A, 3D, 3G, 3L). The charac- having a high concentration of the dibenzodioxocin substructure of teristic dotted appearance of black nanoparticles disappears entirely lignin in birch (Kukkola et al., 2004). Lignin was detected through- (Fig. 3G, L) or partly (Fig. 3K), and the pit membranes show a more out the secondary walls making up pit borders of poplar, including homogeneous, blurry appearance in PLC treated samples. the lumen-­facing and pit-­chamber-­facing surfaces (Herbette et al., 2015). While the middle lamella and primary wall are composed of Energy-­dispersive X-­ray spectroscopy (EDX) more than 50% lignin, about 20% of the secondary walls in vessels is lignin (Donaldson, 2001) As expected from cell walls containing polysaccharides, EDX Interestingly, a very thin, continuous layer of lipids was ob- spectra for the surfaces of pit membranes showed high peaks served on vessel lumen surfaces in dye-­injected vessels of all spe- for carbon (0.30 keV) and oxygen (0.55 keV) for all four species cies (Fig. 1D, H, L, P, T D). The brightness of that layer was greatly (Fig. 4; Liriodendron data were not available). All spectra also enhanced by treatment with phospholipase C (Fig. 2E, G), which showed clear peaks for potassium (3.30 keV), calcium (0.371 cleaves phospholipids into their lipophilic and hydrophilic compo- keV), and chlorine in the case of Triadica, Geijera, and Encelia, nents. Therefore, almost certainly, the lipid layer on vessels at least but not Distictis. All these elements are abundant ions in xylem partially consists of phospholipids, which have been found in xy- sap. Distictis, Geijera, and Encelia also showed phosphorus peaks lem sap (Gonorazky et al., 2012; Schenk et al., 2017). Under TEM, (2.01 keV) in intervessel pit membranes (Fig. 4A, C), but either this same lipid layer along vessel walls appears broken up into black no or a very small, insignificant phosphorus peak in fiber or vessel OsO4-­stained nanoparticles (Fig. 3C, H, J), as seen in previous stud- wall area without bordered pit membrane (Fig. 4B, D). A phos- ies for a wide variety of vascular plants (Pittermann et al., 2010; phorus peak was unclear in intervessel pit membranes of Triadica, Tixier et al., 2013; Brodersen et al., 2014; Dute et al., 2014; Li et al., which showed an aluminum peak instead that did not appear in 2016; Schenk et al., 2017). This aggregation of lipids into nanopar- any other species. Encelia was the only species to show a clear sul- ticles is unlikely to be the configuration under natural, fully hy- fur peak (Fig. 4A, B). drated conditions. Preparation of xylem samples for TEM involves February 2018, Volume 105 • Schenk et al.—The nature of vessel surfaces in angiosperm xylem • 179

FIGURE 3. Transmission electron micrographs of intervessel pits and their pit membranes after fixation with glutaraldehyde and osmium tetroxide. Samples were either untreated (A–C, E–F, H–J) or treated with phospholipase C (D, G, K, L; marked with PLC in upper right corner), which cleaves phos- pholipids into water-­insoluble oil and water-­soluble products. All bordered pits and pit membranes have an axial orientation. A bordered pit with a developing pit membrane before cell death is shown in (A). Dark, electron dense pit membranes with nanoparticles occur in the non-PLC­ treated samples (C, E, F, H, I, J). Pit membranes are more homogeneously grey to dark, with little to no granular structure in PLC-treated­ samples (D, G, arrows in K, L). Db = Distictis buccinatoria, Ef = Encelia farinosa, Gp = Geijera parviflora, Ts = Triadica sebifera. propanol dehydration and ethanol/uranyl acetate treatment, which during dehydration. Lipid micelles are likely to be altered and re- could have strong effects on phospholipids lining lumen surfaces versed, causing the hydrophilic head groups to face inward and fatty in the apoplast, because ethanol is well known to affect structures acids to face outward (Correa et al., 2012). Dehydration therefore made up of phospholipids (Gurtovenko and Anwar, 2009). Lipids would explain the broken-­up nature of the lipid layer along sur- lining hydrophobic surfaces, with their hydrophilic head groups faces under TEM, which suggest that microscope techniques that facing the sap, would probably be detached from those surfaces preserve hydration are better tools for studying the ultrastructure 180 • American Journal of Botany

FIGURE 4. Emission spectra based on SEM-EDX­ analysis of intervessel pit membranes (A, C) and fiber wall area without bordered pit membranes (B, D). High peaks for carbon (C) and oxygen (O) and clear peaks for potassium (K), calcium (Ca) and chlorine (Cl) were present. Phosphorus (P) peaks occurred for intervessel pit membranes of Encelia farinosa (A), and Geijera parviflora (E), but were very small or absent in fiber walls (B, D) of these spe- cies. Encelia farinosa had a peak for sulfur (S). Peak heights denote the relative amount of the elements within a sample, but they cannot be compared directly between samples. of this layer. Super-resolution­ microscopy would thus be required species studied, about 250 nm when measured using TEM, whereas (Schubert, 2017). the other four species have pit membranes thicker than 500 nm No continuous layer of proteins was observed on any vessel lu- (Schenk et al., 2017). Pit membranes of Liriodendron were hardly men surfaces, but most species showed some evidence for protein-­ visible in any transverse section and not clearly marked with any containing structures close to vessel walls (e.g., Fig. 1C, K, S). These fluorophore. Despite the limited resolution, the infiltration approach do not appear to be surface features but may be proteins carried revealed some very clear patterns for the surface chemistry of pit in xylem sap, which are often quite abundant (Buhtz et al., 2004). membranes. Not surprisingly, intervessel and vessel-­parenchyma There was no evidence for pectins on any vessel lumen surface pit membranes were strongly marked for cellulose with Direct Red (Fig. 2H–K). 23 (Fig. 1B, E, F, I, J, M, N, Q, R), confirming the previous consen- sus that these structures consist mainly of cellulose (Choat et al., Surfaces of bordered pits 2008). There was no evidence for lignin autofluorescence from any pit membranes. Lignin has been reported for mature intervessel pit Standard confocal microscopy is at its resolution limit for imag- membranes of a few angiosperms (Schmitz et al., 2012; Herbette ing pit membranes, and this limitation was apparent especially et al., 2015), but those reports are not conclusive and could have for Liriodendron, which has the thinnest pit membranes of all five been caused by accumulation of other phenolic molecules in pit February 2018, Volume 105 • Schenk et al.—The nature of vessel surfaces in angiosperm xylem • 181

membranes. Phenolics are known to occur in xylem sap and have potassium, and calcium, chloride, phosphorus, and sulfur are been attributed to pathogen defense (Potter and Fagerson, 1992; commonly found in xylem sap. The phosphorus peak observed in Wallis and Chen, 2012). Distictis, Geijera, and Encelia could be from inorganic phosphate or Except for the outer rim, the so-­called annulus, intervessel pit organic molecules that contain phosphate groups, including phos- membranes also did not contain any pectins (Fig. 2H–K), again pholipids (Gonorazky et al., 2012; Schenk et al., 2017) or phospho- confirming the findings of multiple previous studies (Wydra and choline (Martin and Tolbert, 1983). The sulfur peak in Encelia was Beri, 2007; Arend et al., 2008; Plavcová and Hacke, 2011; Kim and remarkable for being apparent only in that one species; it could stem Daniel, 2013; Herbette et al., 2015; Klepsch et al., 2016). Images of from inorganic sulfate or possibly from proteins. The pronounced vessel-­parenchyma pits are inconclusive for pectins, because pit aluminum peak in Triadica was a surprise, because the species is membranes cannot be distinguished from protective layers at that not known to be an aluminum accumulator. Some species formerly resolution (Fig. 2H–K), but some of the pectin signal likely orig- placed in the Euphorbiaceae accumulate aluminum (Jansen et al., inates from the vessel-­parenchyma pit membrane itself, as shown 2002), but none of these are related to Triadica. previously for peach and poplar (Wisniewski and Davis, 1995; Arend et al., 2008). Possible functional implications of the findings Proteins were evident in all intervessel pits, except for Liriodendron. Due to resolution limits, it is not easy to see whether The most obvious implication of vessel lumen surface chemistry is proteins coat the pit membranes or are embedded inside the mem- for the nature of interactions with water molecules. Our findings branes, but faint, central black lines in pit chamber images of suggest that the lumen surface consists of a mixture of cellulose and Distictis (Fig. 1O) and Geijera (Fig. 1K) support coating, while the lignin, consistent with previous studies (Saka and Goring, 1985; strong marking of entire pit membranes in Triadica suggest that the Fromm et al., 2003; Kukkola et al., 2004; Gierlinger, 2014; Herbette proteins may be embedded in those membranes (Fig. 1G). The res- et al., 2015). This result implies that the lumen surface is neither olution in the Encelia image (Fig. 1S) is insufficient for a conclusion. completely hydrophilic (i.e., zero contact angle) nor completely The accumulation of proteins at intervessel pit membranes is hardly hydrophobic (contact angle >90°). Previous observation of contact surprising because proteins are quite abundant in xylem sap (Buhtz angles between water and lumen surfaces of xylem conduits range et al., 2004) and likely to accumulate at pit membranes (Neumann from 40 to 90° with the most commonly observed values between et al., 2010). The proteins observed in intervessel pit membranes of 40 and 55° (Schneider et al., 2000; Zwieniecki and Holbrook, 2000; Triadica are likely to be part of the thick membrane coatings ob- van Ieperen et al., 2001; Brodersen et al., 2010; Kim and Lee, 2010; served in this species via atomic force microscopy (Pesacreta et al., Lee et al., 2013; McCully et al., 2014; Knipfer et al., 2016). Purified 2005). It is unclear whether these thick coatings may be unique to lignin has a contact angle somewhere between 46 and 56° (Notley this species or perhaps the family Euphorbiaceae and what their and Norgren, 2010). Some have reported a pronounced variabil- functions may be. They certainly do not consist of pectin (Fig. 2A) ity of wetting angles either spatially or depending on whether the as suggested by Pesacreta et al. (2005). water was advancing on the surface or receding (Brodersen et al., Probably the most remarkable finding is the observation of 2010; Kim and Lee, 2010; Lee et al., 2013; McCully et al., 2014; abundant lipids on intervessel pit chamber walls and coated onto Knipfer et al., 2016), which may have been caused by the presence intervessel pit membranes (Fig. 1D, H, L, P, T). Lipids were also of surfactants on the lumen surfaces (McCully et al., 2014). Contact observed in locations that are likely to be vessel-­parenchyma pits, angles matter for xylem refilling after embolism (Zwieniecki and but the images were not focused on those locations, so the evi- Holbrook, 2000; Rolland et al., 2015), but in functional xylem, con- dence for those pits is somewhat more ambiguous. All four spe- tact angle also affects heterogeneous gas bubble nucleation on sur- cies (Liriodendron was not available) show black pit membranes faces (Schenk et al., 2017). This effect is relatively minor if the angle clearly coated with lipids on both sides. Phospholipase C treatment is much below 90° (Wheeler and Stroock, 2009); for example, a 45° enhanced the fluorescent FM 1-43­ signal in pit chamber (Fig. 2E, contact angle would lower the energy requirement for nucleating a G), indicating that these lipids include phospholipids. The confocal bubble on a surface compared to a perfectly hydrophilic surface by images do not show lipids inside the mostly pillow-shaped­ interves- only about 6% (Blander and Katz, 1975). sel pit membranes, which raises the possibility that the lipid nan- Contact angles in pit membrane pores have a much stronger oparticles observed inside pit membranes via TEM (Fig. 4) could effect on bubble formation via air seeding through the pores be caused by the ethanol dehydration treatment for TEM, possibly (Zimmermann, 1983; Schenk et al., 2015). Our finding that pit shrinking the pit membranes from the apparent thick pillow shapes membranes consist mainly of cellulose support the widespread as- seen under confocal microscopy to the more or less straight shapes sumption that the contact angle in these pores may be close to zero seen under TEM, and drawing lipids into the drying pores of the (Crombie et al., 1985; Comstock and Sperry, 2000; Choat et al., 2003; membranes. The other possibility is that the lipid concentration in- Cochard et al., 2009; Plavcová et al., 2011; Brodersen et al., 2014; side membranes was too low to create fluorescent signals or that Capron et al., 2014; Johnson et al., 2014), but lipids in pit membrane the fluorophore could not penetrate membrane pores. Previously, pores could affect the contact angles there. Additionally, the surface we initially proposed that lipids would not penetrate into pit mem- chemistry of nanopores has major impacts on liquid flow through branes (Schenk et al., 2015) and then concluded that they do, based the pores (Calabrò et al., 2013; Gruener et al., 2016), which caused on TEM observations (Schenk et al., 2017). Because of the reso- Meyra et al. (2007) to suggest that pit membrane pores could not be lution limits of our current confocal studies, the question remains completely hydrophilic or hardly any flow would be possible due to open and will require super-resolution­ microscopy of hydrated xy- the high viscosity of water in the boundary layer of a hydrophilic lem to resolve. surface. Our current study throws little light on that question. Both The elemental composition of pit membranes (Fig. 4) had few proteins and lipids could occur in pit membrane pores (Fig. 1G, H, surprises; carbon and oxygen are components of cellulose and K, L, O, P, S, T) and affect the flow characteristics there. 182 • American Journal of Botany

The finding of abundant lipids on vessel lumen surfaces and pit ACKNOWLEDGEMENTS membranes raises the question of their effects on surface tension (Schenk et al., 2015, 2017). A coating of phospholipids on hydro- The research was funded by an EAGER grant from the National phobic vessel walls would render the surface hydrophilic, but it Science Foundation (IOS-­1558108). Many thanks to two anony- would do so while greatly lowering surface tension. The energy re- mous reviewers for their helpful comments, Steve Karl (Nano-­ and quirement for nucleating a bubble on a surface is a function of the Micro-­Visualization Lab, Cal State Fullerton) for help with the con- surface tension cubed (Blander and Katz, 1975; Chen et al., 2016; focal imaging, and David Walsh and Arjen Sein at DSM for provid- Schenk et al., 2017). For example, the reduction of surface tension ing the Purifine PLC. The authors thank the Electron Microscopy from that of pure water (72 mJ·m−2) to a third of that (24 mJ·m−2), Section of Ulm University for preparing TEM samples and Dr. A. a typical equilibrium surface tension for phospholipids (Lee et al., Ziegler for assistance with the SEM-­EDX analyses. 2001), would reduce the energy requirement for bubble nucleation by a factor of 27. This reduction would make surface bubble nucle- LITERATURE CITED ation vastly more likely than considered before. Schenk et al. (2017) provided a hypothesis for why resulting surfactant-coated­ bubbles Anderson, C. T., A. Carroll, L. Akhmetova, and C. Somerville. 2010. Real-­time may not expand into embolisms: Low surface tension causes much imaging of cellulose reorientation during cell wall expansion in Arabidopsis smaller bubbles to form (Winkel et al., 2004), which are stable under roots. Plant Physiology 152: 787–796. negative pressure. As such, lipids are hypothesized to prevent con- APG III. 2009. An update of the Angiosperm Phylogeny Group classification for stant hydraulic failure when xylem transport system experiences the orders and families of flowering plants: APG III. Botanical Journal of the negative pressure. The main point here is that the lipids clearly exist Linnean Society 161: 105–121. on inner walls of conduits and therefore require an explanation that Arend, M., M. Muninger, and J. Fromm. 2008. Unique occurrence of pectin-­ is consistent with the massive evidence supporting the cohesion– like fibrillar cell wall deposits in xylem fibres of poplar. Plant Biology 10: tension theory (Dixon and Joly, 1895; Dixon, 1914). 763–770. No pectin is found in intervessel pit membranes, except on the Askenasy, E. 1895. Ueber das Saftsteigen. Verhandlungen des Naturhistorisch-­ outer edge of the membrane (Fig. 2). Because this finding agrees medizinischen Vereins zu Heidelberg, N.F. 5: 325–345. Belova, V., D. A. Gorin, D. G. Shchukin, and H. Möhwald. 2011. 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