Protocol for Isolation and Culture of Protoplast (With Diagram)

In the below mentioned article, we will discuss about the protocol for isolation and culture of protoplast.

Protoplasts can be prepared from a variety of tissue but among them leaf mesophyll tissue from a wide range of plants has been proved to be the most ideal source of plant material for protoplast isolation (Fig 12.5).

Leaves of Nico­tiana tabacum is a highly standardized material for easy entry into the art of protoplast isola­tion and culture.

Now-a-days the mixed enzyme method (single step) is very popular and this procedure is followed as routine work in most of the laboratories of the world.

The protocol for isolation of protoplast from the mesophyll cells of tobacco using mixed enzyme method and its culture are described below:

Isolation of Protoplast:

1. Young fully expanded leaves from the up­per part of 7-8 weeks old plants growing in a greenhouse are detached and leaves are washed thoroughly with tap water.

2. Surface sterilization of leaves is done by first immersing in 70% ethanol for 60 seconds followed by dipping into 0.4-0.5% sodium hypochlorite solution for 30 minutes. For this purpose, a sterile casserole dish is used as a sterilizing container. Sterilization is done in front of laminar air flow.

3. After 30 minutes, the sterilant is poured off and leaves are washed aseptically 3-4 times with autoclaved distilled water to remove every trace of hypochlorite.

4. With the help of long sterilized forceps (8 inches), one leaf is transferred on a steril­ized floor tile. The impermeable lower epi­dermis of the surface sterilized leaves is peeled off as completely as possible. During this process, a sterilized fine jeweler’s force is inserted into a junction of the midrib and a lateral vein and the epidermis, is care­fully peeled away at an angle to the main axis of the leaf.

N.B.:

Where peeling of the leaf is not possible, slicing of the leaf into thin strips may be suffi­cient to allow entry of the enzymes through the cut edges of the strip.

5. Peeled leaf pieces are placed lower surface down onto 30 ml sterilized CPW 13M*solution in a 14 cm petridish. When the liquid surface is completely covered with peeled leaf pieces, then the CPW 13M solution is pipetted off from the beneath of leaf pieces. The CPW 13M solution is replaced by bac­terial filter sterilized solution of enzyme containing 2% cellulase (Onozuka R1O), 0.5% macerozyme in 13% manitol added CPW (pH 5.5).

6. Leaf pieces in enzyme solution are incubat­ed in the dark at 24-26°C for 16-18 hrs.

7. Without disturbing the digested leaf pieces the enzyme solution is gently replaced by CPW 13M. Then digested leaf pieces are gently agitated and squeezed with sterile fine forceps to facilitate the release of the protoplast. The protoplast suspension is then allowed to pass through a 60 µ-80µ nylon mesh to remove the larger pieces of undigested tissue.

8. The filtrate is transferred to screw-capped centrifuge tube and is spinned for 5 minutes at 100 g.

9. The protoplasts form the pellet. The super­natant is pipetted off and the pellet is re- suspended in CPW 21S solution. It is again centrifuged for 5-7 minutes at 200 g. The viable protoplasts will float at the top surface of CPW 21S in the form of dark green band while the remaining cells and debris will sink at the bottom of the tube.

10. The viable protoplasts are collected from the surface and are re-suspended again in CPW 13M to remove the sucrose. Centrifugations are repeated two-three times for washing.

11. Finally the protoplasts are suspended in measured volume of liquid Nagata and Tabeke medium (1971) supplemented with NAA (3 mg/L), 6-BAP (1 mg/L) and 13% mannitol.

Isolation of Protoplast from Cell Suspension Culture:

Rapidly growing cell suspension cultures are the most suitable material for protoplast isola­tion. A new cell suspension does not yield many protoplast until it has been sub-cultured at least twice. For protoplast isolation, suspension cul­tures are generally harvested at its early expo­nential growth phase or log phase.

Older suspension cultures have a tendency to form elongated giant cells with thick wall. So it is very difficult to isolate the protoplast from such culture. Again, the presence of large num­ber of cell aggregates in suspension culture is not desirable for the isolation of protoplast.

Addi­tion of colchicine and some chelating chemicals in suspension culture generally prevents the for­mation of cell aggregates. Sometimes very low concentration of cellulase (0.1%) is added in cell suspension culture two days before use to dis­courage the formation of thick wall.

Step 1:

Filtering of cell suspension:

The harvested cell suspension is passed through a coarse nylon sieve so that filtrate contains single cells as well as very small cell clumps.

Step 2:

Preparation of liquid medium free cells for plasmolysis:

The filtrate is allowed to settle out of the medium. Most of the medium is decanted off and the cells are transferred (by pouring) to a flask. Using the Pasteur pipette, all of the cul­ture medium is removed. This is best achieved by drawing off medium from the base of the cell layer.

Step 3:

Pre-plasmolysis:

The cells are suspended in CPW13M solution for 1 hr. After 1 hr., the plasmolyticum is pipetted off.

Step 4:

Enzyme incubation:

The enzyme solution is added to the cells. The flask containing cells and enzyme are placed on the platform of a slow­ly rotating gyratory shaker (ca 30-40 rpm) for standardized period (4-6 hrs.).

Step 5:

Washing and purification:

Protoplast sus­pension is filtered through 60/i-100/i stainless steel sieve to remove the larger debris. The filtrate is transferred into centrifuge tubes and is spinned at 80 g for 5-10 minutes so as to sediment the protoplasts and then supernatant is pipetted off. The pellet is re-suspended in CPW 21S solution.

The protoplast suspension is again spinned at 100 g for 5-7 minutes. Viable and de­bris free protoplasts are collected at the surface. The protoplasts are washed with CPW 13M by repeated centrifugation and finally protoplasts are re-suspended in measured volume of liquid culture medium. Finally, the yield of protoplast is measured by counting in haemocytometer.

Sample Protocol—Isolation of Protoplast from Cell Suspension Culture of Daucus carota:

1. Allow the cells to settle out of the medium and remove the supernatant with Pasteur pipette.

2. Add 20-30 ml of enzyme solution (Onozuka R10 cellulase – 2% and hemicellulose – 1% in 10% mannitol added CPW solution, pH 5.5) and incubate the cells at room tem­perature on the platform of slowly rotating gyratory shaker 30 rpm) for 4 hrs.

3. Filter protoplast suspension through 67 µ stainless steel sieve and transfer the filtrate in centrifuged tube. Spin at 80 g for 5 min.

4. Discard in supernatant and re-suspend in CPW 21S. Spin 100 g for 5-7 minutes.

5. Collect the protoplast from the surface of CPW 21S solution.

6. Wash the protoplast with CPW 13M solu­tion by repeated centrifugation (3-4 times).

7. Finally transfer the protoplast to a mea­sured volume of liquid B₅ medium supple­mented with 2, 4-D(l mg/L), kinetin (2 mg/L) and casein hydrolysate (1 gm./1).

Isolation of Haploid Protoplast:

Pollen mother cell (PMC), pollen tetrad (PT) and mature pollen are the natural source of haploid cells. Since they are produced in large number in anther and is very easy to squeeze out from anther, they are suitable material for the isolation of protoplast. Egg cell is also haploid cell, but generally single egg cell is produced per ovule and is very difficult to separate them from complex tissue integration.

Isolation and yield of protoplast from hap­loid male cells depends upon the composition of cell wall. The cell wall of PMC and PT is made up of callose (un-branched 1-3 glucan) whereas the exine of mature pollen is coated with sporopollenin which is very difficult to digest by en­zymes. The freshly isolated protoplast from ma­ture pollen tends to fuse spontaneously to form multinucleate giant protoplast.

Therefore, isola­tion and culture of protoplast have yielded only limited success. Better preparation of proto­plasts is obtained from pollen mother cell or poll­en tetrad than the mature pollen. The isolated protoplast from haploid cell is known haploid protoplast or gametoplast. Isolation and culture of haploid protoplast are useful for the induction mutation and to study the effect of irradiation as well as chemical mutagens.

Isolation of Protoplast from PMC and PT:

To ascertain the presence of PMC and PT in anther in relation to flower bud size, the an­thers are smeared with 2% acetocarmine before using them. Like leaf mesophyll tissue and cell suspension culture, the isolation of haploid pro­toplast is also multistep process.

Approximately, the anticipated size of an­ther containing PMC and PT are aseptically re­moved with fine forceps. The content of this anther are squeezed out with the help of glass spatula. The pollen mother cell and the pollen tetrad come out as a milky fluid. It is treated with 0.75-1% helicase enzyme in 8-10% sucrose for about 30-45 minutes.

Helicase is obtained from snail gut. Sometimes Zymolase, an enzyme isolated from Arthobactor luteus is used for the isolation of protoplast from pollen tetrad. After incubation enzyme is carefully replaced by 10% sucrose and the protoplasts are allowed to settle. Protoplasts are rinsed by conventional washing medium. Finally, protoplasts are cultured in ei­ther liquid or solid medium.

Culture of Protoplast:

Isolated protoplast can be cultured in sev­eral ways of which agar embedding technique in small petridish is commonly followed. In this technique, protoplast suspension is mixed with equal volume of melted 1.6% ‘Difco’ agarified medium (37° C) and the protoplast-agar mixture are poured into small petridish.

In petridishes, embedding of protoplasts in solid agar medium is known as plating of protoplasts. The plated pro­toplast can be handled very easily and the agar medium provides a good support to the proto­plast. In situ developmental stages of embed­ded protoplast can be studied under compound microscope. Besides this, separated clones de­rived from individual protoplast can be moni­tored.

The method is described below:

1. The protoplasts in liquid NT medium* are counted with the help of haemocytometer. The protoplast deficit is adjusted to 1 x 10⁵ to 2 x 10⁵ protoplast/ml.

2. Agar solidified (1.6% ‘Difco’ agar) NT me­dium is melted.

3. The tight lid of Falcon plastic petridish (35 mm diameter 5 mm thickness) is opened and 1.5 ml of protoplast suspension is taken. To this equal aliquot of melted agar medium is added when it cools down at 37°C to 40°C (Fig 12.6).

4. The lid is quickly replaced tightly and the whole dish is swirled gently to disperse the protoplast-agar medium mixture uniformly throughout the dish.

5. The medium is allowed to solidify. The petridish is then inverted.

6. The culture is incubated at 25° C with 500 lux illumination (16 hrs. light) initially.

7. The cultures are sub-cultured periodically in the same solid medium (0.8% agar) with gradually reducing mannitol.

Several other methods have been described for the culture of protoplasts, such as droplet cul­ture, Co-culture, feeder layer, hanging droplets and immobilized/bead culture.

Droplet Culture:

Suspending protoplasts in liquid culture media are placed on petri dishes in the form of droplet (Fig 12.7) with the help of micropipette. This method enables the subsequent microscopic examination of protoplast development. In this method, cultured protoplast clump together at the centre of droplets.

Co-culture:

Sometimes a reliable fast growing proto­plast is mixed in varying ratio with the less fast growing protoplast. The mixed protoplasts are plated in solid medium as described previously. The fast growing protoplast presumably provides some growth factors which induces the growth and development of the desirable protoplasts. This is known as co-culture technique.

Feeder Layer Technique:

Fast growing protoplasts are sometimes made initotically blocked protoplast by low doses (1-2 Krad) of X-ray treatment. Such irradiated protoplasts are plated with agar medium. Upon this thin solidified layer of irradiated protoplast, desirable protoplasts are again plated at a low density with agar medium.

As a result, it makes two agar layers containing irradiated protoplast in lower layer and desirable protoplast in upper layer. The lower irradiated protoplast is known as feeder layer which improves the growth and development of normal protoplasts even at lower density (Fig 12.7).

Hanging Droplet Method:

Culture of protoplasts in an inverted liq­uid droplet (0.25-0.50 µl) is known as hanging droplet method. Each droplet contains very small group of protoplasts. A number of droplets are generally placed on the inner surface of the lid of a petri dish.

Very thin layer of water is gen­erally kept on the lower part of the petri dish to make a humid condition inside the petri dish as well as to prevent the dessication of the droplets. This technique facilitates to observe the devel­opment of protoplast under microscope. Proto­plasts also gets better aeration as they go down to the hanging surface of the droplets.

Bead Culture:

Sometimes protoplast suspension are mixed with several polymer like alginate, carrageenan etc. as well as melted standard difco agar. Small beads are made by dripping the mixture into liq­uid medium. After that, beads in liquid medium are put on moving shaker.

Entrapped proto­plasts culture have shown several advantage over static liquid culture or slowly moving liquid cul­ture where the protoplast suffers the mechanical breakage. This technique increases the mechan­ical stability, aeration and viability with bioche­mical activity.

Tests for Viability of Protoplast:

Viability of protoplasts after isolation and during culture in liquid medium is very impor­tant. Cell wall formation, cell division, callus formation etc. depend upon the viability of pro­toplast. The most frequently used staining methods for assessing protoplast viability are fluorescein diacetate (FDA), phenosafranine. FDA dissolv­ed in 5.0 mg/ml acetone is added to the proto­plast culture at 0.01% final concentration. The chlorophyll from broken protoplasts fluoresces red. Therefore, the percentage of viable proto­plasts in a preparation can be easily calculated.

Phenosafranin, also used at a final concen­tration of 0.01% is specific for dead protoplast. As soon as the stain is mixed with protoplast preparation, the inviable protoplasts stain red and viable protoplasts remain unstained.

Wall Formation, Cell Division and Callus Formation:

The viable protoplast in culture regener­ates its own wall around them. Once the wall is formed, the protoplast becomes essentially a regenerated cell. Depending upon the species, the protoplast remains in naked condition hardly for 10 minutes or a day. Generally protoplast be­gins to deposit cellulose micro-fibril immediately after washing and enzyme removal.

Cell mem­brane of newly isolated protoplast contains pro­truding microtubules that function in the orien­tation of newly synthesized cellulose micro-fibrils. The rate and regularity of cell wall regeneration depend on the plant species and the state of dif­ferentiation of the donor cell used for protoplast isolation.

Calcafluor white (CFW) is the most com­monly used stain to detect the onset of cell wall regeneration. CFW binds to the—linked glucosides in the newly synthesized cell wall. Opti­mum staining is achieved when 0.1 ml of proto­plasts is mixed with 5.0 pi of a 0.1% v/v solution CFW. Cell wall synthesis is observed by a ring of white fluorescence around the plasma membrane.

Cell wall regeneration is prerequisite for nu­clear and cell division. After the formation of cell wall, the walled cells expand and divide into two cells’. At this stage it looks like ‘8’. Cell division stages can also be stained using CFW. In most of cases, first cell division usually takes place with 2-7 days of culture.

After the first division, each daughter cell divides into two cells. Repeated di­vision results the formation of cell clump or cell aggregates. All the cells derived from the proto­plasts do not divide and form the cell colonies.

Therefore, the percentage of cells which give rises cell colonies, to known as plating (colony form­ing) efficiency. Several factors such as genotype of the donor plant, culture medium, hormones as well as physical factors are important for the division of protoplast and callus formation.

The small callus mass can be handled in the conventional manner which means that it can be sub cultured at regular interval and can be used for organogenesis. For sub-culturing, the plate containing dividing protoplasts are sliced into several agar blocks. Each block is transferred to the surface of fresh medium. The plasmolyticum level in the culture medium is progressively re­duced to zero by repeated sub-culturing.

Plant Regeneration:

The ultimate objective in protoplast cul­ture is the reconstruction of plant from the sin­gle protoplast. The strategy for plant regener­ation has been to recover rapidly growing cal­lus from protoplasts and to transfer the callus to a species specific regeneration medium (Fig 12.8). It is generally noted that plant regener­ation occurs very easily in some plant species while others are recalcitrant.

Plant regeneration from protoplast derived callus tissue have been reported mainly from solanaceous species. It in­cludes 17 Nicotiana species, 6 Petunia species and 6 Solanum species. In recent years the list of non-solanaceous species capable of plant regen­eration from protoplasts has been steadily ex­panding. The list contains several species mono- cot and dicot including carrot, endive, cassava, alfalfa, millet, clover, rapeseed, asparagus, cab­bage, citrus etc.