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Published in Issue No. 141, page 56 to 60 - (19761) characters

Use of microtubers for slow-growth in vitro conservation of potato germplasm

J. Gopal  Anjali Chamail  Debabrata Sarkar  


The cultivated potato, Solanum tuberosum L., is a tetraploid (2n = 4x = 48) and exhibits complex tetrasomic inheritance. It is highly heterozygous and segregates on sexual reproduction. Elite parental lines and cultivars of potato are thus maintained through vegetative propagation in order to maintain their genetic integrity. Maintenance of potato germplasm through field clonal propagation is time-consuming, requires large amounts of space and is labour-intensive. This also exposes the plants to disease, pests, and risks of loss due to abiotic stresses and natural calamities (Withers et al. 1990). Therefore, throughout the world, potato gene banks prefer to conserve elite parental lines and clones as in vitro propagated microplants under disease-free tissue culture conditions (Westcott et al. 1977; Golmirzaie et al. 1999; Gopal et al. 2002). When grown under optimum propagation conditions (MS medium with 30 g l−1 sucrose 16-h photoperiod 22–25°C), the microplants require sub-culturing after 4–8 weeks. In order to reduce the frequency of sub-culturing, growth of the microplants is restricted by employing growth retardants or osmotic stress in combination with a reduced energy source, low temperatures, low light intensity and varied photoperiod (Westcott 1981a,b; Estrada et al. 1983; Ishige 1995; Siddiqui et al. 1996; Lopez-Delgado et al. 1998). The use of low temperatures (6–8°C) and 16-h photoperiod (15–30 µmol m −2 s−1 light intensity from cool white fluorescent lamps) is almost universal in potato gene banks for conservation.

Microplants for in vitro conservation of germplasm are usually raised by using shoot tips or nodal cuttings as the explant. These shoot cuttings or microplants can also be induced to produce microtubers by incubating them under suitable conditions (Wang and Hu 1982; Estrada et al. 1986; Gopal et al. 1998). Though microtubers are convenient for handling, storage and transport of germplasm (Tovar et al. 1985; Kwiatkowski et al. 1988; Thieme 1992), their use for conservation of germplasm is not common.

In a previous study, we found that microtubers induced on MS medium supplemented with 60–80 g l−1 sucrose and 10 mg l−1 of BA (N6-benzyladenine) could be stored at 6–8 °C in diffused light for 12 months (Gopal et al. 2004). In order to increase the shelf-life of microtubers, efforts were made to enhance microtuber dormancy by adding abscisic acid ( ABA) to the medium, as ABA is known to be generally higher in dormant tubers (Korableva et al. 1980; van den Berg et al. 1991 Cvikrova et al. 1994; Suttle and Hultstrand 1994). However, an exogenous supply of ABA adversely affected both microtuber production and microtuber dormancy (Gopal et al. 2004).

In the present study, we explored the use of microtubers for in vitro conservation of germplasm on a slow-growth medium. Both freshly harvested and stored microtubers of three genotypes that had been induced using varying concentrations of sucrose and ABA were tested.

Materials and methods

Plant material

Three potato (Solanum tuberosum L.) varieties of differing maturity type were used for the study; ‘Kufri Chandramukhi’ (early), ‘Kufri Badshah’ (medium) and ‘Kufri Sindhuri’ (late). These varieties vary in tuber dormancy, tuber size, number of tubers, tuber dry matter and other characteristics. (Gopal and Kang 1988; Gopal et al. 1992)

In vitro microtuber production

Single nodal cuttings (SNCs) were dissected from 6–8 week old axenic plantlets that were routinely propagated and maintained on semi-solid (7g l−1 agar) MS (Murashige and Skoog 1962) medium supplemented with 30 g l−1 sucrose (adjusted to pH 5.8 before autoclaving), under standard culture conditions (16-h photoperiod; 40 µmol m−2 s−1 light intensity and 24 ± 1°C). The SNCs were sub-cultured on MS medium supplemented with 10g l−1 of BA and different combinations of three concentrations of sucrose (40, 60 and 80g l−1) and two concentrations of ABA (0 and 8µM). Three SNCs were cultured in each culture tube (25 × 150 mm) containing 15 ml of liquid medium. The culture tubes were closed with polypropylene caps and sealed with Parafilm M TM (American National Can Company, USA), and incubated under continuous dark at 19 ± 1°C in a culture room. After 90 days, when no further microtuber production was observed in any treatment, microtubers were harvested.

Conservation using microtubers

Harvested microtubers from each treatment were surface-disinfected by dipping in a 0.05% aqueous solution of mercuric chloride for 5 min, and divided into two lots. One lot was cultured on slow-growth conservation medium (MS medium with 40 g l−1 sucrose and 20 g l−1 mannitol and 7 g l−1 agar, with pH adjusted to 5.8 before autoclaving) and kept at 6 ± 1°C in a walk-in-chamber under a 16-h photoperiod (from cool white fluorescent lamps, approx. 20 µmol m−2 s−1 light intensity). The Central Potato Research Institute, Shimla, India routinely uses these conditions and this medium at this Institute for in vitro conservation of potato germplasm as microplants raised from nodal cuttings.

The second lot was stored in Petri dishes sealed with parafilm and kept at 6 ± 1°C in the walk-in-chamber under continuous light (from cool white fluorescent lamps, approx. 20 µmol m−2 s−1 light intensity) for 9 months. After this 9 months’ storage, these microtubers were also cultured on slow-growth conservation medium and incubated under conditions similar to the first lot.

The experiment was conducted in a completely randomized factorial design [2 types of microtubers (fresh vs. stored for 9 months) × 3 genotypes × 3 sucrose levels × 2 ABA levels] with six replicate culture tubes. Three microtubers were cultured in each culture tube (25 × 150 mm) containing 15 ml of slow-growth conservation medium (composition as above). Culture tubes were closed with polypropylene caps and sealed with Parafilm M TM (American National Can Company, USA). When possible, microtubers of the same size (average microtuber weight 90–110 mg) were used for the study.

Data recording and statistical analysis

Cultures were checked at 12, 15 and 18 month for the percentage of microplant survival and microplant condition (on a visual 0–5 preference scale: 0 = dead, 1 = very poor, to 5 = very good) for suitability for sub-culturing. Observations were also recorded on the presence, or absence, of aerial roots, microtubers or phenotypic abnormalities. The data on percentage microplant survival were transformed into arcsines and those of microplant condition into square roots (x + 0.5). Both non-transformed and transformed data were analyzed using standard procedures (Steel and Torrie 1980). CPCS1 statistical software ( PAU, Ludhiana) was used for the purpose.

Results and discussion

Analyses of transformed and non-transformed data led to similar results and conclusions, hence, for simplicity, only results based on non-transformed data are presented here.

Analysis of variance showed that the differences for microplant survival were non-significant at 12 and 15 months of conservation, and significant at 18 months of conservation due to the types of microtubers (fresh vs. stored) and genotypes. On the other hand, microplant condition score varied significantly at 12 months of conservation and became non-significant due to sucrose level at 15 months of conservation, and due to ABA and sucrose levels at 18 months of conservation. All interactions except microtuber type × genotype and microtuber type × ABA level × sucrose level were non-significant both for microplant survival and microplant condition at all the conservation periods (Table 1). A few other interactions, however, were significant only for microplant condition at one or the other conservation period.

Mean microplant survival showed that both fresh and stored microtubers had little mortality up to 15 months including the 9 months of storage in the case of stored microtubers. However, at 18 months of conservation, microplant survival was only 30% for the stored microtubers as compared with 79% when fresh microtubers were used (Table 2). At this conservation period, genotypic differences were also significant; mean microplant survival was 75% in ‘Kufri Sindhuri’ as compared with 44% in ‘Kufri Badshah’ and ‘Kufri Chandramukhi’. The condition of the microplants also deteriorated with an increase in the conservation period (Table 3). Microplants raised from stored microtubers were poorer than those raised from fresh microtubers at all conservation periods. The upper limit of conservation period for having good microplant condition (score = 4) was 15 months, but only when microplants were raised from fresh microtubers (Table 3).

The stored microtubers were used in the present study with the idea of prolonging the total conservation period, anticipating that microplants raised from stored microtubers would last for the same length of time as those raised from fresh microtubers after they had been placed on the conservation medium. However, the findings did not meet this expectation. The maximum possible conservation period for stored microtubers was 12 months (9 months in storage plus 3 months on the conservation medium (Table 3); after which the microplant condition score was below average (3 on 1–5 scale), indicating that the microplants would not be in good enough condition for subsequent regeneration or sub-culturing. This may be due to the fact that stored microtubers became senile during the 9 month storage period; culturing them on slow-growth conservation media thus resulted in poor microplant growth.

On the other hand, fresh microtubers produced vigorous microplants on sprouting and their growth could be better sustained by the medium, hence it would be advisable to use fresh microtubers for slow-growth conservation as those could be conserved satisfactorily up to 15 months with a mean microplant condition score of 3.93 (≈4 = ‘good condition’). Conservation beyond this period is not advisable because microplant survival at 18 months of conservation was as low as 44% in some genotypes (e.g. in ‘Kufri Badshah’ and ‘Kufri Chandramukhi’). Among the three genotypes studied, ‘Kufri Sindhuri’ has the longest maturity period under field conditions (Gopal and Kang 1988). The highest microplant survival at 18 months of conservation in ‘Kufri Sindhhuri’ thus indicates that in vitro and in vivo responses for foliage maturity are related to each other. Such relationships have also been observed for other characteristics (Gopal and Minocha 1998; Gopal et al. 2004).

Microtubers produced in the absence of ABA resulted in microplants with better condition. This may be because microtubers produced in the presence of ABA were of smaller size (Gopal et al. 2004). Thus ABA should not be used in the microtuber induction medium.

The effect of sucrose level on both microplant survival and microplant condition score was non-significant. The use of a higher concentration of sucrose is recommended as it promotes microtuberization (Wang and Hu 1982; Hussey and Stacey 1984; Gopal et al. 1998), and thus would produce more microtubers of bigger size.

The medium used in the present study was also supplemented with BA (10 g l−1) for promoting tuberization (Palmer and Smith 1969; Wang and Hu 1982; Hussey and Stacey 1984; Gopal et al 1998). The standard MS medium used for microtuber production in our laboratory contains 80 g l−1 sucrose and 10 g l−1 BA. BA is the normal component of microtuberization media in other laboratories including the International Potato Center, Lima, Peru (Estrada et al. 1986) because of the proven beneficial effect of cytokinins on microtuberization.

In general, the microplants raised from microtubers had normal growth with thick stems and broad leaves, whereas those raised by using nodal cuttings as explants had stunted growth with thin stems and small, or no, leaves. Although the percentage survival in some genotypes was as low as 44% at 18 months of conservation, surviving microplants raised from freshly harvested microtubers had produced a few new microtubers by this time. This may be due to stored resources present in the fresh microtubers (Garner and Blake 1989), that were not present in microplants raised from nodal cuttings and that had almost been exhausted in the stored microtubers before these were cultured. This indicates the possibility of developing a microtuber-to-microtuber cycle for conservation of potato germplasm using freshly harvested microtubers as explants (Kwiatkowski et al. 1988). The feasibility of this option is being investigated. To our knowledge, this is the first study of its kind wherein the effect of varying concentrations of sucrose and ABA on freshly harvested and stored microtubers has been tested for slow-growth conservation of germplasm.


The study was funded by the National Agricultural Technology Project (NATP) on Plant Biodiversity, Indian Council of Agricultural Research (ICAR), New Delhi, India.


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