Micropropagation

Abstract

Clonal propagation of plants, which refers to multiplication of genetically identical individuals by asexual methods of regeneration from somatic tissues or organs, is a common practice in horticulture and forestry to preserve the desirable characters of selected genotypes or varieties in the progeny. Traditionally, it is achieved by cuttings, layering, splitting, grafting and so on. However, for many plants, especially the orchids and tree species, these methods are either very difficult or painfully slow. Since mid 1960s tissue culture has become an established industrial technology being widely used the world over to multiply orchids, other ornamentals, and fruit and forest tree species. In vitro clonal propagation of plants is popularly called micropropagation because of the miniaturization of the process. The major advantages of micropropagation over the conventional methods of clonal propagation are that it is much faster and goes on round the year, protected from pests, pathogens and vagaries of nature. This chapter describes the techniques of micropropagation, the factors that affect the various stages of micropropagation, and commercial aspects of micropropgation and discusses some of the common problems associated with it. The chapter is annexed with protocols and media composition for the micropropagation of selected crops.

Keywords

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Appendix

• 1. Protocol for micropropagation of the orchid—Phalaenopsis (after Paek et al. 2011)

  • (i) Excise 3–4 cm segments from floral stalk at the 3–5 open flowers stage, each with single lateral bud. Surface sterilize the segments with 3 % sodium hypochlorite solution containing one drop of Tween 20 for 10 min. Rinse the segments three times in sterile water, trim-off the bleached ends of the sections and plant them on VW medium (Table 17.1) containing 2 % sucrose, 20 % coconut water (CW), 3 mg L⁻¹ BAP or 0.1 mg L⁻¹ TDZ, and 1 % agar.¹

  • (ii) Incubate the cultures at 26–28 °C under 16 h photoperiod and 30 μmol m⁻² s⁻¹ irradiance.

  • (iii) After 1–2 months, when 2–3 leaves have developed, excise the youngest leaf and cut 5–7 one mm thick sections from its proximal end. Soak the segments in half strength MS liquid medium for 2 h.

  • (iv) Plant the sections on half strength MS medium + 2 mg L⁻¹ TDZ or 10 mg L⁻¹ BAP + 10 mg L⁻¹ adenine sulphate + 20 % CW +2 % sucrose + 0.23 % Gelrite, and pH adjusted to 5.7.

  • (v) After incubation in dark at 27 °C for 1 week, transfer the cultures to 25 °C under 16 h photoperiod and 20 μmol m⁻² s⁻¹ irradiance for 6 weeks.

  • (vi) Excise the PLBs from leaf segments, trim their apical portion to remove the small shoot and leaf and transfer them (20 pieces per 10 cm Petri plate containing 25 ml of medium) to protocorm multiplication medium (1 g L⁻¹ each of Hyponex- N:P:K- 6.5:6:19 and Hyponex- N:P:K 20:20:20 + 2 g L⁻¹ Peptone + 10 % CW + 30 g L⁻¹ homogenate of unsprouted potatoes + 0.05 % activated charcoal (AC) + 0.8 % Agar, with pH adjusted to 5.5).

  • (vii) Subculture the protocorms every 4 weeks as above. The protocorms can be cut into 2–4 longitudinal sections to enhance the rate of multiplication (10–20 protocorms per piece after 4 weeks). Since the basal tissue of protocorms shows genetic instability it should be discarded. Also discard the small, yellow protocorms at the time of subculture to avoid formation of abnormal plants. Do not extend the multiplication of PLBs from one explant beyond a year.

  • (viii) Maintain the cultures at 25 °C under 16 h photoperiod and 30 μmol m⁻² s⁻¹ irradiance for 4 weeks.

  • (ix) Once the PLBs develop the first leaf transfer them to 1st TP medium (1 g L⁻¹ each of Hyponex- N:P:K- 6.5:6:19 and Hyponex- N:P:K 20:20:20 + 2 g L⁻¹ Peptone + 10 % CW + 0.05 % AC + 0.8 % Agar, with pH adjusted to 5.5) as for symbiotic seed germination and subsequently to grow.

• 2. Micropropagation protocol for Musa accuminata (after Singh et al. 2011)

  • (i) Remove sword suckers from healthy disease-free stock plants. Cut it to expose the shoot tip of 10 cm³ and trim further to about 3 cm diameter and 5 cm length without damaging the meristem.

  • (ii) Wash the shoot tip thoroughly in tap water, and surface sterilize with 0.1 % mercuric chloride for 10 min. Wash thoroughly under running tap water.

  • (iii) Carefully remove the outer 1–2 leaves and trim the corm base with sharp sterilized scalpel. Wash it with sterile distilled water three times under aseptic conditions, and disinfect with 5 % sodium hypochlorite followed by 0.1 % mercuric chloride, each for 15 min.

  • (iv) Wash three times in sterile distilled water and trim to remove 1–2 mm of peripheral tissue of the corm.

  • (v) Transfer the shoot tip to culture initiation medium (Table 17.3; 30 ml medium in 250 ml glass jar container), and incubate the cultures at 24–26 °C, under 16 h photoperiod and light intensity of 1500–3000 lux.

    Table 17.3 Culture media used at different stages of Banana micropropagation at the National Research Centre of Banana, India (after Singh et al. 2011)

  • (vi) As the explant turns green after 20–25 days remove the shoot tip from the jar, trim to remove the outer dead tissue and peel off 1–2 leaves to the base until fresh meristem tip gets exposed. Give two gentle cross incisions to the meristem tip and transfer it to multiplication medium (Table 17.3).

  • (vii) In 20–15 days the meristem would have produced a cluster of proliferating buds, and 1–3 axillary buds would have regenerated from the basal part of the explant around the central apical meristem.

  • (viii) In subsequent 3–4 weekly subcultures trim to remove the dead basal tissue and cut the shoot cluster into 3–4 pieces, each with up to four shoot buds and transfer to fresh multiplication medium.

  • (ix) After 5–6 cycles of shoot multiplication transfer the proliferating shoots to (Table 17.3) rooting medium. Proliferating shoots can also be rooted in polybags under greenhouse conditions to reduce the cost and better establishment of the plants ex vitro.

  • (x) For hardening, transfer the plants from culture vessels and leave exposed in a room at ambient temperature for 4–6 days.

  • (xi) Wash the plants (8 cm shoots with 3–4 ramified roots) to remove the agar medium and plant individually in micropots in a protray. Keep them under high humidity (90–95 %) and diffuse light for 6–8 days. Reduce the humidity and raise the light intensity gradually.

  • (xii) After 5–6 days remove the plants from micropots, dip in fungicide solution (0.1 % Bavastin) and plant in polybags and maintain under 70 % RH, which is gradually reduced to 40 %.

• 3. Micropropagation protocol for Gladiolus (after Dantu and Bhojwani 1992)

  • (i) Dehusk the corm and scoop out individual axillary buds along with corm tissue. Trim the corm tissue to 5 mm³.

  • (ii) Rinse the excised buds in 90 % ethanol, air dry for 15 min, dip in a 0.5 % solution of Cetavlon or any other surfactant for 5 min and wash thoroughly under running water for 60 min. Surface sterilize the buds in a 2.5 % (w/v) solution of sodium hypochlorite for 15 min and give three quick rinses in sterile distilled water under aseptic conditions.

  • (iii) Trim the peripheries of the corm tissue and plant the bud on MS + 0.5 mg L⁻¹ BAP (BAP concentration might vary with cultivar) and incubate the cultures at 25 °C in continuous light of 10 W m⁻² s⁻¹ irradiance.

  • (iv) After two passages of 4 weeks each on the same medium as in step (iii), when multiple shoot buds have developed, cut the bud cluster into pieces, each bearing at least five buds and transfer them individually to shoot multiplication medium (Table 17.4).

    Table 17.4 Gladiolus shoot multiplication medium (after Dantu and Bhojwani 1992)

  • (v) At the end of 4 weeks again cut the shoot clusters into small propagules each bearing about 10 buds and transfer them individually to fresh shoot multiplication medium (Table 17.4). At this stage the cultures are incubated at 25 °C in complete dark.

  • (vi) For shoot elongation, cut the shoot clusters into small clumps each bearing about 10 shoots and transfer them to liquid MS medium containing 0.5 mg L⁻¹ BAP. Incubate the cultures as in step (iii).

  • (vii) After 4 weeks separate individual shoots (ca 10 cm long) transfer them individually to MS liquid medium with 6–10 % sucrose for corm formation. Incubate the cultures in continuous light. Corms should be ready for harvest in 10–12 weeks.

  • (viii) Remove the corms, wipe off the medium, and store them at 5 °C for 6–8 weeks.

  • (ix) Sow the corms directly in the field following the standard horticultural practice. The corm should sprout within a fortnight.

• 4. Micropropagation protocol for Ficus lyrata (after Jona and Gribaudo 1988)

  • (i) Select fully opened but not completely expanded leaf that is still very soft as the source of explants. Bring the leaves to the laboratory for further preparation. Care should be taken that the mother plant is growing in bright conditions and is healthy.

  • (ii) Wipe the whole leaves with 70 % ethanol, and allow it to dry. Dip the leaves in water containing 2–3 drops of Savlon for about 5 min followed by thorough washing under running tap water for about 15 min. Place the leaf on a Petri plate and remove the margins on both side of the mid rib by cutting with a sharp blade leaving about 1–1.5 cm of margin on both sides of the mid rib. Cut the mid rib portion of the leaf into 1 cm long pieces resulting in 1 × 1.5 cm ready to use leaf explants.

  • (iii) Sterilize the leaf pieces in 1 % sodium hypochlorite solution for about 12 min. Remove all traces of hypochlorite by washing 3–4 times with sterile distilled water in a laminar air flow hood.

  • (iv) Inoculate the leaf pieces on the initiation medium, consisting of Nitsch and Nitsch (1969) basal medium + 24 μM BAP + 24 μM Kn + 24 μM 2iP + 12.5 μM IBA in culture tubes (Table 17.5). Incubate the cultures at 24 ± 1 °C and continuous light. Leave the cultures undisturbed for almost 3 months. If medium in the culture tubes is sufficient there is no need of transfer to fresh medium.

    Table 17.5 Culture media for adventitious shoot differentiation from leaf discs (shoot multiplication) and rooting of the shoots (rooting) of Ficus lyrata (after Jona and Gribaudo 1987)

  • (v) At the end of 3 months tiny adventitious shoots could be seen arising all along the margins. In another fortnight these shoots grow slightly big. At this stage, the leaf explants along with the young shoots are transferred to fresh medium to allow the shoots to grow.

  • (vi) The cluster of shoots is cut into small clumps and inoculated into fresh medium of the same composition for shoot multiplication.

  • (vii) Individual shoots, about 5–6 cm long, are transferred to the rooting medium (Table 17.5).

  • (viii) Rooted shoots are acclimatized in a net house.