Microbead encapsulation of living plant protoplasts: A new tool for the handling of single plant cells.

PREMISE OF THE STUDY: Understanding plant cell biomechanics has been hampered by a lack of appropriate experimental tools. Here we introduce a protocol for the incorporation of individual plant protoplasts into precisely sized agarose microbeads. This technology may lead to new ways to manipulate the physical and chemical microenvironment of individual plant cells. METHODS AND
RESULTS: Living protoplasts obtained from BY-2 tobacco suspension cultures were continuously incorporated into a stream of agarose microdroplets, collected in cooled mineral oil as gelled microbeads, and then transferred into liquid MS medium for culture. In this first report, we show that spherical microbeads containing living protoplasts can be easily generated in quantity and that these encapsulated cells continue to grow and divide.
CONCLUSIONS: Microbead encapsulation of protoplasts affords the opportunity to precisely control the physical microenvironment of individual plant cells. Ultimately, this method may help facilitate novel studies in plant biomechanics.

Although the likely significance of cell and tissue mechanics in plant development has been appreciated for many years, the study of cellular micromechanics at the level of the individual cell has proven to be problematic. The universal presence of the cellulosic cell wall and the apoplastic continuity that it provides endows plant tissues with a unique level of mechanical coupling. In principle, this makes it possible for plant tissues to transmit stress-mechanical information precisely and instantaneously over multicellular distances. However, the same apoplastic continuity that makes stress-mechanical signaling attractive as a possible developmental effector also makes it difficult to interpret responses and isolate mechanical variables at the level of the individual cell.

With the advent of droplet microfluidics, it is now possible to manipulate individual cells in novel ways, potentially revealing levels of developmental control that have previously been experimentally inaccessible. In this study, we present a reliable procedure for capturing large numbers of individual plant protoplasts in mechanically isotropic hydrogel microbeads, thereby isolating them from the physical influence of neighboring cells and allowing them to regenerate their walls and proceed through cell division in a precisely controlled physical environment. A detailed protocol is provided as Appendix 1.

Microfluidic devices designed to facilitate the handling and analysis of individual cells are now becoming available and have already been used to encapsulate animal cells (Kumachev et al., 2011), but in studies with plant cells they have found only limited use (Agudelo et al., 2012; Ghanbari et al., 2014). Here we show that droplet microfluidic systems are capable of rapidly and efficiently capturing large numbers of individual plant protoplasts in precisely sized spherical hydrogel beads, providing plant scientists with new ways of dissecting the biophysical background of plant development.

METHODS AND RESULTS

Protoplast isolation

Protoplasts were obtained from 4-d-old BY-2 tobacco (Nicotiana tabacum L. cv. BY-2) suspension cultures grown at 27°C in Murashige and Skoog (MS) basal salts (Caisson Laboratories, Smithfield, Utah, USA), 3% sucrose (Fisher Scientific, Fair Lawn, New Jersey, USA), 100 mg/L myo-inositol (Alfa Aesar, Ward Hill, Massachusetts, USA), 1 mg/L thiamine (Fisher Scientific), and 0.2 mg/L 2,4-dichlorophenoxyacetic acid (2,4-D; Caisson Laboratories). Suspension-cultured cells were removed from the culture medium and then enzymatically stripped of their cell walls in a digestion medium consisting of 0.5% (w/v) Onozuka RS, 0.5% (w/v) Onozuka R-10, 0.1% (w/v) Macerozyme R-10 (all from Yakult Pharmaceuticals Co. Ltd., Kunitachi-Shi, Tokyo, Japan), and 0.1% Pectolyase Y-23 (MP Biomedicals LLC, Solon, Ohio, USA). The digestion medium was made up in 5.2% mannitol (Acros Organics, Morris Plains, New Jersey, USA), 0.05% MgCl2 (Sigma-Aldrich, St. Louis, Missouri, USA), and 0.2% MES buffer (Sigma-Aldrich) at pH 5.8. Suspension culture BY-2 cells were digested for 3 h at 27°C on a shaker table. Protoplasts were separated from the digestion medium by centrifugation at 100 × g for 7 min. After being washed twice in a solution containing 5.5% mannitol and 0.05% MgCl2 at pH 5.8 (Medium 1) protoplasts were suspended in a culture medium containing 0.44% Caisson MS medium (Caisson Laboratories), 3% sucrose, 2.7% mannitol, and 0.2 mg/L 2,4-D and allowed to rest (7°C for 8 h followed by 12 h at room temperature). Before droplet encapsulation, cells were pelleted by centrifugation at 100 × g for 7 min and then resuspended in MS medium for droplet encapsulation.

Microbead production

Agarose microbeads were generated using a microfluidic droplet system manufactured by Dolomite Microfluidics (The Dolomite Centre Ltd., Royston, United Kingdom). Water-based (agarose) droplets were formed as the discontinuous phase in a 2-reagent, 4-channel, glass microfluidic junction chip. The continuous phase was a light mineral oil (Sigma-Aldrich M5310) with 4% Span 80 (Fluka Analytical, St. Louis, Missouri, USA) added to prevent droplet coalescence. Flow rates of the three component fluids were adjusted with three dedicated Dolomite P-Pumps independently controlled by proprietary Dolomite software. The fluids were fed into the droplet chip through microbore polytetrafluoroethylene (PTFE) tubing. The two outer channels were reserved for the mineral oil continuous phase (Fig. 1). The two central channels were functionally separated, with one channel reserved for live protoplasts at an approximate density of 5.0 × 106/mL in MS medium containing 0.44% Caisson MS medium, 3% sucrose, 2% mannitol, and 0.5% high-molecular-weight dextran sulfate. The second channel was fed from a warmed reservoir of 1.5% low-gel-temperature agarose (Sigma-Aldrich) made up in 0.44% Caisson MS medium, 3% sucrose, and 2% mannitol, and maintained at a temperature of 34°C. All reagents were sterile or prefiltered through 0.22-μm syringe filters.

Fig. 1.

Schematic of the microdroplet system. Three pumps store solutions in pressure-controlled chambers and drive fluid flow during droplet formation (top). Microbore PTFE tubing (dotted lines) carries fluids from the pressure pumps to the 2-reagent droplet chip. Droplet production occurs at the droplet chip junction where the channels intersect. Agarose (blue) and cells (black) meet immediately before being cleaved into droplets by continuous oil flow (orange). Liquid microdroplets exit the microchip into a cooled mineral oil bath where they solidify.

With appropriate flow control, a stream of monodisperse agarose droplets forms at the chip junction where the aqueous and oil phases intersect (Video 1; Fig. 1). Droplet diameter can be increased or decreased by adjusting the flow rates of the continuous and discontinuous phases. Liquid droplets exit the chip and stream downward into a cooled mineral oil bath (Sigma light mineral oil with 4% Span 80), where they solidify into gelled beads (Fig. 1).

The production of uniformly sized microdroplets at the junction of the droplet chip. Also seen is the problematic coalescence of droplets caused by occasional cell clumps. This video is an MPEG file (.mpg) and can be viewed here with QuickTime or Windows Media Player, or it can be viewed from the Botanical Society of America’s YouTube channel.

Microbeads were separated from the mineral oil by centrifugation into a sublayer of liquid MS culture medium and then filtered through an 88-μm nylon mesh to remove coalesced droplets. Finally, agarose microbeads containing live protoplasts were suspended in liquid culture medium containing 0.44% Caisson MS medium, 3% sucrose, 1.6% mannitol, and 0.2 mg/L 2,4-D (pH 5.8). The growth medium was supplemented with filtered medium from a rapidly growing BY-2 cell suspension culture. New cell wall synthesis was confirmed by adding a drop of 0.1% Calcofluor (Fluorescent Brightener 28; Sigma-Aldrich) to a microbead slide preparation.

Consistently sized (ca. 60 μm), spherical hydrogel microbeads were successfully generated at a rate of approximately 130 beads per second. Individual protoplasts were successfully encapsulated with good viability as determined by cytoplasmic streaming activity shortly after encapsulation (Fig. 2A). Cellulose staining with Calcofluor White showed that 32 h after protoplast release and 20 h after cells were encapsulated all living cells had begun to regenerate a thin cell wall (Fig. 2D). Living cells proceeded to elongate and divide, eventually bursting the agarose microbead in which they had been encapsulated (Fig. 2B). Immediately following droplet formation, approximately 25% of generated droplets contained protoplasts.

Fig. 2.

BY-2 plant cells in various stages following encapsulation in agarose microbeads. (A) Phase-contrast image of BY-2 protoplasts in solidified microbeads suspended in MS medium immediately after encapsulation. (B) Phase-contrast image of a BY-2 cell breaking through the agarose microbead 6 d after encapsulation. (C) Nomarski differential interference contrast image of an individual BY-2 cell 20 h after encapsulation and 32 h after the removal of its cell wall. (D) Calcofluor White fluorescence of the respective cell in Fig. 2C showing regeneration of a thin surrounding cell wall. Scale bars = 100 μm (2A, 2B) and 50 μm (2C, 2D).

CONCLUSIONS

In order for the study of plant biomechanics to keep pace with rapid advances in our understanding of the molecular controls underlying plant development, we need to be able to isolate mechanical and physical inputs at the cellular level. Studies conducted at the tissue level are inherently limited by the stresses and strains that pervade and organize all multicellular plant tissues. With the technique described here, it is possible to capture individual plant protoplasts in physically isotropic, hydrogel microbeads whose mechanical properties can be controlled.

The potential for this technique to be used as a means of manipulating the mechanical forces acting on individual plant cells in culture suggests that it could facilitate novel studies. Additionally, with ongoing rapid development in the fields of microfluidics and hydrogel engineering, it seems that these techniques are poised to improve over time. We anticipate that future work in these areas will lead to improvements in the capture efficiency of live cells, and to the investigation of other hydrogel environments with different material properties.

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