Low-Cost In Vitro Options for Banana Mutation Breeding

Abstract

Mutation fixation of irradiated banana cultures is achieved through at least three generation advancements by in vitro subculturing. The in vitro culture is a technique which allows rapid multiplication of plantlets within a short time and which often relies highly on expensive inputs that are almost unaffordable in many developing countries. This chapter highlights some easily affordable options that can be adopted for in vitro propagation and weaning of tissue-cultured banana plantlets and other horticultural crops. The presented options provide resource restricted laboratories opportunities to coopperate with irradiation facilities for mutation induction. Thus, when applied to any locally selected banana variety, the low-cost in vitro methods allow an efficient mutagenesis process to improve local accessions. Low-cost alternatives adopted to carry out in vitro mutagenesis activities in the current FAO/IAEA project are presented, by using as baseline other cheaper options developed and adapted through a locally funded project (supported by the Mauritius Research and Innovation Council).

1 Introduction

Banana breeding of popular varieties through conventional methods is a long and tedious process, challenged by low female fertility, polyploidy and heterozygosity (Jenny et al. 2002; Ortiz 2013). Unlike seeded species, the classical cross-breeding for genetic improvement of the mostly sterile and seedless banana can last over several years. This has prompted many countries to have recourse to mutation breeding as an alternative method for the improvement of banana.

Earliest studies on the effect of radiation on plant development dates back to 1928 by Lewis Stadler (Stadler 1928; FAO/IAEA 2011) with groundwork on induced mutation breeding of banana starting in the early 1970’s by De Guzman, Ubalde and Del Rosario (De Guzman et al. 1976). Since then, significant advancement has been achieved in mutation breeding of banana through use of physical mutagens (X- or gamma rays) or chemical mutagens (ethyl methanesulphonate, EMS) (FAO/IAEA 1995). Successful applications of mutation in genetic improvement of banana include the development of an early flowering mutant of banana cv. Grande Naine, named ‘Novaria’ by F. J. Novak (Mak et al. 1996) and an improved local variety named “Al Beely” banana (FAO/IAEA 2011). Furthermore, Jain et al. (2011) reported on ten banana mutants/variants that were identified as promising with improved traits such as earliness, improved bunch and fruit size, reduced plant height or tolerance to Fusarium wilt.

Countries facing important banana pest and disease pressures can have access to improved genotypes such as FHIA hybrids developed by the Honduran Foundation for Agricultural Research, however, many countries still opt to improve their popular locally adapted variety. Mauritius is one such country which embarked on mutation breeding of its popular banana varieties through the joint support of the FAO and IAEA. A first project aimed at inducing tolerance/resistance to Panama disease in a highly prized local dessert type banana, namely ‘Gingeli’ banana (AAB genome) from year 2004 to 2015 using gamma rays on in vitro cultures. In the absence of hotspots, the generated mutant lines were screened at greenhouse level and the surviving lines were multiplied for field testing. Till date, no promising line has been identified. In another FAO/IAEA funded project, mutagenic treatments were performed on three other popular varieties, the ‘Mamoul’ (ABB), ‘Ollier’ (AAA) and D. Cavendish banana (AAA) with aim to induce tolerance to Fusarium wilt race 1 (in ‘Mamoul’) and race 4 in the others.

In vitro mutation breeding is a powerful tool for induction of desired traits. However, the restricted access to irradiation and in vitro facilities remains among the main withdrawal factors for these countries to embark on banana breeding using plant biotechnology techniques. The Plant and Breeding Genetics laboratory of the FAO/IAEA in Seibersdorf, Austria is one such laboratory that supports developing countries by irradiating plant materials, free-of-charge. Similarly, Mauritius can also extend physical mutagenic treatments to neighboring countries.

In vitro culture methods are used in banana mutation breeding whereby emerging buds from the mutagenized cultures are isolated and brought back into culture, through a series of up to four subcultures (generations) (M₁V₀ to M₁V₄) which then allows the mutation to be fixed in the genome (in vitro mutagenesis) (Roux 2004). The in vitro tissue culture (TC) is often associated with high production costs as it relies on aseptic conditions, high quality chemicals, high-energy and expensive glassware which restrict its broad application in plant breeding. To make TC affordable without compromising the quality of the plants, several low-cost alternative components of tissue culture have been considered and adopted (Kodym and Zapata-Arias 2001; FAO/IAEA 2004; Ogero et al. 2012; Datta et al. 2017).

A minimum, recommended number of shoot tips (500–1000) can be irradiated and if managed in batches, the number of plants generated by the M₁V₄ population can be easily handled using the low-cost approach. The sections below highlight the low-budget approach of in vitro banana mutation breeding.

1.1 Low-Budget In Vitro Options

1.1.1 Infrastructure

A typical TC laboratory consists of rooms dedicated to each aspect of in vitro culture: A first room (usually the first entry point) is for the reception and handling of mother plants received from the field. The other rooms include the media preparation room, the transfer room with laminar flow cabinets which serve as an aseptic area for handling of in vitro cultures and a growth room with controlled temperature and light for growth of cultures. In a low-cost situation, a simple large room or two-rooms can be partitioned to accommodate the above areas while respecting the flow of each section and ensuring that the aseptic area is against the flow of air.

1.1.2 Consumables and Equipment

Banana cultures are mainly established in Murashige and Skoog (MS media) or modified MS media supplemented with auxin and cytokinin. The media can be prepared by using powdered form of ready-made MS media or by mixing aliquots of four stock solutions of macronutrients, micronutrients, iron and vitamins (see Annex 10.1). The latter is more accessible and relatively cheaper for developing countries. High quality analytical grade chemicals are expensive and not a requisite for routine TC. On the other hand, commercial grade chemicals of lower purity can be easily procured from school suppliers that provide such chemicals as teaching materials and their use can lead to up to 70% savings (Dussoruth et al. unpublished). In a study, Gitonga et al. (2010) reduced the cost of producing tissue culture banana seedlings by 93.9% by using alternative nutrient sources. Similarly, Ogero et al. (2012) successfully realized cost reduction of over 80–95% by substituting conventional MS compounds by locally available fertilizers (for macro/micronutrients), over-the-counter vitamins (for vitamins) and seaweeds or agricultural plant hormones (for plant growth regulators, PGR’s). However, Dussoruth et al. (unpublished) found that the quality of agricultural fertilizers had significant effect on in vitro plants. Aliquots of field fertilizers used to make the MS media did not dissolve well, leaving behind deposits of impurities that reduced the effective percentage of each element in the media. Moreover, the explants inoculated in these media died due to phytotoxicity. On the other hand, MS media made up with hydroponic salts did not cause phytotoxicity but significantly reduced the multiplication rate of banana plants. Thus, all substitutions that aim in cost reduction need to be fine-tuned as these can affect the quantity and quality of the plants.

Gelling agents and high-grade sucrose constitute the most expensive components of the culture media (Sahu and Sahu 2013). The gelling agents account for over 70% of media costs while the media chemicals contribute to 5–15% of production costs (Prakash 1993). Dussoruth et al. (unpublished) also noted that when both high-quality chemicals and analytical grade sucrose are used for MS media, sucrose becomes the largest components of MS media (63–78%) and this is reduced to about 3–8% when replaced by household sugarcane-based sugar (see Table 10.1). On the other hand, when commercial grade chemicals and household sugar are used, the gelling agents account to approximately 70–80% of media cost (Dussoruth et al. unpublished). As the media chemicals (nutrients and PGR’s) contribute to only about 10–20% of overall cost of 1 L media (see Table 10.1), a significant cost reduction in media is possible mainly by substituting the expensive high-quality gelling agents and analytical sucrose by cheaper commercial substitutes. It is now standard practice by many laboratories to use commercial sugar in media preparation.

Table 10.1 Percentage cost contribution of each component in MS media depending on quality of chemicals and quality of gelling agents used

Banana cultures can be generated in solid, semi-solid or liquid media. The liquid media allows a more rapid proliferation (Ahloowalia and Prakash 2004) but relies on expensive orbital shaker at around 60–80 rpm for media aeration to prevent asphyxia of the explants. The orbital shaker can, however, be replaced by a locally mounted shaker. In the absence of a shaker, the cultures can be multiplied in stationary liquid media by using supports as interface (matrix), which serve as an intermediate layer between the liquid and explants. Cheaper matrices such as marbles (beads), paper shreds, glass wool, filter paper bridges, cotton fibers among others were used by several researchers as reported by Prakash et al. (2004) and Datta et al. (2017). Dussoruth et al. (unpublished) assessed the effectiveness of sugarcane based bagasse, cotton wool, paper shreds, tissue-paper, marbles (see Fig. 10.1) for micropropagation of banana and African violet (an ornamental, Saintpaulia ionantha). Tissue-paper and cotton wool were as effective as phytagel or agar and agitated liquid, but the disadvantage was that explants, including the roots, were strongly attached to the matrix. The size of the marbles are important because large spaces among the beads can lead to drowning of explants.

Fig. 10.1

Use of low-cost physical support matrices for growth of cultures. The matrices (from left: cotton, cotton, tea paper, marble, liquid-agitated, bagasse) and the banana cultures, 4 weeks after subculture

As shakers/agitators add to production costs, many laboratories opt for semi-solid or solid media by using gelling agents which contribute to the viscosity of the medium. The solidified media acts as an interface allowing the explants to be seated in the media while uptaking necessary nutrients for growth and development. Conventional gelling agents are agar, agarose, and gellan gum (marketed under trade names such as phytagel or gelrite) (Prakash et al. 2004). The gellan gums produce transparent media, allow easy detection of contamination and are of higher quality than agar, however they are often expensive. In Mauritius, the gelrite and phytagel cost around 1200–1600 €/kg and 500–700 €/kg respectively while agar costs 90–400 €/kg, depending on the purity. To reduce the cost of media, cheaper sources of phytagel can be mixed with agar. Agar is the most widely used gelling agent, since it is usually unnecessary for high purity agar in large-scale micropropagation (Prakash et al. 2004). Extensive research has been done into cheaper alternatives to replace agar and gellan gums or in combination with other gelling agents (Puchooa et al. 1999; Wilson and Tenkouano 2020 and as compiled by Prakash et al. 2004; Datta et al. 2017).

Dussoruth et al. (unpublished) tested household tapioca pearls (locally called as ‘sagoo’), rice flour, cornstarch and arrowroot as alternative gelling agents, of which rice and tapioca were least suitable. Arrowroot gel led to improved plant development compared to agar. However, the 100% cornstarch-based or arrowroot-based media is firm and the opaque white to grayish-white color make difficult to detect any contamination (see Fig. 10.2). Addition of agar (1:1 ratio) softened the media while allowing explant growth and development. To allow detection of contamination during the initial stages, it is advisable to avoid the opaque media and use mainly agar as gelling agent. An agar or phytagel-based medium can be cooked in a microwave to produce a homogeneous medium prior to dispensing in culture jars, while the household gelling agents need to be carefully cooked in a pan on a stove with continuous stirring, to avoid lumps.

Fig. 10.2

Effects of different low-cost gelling agents and physical matrices. Effect of (from left to the right) agar, arrow root, cornstarch, arrowroot plus agar and cornstarch plus agar on banana shoot proliferation. Addition of agar to the low-cost gelling agents improved the structure of the media

The protocol in ‘Methodology’ section describes the banana micropropagation on modified MS media solidified with agar-cornflour (1:1) and agar-arrowroot (1:1). Growth of cultures was comparable to those grown in agar-based media (see Fig. 10.2). Cost of cornstarch and arrowroot was around 4 €/kg which is only 0.1% of the cost of agar. As 80–100 g of the cornstarch and arrowroot is used per litre of MS media compared to 2.5 g/l phytagel and 8.0 g/l agar, this represented a respective 60% and 80% savings over agar and phytagel.

Water is another main component which is used to make stock solutions and the culture media. Conventionally, distilled, doubled distilled or de-ionized water is used (Ahloowalia and Prakash 2004). These are expensive and can further significantly increase the cost of production if they are operated using electricity. As reported by Ahloowalia and Prakash (2004), several researchers used cheaper options such as operating distillatory with gas or altogether replacing distilled/deionized water with cheaper alternatives like tap, rain or bottled water. Experiments using distilled/deionized, tap, bottled and household-filtered water showed that cultures were successfully produced using filtered water (see Fig. 10.3). Filtered water was then adopted as the most economic source for preparation of both media and stock solutions used in this project. The unit cost of a home-scale filtration unit ranged from € 50 to € 125, depending on brand and quality compared to about € 5000 for a deionizer/distillatory unit leading to a cost reduction of over 95%.

Fig. 10.3

Effect of five water sources (from left to the right tap, bottled, household-scale filtered, deionized and double-distilled) used in media preparation. Performance of banana cultures in media made using filtered water was comparable to those from deionized water

Big commercial laboratories use high quality glassware that tend to be costly and fragile. Similarly, chemicals or stock solutions which are accurately measured during media preparation using expensive equipment can be approximate to the nearest amount. Over 90% cost savings can be made by using easily available, hardy and cheap wares such as syringes (for dispensing on aliquots of 1 ml to 50 ml) or plastic cylinders for larger volumes. These can easily be procured over the counter from school supply shops or medical stores. Access to culture vessels, namely ‘Magenta’ commonly used for TC may not be easily found in developing countries, alternatively recycled jam jars can be used. Cheap polypropylene (PP) lids can be used to replace the metal caps with added advantage that they do not rust and allow light to reach the cultures. Additionally, cling film or sterile PP plastic films can also be placed around the mouth of the jar and held tight using rubber bands (see Fig. 10.4). This method is not recommended, as wrapping the cover/film around the mouth of the jar takes more time than closing with a lid. It however, remains an option for those not having access to required lids. The Guangdong Academy of Agricultural Sciences (GAAS) uses a special PP pouch (see Fig. 10.4) for growing cultures both under natural and artificial LED lights.

Fig. 10.4

Options for culture vessels and covers. From left: jar covered with polypropylene (PP) cap, conical flask covered with an aluminium and jam jar with metal cap

While most activities can be easily handled using cheaper alternatives, some steps such as pH testing, require a closer monitoring. Some laboratories used pH indicator paper as low-cost option, however this method is not very accurate (Ahloowalia and Prakash 2004) because slight changes in pH are not easily detected through the indicator paper. Alternatively, portable hand-held cheaper pH meters can be used and they cost about 10% of analytical high accuracy pH meters. The expensive magnetic stirrers (€ 300 to € 900) can equally be substituted by simple ones at less than € 50.

An aseptic condition is a major requisite in any TC activity. Wares and media are best sterilized using autoclave, which can be costly. Commercial laboratories working with large volumes can afford autoclaves but this can be an expensive option for laboratories with small turn-over rates. Stericlave (portable medical pressure steam sterilizer) of above 40 l, which cost about 30% of a conventional autoclave is an affordable option (see Fig. 10.5a). Similarly a pressure cooker (20–25 l volume) which cost about 6–10% of an average autoclave can also be used (see Fig. 10.5b). Effective steam sterilization can be achieved through automatic setting of autoclaves at a pressure of 15 psi (1.05 kg/sq.cm) along with a temperature of 121 ° C for at least 30 min. On the other hand, the pressure cooker needs closer monitoring as overcooking can lead to caramelization of the medium (see ‘Methods’ section). Another point to consider is that a filled pressure cooker can be very heavy to lift and thus the height of the table, where the stove will be placed to heat the cooker, may need to be adjusted for easy handling. A pressure cooker is an easily available option for small laboratories where for example in a 20 l capacity pressure cooker, only about thirty 200–250 ml capacity culture (jam) jars can be stacked. As each jam jar is filled with about 30 ml medium, a total of only 1 l media can be sterilized at a time in a 20 l capacity cooker.

Fig. 10.5

(a) Stericlave. (b) Pressure cooker. (c) Low-cost, glass hood

In vitro culture manipulation is done inside a laminar hood in a transfer room under aseptic conditions. Instead of using high purity ethanol, commercial ethanol can be equally used to clean working surface and a flame to sterilize the scalpels and forceps. In the absence of glass beads sterilizer, the sterilization can easily be done by dipping the working ends of the tools in alcohol and then immediately flaming them over an alcohol lamp or gas burner. A laminar flow cabinet allows handling of cultures in a sterile environment and the cheapest ones varies around 5000–6000 €. If the laboratory intends to undertake TC works for many years, then it is advisable to invest in a laminar hood. Otherwise, if the hood is needed only to allow generation advancement of the mutated population, a glass hood can be designed such that the opening is wide enough only to allow easy manual handling of the cultures. The glass hood can be cuboidal about 1 m wide, 40 cm deep and 60 cm high with an opening on one side (of about 30–40 cm from base) (see Fig. 10.5c). The hood should be placed against flow of air and surface sterilized regularly.

After placing the explant in the media, the jar is immediately closed with the cover to prevent microbial contamination. A layer of parafilm is often additionally wrapped around the base of the cover to ensure that the entry point for any microbes is blocked. The parafilm, however, is relatively expensive and can be easily substituted by equally effective food wrap (cling films), which thus allows a 90% savings over parafilm.

1.1.3 Light and Energy

The cultures are conventionally placed on shelves illuminated with cool light fluorescent tubes (conventionally at 1200–2000 lux) (Ahloowalia and Prakash 2004) under a 16 h:8 h light/dark cycle at 25 ± 2 °C. Artificial lighting of cultures accounts to nearly 60% production cost (Ahloowalia and Savangikar 2004). However, many in vitro growing plants can tolerate wide fluctuations with temperatures (Ahloowalia and Savangikar 2004) as high as 30 °C and as low as 10 °C with improved plant growth (Kodym et al. 2001). Research works have also been reported on the performance of in vitro cultures under natural light using solatube (Kodym et al. 2001), nethouse, light-emitting diodes (LEDs) (Datta et al. 2017), domelight or diffused light (Ahloowalia and Savangikar 2004) and sidewise lighting system rather than downward illumination of culture racks (Datta et al. 2017).

In this project, all cultures were grown under natural light from either a solatube (purchased at around 1200 €) (see Fig. 10.6) or from a 1 m² domelight made of superior quality acrylic (purchased at around 120 €), that were fixed on the roof of a 9 m² room. In a separate study, cultures were grown in a netcloth shed that was lined with plastic running from the roof to the sides (see Fig. 10.6), and also in a room, receiving diffused light through the windows.

Fig. 10.6

Alternative sources of natural light that can be potentially used for micropropagation. Left image: Shed, lined up with plastic and netcloth, which can be rolled up depending on heat accumulation. Right image: Solatube directs diffused light into the room, without heating the room

Alternative culture growing conditions are summarized in Table 10.2. The solatube redirects the daylight through reflective tubing without heating the room and can illuminate an area of 3–5 m² (Kodym et al. 2001) whereas the domelight can heat up the room, requiring the window to be opened for air flow.

Table 10.2 Comparison of radiation and temperature of the different environment in Mauritus

Depending on the season, the number of buds produced per explant in diffused light was reduced or comparable to conventional controlled conditions. The growth rate was also slower under diffused light. Conditions with the solatube were more stable, while the domelight’s conditions were directly dependent on the weather. Proliferation rates were better in rooms lit by the solatube than by the domelight. While no electricity is involved in use of natural light, each system has its own advantages and constraints and need to be optimized prior to application (see Fig. 10.7).

Fig. 10.7

Response of banana cultures under different growing conditions, from left to the right: control, diffused light (2 plants), shed. Those grown under diffused light were comparable to control, but with slower rate. In shed, mortality and scorching were high. Remaining explants developed but with rate of development depending on the season. Development of rooted plantlets was also possible under diffused light

In the shed, the rate of development was not uniform. Mortality was relatively high (50–80%) depending on the outside conditions. The afternoon sun, especially in summer, led to intense scorching due to hot microclimate within the jars. This can be reduced by covering the west side of shed with shade netting. The other problem associated with growing cultures in a shed was the accumulation of dusts which intensified risks of contamination when brought back in the transfer room. However, cultures at rooting stage can be grown in such conditions as the plants can be sent to nursery for hardening.

In a commercial micropropagation laboratory, where the volume and quality of planting materials to be produced are crucial, there is often little concern about using cheaper options. The above low-cost substitutes have been proposed for countries with restricted or no TC facilities so that they can proceed with in vitro mutagenesis to improve any local banana cultivar for which conventional breeding can be too lengthy or complex. The low-cost alternatives can as well be included in a routine TC where up to 80% cost reduction can be achieved (see Table 10.3).

Table 10.3 Cost savings through adoption of low-cost substitutes of the TC components

1.2 In Vitro Mutagenesis

In vitro methods as described by Vuylsteke (1998) and Lopez et al. (2017), can be applied to mass multiply the desired local accessions to generate enough explants that can then be sent to another laboratory for irradiation. This section describes only the physical mutagen (gamma rays) which was applied on excised banana shoot tips. In order to reduce risks of chimerism, the explants are trimmed to produce shoot-tips of about 2–3 mm length.

After irradiation, the mutation is fixed in the genome by carrying out three subcultures (the whole process is referred to as generation advancement). In order to dissolve potential chimeric sectors during each subculture, longitudinal sections are made through the explant to select buds or propagules (see Fig. 10.8). The mother explant can be further subcultured to recover the main meristem mutation after homohiston formation.

Fig. 10.8

Explant with buds after incubation ready for generation advancement. The arrows indicate buds/propagules that will be carried over in the next generation after subculture. Longitudinal sections are made such that the propagules with part of mother tissue is transferred to fresh media

In practice, however, it is often difficult to dissociate the buds from the mother explant. In such cases, routine subculturing/decapitating the main explant is practiced to promote shoot proliferation. After irradiation, the cultures are referred to as the M₁V₀ generation and the buds thus produced (after 4–6 weeks of incubation) are subcultured and make the population of M₁V₁ individuals and this process is repeated after each incubation period until the population of M₁V₃ is reached. The resultant M₁V₃ population is placed in regeneration medium (usually of half strength macronutrients and without any PGR) for the buds/shoot to regenerate into plantlets (or referred to as putative mutant line). These individual mutant lines might be clonal propagated for multiple traits screening, particularly abiotic or biotic stress which usually uses destructive methods. Moreover, the development of multiple clones per individual is also important as avoid loss of the mutant lines during the weaning of ex vitro plantlets.

1.3 Weaning of Mutant Lines

Prior to screening the mutant lines for any traits of interest, such as to Fusarium wilt using the double-tray system or field tests, the rooted plantlets are hardened so that they adapt to the outside environment where the light intensity (4000–12,000 lux) is higher compared to growth rooms that are artificially illuminated (1200–2000 lux) and the relative humidity lower (40–80% v/s 98–100%) (Ahloowalia and Savangikar 2004). Weaning is preferably done gradually, starting with high humidity and partial shade which is then gradually decreased. Commercial laboratories rely on greenhouses with sections of (i) partial shading (70–80%) and intermittent misting, (ii) 40–50% shade with regular irrigation/overhead fine irrigation and (iii) 20% shade to full sunlight with irrigation. In order to support developing countries, an easy method of weaning is proposed to handle the mutant lines in batches.

The high humidity in the weaning stage is created by placing the plantlets on a table and covering them with a frame, lined with clear polyethylene sheeting, and regularly watering the plants (or mist using a spray bottle) (see Fig. 10.9). The partial shade is provided by covering the structure with shadecloth (70–80% shade). The plastic sheet is removed after about 2–3 weeks to allow the plants to adjust to ambient relative humidity. The hardening media should be freely draining and relatively rich. Jhurree-Dussoruth and Kallydin (2011) successfully hardened banana plantlets in media consisting of sterile soil and manure (see Fig. 10.9). The addition of perlite improves the porosity of the medium and is important mostly in the first stage hardening.

Fig. 10.9

Weaning and hardening stages. (a) & (b) Simple structures constructed out of metal framework, lined with polyethylene sheeting. During the hardening, the sheet covers the whole structure and is gradually lifted after 2–3 weeks, to allow plantlets to adapt to ambient humidity. (c) Testing of most appropriate potting media from locally available materials. Sterile mix of manure and soil (1:1) allowed rapid development of plants 4 weeks after weaning. (d) Fully acclimatized plants ready for transfer to the experimental plot

In this chapter, a low-budget TC and weaning procedure are presented to allow laboratories with limited resources to proceed with banana mutation breeding after an outsourced irradiation. Banana cultures in this project were irradiated using local facilities, but an additional section has been included to guide those who need to seek an international support for irradiation.

2 Materials

2.1 Preparation of Media

• 1. MS stock solutions I to IV made as per Annex 10.1 (stored in refrigerator).

• 2. Gelling agents (technical grade agar, commercial grade cornstarch and arrowroot starch).

• 3. Granulated table sugar.

• 4. Jam jars (250 ml) with metal caps.

• 5. Stove, pan and ladle.

• 6. Pressure cooker/stericlave.

• 7. Top loading weighing balance (1- 200 g, (2 d.p)).

• 8. Portable pH meter, calibrated with dilute (0.1 N NaOH and 0.1 N HCl for pH adjustment).

• 9. Magnetic stirrer with magnets.

• 10. Plastic, graduated measuring cylinder (100 ml).

• 11. Graduated 2 l plastic beaker.

• 12. Syringes (2 ml, 5 ml, 25 ml or 50 ml).

• 13. Rod.

• 14. Kitchen jug for dispensing of cooked media.

• 15. Refrigerator.

2.2 Shoot-Tip Establishment

• 1. Suckers of desired variety.

• 2. Knife.

• 3. Detergent dish soap in water.

• 4. Wetting agent (Tween or Teepol).

• 5. Broad spectrum fungicide.

• 6. Commercial bleach (at 2%).

• 7. Sterile water.

• 8. Magnetic stirrer/magnets.

• 9. Plastic container (1 l).

• 10. Glass jar (250 ml) with caps.

• 11. Laminar air-flow cabinet or glass hood.

• 12. Household alcohol.

• 13. Alcohol lamp.

• 14. Cotton roll.

• 15. Scalpel and sterile blades.

• 16. Forceps, medium (20 cm) and long (30 cm).

• 17. Culture jar with low-cost gelling agent-based MS medium (supplemented with BAP at 5 mg/l or 2 mg/l).

• 18. Smooth, white/off-white ceramic tile (15 × 15 cm).

• 19. Growth Chamber.

2.3 Shoot Tip Multiplication/Generation Advancement

• 1. Scalpel and removable sterile blades.

• 2. Forceps: medium (about 20 cm) and long (about 30 cm).

• 3. Smooth, white/off-white/creamy ceramic tile (15 × 15 cm).

• 4. Recycled newspaper.

• 5. Aluminium foil.

• 6. Household alcohol.

• 7. Alcohol lamp.

• 8. Cotton roll.

• 9. Laminar air-flow cabinet or glass hood.

• 10. Growth chamber.

• 11. Culture jar with low-cost gelling agent based MS medium (supplemented with BAP at 5 mg/l).

2.4 Growing of Cultures

• 1. Shelves (meshed to allow natural light to reach lower shelves).

• 2. Naturally lit room (either ‘solatube’ or ‘domelight’).

• 3. Minimum/maximum thermometer.

2.5 Mutation Breeding/Generation Advancement

• 1. Banana proliferating tissues (raised from shoot-tip, after fourth subculture onwards).

• 2. Irradiated banana cultures (stage M₁V₀).

2.6 Rooting

• 1. As in above Sect. 2.3, except item no. 11.

• 2. Culture jar with low-cost gelling agent-based MS medium (no PGR).

2.7 Weaning

• 1. Seedling trays.

• 2. Beaker or bucket for washing off of gels.

• 3. Clean pair of scissors.

• 4. Perlite, peat, sterile commercial soil, sterile farmyard manure.

• 5. Fertiliser (slow release or complex fertiliser such as N/P/K, 13:13:20:2).

• 6. Potting bags (2 l).

• 7. Water for spraying.

• 8. Spray bottle.

• 9. Wooden or metal-frame structure (5 m × 2 m × 1 m high) lined with clear polythene sheet.

• 10. Shade cloth (70–80% and 40–50% shade) to be placed on roof of shed.

• 11. Bench to place trays or pots.

3 Methods

3.1 Preparation of Stock Solutions

• 1. Prepare stock solution of plant growth regulator (PGR) at the rate of 0.1 g/l water for each PGR: 6-Benzylaminopurine (BAP) and Indole-3-acetic acid (IAA) (see Note 1).

• 2. Refrigerate until use.

3.2 Preparation of Multiplication Medium

• 1. In a beaker, add 100 ml of stock I, 1 ml of stock II, 5 ml of stock III and IV.

• 2. Add 50 ml of BAP, 2 ml of IAA.

• 3. Add 500 ml filtered water, add 30 g sucrose, stir until sugar is dissolved.

• 4. Pour in 1 l measuring cylinder or leave graduated beaker. Make up to almost 1 l.

• 5. Adjust pH to 5.8.

• 6. Pour in saucepan, select any of the two gelling agents

• (a) Agar/cornflour gelling agent: Add 40 g cornflour and 4 g agar (media are labelled as ‘AC5’ representing the 5 mg/l BAP)

• (b) Agar/arrow root gelling agent: Add 40 g arrow root and 4 g agar (media are labelled as ‘AA5’ representing the 5 mg/l BAP)

• 7. Stir well until there are no clumps.

• 8. Cook on slow flame until a semi liquid consistency is obtained.

• 9. Dispense 30 ml of media in culture jars and close jars immediately.

• 10. Sterilise in:

  • (a) Stericlave set at 121 °C at a pressure of 15 psi for 20–25 min

  • (b) Pressure cooker as follows:

    • Place a perforated plate (on a stand, with gap of about 4–6 cm) inside cooker, add tap water just below the rim

    • Place jars on plate, cover and heat cooker on medium flame until first whistle

    • Leave on medium flame for 25–30 min after first whistle

• 11. Allow to cool down, remove from cooker, close caps tightly and store at room temperature until use.

3.3 Preparation of Rooting Medium

• 1. Proceed as above (Sect. 3.2), except in step 1 use only 50 ml aliquot of Stock I, and in step 2 no plant growth regulator is added. The media are labelled as AC0 or AA0, respectively for the agar/cornflour and agar/arrow root gelling agents (see Note 2).

3.4 Sterilization of Tools

• 1. Wrap one long forceps, one medium forceps and scalpel in aluminium foil.

• 2. Wrap working tile in newspaper and then with aluminium foil (to save on foil, newspaper is used).

• 3. Sterilize as above (Sect. 3.2, point 10).

3.5 Sterilization of Working Surface

Prior to all works under laminar or hood, spray household alcohol to whole working table and side of hood with cotton.

3.6 Establishing Banana Cultures

This section highlights steps from culture initiation until cultures reach stage for mutagenesis. Details can be obtained in Vuylsteke (1998).

3.6.1 Culture Initiation

• 1. Select and uproot suckers from elite mother plants (take care to avoid damaging base which contains the meristem).

• 2. Wash thoroughly outside the lab and using knife, remove outer leaves to produce 5–7 cm tall and 3 cm wide block of tissue resting on 3 cm tall corm (base) (now referred to as explants). Bring inside the laboratory.

• 3. Wash explant in running water with washing detergent for 10 min.

• 4. Using sharp knife, trim explant to smaller size (3 cm tall and 0.7 cm wide block) of tissue on 1 cm tall corm.

• 5. Place explants in a beaker of water containing a broad-spectrum fungicide (0.5%) and a few drops of wetting agent and allow stirring for 5 min (see Note 3).

• 6. Bring the beaker under sterile hood. Ensure dissecting tools, sterile working tile, dipping alcohol and flame are ready for regular sterilization of the dissecting instruments.

• 7. Surface disinfect the explants in diluted household bleach (2%) for 15 min with a few drops of wetting agent and swirl frequently.

• 8. Decant bleach and reduce explant to size of about 1.5 cm tall using scalpel and sterile medium forceps. The aim is to remove tissues damaged by bleach and excise inner tissues containing the meristem.

• 9. Carry out three rinses in sterile filtered water for 5, 10, 15 min respectively (swirl frequently inside hood).

• 10. Further trim explants to about 0.5–1 cm tall that contain the meristem with corm. Trimming is done on the sterile tile using sterile dissecting tools (see Note 4).

• 11. Transfer (using long forceps) to initiation MS-based medium supplemented with 5 mg/l BAP+ 0.175 mg/l IAA. Media are labelled as MS5 (see Note 5).

• 12. Transfer jars to growth room. Cover jars with black cover for 3–4 days until explant turn green.

3.6.2 Multiplication of Shoot-Tip Cultures

• 1. After 2–3 weeks, bring jar to Transfer Room. Trim the brown/black/oxidised parts and base and transfer the explant into fresh multiplication medium. Bigger explants can also be cut longitudinally through the apex. This is referred to as fist subculture (s1).

• 2. In first subcultures, no new buds will be seen. Buds often start to be produced after the second subculture. During subculture, the original explants can be decapitated if they are big. They can also be split longitudinally. At that stage it is not necessary to cut off each single bud/sprout and inoculate separately, they can be transferred as small clumps.

• 3. During repeated subculture, split the bigger explants longitudinally and subculture each part. Depending on multiplication rate, repeat subculturing every 5–7 weeks, to promote rapid multiplication (see Note 6).

• 4. After repeated subculture of explants (often by 4th to 5th, depending on variety), the multiplication rate will increase rapidly, and explants will enter the log phase of multiplication (see Fig. 10.10). Select explants from these stages for irradiation.

Fig. 10.10

Number of buds produced from explant after each subculture (4–6 weeks) in banana (var. Dwarf Cavendish, AAA) at different BAP levels. The explants entered the rapid phase of multiplication after the 4th subculture. Explants for irradiation were selected among the population of explants from the 4th or 5th subculture onwards

3.6.3 Shoot-Tip for Mutagenesis – Dispatch and Reception

• 1. In order to reduce effects of chimerism in shoot-tip mutation breeding, the explants are trimmed to 4–5 mm length leaving as little vegetative tissue around the meristem as possible.

• 2. When preparing the bulk for irradiation, inoculate in small batches, as the trimmed explants can easily dehydrate if left on working surface during this process.

• 3. The explants are preferably inoculated on multiplication (solid) media (see Note 7).

• 4. It is advisable to carry out a radiosensitivity test to determine the optimum dose for bulk irradiation. Thus, this may necessitate that a first batch is sent beforehand for the radiosensitivity test.

• 5. The cultures should be carefully sealed and packed upright (see Note 8, Fig. 10.11).

• 6. Upon reception of the irradiated in vitro cultures, wipe jars with ethanol to surface sterilise culture.

Fig. 10.11

Sending banana cultures for irradiation to an international or regional facility. (a) Sealed culture box with banana shoot tips. (b) The seal should remain intact if arrangements have been made for irradiation without opening the culture vessel. (c) Culture vessels placed tightly in vertical position. (d) Box ready for dispatch

3.7 Generation Advancement Under Low-Cost Conditions

• 1. Transfer the irradiated explants into fresh multiplication medium. This stage is labelled as M₁V₀. All explants from the same variety can be bulked as they were irradiated with the same optimum dose. No additional nomenclature is adopted.

• 2. Maintain cultures in growth chambers under natural light and ambient temperature.

• 3. After 5–7 weeks (or until buds are produced), transfer the buds into fresh multiplication medium. Label the individuals in this population as M₁V₁ (refer to Sect. 1.2 and see Fig. 10.8). In order to increase possibility of having mutant lines that differ, isolate each bud and transfer. When buds cannot be dissociated from the original explant or when buds are compact around the explant, then split the explant longitudinally and advance the next generation.

• 4. Repeat this isolation of buds for two more times every 5–7 weeks until M₁V₃ generation is reached.

3.8 Regeneration of Mutant Lines

• 1. Transfer M₁V₃ cultures to rooting medium without growth regulator (AC0 or AA0). Low-cost medium can be used (agar/cornstarch or agar/arrowroot).

• 2. Maintain under growing conditions for 4–7 weeks or until the shoots develop roots (see Note 9).

3.9 Low-Cost Weaning

This stage concerns post-flask low-cost hardening of the in vitro derived plantlets.

• 1. Gently remove plants from jars and wash off all gel under running water.

• 2. Trim back yellow/dead leaves, dead roots and extra-long roots. Leave only soft green leaves and white primary roots.

• 3. Plug plantlets in plastic seedling trays filled with autoclaved potting media containing peat and perlite (1:1) or sterile:manure:perlite (1:1:1).

• 4. Keep the trays with the plantlets under partial shade (70–80%) and high humidity (70–90%). The humidity is maintained either through a misting system or by covering the trays with a plastic sheet and regularly watering the plantlets (or misting using a spray bottle).

• 5. After 2–3 weeks, gradually remove the polythene sheet to allow plants to adapt to ambient temperature and relative humidity. Leave the plants for further 2 weeks under same shade conditions.

• 6. After 3–4 weeks when the plants have reached an average of 10–15 cm, they are ready for further hardening at higher light intensities (50–40% shade) at ambient relative humidity. The plants can be either potted individually for field planting or potted in groups into a deeper, rigid potting box with a depth of at least 15 cm for immediate screening, as follows:

  • (a) Hardening for field planting: Individually transfer plugged plants into planting pots (1.5–2 l) containing autoclaved potting mix (soil:manure:perlite:peat at 1:1:1:1 or soil:manure:perlite at 1:1:1). Keep plants under 50–40% shade for about 1 month and later at full sunlight or 30% shade until they reach a height of about 30 cm for field planting. Use a slow release to fertilise the plants, otherwise use a locally available complex fertiliser high in nitrogen at the rate of ½ teaspoon per pot to avoid phytotoxicity (at 5 weeks interval).

  • (b) For nursery level screening of potted plants: Transfer plugged plants in large potting boats with bottom tray to collect exudates. Use same potting media as used above. The plants can be used immediately for screening using an adapted low-cost modified double-tray system and applying same practices as highlighted in this book.

4 Notes

• 1. For ease of preparation of the modified MS media from the salts (Annex 10.1), it is advisable to prepare separately stock solutions of macronutrients, micronutrients, vitamins and iron salts.

• 2. The ½ strength of macronutrients save on mineral salt.

• 3. Conventionally 2 min swirling with 95% absolute alcohol can be done if available.

• 4. During manipulations, usual sterile methods of working near flame and regular flaming of dissecting tools after dipping in alcohol.

• 5. To save on PGR, use lower concentration of BAP (2 mg/l) and transfer to required BAP level once no contamination is observed.

• 6. Multiplication rate will depend on conditions under which plants are growing.

• 7. In order to prevent entry of diseases some countries, like Mauritius, have strict importation guidelines. Thus, in such cases the vessels are dispatched, irradiated and returned without opening the jar (see Fig. 10.11). Where the quarantine conditions for banana is less strict, the cultures can be sent in any type of container for transfer into appropriate vessels for irradiation.

  If irradiation is done locally then the explants are gently placed horizontal on the media, so as maximise chance of irradiation. Sucrose can be excluded.

  • (a) For long distance (or international) transfers, the explants need to grow, and they can be gently pushed into the medium (at an angle), so that the latter receive nutrients for growth during transit.

  • (b) This aspect is handled depending on quarantine exigencies of the country requiring the service (see Table 10.4).

• 8. Prior to dispatch all necessary plant import/export formalities for international transaction need to be finalized. The exporting country need to provide necessary phytosanitary certificates that satisfy the plant import conditions of country receiving the consignment. An agreement has often to be finalized to ensure that irradiated cultures are received with minimum delay.

• 9. As from this point the mutant line is ready for either field screening (for agronomic performance and response to field biotic stresses) or simple greenhouse screening for FW disease.

Table 10.4 Conditions associated with outsourcing for mutagenic treatments

Annexure

Annex 10.1 Composition per litre of the modified Murashige and Skoog (1962) and preparation of stock solution