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Fluid phase endocytic uptake of external solutes in plant cells was further substantiated using artificial polystyrene nano-spheres (40 nm) and CdSe/ZnS quantum dots (20 nm). Both types of artificial nano-particles were taken up by sycamore-cultured cells. However, whereas polystyrene nano-spheres were delivered to the central vacuole, CdSe/ZnS nano-dots were sequestered into cytoplasmic vesicular structures. Using dextran-Texas Red (m.w. 3,000; d-TR) as additional marker, confocal micrographs confirmed the distinct topographic distribution of CdSe/ZnS quantum dots within the cell. Initially, d-TR and CdSe/ZnS quantum dots colocalized within cytoplasmic vesicles. After 18 h incubation, d-TR was distinctly localized in the vacuole whereas CdSe/ZnS quantum dots remained sequestered in cytoplasmic membranous compartments. The data provide a first evidence for the rapid distribution of solutes taken up by endocytosis to distinct intracellular compartments.
One of the distinctive properties of the endocytic pathway for solute uptake described for sycamore cultured cells1,2 and Citrus juice cells3 is the apparent indiscriminate trapping and internalization of external solutes. Whether using the small ionic markers Lucifer Yellow-CH (LY) and Alexa-488, or the larger neutral dextran-Texas Red (d-TR; m.w. 3,000), internalized soluble markers, which mainly occurred in the presence of sucrose, were eventually delivered to the central vacuole. The vacuole was also the final destination for fluorescent endocytic markers in experiments performed with tobacco suspension cultures4 and with onion root5 apices. In our studies,1,3 the distinct stimulation by sucrose confers the endocytic system characteristics of receptor-mediated endocytosis, whereas the general trapping and transport of all other solutes is more distinctive of non-receptor-mediated fluid phase endocytosis.6,7 This situation is further complicated by skepticisms arising from reports in which small ionic molecules such as LY allegedly traverse the plasma membrane through ion channels as suggested by experiments using probenecid and other potent anion channel transport inhibitors,8,9 and by uptake into the endosome/lysosome following micro-injection of dyes into the cytosol.10
Recent advances in plant endocytosis11,12 have implied the presence of various distribution routes and centers that operate according to the nature of the trapped solutes. Our initial determinations have exposed minimal degree of selectivity and intracellular distribution, although the presence of intermediate structures were inferred from scanning fluorescent micrographs after three hours incubation in LY (see Fig. 6 in ref. 1). To advance our understanding of endocytic solute uptake in plant cells, we proceeded to investigate several key aspects of the endocytic pathway that remained unresolved following our initial studies. First, can it be unmistakably demonstrated that external markers are taken up by fluid-phase endocytosis and not by ion channels, membrane-bound carriers, or by the conformational coupling of corresponding channels on different membrane systems?13,14 Second, is fluid-phase endocytosis in plant cells exclusively for photoassimilate uptake and other solutes excluded? Third, are all indiscriminately trapped solutes eventually delivered to the vacuole? To resolve these crucial issues, we used sycamore-cultured cells, polystyrene nano-spheres, polyethylene coated CdSe/ZnS quantum dots, and fluorescent dextran (m.w. 3,000 tagged with Texas Red) coupled with electron and confocal microscopy.
Sycamore (Acer pseudoplatanus) cells were cultured in a medium supplemented with sucrose (complete medium) as described in ref. 15 in continuously agitated 250 ml flasks (200 rev/min at 28°C). Cell samples were obtained from 6- to 7-d-old cultures during rapid growth phase.16 Starved cells were prepared by transferring 5-to 6-d-old culture cells into equal volume of culture medium without sucrose (starving medium) for 24 h.
Protoplasts from sycamore-cultured cells were prepared using a cell wall hydrolytic solution as described in ref. 15. After incubation in hydrolytic medium, protoplasts were washed thoroughly with a solution similar to the hydrolytic medium but without cell degrading enzymes.
Protoplasts were incubated for 12 h in sucrose containing culture medium15 supplemented with 4 × 1014 fluorescent polystyrene beads/ml (average diameter 40 nm and internally labeled with Texas Red; FluoSpheres® carboxylate-modified microspheres F-8786, Molecular Probes, Eugene, OR, USA). Microspheres were desalted by ultracentrifugation on Microcon YM-10 centrifugal filters (m.w.co 10,000, Millipore Corporation, Bedford, MA, USA) and resuspended in 1 ml incubation medium supplemented with 0.01% bovine serum albumin. The suspension was sonicated for five minutes and added to 4 ml of a solution containing protoplasts.
Approximately 100 mg fresh weight of sycamore cultured cells were incubated in 1 ml of culture medium containing 1014 CdSe/ZnS 20 nm quantum dots/ml (PEGylated derivatized-polyacylacrylic acid coated cadmium selenide/zinc sulfide core/shell quantum dots, Q tracker 565 nontargeted quantum dots; Quantum Dot Corporation, Hayward, CA, USA) with or without fluorescent dextran (m.w. 3,000) tagged with Texas Red (d-TR; from Molecular Probes, Eugene, OR, USA) at a final concentration of 1.0 mg·ml−1. At determined times (indicated in text according to individual experiments), cells were washed three times with culture medium without markers and protoplasts prepared as described above. Observations of live protoplasts were performed using a Leica TCS SL confocal microscope (Leica, Heidelberg, Germany).
For electron microscopy, protoplasts, incubated for 12 h in a solution containing fluorescent microspheres, were centrifuged at 70 RCF for three minutes and washed five times with fresh incubation medium without spheres. The pellet was finally resuspended in 3 ml of incubation medium supplemented with 4% glutaraldehyde and 0.1 M cacodylate buffer, and incubated for 24 h at 4°C. After washing in phosphate buffer (0.1 M, pH 7.0) the fixed protoplasts were stained with 1% osmium in 0.1 M phosphate buffer for one hour at 4°C. Cells were washed twice in buffer, immobilized in 4% agarose and embedded in Spurr's resin. Immobilized cells were sectioned and stained in uranyl acetate/lead citrate buffer prior to viewing in a Philips 201 TEM.
Our initial attempt to examine the possible uptake of nano-particles as a means to verify fluid phase endocytosis in plant cells was hindered by the severe clumping of polystyrene beads and by their tenacious adherence to the cell walls. When observed under the microscope, cells were coated with intense red fluorescence obstructing any view of internal details. The attachment of beads to the cells also prevented hydrolysis of cell walls in order to generate protoplasts for better visualization. To overcome this obstacle, we first generated protoplasts and incubated them overnight in growing medium supplemented with 4 × 1014 polystyrene beads/ml. Figure 1A–D shows the presence of polystyrene particles inside the vacuole as well as outside several cells under two magnifications. These particles are unmistakably distinct from any other natural occurring vacuolar structures present in control cells (Fig. 1E). In fact, the vacuole of untreated cells appeared uniformly homogeneous and devoid of any recognizable particles.
When examined at higher magnification, particles, whether inside or outside the cell (Fig. 1A–D), looked fairly consistent and identical to control beads fixed in agar and treated similarly (Fig. 1F). The variation in size between particles likely resulted from the sectioning along different planes of the spheres randomly located within the vacuole. It is worth noting that the average size of these particles in the micrographs appear larger than their original size of 40 nm, however, this could be attributed to swelling during fixation and handling of the tissue for electron microscopy since control beads also appeared to be larger in size after handling (Fig. 1F). On several occasions, nano-spheres were observed in locations other than the central vacuole (Fig. 1G), but it became difficult to distinguish and impractical to attempt defining such cellular structures.
The lack of evident cellular organization associated with the relatively large polystyrene beads and the restrictions imposed by the use of protoplasts prompted further investigations using walled cells and smaller fluorescent polyethylene glycol (PEG)-coated 20 nm CdSe/ZnS quantum dots17 as fluid-phase endocytosis marker. After two hours incubation in the presence of sucrose, confocal microscopy revealed the presence of CdSe/ZnS quantum dots green fluorescence within the cytoplasm (Fig. 2A and B). The numerous brightly and compact fluorescent structures appear scattered throughout the cortical and perinuclear region. After prolonged incubation (18 h) green fluorescence was still evident, but at this time, CdSe/ZnS quantum dots were mostly contained in larger spherical organelles (Fig. 2C and D), although oblong and elongated structures were occasionally observed. At no time, however, fluorescence from the CdSe/ZnS nano-particles was observed in the central vacuole, similar to soybean (Glycine max L.) protoplasts incubated with gold-labeled bovine serum albumin particles.18
In contrast to soluble endocytic markers1–3 and polystyrene beads (Fig. 1), CdSe/ZnS quantum dots were not delivered to the central vacuole, but remained in cytoplasmic organelles even after 18 h incubation (Fig. 2C and D). If trapped indiscriminately along with other particles in the incubation medium, CdSe/Zn quantum dots must be colocalized with other markers at least at early stages of vesicle trapping.
During initial two hours of culture in medium containing CdSe/ZnS quantum dots and d-TR, colocalization of both markers was evident in small structures along the cytoplasm as evidenced by the fluorescent yellow color resulting from the overlapping of red and green fluorescence (Fig. 3). Double labeling images were produced by superimposing fluorescent micrographs taken simultaneously at different exitation/emission wavelengths corresponding to individual endocytic markers. The tight cluster of vesicles in Figure 4 authenticates the early vesicular colocalization of d-TR and CdSe/ZnS quantum dots. In this figure, the yellow fluorescence must have resulted from the colocalization of markers within vesicles as it is highly unlikely that vesicles with separate fluorescent cargoes would be uniformly aligned in pairs along the visual axis as to generate a yellow color. These results showing colocalization of both markers within the same vesicles validate the indiscriminate trapping nature characteristic of fluid phase endocytosis. Most importantly, at four hours of culture, although in apparent close proximity, colors were not always overlapping (Fig. 5), suggesting a rapid distribution of substances taken-up by endocytosis. After 18 h incubation, it was evident that highly systematic distribution had taken place. At this point, the vacuole was highly fluorescent with d-TR (Fig. 6), yet only weakened signs of CdSe/ZnS quantum dots remained, and only within cytoplasmic structures but never in the vacuole.
From the data presented in this communication, several conclusions can be drawn with high degree of certainty. First, the uptake of artificial polystyrene nano-spheres and fluorescent CdSe/ZnS quantum dots constitutes compelling evidence to confirm the presence of fluid phase endocytosis in plant cells as previously suggested using smaller soluble fluorescent markers1,2,3 and gold-labeled BSA.18 The size and nature of the particles guarantees that uptake can not occur through membrane located channels or transporters, or by the juxtaposition of channels between two membranes.13,14 It is highly improbable that these particles would be recognized as natural nutrients, and their transport mediated through existing channels or carriers specialized in transport of single nutrient molecules. Given that uptake of endocytic markers in sycamore cultured cells occurs predominantly in the presence of sucrose,1 fluid phase endocytosis must serve as a means of nutrient uptake. However, it is also quite probable that a low level steady state of endocytosis is continuously present in these cells as a means of membrane recycling or internalization of pectic substances.19 Second, although solutes enter “in mass” within individual vesicles, a mechanism of recognition, separation and redistribution occurs at some early stage in the endocytic process. This conclusion is based on the substantial green fluorescence from CdSe/ZnS quantum dots and d-TR visible in small compartments within the cells at the beginning of the endocytic process (Fig. 3), but the segregated fluorescence visible shortly afterwards (Fig. 5). Third, in plant cells, not all solutes trapped by fluid phase endocytosis are ultimately delivered to the vacuole as evidenced by the conspicuous sequestration of CdSe/ZnS quantum dots into cytoplasmic compartments even after 18 h (Fig. 6). It is tempting to speculate that after segregation of d-TR and CdSe/ZnS quantum dots, the latter is extruded back to the medium, whereas the former is transported to the vacuole. If CdSe/ZnS quantum dots were to continuously accumulate in some cellular compartment, higher fluorescence intensity would be visible at 18 h than at two hours. Recycling of endocytic probes back to the medium has been reported in animal cells.20 Aside of reaffirming the existence of diverse storage compartments,21 the data also demonstrate, for the first time, the delivery of trapped particles to different cell compartments and the use functionally diverse vesicles during fluid-phase endocytosis in plant cells.
We thank Diann Achor, University of Florida's Citrus Research Center for her microscopy work.
Previously published online as a Plant Signaling & Behavior E-publication: http://www.landesbioscience.com/journals/psb/abstract.php?id=3142