Introduction
The extracellular matrix (ECM) of plants is composed of highly complex domains of biopolymers, enzymes, ions and water that integrate into diverse architectural designs and form an effective interface between the cell surface and external environment. The matrix also dynamically modulates throughout the life of the cell in response to both internal genetic programs and external signals/stresses. The plant ECM may be dissected into two domains, the cell wall, and “components deposited onto the cell wall surface” and/or secreted beyond the wall. The cell wall can be described as a fiber reinforced hydrogel (Ali and Traas, 2016) consisting primarily of polysaccharides and proteoglycans. The main load-bearing component of the wall is crystalline cellulose organized in approximately 3 nm-wide microfibrils that are cross-linked by, and embedded in, a matrix of neutral polysaccharides (hemicelluloses), acidic polysaccharides (pectins) and proteoglycans (extensins, AGPs) (Braidwood et al., 2014; Cosgrove, 2014). The cell wall most often contains various ions that contribute to its structural integrity (Shomer et al., 2003) and in secondary cell walls, lignins generally impregnate the wall infrastructure (Marriott et al., 2016; Özparpucu et al., 2017). External to the wall exists a highly diverse assortment of biochemical components including lipid cuticles (Yeats and Rose, 2013), post-germination seed gels (Francoz et al., 2015) and root mucilages (Walker et al., 2003). The functions of the ECM are of paramount importance to the plant as it provides physical and chemical protection, a communication conduit between cells and their biotic/abiotic surroundings, a means to hold water (i.e., desiccation avoidance), a turgor-balancing control for cell expansion and a highway for translocation of water and minerals (Lionetti and Métraux, 2014; Bidhendi and Geitmann, 2016; Chebli and Geitmann, 2017). It is therefore not surprising that plant cells expend large amounts of photosynthetic carbon and devote up to 30% of their genetic machinery to the synthesis of the ECM.
Many components of the ECM found in embryophytes or “land plants” are also found in the charophyte green algae (or basal Streptophytes), i.e., the group of extant green algae whose ancestors gave rise to land plants between 450 and 500 million years ago (Becker and Marin, 2009; Leliaert et al., 2012; Delwiche and Cooper, 2015). Recent biochemical and immuno-binding studies have demonstrated the presence of cellulose, xyloglucans, ß,1-3/1-4 glucans, pectins, extensin, AGP and even lignin-like components in the cell walls of charophytes (Sørensen et al., 2011; Domozych et al., 2012; De Vries et al., 2017; Herburger et al., 2018). Additionally, many charophytes secrete large amounts of complex extracellular polysaccharides (EPS) beyond their cell walls (Kiemle et al., 2007) that are involved in photomotility, substrate attachment, anti-desiccation and biofilm formation (Boney, 1981; Oertel et al., 2004; Domozych et al., 2005; Domozych and Domozych, 2008). However, we are only just beginning to decipher the detailed structure, location and function of the specific components of the charophyte ECM. Further studies would yield valuable insight into the molecular and cellular adaptations that may have occurred during the emergence of green plants onto land and the subsequent proliferation of plants into diverse terrestrial habitats. Furthermore, many charophytes are very attractive model organisms for ECM studies as their small size, rapid growth rates and ease in laboratory culture manipulation provide superb systems for experimental analyses at all levels ranging from transcriptome analysis (Rippin et al., 2017; Van De Poel et al., 2016) to cell wall composition (Brosch-Salomon et al., 1998; Lütz-Meindl and Brosch-Salomon, 2000; Eder and Lütz-Meindl, 2008; Eder et al., 2008; Proseus and Boyer, 2008; Domozych, 2014; Domozych et al., 2014, 2016; Boyer, 2016; Lütz-Meindl, 2016) and ecology.
Adhesion is a phenomenon that is essential for most living organisms. This process is central to a diverse array of biological processes ranging from the establishment of multicellularity (Grosberg and Strathmann, 2007; Olson and Nedelcu, 2016) to the formation of a sessile habit (Morrison et al., 2009) and to the development of biofilms (Jain et al., 2007). While some adhesion phenomena require specialized cells for sustaining close contact with a substrate (e.g., rhizoids), most are directly associated with the structural architecture and dynamic modulation of the ECM. Cell adhesion molecules or CAMs are the specific components of the ECM, typically proteins or polysaccharides that are integral to the adhesion process. Their unique biochemical and structural properties are often precisely designed for interaction with a particular substrate. In plants, CAMs are components of the ECM that work in close concert with a substrate, cell wall polymers, the plasma membrane and the cortical cytoskeleton/cytoplasm (Baluška et al., 2003; Huang et al., 2016; Langhans et al., 2017). Adhesion phenomena are also commonly found in charophytes and are perhaps best exemplified by the cell/thallus establishing close contact with, and attaching to, a stationary substrate. This is a critical step for transition from the planktonic to sessile state, maintaining an optimal position for light acquisition for photosynthesis and/or living in a complex biofilm community in a frequently changing habitat like an ephemeral wetland, i.e., common homes of both modern day and ancient charophytes.
Recent studies have demonstrated that AGPs are integral to many adhesion-based mechanisms (Groot et al., 2003; Bowling and Vaughn, 2008; Huang et al., 2016). AGPs represent a diverse group of highly glycosylated proteins that are found in the cell wall, the apoplastic space and/or the medium surrounding plant cell. They are widely distributed throughout the major taxa of land plants (Šamaj et al., 1999b; Bartels and Classen, 2017; Bartels et al., 2017; Johnson et al., 2017b; Ma et al., 2017). AGPs belong to a large group of hydroxyproline (hyp)-containing proteins and possess a protein backbone consisting of repeating subunits that contain hyp/proline (pro), alanine (ala), and serine (ser)/threonine (thr) (Johnson et al., 2003, 2017a,b; Lamport et al., 2014, 2018; Herve et al., 2016; Bartels et al., 2017). The hyp is often O-glycosylated with ß 1–3 or ß1–6 or ß 3–6 galactan chains to which are attached other sugars including glucuronic acid. The carbohydrate component of AGPs is remarkably diverse amongst different taxa and often constitutes up to 99% of the whole molecule. Recent work has also shown that AGPs may be structurally and/or functionally associated with other polymers leading to the supposition that they may be part of large polymer domains of the cell wall that include pectins and xylans (Tan et al., 2012, 2013). The functions of AGPs are many and diverse. In addition to their roles in adhesion (Šamaj et al., 1999a,c; Huang et al., 2016; Langhans et al., 2017), they also contribute to cell expansion (Yang et al., 2007) pollen tube dynamics during fertilization (Nguema-Ona et al., 2012; Pereira et al., 2015, 2016; Lamport et al., 2018) cell expansion (van Hengel and Roberts, 2002; Seifert and Roberts, 2007), calcium dynamics during development (Majewska-Sawka and Nothnagel, 2000; Oxley and Bacic, 2000; Lamport and Varnai, 2012; Lamport et al., 2018), salt tolerance (Olmos et al., 2017), and many others (Seifert and Roberts, 2007; Ellis et al., 2010).
AGP-like proteoglycans have been demonstrated in charophytes (Sørensen et al., 2011; Ruiz-May et al., 2018) including desmids (Lütz-Meindl and Brosch-Salomon, 2000; Domozych et al., 2007; Eder et al., 2008) and Chara (Domozych et al., 2010, 2009) and have been implicated in various cellular events including adhesion. However, knowledge of their distribution amongst the various charophyte taxa and inclusive morphotypes is limited and their roles in adhesion are poorly resolved. In this study, we examined five taxa from three major groups of charophytes, the early divergent Chlorokybales and the late divergent Coleochaetales and Zygnematales that exhibit distinct adhesion phenomena. Employing a variety of labeling protocols and experimental techniques, we show that AGP-like macromolecules are involved in various adhesion phenomena.
Materials and Methods
General
Live cultures of algae were maintained at 20oC, 16 h light/8 h dark with 74 μmol photons m-2 s-1 of cool white fluorescent light in liquid cultures containing the following media: Penium margaritaceum Skidmore College Collection: SKD-8 (Woods Hole Medium with 5% soil extract; Domozych et al., 2017), Chlorokybus atmophyticus UTEX 2591 (Woods Hole Medium with 10% soil extract), Zygnema circumcarinatum SAG 2419 (Herburger et al., 2019, for this study grown in 3N BBM1), Spirogyra sp. Carolina Biological 152525 (3N BBM) and Coleochaete orbicularis UTEX 2651 (Woods Hole medium with 5% soil extract and 1% peat extract). Cells were harvested for labeling and experiments 10–14 days after subculturing, in the case of Zygnema also older cultures (3–6 months) were used for transmission electron microscopy. Cells were concentrated by centrifugation at 700–1,000 ×g for 1 min. Washing consisted of resuspending centrifuged pellets in fresh growth medium, shaking and recentrifuging. This step was repeated three times before labeling.
Wound Response of Rhizoids
Spirogyra sp. filaments were removed from culture and placed on the bottom of a sterile plastic petri dish. The filaments were chopped to small fragments with a sterile razor blade. Masses of chopped filaments were then added to a sterile petri dish containing 3N BBM with sterile 22 × 22 mm coverslips lining the bottom. The petri dishes were cultured as described above. Within 24 h rhizoids emerged from the wounded filaments and attached to the coverslips. The coverslips containing the rhizoids were used for labeling. A similar wounding protocol was employed for Zygnema circumcarinatum but no rhizoids or adhesion to coverslips were observed.
Fluoresbrite Bead Labeling
The following protocols were employed in order to screen for adhesive ECM components. Cells/thalli that were either attached to a surface (e.g., glass coverslip, plastic petri dish) or planktonic were collected and washed with fresh growth medium in order to remove any pre-existing materials from the cell surface that might interfere with subsequent experiments. They were then incubated in a solution of 50 μL 0.5 μm Fluoresbrite beads (Polysciences, United States)/mL growth medium for 15 min with gently shaking. Cells/thalli or substrates with attached algae were washed 3× with fresh growth medium to remove excess beads. Cells or substrates were mounted on glass slides and viewed with wide field fluorescence labeling (WFLM) equipped with a FITC filter set. For Spirogyra, coverslips containing rhizoids were collected, washed in a stream of fresh growth medium and then incubated in the Fluoresbrite solution. Cells were also placed on plastic petri dishes (Penium, Chlorokybus) were allowed to settle onto the surface. They were then labeled with the Fluoresbrite beads or cells were gently removed with a fine stream of growth medium and the sites of attachment were incubated in drops of the beads. All samples were viewed with an inverted light microscopy (LM) equipped with a FITC filter set.
ß-Glucosyl-Yariv Reagent Labeling and Growth Inhibition Analyses
Washed cells or coverslips were incubated for 60 min in 10 and 20 μM ß-glucosyl-Yariv (BioSupplies, United States), washed with growth medium and examined with LM. As a control, algae were labeled with β-galactosyl Yariv and imaged. This reagent does not bind to AGP. For inhibition experiments, cells or wound induced filaments were cultured in growth medium containing 20 μM ß-glucosyl-Yariv (or β-galactosyl Yariv as a control). Cells were collected 24, 36, and 96 h after treatment and incubated with Fluoresbrite beads or allowed to settle on the base of a plastic petri dish or glass coverslip to observe adhesion properties. For Spirogyra sp. rhizoid analysis, cut filaments were placed in petri dishes with coverslips as described above and cultured in the Yariv reagents. Examination of adhesion efficacy was then monitored by LM.
Immunofluorescence Labeling
Harvested cells were washed three times with fresh growth medium and labeled for immunofluorescence as described in Domozych et al. (2014, 2017). The primary antibodies used were obtained from Plant Probes (Leeds, United Kingdom2) and included JIM5 (specificity: Homogalacturonan, HG, with low degree of esterification), JIM13 (sp: β-D-GlcpA-(1 → 3)-α-d-Galp A-(1 → 2)-l-Rha), JIM8 (sp: AGP), LM2 (AGP with ß-glucuronic acid), and LM6 (sp: (1 → 5)-alpha-arabinan/AGP epitopes). Primary antibodies were diluted 1/10 with growth medium before labeling of the algae. The secondary antibody used was goat-anti-rat TRITC (Sigma Chem. St. Louis, MO, United States) diluted 1/75 with growth medium. Control labeling was performed without primary antibody application. For Spirogyra rhizoids, coverslips containing rhizoids (above) were labeled by placing drops of antibody and washes onto the surface of the coverslips for the times indicated above.
For quantification of fluorescence signal of JIM13, we use three independent biological replicates to measure the fluorescence intensity using the transect function of the Olympus Fluoview 300 CLSM software. The fluorescence signal was estimated using six random segments in each biological sample. Equal threshold level was applied for each replicate to allow comparison among taxa. Data were analyzed with ANOVA in JMP version 12.0. Differences in the mean of fluorescence intensity signals were statistically tested using Tukey’s test (significance P < 0.05).
Light and Confocal Laser Scanning Microscopy
For WFLM, samples were viewed with an IX-83 LM (Olympus) or BX 60 LM (Olympus). Samples for confocal laser scanning microscopy or CLSM were imaged with an Olympus Fluoview 300 CLSM. For TRITC imaging, we used the HeNe-G (Green/Red) laser, with 568nm excitation and 50% laser intensity. For samples treated with FITC and chlorophyll autofluorescence, we used the Ar laser with 488nm excitation and 10% laser intensity. For both, we used U-PLAN FL 20X/0.50 or UplanApo 60X/1.40 oil objectives. For samples analyzed with the CLSM, we chose the pseudo-colors for fluorescent signals that best high the labeling.
Immunogold Labeling
Harvested cells were cryofixed, freeze substituted and processed for transmission electron microscope (TEM) using the methods described in Domozych et al. (2014). Spray freezing into liquid propane was employed for Penium and Chlorokybus. For multicellular taxa (Spirogyra, Coleochaete) samples were placed on 1 × 1 cm aluminum sheets, quickly blotted with filter paper to remove most growth medium and plunge frozen into liquid propane cooled with liquid nitrogen. Zygnema was frozen by a Leica EMPACT High pressure freezing (HPF) device and freeze substituted (Leica EM AFS) in 0.05% Ur-acetate and 0.1% OsO4 and embedded in London resin (LR) white according to Herburger and Holzinger (2015) and Holzinger and Pichrtová, 2016. For Spirogyra rhizoids, chopped filament fragments (see above) were allowed to settle onto sterile nitrocellulose sheets immersed in growth medium in a sterile petri dish. After rhizoid formation the sheets were placed on aluminum sheets as above and plunge frozen. 60 nm sections were cut with a Leica Ultramicrotome and collected on Formvar-coated nickel grids. Immunogold labeling followed the protocol of Domozych et al. (2017) or Herburger and Holzinger (2015) for Zygnema. Samples were imaged on a Zeiss Libra 120 TEM.
ECM Isolation
For cell wall studies, cells were harvested by centrifugation 1,000 ×g for 1 min. The cell pellet was washed three times in fresh growth medium and re-centrifuged. Except for Penium, Alcohol Insoluble Residues (AIR) of cell walls were obtained using the technique of Foster et al. (2010). Penium cell walls were isolated via sonication/centrifugation method of Domozych et al. (2014). For EPS isolation, the growth medium of 10–14 day old cultures of Penium, Chlorokybus and Zygnema was collected by centrifugation at 3,000 ×g for 5 min to remove particulate materials. The supernatant was then recentrifuged at 10,000 ×g for 5 min and the supernatant was dialyzed against 4 l of distilled H2O (dialysis tubing cut off mw 3,000). The water was changed every 8 h for 2 consecutive days. The dialyzed supernatant was then collected and freeze dried.
Protein Electrophoresis and Western Blotting
Freeze-dried samples were dissolved in deionized water (1 mg/mL) and mixed with 2× Laemmli Sample Buffer (Bio-Rad, Hercules, CA, United States) with 5% (v/v) β-mercaptoethanol. The samples were vortexed, boiled in a water bath for a total of 2 min, cooled on ice, and centrifuged at 12,000 rpm for 2 min. Twelve microliter aliquots of samples were loaded into wells of a Mini-Protean® TGXTM gel 4–20% (Bio-Rad, Hercules, CA, United States). Twelve microliter of the Precision Plus Protein unstained standards (Bio-Rad; cat #161-0363) were loaded as a control. Electrophoresis was performed in a Tris/Glycine/SDS buffer (Bio-Rad, Hercules, CA, United States) at 100 V, 1 A for 5 min, and then at 125 V, 1.25 A for 55 min.
The separated proteins were transferred onto a nitrocellulose membrane (NitroBind 0.45 μm, GVS North America) by electroblotting using a Trans-Blot® SD Semi-dry Electrophoretic Transfer Cell (Bio-Rad, Hercules, CA, United States) at 12 V, 0.3 A, 2 h in Towbin transfer buffer (25 mM Tris, 192 mM glycine, 20 % (v/v) methanol, pH 8.3). The gel was placed in a box, and with constant gentle shaking, washed with water 3 × for 5 min, developed with Bio-SafeTM Coomassie G-250 stain (Bio-Rad, Hercules, CA, United States) for 1 h, washed with water 3 × for 5 min, and de-stained overnight in water, until photographed.
The membrane was transferred to a plastic box. The Western blot steps were performed at RT, with constant gentle shaking. The membrane was incubated with a blocking solution containing 0.5% (w/v) blotting grade blocker (Bio-Rad) in PBST for 30 min, washed with PBST 3× for 5 min, and incubated overnight with 1:10 solution of JIM13 mAb with PBST. After a 5 min wash for 3× with PBST, the membrane was blocked, rewashed, and incubated for 90 min with a 1:50 solution of goat anti-rat IgG-peroxidase antibody (Sigma, St. Louis, MO, United States). After washed for 10 min 3× in PBS, the membrane was developed with a colorimetric detection Amplified Opti-4CNTM Detection Kit (Sigma, St. Louis, MO, United States) and photographed.
Conclusion
Adhesion is a common phenomenon exhibited by charophytes. To be able to stick on a surface is critical for many functions ranging from holding cells together in a multicellular thallus and more often for attaching to substrates. A common attribute of most early divergent organisms is adhesion to a substrate as they establish and maintain their existence in a biofilm, i.e., an interactive community of diverse taxa that enhances growth and development. Fluorescence quantification of JIM13 labeling yield significant differences between two filamentous members of Zygnematophyceae; Spiorogyra with the lowest signal, while Zygnema showed the highest signal. These results can possibly be interpreted by the ecological background where Zygnema forming extensive mats (e.g., Holzinger et al., 2009) might benefit from sticking the filaments together, a feature not necessary in the strictly aquatic Spiorgyra. From the other examined algae, Chlorokybus, Penium, and Coleochaete, only the latter was significantly different from Zygnema in the fluorescence intensity.
For extant and ancient charophytes, a rapid and effective adhesion mechanism would be especially critical in highly changeable wetlands. Additionally, adhesion of the thallus to a stable position on a substrate would enhance light absorption for photosynthesis and also create a direct physical conduit for capillary water movement from the substrate that would combat desiccation. Recent work has shown that charophytes live in complex biofilm communities (Domozych and Domozych, 2008) and a rapid adhesion mechanism would be critical for establishment of these communities. Further work will be required to elucidate the biology of extant charophyte-associated biofilms that would also provide insight into the role of these communities in the invasion of ancient terrestrial habitats.
This study demonstrates that the adhesion mechanism in various charophyte taxa is a result of the positioning of CAMs in the ECM at specific sites of the thallus or cell. This includes the sheath surrounding rhizoids or the thallus and the outer layer of the cell wall. After surveying a diverse taxonomic assemblage of charophytes, we provide evidence that indicates that AGPs constitute one part of the components that are secreted and used in adhesion phenomena. Thus, in this study, we suggest that AGPs participate in adhesion. AGPs are just one group of the large assortment of hyp-containing proteoglycans that have been found in all plant groups (Lee et al., 2006; Huang et al., 2013) and even other eukaryotes (Herve et al., 2016). The presence of AGPs in the taxa studied here and in the wall pores of the desmid, Pleurotaenium (Domozych et al., 2007) show that AGPs are employed at very different geographic zones and during different developmental stages of charophytes. It might be also surmised then that AGPs also evolved to play key roles in adhesion and other functions in land plants. Further work into the biochemistry and molecular biology of charophyte AGPs will be necessary for understanding the functional and evolutionary significance of these proteoglycans.