Cloning of Plant Cells: Isolation and Culture of Single Cells

A clone of cells consists of all the cells derived through mitosis from a single cell, and the process of obtaining a clone is called cloning.

Therefore, all the cells of a clone are expected to be identical with each other in their genotype and karyotype (chromosome constitution), and other attributes, except for the changes that may arise afresh during and after cloning. Cloning is based on single cells separated from tissues and cultured in a manner to allow separate recovery of the cell mass derived from them.

Isolation of Single Cells:

Single cells can be isolated from plant organs, particularly leaf, either by mechanical or enzymatic means or they can be separated from suspension cultures. For mechanical isolation, leaves are cut into small (1 cm²), pieces, ground in a suitable culture medium using pestle and mortar, the resulting homogenate is filtered through muslin cloth, and the cells are pelletted out by centrifugation.
In the enzymatic method, the lower epidermis of leaves is peeled off and the leaves are cut into moderate pieces (4 cm²), which are incubated in a macerozyme (0.5%) or pectinase solution. Partial vacuum may be used to facilitate the entry of enzyme solution into the tissue, and the solution may be changed during the treatment.
The enzyme treatment tends to weaken cell walls; therefore, a suitable osmoticum (e.g., 0.3 M mannitol) is added to the enzyme and culture medium solutions. Single cells can also be isolated from suspension cultures by suitably filtering out cell clumps and harvesting the cells by centrifugation. A fine suspension of cells is usually obtained particularly when friable calli are used for the initiation of suspension cultures.

Culture of Single Cells:

Single cells can be cultured using the following techniques: (1) filter paper raft-nurse tissue technique, (2) microchamber technique, (3) microdrop method, (4) Bergman’s plating technique and (5) thin layer liquid medium. Isolated single cells fail to divide in normal tissue culture media. Therefore, either a nurse tissue or a conditioned medium is used for their culture.
A conditioned medium is a medium in which plant cells have been grown at a high cell density for about 24-48 hr, after which the cells had been filtered out. The conditioned medium is enriched by the various molecules secreted by the cultured plant cells, which sustain proliferation and development of the cells cultured in such a medium at a rather low density.
Plant cells in culture release proteins and carbohydrates, some of which function as signalling molecules, while some others can sustain embryogenesis and somatic embryo (SE) development. For example, a pentapeptide isolated from rice cell-conditioned medium stimulates mitosis in Asparagus officinal is mesophyll cells cultured in vitro.
Similarly, proteins and oligosaccharides secreted by plant cells are known to promote embryonic development in carrot and formation of shoots and flowers in tobacco. Further, glycosylated proteins (including AGPs) secreted by barley microspores sustain cell growth and promote maize zygote differentiation into embryo.

Filter Paper Raft-Nurse Tissue Technique:

Single cells are placed on small pieces (8×8 mm) of filter paper (sterilized), which themselves are placed on top of established callus cultures several days in advance. This allows the filter papers to be wetted by the exudates from callus tissue. The single cells placed on the filter papers (Fig. 8.4A) derive their nutrition from the callus exudates diffusing through the filters. The cells divide and form macroscopic colonies on the filters; the colonies are then isolated and cultured.

Micro-chamber Technique:

A micro-chamber can be created either by using a microscope slide and coverslips (the latter are held in place by sterile mineral oil), or by a cavity slide (Fig. 8.4B, C). Single cells are suspended in conditioned medium, and a drop of medium having a single cell is placed in the micro-chamber, which is covered with a coverslip, (Fig. 8.4B). In case of cavity slide, the drop is placed onto a coverslip, which is then inverted into the slide cavity. Micro-chambers allow microscopic observation, and they can be kept in a Petri dish for incubation.

Microdrop Method:

A specially designed dish, Cuprak dish, having a smaller outer chamber (to be filled with sterile distilled water to avoid desiccation of cells) and a larger inner chamber (having several numbered microwells) is employed. Microdrops of 0.25-0.5 µ1 are distributed in the microwells and the dish is sealed with parafilm. Cell density in the medium is so adjusted as to give, on an average, one cell per droplet (it works out as 2-4 x 10³ cells /ml). This method has been successfully used for protoplasts and should work with single cells as well.

Bergmann’s Plating Technique:

In this widely used technique, cells are suspended in a liquid medium at a cell density that is twice the desired density in the plate. Sterilized agar (Ca. 1%) medium is kept melted in a water bath at 35°C. Equal volumes of the liquid medium containing cells and the agar medium are mixed thoroughly and quickly spread in Ca. 1 mm thick layer in a Petri dish. The cells remain embedded in the soft agar medium and are observable under a microscope; when macroscopic colonies develop they are isolated and cultured separately.

Thin Layer Liquid Medium Culture:

Cells can be placed in a thin layer of liquid medium to allow adequate aeration. Since cells are not fixed in position, it is not possible to follow up on individual cells during culture. This technique is common for protoplast culture.
It is important to (i) culture the single cells in dark and keep microscopic observations to the minimum since light has a detrimental effect on cell proliferation. In addition, (ii) either a conditioned medium or a suitably enriched medium should be used since standard tissue culture media are unsuitable for single cell culture. Some highly enriched synthetic media permit the culture of as few as 25-50 cells/ml, while some media supplemented with casamino acids and coconut milk support cell division at 1-2 cells/ml. It is postulated that some bio-chemicals essential for cell division leach into the culture medium when they are sub-cultured; this produces the lag phase.
Cells begin to divide only when an equilibrium is established for these metabolites between the medium and the cells; this happens much later at lower cell densities, and below a critical cell density it may not be achieved. For this reason, conditioned or specially enriched media are required for culture of cells at low densities.

Cell Viability Test:

Cell viability can be determined by any one of the following approaches: (1) phase contrast microscopy, and staining with (2) 2, 3, 5-triphenyltetrazolium chloride (TTC), (3) fluorescein diacetate (FDA) and (4) Evan’s blue.
Live cells show cytoplasmic streaming and a well defined healthy nucleus, which are easily observable with a phase contrast microscope or even a light microscope. Cell masses can be stained with 1-2% solution of TTC, which is reduced by living cells to formazon that stains the cells red. Formazon can be extracted and measured with a spectrophotometer to give a quantitative estimate of viability, but it is not suitable for single or few cells.
Cells are treated with 0.01% solution of FDA. Live cells cleave FDA by esterase activity and produce fluorescein, which cannot move across plasma membrane. With UV exposure, fluorescein gives green fluorescence so that live cells appear green, while dead cells do not fluoresce. Evan’s blue (0.025%) is not taken up by live cells, while it freely enters into damaged dead cells. Therefore, all cells that take up stain are dead. Evan’s blue is usually used in conjunction with FDA.