Material:

• Green leaf (mesophyll) and cultured cells (albino) of N. tabacum
• PEG fusion solution contains:
  • 22.5% (w/v) PEG molecular weight 6000
  • 1.8% (w/v) sucrose
  • 154 mg/l CaCl2·2H2O
  • 9.52 mg/l KH2PO4
  • pH 5.8 (adjust with 1M KOH or IN HCl, autoclaved and stored in dark at 4°C)

Protocol:

• In continuation with protoplast isolation experiment, perform the fusion experiment.
• Take 4–6 ml of freshly isolated protoplast suspension from two sources in protoplast washing media CPW salt medium –13 % mannitol (CPW13M) with a density of 5 × 105/ml.
• Use a Pasteur pipette to place a small drop (~50l) of sterile silicone on to the center of a plastic petri dish (35 mm) and gently lower a cover glass on to the drop of silicone fluid.
• Pipette ~150l of protoplast suspension directly on to the middle of cover glass and allow it to settle for 10–15 min. (Protoplasts suspended in CPW13M for convenience are mixed in screw capped centrifuge tube in a 1:1 ratio).
• Add 450l of PEG fusion solution drop-wise to the edge of the protoplast culture, placing the last drop in the centre by using a Pasteur pipette.
• Leave the protoplasts undisturbed for 20–40 min at room temperature. Then add one drop of washing solution very gradually whilst withdrawing some of the fusion solution from the coalesced drops.
• Repeat this procedure at 5 min intervals for the next 20 min. During this procedure it is important that the protoplasts are subjected to minimal disturbance. Washing solution: CPW13M medium to which 0.74g/l CaCl2·2H2O is added (sterile).
• Five min after the last wash, carefully suck off the solution with a Pasteur pipette leaving the protoplasts with a thin film of medium and replace it with culture medium (MS protoplast medium with 9% mannitol). Wash at least 3 times with culture medium.
•  For the culture of protoplasts flood the cover glass with culture medium and scatter several drops of medium on the base of petri dish prior to sealing with parafilm.
• Incubate the cultures initially in the dark for 24 h at 25°C and then transfer to white light (1000 lux).
•  The fusion products can be easily observed using an inverted microscope.
• Subsequently transfer the colonies for regeneration.
• Test the hybridity of fusion product and somatic hybrid as per equipment available in the lab, which has been explained in the text.

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1. Plant materials- leaf of Nicotiana tabacum
2. Cell and protoplast washing (CPW) solution
3. 10% sodium hypochlorite
4. Tween 20
5. 13% mannitol-inorganic salts
6. Enzyme mixture: macerozyme (0.1–0.5%) and cellulase (0.5–1%)
7. Protoplast culture media: Kao and Michayluk media
8. Pasteur pipette
9. petri dish

Procedure: Isolation of protoplast from Nicotiana tabacum

• Isolate fully expanded leaves including petiole.
• Use 10% sodium hypochlorite solution containing two drops of Tween 20 for 10 min to sterilize it.
• Wash 3–4 times with sterile de-ionized water and blot dry on sterile tissue paper.
• With fine forceps, peel the lower epidermis from sterilized leaves and cut into pieces by using fine scalpel.
• Transfer the leaf pieces (peeled areas only) with lower surface down into 30 ml of 13% mannitol-inorganic salts of cell and protoplast washing media (CPW) solution contained in a petri dish for 30 min–1 hr for plasmolysis.

• Composition of Cell and protoplast washing (CPW) solution:
  • Ingredients                        mg/litre
  • KH2PO4—————————–27.2
  •  KNO3——————————–101.0
  •  CaCl2·2H2O———————- 1480.0
  • MgSO4·7H2O———————-246.0
  • KI ————————————–0.16
  • CuSO4·5H2O———————–0.025   
  • *pH maintained at 5.8

• Take out the mannitol-CPW salt solution with a Pasteur pipette and replace it with filter sterilized enzyme mixture( macerozyme and cellulase)in 13% mannitol solution (~ 20 ml).
• Gently stir the leaf pieces with sterile Pasteur pipette to enables the separation of protoplasts, pushing the larger pieces of leaf material to one side and keeping the petri dish at an angle of 15°C. Or, remove the larger debris by filtering through a 60–80 μm mesh.
• Transfer the filtrate to a screw cap centrifuge tube.
• Centrifuge at 100x g for 5–10 min to sediment the protoplasts.
• Remove the supernatant and resuspend the protoplast pellet in CPW + 21% sucrose (prepared in CPW) and gently spread the protoplasts in the solution.
• Centrifuge again at 100x g for 10 min. The viable protoplasts will float to the surface of the sucrose solution while the remaining cells and debris will sink to the bottom of tube.
• Remove the band of protoplasts from the top with a Pasteur pipette and transfer into another centrifuge tube.
• Resuspend in CPW + 10% mannitol. Centrifuge again at 100x g for 10 min to separate the contaminating debris.
• Repeat the washing procedure at least 3 times.
• After the final washing, add enough MS protoplast culture medium with 9% mannitol to achieve a protoplast density of ~ 5 × 104/ ml.
• Kao and Michayluk media has been widely used for protoplast culture. Plate the protoplasts as small droplets (100–150 μl) or a thin layer in small petri dishes for 24h at 25°C.
• Keep the cultures at low light intensity of 500 lux for the next 2 days and then for rest of the experiment at 2000 lux.
• Protoplasts form cell walls and begin to divide after 3–5 days.
• Protoplasts may also be cultured in an agarose medium by mixing 1.5 ml of protoplast suspension with an equal volume of 1.2% agarose medium at 45°C in 35 mm petri dish.
• Cultures continue to grow in the media and the first colonies become visible after 3–4 weeks.
• Transfer the colonies to new medium with a reduced mannitol level.
• Transfer the colonies to a suitable medium for embryo differentiation and plantlet regeneration.

Protocol for protoplast isolation

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