Parasexual Hybridization

Abstract

Protoplasts are single cells that have been stripped of their wall exposing the plasmalemma. A special property of the protoplasts is that when brought into close contact they tend to fuse with each other irrespective of the sources of the protoplasts. The technique of fusion of isolated protoplasts from somatic cells and regeneration of hybrid plants from the fusion products, called somatic hybridization or parasexual hybridization completely bypasses the sex and, thus, allows combining genomes of two desirable parents, irrespective of their taxonomic relationship. The technique of cell fusion has been applied to produce a large number of intergeneric, intertribal or interfamily symmetric allotetraploid somatic hybrids, asymmetric hybrids and somatic hybrids. A unique advantage of parasexual hybridization is in creating new combinations of nuclear and/or cytoplasmic genomes, leading to enrichment of the current gene pool. According to Liu et al (2005) 90 intergeneric somatic hybrids have been obtained. This chapter describes the techniques of enzymatic isolation of protoplasts in large quantities, their fusion chemically and electrically, regeneration of somatic hybrids from the fused somatic cells, and application of parasexual hybridization.

Appendix

• 1. Isolation and culture of hypocotyl protoplasts from aseptic seedlings of Brassica napus cv isuzu. (after Chuong et al. 1985)

  • (i) Raise aseptic seedlings of B. napus on a medium containing 0.2 % sucrose and 0.8 % agar, in the dark, at 25 °C.

  • (ii) Excise 150–200 hypocotyl hooks (2–3 cm long) from 2 days old seedlings and plasmolyze them for 1 h in CPW 13 M solution¹ supplemented with (in mg L⁻¹) ampicillin (400), gentamycin (10) and tetracyclin (10).

  • (iii) Transfer the hypocotyl pieces to the enzyme solution containing 2 % Rhozyme HP-150, 4 % meicelase, and 0.3 % macerozyme R-10 in CPW 13 M in a Petri dish and incubate at 25 °C in dark on a shaker at 60 rpm.

  • (iv) After 12 h, gently agitate the enzyme mixture by pumping in and out of a pipette several times to enhance the release of the protoplasts.

  • (v) Filter the enzyme mixture through two layers of a nylon mesh (60 μm pore size on top of 44 μm).

  • (vi) Transfer the filtrate to a centrifuge tube and spin at 100 g for 3 min.

  • (vii) Suspend the pellet in CPW 13 M and spin again. Repeat this washing process three times.

  • (viii) Finally, suspend the protoplast pellet, at a density of 2 × 10⁵ protoplasts ml⁻¹ in Lichter’s medium,² supplemented with 13 % w/v sucrose, 5 g L⁻¹ Ficoll 400, 0.5 mg L⁻¹ BAP, 1 mg L⁻¹ NAA, 0.5 mg L⁻¹ 2,4-D with pH set at 5.7. Plate them in Petri dish (2.5 ml of protoplast suspension per 15 × 60 mm Petri dish) and incubate in dark at 25 °C.

  • (ix) After 4–6 weeks, transfer the floating microcalli to MS medium containing 200 mg L⁻¹ casein hydrolysate, 5 mg L⁻¹ BAP, 0.5 mg L⁻¹ NAA and 0.6 % agarose, with pH set at 5.7.

  • (x) After 3–4 weeks shoot buds may differentiate.

• 2. Isolation and culture of protoplasts from embryogenic suspension cultures of rice (Oryza sativa cv. Taipei 309). (after Davey et al. 2010)

  Isolation

  • (i) Surface sterilize the dehusked seeds of rice and, after three washings in sterile distilled water, culture them on LS2.5 medium (Linsmaier and Skoog’s medium) containing 1.0 mg L⁻¹ thiamine HCl and 2.5 mg L⁻¹ 2,4-D, and gelled with 0.8 % SeaKem Le agarose (FMC BioProducts, USA). Seal the dishes with Nescofilm or Parafilm and incubate in the dark at 28 ± 2 °C.

  • (ii) After 28 days, excise yellow/white compact and nodular embryogenic callus and transfer ca 1 g aliquots to fresh LS2.5 medium of the original composition.

  • (iii) Transfer 1–2 g (F.W.) of the embryogenic callus to 100 ml Erlenmeyer flasks each containing 25 ml of LS2.5 liquid medium and incubate the cultures on an orbital shaker (120 rpm) in the dark, at 27 °C.

  • (iv) Every 5 days remove the flasks from the shaker and allow the cells to settle. Carefully remove 80 % of the spent medium by decanting or using a sterile pipette, avoiding any loss of cells and reduction in cell density, and replenish it with fresh medium.

  • (v) After 42 days, transfer the cell suspensions to 250 ml flasks and add 15 ml of fresh medium to each flask. Every 5 days, allow the cells to settle, remove 30 ml of spent medium and replace it with fresh medium.

  • (vi) At the third subculture, pass the cell suspensions through a sieve of 500 μm pore size. Discard the large aggregates and retain the suspensions.

  • (vii) After 90–120 days, transfer the cells to the same volume of AA2³ liquid medium. Transfer the cultures every 7 days to new liquid medium by mixing 1 volume of cell suspension with three volumes of new medium.

  • (viii) Filter the cell suspension through a nylon sieve of pore size 500 μm into a preweighed 9 cm Petri dish and pipette out the liquid medium, leaving the cells in the dish. Reweigh the Petri dish and add appropriate volume of enzyme mixture (10 ml of enzyme solution g⁻¹ FW of cells), containing 0.3 % (w/v) Cellulase RS, 0.03 % (w/v) Pectolyase Y23 and 0.05 mM MES in CPW13 M solution, pH 5.6.

  • (ix) Seal the Petri dish with Nescofilm or Parafilm M and incubate them on an orbital shaker at slow speed (30 rpm), for 16 h in the dark, at 27 ± 2 °C.

  • (x) Filter the protoplast suspension through sieves of 64, 45, and 30 μm pore size to remove undigested cell clumps, and transfer the protoplast suspension to a sterile 15 ml centrifuge tube. Wash the protoplasts three times by gentle centrifugation (at 80 g for 10 min each) and resuspension in CPW 13 M solution.

  • (xi) Suspend the protoplasts in a known volume (e.g. 10 ml) of CPW13 M solution and count protoplasts using a hemocytometer. Stain an aliquot of the protoplast suspension with Tinopal or Calcofluor White to confirm that the cell walls have been digested completely.

  • (xii) Check the viability of the protoplasts by staining with fluorescein diacetate and observing under UV illumination.

  • (xiii) Resuspend the protoplasts in normal strength liquid K8p medium (Gilmour et al. 1989) + 2,4-D (3 mg L⁻¹) at a density of 5.0 × 10⁵ ml⁻¹ in 15 ml screw-capped centrifuge tubes. Heat shock the protoplasts by placing the tubes in a water bath at 45 °C and, after 5 min, plunge the tubes into ice for 30 s.

  • (xiv) Pellet the protoplasts by centrifugation at 80 g. Remove the supernatant and resuspend the pelleted protoplasts in fresh K8p liquid medium (normal strength). Repeat this procedure. Pellet the protoplasts.

  • (xv) Mix equal volumes of liquid K8p medium (double strength) with 2.4 % SeaPlaque agarose at 40 °C. Carefully resuspend the protoplasts at a density of 3.5 × 10⁵ ml⁻¹ in the resulting K8p agarose culture medium.

  • (xvi) Immediately dispense 2 ml aliquots of the protoplast suspension in K8p agarose medium into 3.5 cm diameter Petri dishes. Allow the medium with the suspended protoplasts to gel for at least 1 h. Seal the dishes with Nescofilm or Parafilm M and incubate the cultures in the dark at 27 ± 2 °C.

  • (xvii) After 14 days, divide the agarose layers from each dish into quarters with a sterile scalpel. Transfer each quarter to a separate 5 cm diameter Petri dish. Add 3 ml of K8p liquid medium (normal strength) to each dish. Incubate the cultures in the dark at 27 ± 1 °C until cell colonies develop from the embedded protoplasts.

  • (xviii) Mix MSKN (MS + 2 mg L⁻¹ NAA + 0.5 mg L⁻¹ zeatin +3 % sucrose) liquid medium (double strength) with 2.4 % SeaPlaque agarose at 40 °C. Immediately dispense 20 ml aliquots of the medium into 9 cm diameter Petri dishes. Allow the medium to gel for at least 1 h. Transfer the protoplast-derived cell colonies to the plate, seal it and incubate as in step (xvi).

  • (xix) Mix MSKN liquid medium (double strength) with 0.8 % SeaKem Le agarose (8 g L⁻¹) at 40 °C. Immediately dispense 20 ml aliquots into autoclaved screw-capped 50 ml glass jars. From 7 to 14 days-old-cultures in step (xviii), transfer somatic embryo-derived shoots with coleoptiles and roots to the jars (one shoot per jar). Incubate at 25 ± 1 °C in the light (50 μmol m⁻² s⁻¹, 16 h photoperiod, ‘Daylight’ fluorescent tubes).

  • (xx) Remove rooted plants from the jars and gently wash their roots free of semisolid culture medium. Transfer the plants to compost in 9 cm diameter pots, water the plants and cover with polythene bags. Stand the pots in trays containing water to a depth of about 10 cm in a controlled environment room (27 ± 2 °C with 12 h photoperiod, 180 μmol m⁻² s⁻¹, ‘Daylight’ fluorescent tubes).

  • (xxi) After 7 days, remove one corner from each bag and a second corner 3 days later. Continue to open gradually the top of the bags during the next 10 days. Remove the bags after 21 days.

  • (xxii) Maintain the protoplast-derived plants in a controlled environment room at 27 ± 1 °C with an 18 h photoperiod provided by mercury vapour lamps (310 μmol m⁻² s⁻¹). Transfer to glasshouse/field conditions as appropriate.

• 3. PEG (Polyethylene glycol)-induced fusion of protoplasts (the protocol may be slightly modified to suite the system under investigation). (after Kao 1976)

  • (i) Mix freshly isolated protoplasts from the two parental sources (while still in the enzyme solution) in a ratio of 1:1. Filter the suspension through a sieve of pore size 62 μm, collect the filtrate in a centrifuge tube and spin at 50 g for 6 min to sediment the protoplasts.

  • (ii) Remove the supernatant with a Pasteur pipette and wash the protoplasts with 10 ml of solution-I (500 mM glucose, 0.7 mM KH₂PO₄.H₂O and 3.5 mM CaCl₂.2H₂O, pH 5.5).

  • (iii) Resuspend the washed protoplasts in solution-I to make a suspension with 4–5 % (v/v) protoplasts ml⁻¹.

  • (iv) Pipette 4–6 drops (40 μl) of protoplast suspension onto a Petri dish (6 cm diameter) and leave it undisturbed for 5–10 min, so that the protoplasts settle on the bottom of the Petri plate.

  • (v) Add, drop by drop, 60 μl of the fusion mixture (40 % PEG 1,500, 10.5 mM CaCl₂.2H₂O, 0.3 M glucose, pH 6) to each drop and incubate the plate in the dark at room temperature (24 °C).

  • (vi) After 10–20 min, add two 0.5 ml aliquots of solution-II (50 mM glycine, 50 mM CaCl₂.2H₂O, 300 mM glucose, pH 9–10.5) at 10 min interval. After another 10 min add 1 ml of the protoplast culture medium.

  • (vii) Wash the protoplasts five times at 5 min intervals with the culture medium. At the end of each washing do not remove the entire medium; leave behind a thin film of the medium over the protoplasts and add fresh medium. If the protoplasts of the two parents are distinguishable, it may be possible to identify the heterokaryon formation at this stage.

  • (viii) Finally, add 2–2.5 ml of the culture medium and gently swirl the plate to make a thin layer of the medium. Seal the Petri plate with Parafilm and culture in dark for 4–5 weeks.

• 4. Microelectrofusion of tobacco protoplasts (based on Rákosy-Tican et al. 2001)

  • (i) Take a 33 mm diameter Petri dish and cover its bottom with 1 ml of mineral oil, purified by washing with 1 N HCl and 1 N NaOH, followed by washing with distilled water until the pH of the water is 7.

  • (ii) Using a micropipette of bore diameter of 100 μm, attached to a micromanipulator, place 20 microdroplets (30 nl) of 0.4 M sorbitol in a matrix formation in the center of the bottom of the dish.

  • (iii) Along the wall of the dish place microdroplets of equimolar sorbitol solution containing protoplasts of the two parents separately and blank droplets to maintain humidity inside the dish. The design of the microelectrofusion chamber is depicted in Fig. 14.4.

  • (iv) To each sorbitol droplet inject two protoplasts, one of each parent, using micropipette.

  • (v) Lower the electrodes, with a distance of 500 μm between their tips, into the drops, so that the pair of protoplasts is positioned between their tips. The electrodes, made of entomological needles of 90 μm diameter, are attached to another micromanipulator and sterilized by immersion in 70 % ethanol.

  • (vi) Apply AC electric field of 84 V cm⁻¹ and 750 kHz frequency to align the protoplasts.

  • (vii) After 30–60 s, apply 1–3 rectangular DC pulses of 500–670 V cm⁻¹ of 4 milli seconds to fuse the aligned protoplasts.

  • (viii) Leave the protoplasts undisturbed for 5–10 min to regain a spherical shape. Transfer the fusion product, by micromanipulation, to nurse cultures⁴ for individual cloning (Fig. 14.6), seal the plate with Parafilm and incubate in dark at 26 °C.

  • (ix) Transfer 1–2 mm cell colonies to RMO⁵ medium and incubate in light (37.8 μmol m⁻² s⁻¹ and 16 h photoperiod) at 26 °C.

  • (x) For shoot differentiation and elongation transfer the well-developed calli to RM medium containing BAP (1 mg L⁻¹) and NAA (0.1 mg L⁻¹) and gelled with 0.8 % Difco agar. Incubate the cultures in light (3,000 lux).

  • (xi) The shoots are rooted on semi-solid medium with RM salts (the concentrations of KNO₃, NH₄NO₃, and MgSO₄ reduced to one-fifth) and 3 % sucrose. The cultures are incubated at 20 °C, under 16 h photoperiod.