13

II. REVIEW OF LITERATURE

Turmeric (Curcuma longa L.) is an important spice crop valued for its dry rhizomes

which impart colour and flavor to food. India is the home and major producer of this crop.

However, the crop improvement in turmeric is restricted to selection of superior types from the

germplasm collected from different regions of the country. Reproductive sterility of turmeric due

to triploid nature (3n=63) is an impediment for improvement through conventional methods of

breeding.

Tissue culture studies in turmeric

Tissue culture is a term used for the growth of plants or more commonly plant parts in

sterile culture. Micropropagation is a method of propagating plants using very small sized parts

of plants that are grown in sterile culture. It is important commercially and can be used to

introduce several concepts that apply to all of tissue culture.

Morel (1960) was the first to show the potential of clonal propagation in culture by

describing in vitro multiplication of Cymbidium orchids. Several protocorms formed on each

explant, and when divided and transferred, additional protocorms formed. The rate of

multiplication was such that several million plants could be produced from one shoot tip in a

single year. As a result, many of the economically important orchids are now usually propagated

in vitro, although techniques have not been worked out for all orchid types. However, credit for

the development of the micropropagation technology in general goes without a doubt to

Murashige, who showed that many plants, in addition to orchid, could be propagated in vitro. He

14

along with his colleague was also responsible for devising a suitable medium for tissue culture

which is widely used (Murashige and Skoog, 1962).

Micropropagation has been shown to be useful especially in horticultural crops. The first

report of micropropagation of turmeric came from National chemical laboratory, Pune using

vegetative buds. Nadagouda et al. (1978) reported successful rapid clonal multiplication when

buds were cultured on MS medium supplemented with coconut milk and BAP. Successful

micropropagation was also achieved by Shetty et al. (1982) who observed both direct

organogenesis and callus production when rhizome buds were cultured.

Keshavachandran and Khader (1989) studied the response of Co-1 and BSR-1 cultivars

of turmeric for in vitro clonal propagation. Buds of these cultivars were cultured on MS medium

supplemented with 1 mg Kinetin /l, 1 mg BAP/l and 40 g sucrose/l with an average number of

shoots of 2.11 in BSR-1 and 2.5 in Co-1. After 5 weeks, rooted plantlets were transferred to pots

and two weeks later plants were well established. A multiplication ratio of 1:2240 was obtained

in a year through micropropagation in turmeric (Vidya et al., 1989). They reported profuse

rooting on minimal medium. Survival of plantlets was 77 per cent.

Balachandran et al. (1990) carried out in vitro clonal multiplication of Curcuma

domestica, C. aeruginosa, C. caesia and ginger by inoculating excised rhizome buds. For shoot

multiplication, 3.0 mg/l of BAP was found to be optimum for all the species. Rhizome buds of

both genera produced shoots and roots simultaneously and within four weeks, complete plantlets

were formed. They observed that the field established plants were morphologically uniform.

15

In vitro studies carried out at Regional Plant Resource Centre, Bhubaneshwar for PTS 28

and summer cultivar of turmeric indicated that shoot multiplication could be achieved in MS

medium supplemented with 4.0 mg/l BAP, 1.0 mg/l IAA and adenine sulfate at 10ppm. Shoots

successfully rooted on half strength MS medium supplemented with IAA or IBA at 0.25–0.5

mg/l and 2% sucrose (Rout et al., 1995).

At Indian Institute of Spice Research, Calicut, micropropagation was standardized using

the bud explants of turmeric on MS medium supplemented with 1 mg/l BAP and 0.5 mg/l NAA.

Such explants responded readily to culture conditions producing 8-10 shoots in 40 days

(Nirmalbabu, et al., 1993 and 1997). They could establish tissue culture plantlets in soil with 80

percent success. However, their opinion was that micropropagated plantlets take three crop

seasons to produce normal seed rhizome and hence the technique cannot be employed for

commercial multiplication.

Salvi et al. (2002) developed a protocol for in vitro propagation of turmeric cv. Elite

using vegetative buds form the rhizome. Multiple shoots were produced on MS solid medium

supplemented with BAP and NAA. Liquid medium was more favourable for shoot

multiplication. RAPD analysis of regenerated plants using sixteen 10 mer primers did not show

any polymorphism.

Rahman et al. (2004) obtained optimum development of multiple shoots for culture

explants on MS medium with 2.0mg/l BAP. Rooting of shoots was obtained on ½MS medium

with 0.1–1.0 mg/l. More than 70 per cent of transplanted plantlets survived in the field. Ali et al.

16

(2004) could get multiple shoots on MS medium supplemented with only BAP and 0.25 mg/l

kinetin. Rooting was achieved on MS medium with 1.0 mg/l NAA. All plants were

morphologically uniform.

Chan and Thong (2004) observed that medium used for in vitro multiplication of ginger

viz., MS medium with 2.0 mg/l BAP and 2.0 mg/l IBA was also suitable for in vitro propagation

of other Zingiberaceae species. Prathanturaung et al. (2005) studied the effect of thidiazuron on

multiplication of turmeric buds. MS liquid medium supplemented with 7264 micro M

thidiazuron (TDZ) prior to culture was used. Regeneration rate was up to 11.4+ 1.7 shoots /

explant.

Culture medium

Success of in vitro culture depends largely on the choice of nutrient medium, including

its chemical composition and physical form (Murashige, 1974). Several media formulations have

been reported for turmeric shoot tip culture but nearly half of them are modified MS media

(Brown et al., 1995). Other popular media include B5 (Gamborg et al., 1968), SH (Schenk and

Hildebrant, 1972), N6 (Chu et al., 1975), and Linsmaier and Skoog (LS) (Linsmaier and Skoog,

1975) media. The culture media vary in both type and concentration of the components, but all

have similar basic components of growth regulators, nitrogen, carbohydrates, inorganic

macronutrients and micronutrients, vitamins and organic additives.

Generally, the cultures are established on a separate initiation medium, which has a lower

concentration of cytokinin than the multiplication medium to which the cultures are subsequently

17

transferred (Jarret et al., 1985, Novak et al., 1989). The composition of initiation, multiplication

and rooting media were used. After autoclaving, the culture medium is stored in a clean dust free

chamber for 1-2 days before use in order to check for any contamination. Bacterial

contamination may be observed, particularly during the rainy season. Use of cefatoxime in the

initiation and subsequent subcultures helps to overcome even latent bacterial contaminations.

Culture initiation

The shoot buds are removed from healthy disease free mother plants for shoot tip culture.

The suckers are cut to expose the shoot tip of 10 cm3 and cut further to about 3 cm diameter and

5 cm length. The explants should be carefully cut to avoid injury to the growing meristem. The

shoot tips are washed in tap water and transferred to a container with 0.1% mercuric chloride for

5 minutes. Then the shoot tips are washed thoroughly under running tap water to remove all

traces of the chemicals. Afterwards, the rhizomatous buds are washed three times in sterile water

in aseptic condition (under laminar air flow) disinfected with 5% sodium hypochlorite and later

with 0.1% mercuric chloride each for 5 minutes. To avoid bacterial contamination, use of

cefataxime (0.1%) in the initiation medium is in vogue in some laboratories.

The outer surface of explants exposed to sterilizing agent is removed and the explants

trimmed using surgical blade to bring the final size to about 2-3 cm length and 1-2 cm diameter.

The explants are inoculated under sterile conditions in 30 ml of initiation medium in a 400 ml

glass jar container. pH is usually maintained at 5.8, which is prone to changes over culture

duration. The optimum incubation temperature should be in the range of 24-26°C. Generally the

light intensity is maintained at 1,500-3,000 lux. Higher levels of 3,000-10,000 lux during later

18

stages improve the survival rate of plantlets upon transfer to soil. Initially, the cultures are

maintained at 16 hrs light/8 hrs dark cycle and once after rooting they are shifted 14 hrs light/10

hrs dark cycles.

Decapitation and wounding of shoot tips are carried out to overcome apical dominance

and to encourage auxiliary bud proliferation. But injuring the apical bud through transverse

sections, either four or eight cuts, is a much preferred method. Injuring the explants encourages

more production of phenols, but it can be kept at minimum using antioxidants like ascorbic acid.

Culture proliferation

First subculture is done after 20-25 days of initiation when the explants turn green in

colour. The cultures are first checked for contamination, in general symptoms of fungal

contamination appear within one week and bacterial contamination symptoms like change of

medium colour and texture or visible colonies appear within one week to one month. For

subculturing, the outer dead tissue from the base of explants is removed and one or two leaf

bases are peeled till the fresh meristematic tip gets exposed. The apical meristem is cut with two

gentle cross incisions and the explants are transferred to subculture medium. During 20-25 days

after the first subculture, the central meristem produces clusters of proliferating buds and one to

three auxiliary buds get regenerated from the basal parts of explants around the central apical

meristem. The number of auxiliary buds developed during first and second subculture range from

1 to 5 depending on genomic constitution of the variety. Among the latter, the number of buds

produced during subculture. Subsequent subculture is done by trimming the tip of emerging

auxiliary buds and removal of dead tissue at the base of explants by gentle scratching. Clusters of

19

proliferating buds develop during third and fourth subculture. For further subculturing, the

explants is cut into three to four pieces and each slice with two to three proliferating clusters is

inoculated to individual culture bottles. This subculture cycle is repeated at 3-4 weeks interval to

increase the proliferation rate. During fourth and fifth subcultures, a single clump contains about

15-25 proliferating shoots. After 5-6 subculture cycles, the proliferated buds are transferred to

rooting medium containing IBA and activated charcoal. After a month, the rooted plantlets are

ready for hardening. To minimize somatic variation, the subculturing is restricted to a maximum

of seven cycles when each bottle contains 25-30 plantlets with well developed shoots and roots.

Experiments have demonstrated that proliferating shoots can be transferred to polybags (10-

20cm size) having rooting media under green house. This reduces cost and enhances better

establishment. Polybag provides enough space for plant growth and natural light enhances the

process of hardening.

German botanist Gottlieb Haberlandt (1902) was the first person to isolate and culture

single cells from the leaves of flowering plants. He used tissue of Lamium purpureum and

Eichhornia crassipes, the epidermis of Ornithogalum and epidermal hairs of Pulmonaria

mollissima. In 1934, Gautheret had cultured cambium cells of tree species like Salix capraea,

Populus nigra on Knop’s solution containing glucose and cysteine hydrochloride and recorded

that they proliferated for a few months. The addition of B-vitamins and IAA considerably

enhanced the growth of Salix cambium. However, the first continuously growing tissue cultures

from carrot root cambium were established by Gautheret in 1939. In the same year White

(1939a) reported the establishment of similar cultures from tumour tissue of the hybrid Nicotiana

glauca N. langsdorffii. Gautheret and White, together with Nobecourt, who had independently

20

reported the establishment of continuously growing cultures of carrot in the same year. Skoog

and Tsui (1951) had demonstrated that in tobacco pith tissue cultures the addition of adenine and

high levels of phosphate increased callus growth and bud formation even in the presence of IAA

which otherwise acted as bud inhibitor.

Biotechnological tools are important in order to select, multiply and conserve the critical

genotypes of plants. In this resurgent era of herbal drugs it is very difficult to make an accurate

assessment of the volume and value of herbal trade in India. According to estimates by the

Ayurvedic Drug Manufacturers Association (ADMA), the current value of trade in Indian system

of medicine (chiefly Ayurveda, Siddha and Unani) and Homeopathy is around Rs.4205 crores,

roughly close to US$ 1 billion (Natesh, 2001). Therefore, the value of plants is also reflected in

the economics of global market which was estimated to be 60 billion US dollars. The Asian

region rich in biological wealth and genetic diversity also has a substantial share in herb trade. It

is surmountable that plant biotechnology has grown from cell technology, specifically plant

tissue culture. Tissue culture is used for conservation of biological diversity for multiplying of

endangered species that have extremely small populations, for species with reproductive

problems and for recovery and reintroduction (Bromwell, 1990). Currently, tissue cultured plants

that have been genetically engineered provide insight into plant molecular biology and gene

regulation. Tissue culture will continue to play a key role in the genetic engineering process for

the foreseeable future, especially in efficient gene transfer and transgenic plant recovery

(Hinchee et al., 1994).

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In vitro cell and tissue culture methodology is envisaged as a mean for germplasm

conservation to ensure the survival of endangered plant species, rapid mass propagation for large

scale revegetation, and for genetic manipulation studies. Combinations of in vitro propagation

techniques (Fay, 1992) and cryopreservation may help in conservation of biodiversity of locally

used medicinal plants. Plant tissue culture techniques are discussed as integrated approaches for

the production of standardized quality phytomedicines, through the mass production of

consistent plant material for physiological characterization, and the analysis of active

ingredients. Protocols for the cloning of some plants such as Catharanthus roseus

(Apocynaceae), Chlorophytum borivilianum (Liliaceae), Datura metel (Solanaceae), and Bacopa

monnieri (Scrophulariaceae) have been developed. The integrated approaches of plant culture

systems will provide the basis for the future development of novel, safe, effective, and high

quality products for consumers. Through the use of biotechnology, desirable genetic traits can be

transferred from one organism to another by transfer of DNA. Many plants with the desirable

DNA can be regenerated from small pieces of the transformed plant tissue. Examples of plants

produced using tissue culture include the large variety of ornamental plants; agricultural crops

such as strawberry, banana, potato, and tomato; and a variety plants.

Micropropagation

Micropropagation is the practice of rapidly multiplying stock plant material to produce

large numbers of progeny plants by using modern plant tissue culture methods.

Micropropagation is the rapid production of high quality, disease free and uniform planting

material. The plants can be multiplied under a controlled environment, anywhere, irrespective of

the season and weather, on a year round basis. Currently, the most popular application of

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micropropagation is the mass clonal multiplication of desirable genotypes of plants. Through

tissue culture, over a million plants can be grown from a small, even microscopic, piece of plant

tissue within 12 months. Such a prolific rate of multiplication cannot be expected by any of the

in vivo methods of clonal propagation. An advantage of tissue culture propagation is that the

shoot bud multiplication cycle is very short (2-6 weeks), each cycle resulting in a logarithmic

increase in the number of shoots. A large number of horticultural, plantation and forest species

are being propagated in vitro on commercial scale.

Micropropagation of various plant species, including many plants, has been described by

many authors during the last two decades (Murashige, 1978; Skirvin et al., 1990; Withers and

Anderson, 1986). From a practical and pharmaceutical point of view, propagation from existing

meristems is not technologically difficult and it yields plants that are genetically identical with

the donor plants (Hu and Wang, 1983). Micropropagation of plants has been achieved through

rapid proliferation of shoot tips and axillary buds in culture. Numerous factors are reported to

influence the success of in vitro propagation of plants and therefore, it is unwise to define any

particular reason for the general micropropagation of plants. The factors that influence

micropropagation of plants have been reviewed by Murashige, 1977; Hussey, 1980, 1983; Hu

and Wang, 1983; Bhagyalakshmi and Singh, 1988; Short and Roberts, 1991.

Micropropagation has become an important part of the commercial propagation of many

plants (Dirr and Heuser, 1987; George and Sherrington, 1984; Zimmerman et al., 1986; Stimart,

23

1986; Fiorino and Loreti, 1987) because of its advantages as a multiplication system (Debergh,

1987; Pierik, 1997; Razdan, 2003). Several techniques for in vitro plant propagation have been

devised, including the induction of axillary and adventitious shoots, the culture of isolated

meristems and plant regeneration by organogenesis and or somatic embryogenesis (Williams and

Maheswaran, 1986; Gautheret, 1983, 1985).

Cytokinin alone in the culture medium induces shoot formation in many plants. MS

medium supplemented with 3.0mg/l BAP was suitable for micropropagation of Ficus benjamina

vars. (Rzepka-Plevnes and Kurek, 2001). Jain (1997) micropropagated Saintpaulia ionantha by

culturing leaf disks on MS medium containing 0.22-0.50μM BAP. In Petunia hybrida, mass

shoot multiplication was achieved on MS medium amended with 2.22μM BAP and 5.7μM IAA

within 4 weeks of culture (Sharma and Mitra, 1976).

Shoot culture has proved to be a widely applicable method of micropropagation, the

appreciation of its potential value developed only slowly, and utilization largely depended on

improvements in tissue culture technology. Robbins (1922) seems to have been the first person

to have successfully cultured excised shoot tips on a medium containing sugar. Shoot tip

explants of between 1.75 and 3.75cm were taken from pea, corn and cotton, and placed in a

liquid medium. White (1933) experimented with small meristem tips (0.1cm) of chickweed

(Stellaria media), but they were only maintained in hanging drops of nutrient solution. Leaf or

flower primordia were observed to develop over a six week period.

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Nasir and Miles (1981) observed that in subcultures of an apple rootstock, some new

shoots arose from callus at the base of the shoot clump; both adventitious and axillary shoots

were produced in Hosta cultures (Papachatzi et al., 1981) and shoot proliferation from some

kinds of potato shoot tips was exclusively from organogenic callus (Roca et al., 1978). Hussey

(1983) has termed cultures providing both adventitious and axillary shoots, ‘mixed cultures’.

Adventitious shoots, particularly those arising indirectly from callus, are not desirable. Hackett

and Anderson (1967) got either single shoots from carnation shoot apices, or else a proliferative

tissue from which shoots were later regenerated. Walkey and Woolfitt (1968) reported a similar

kind of direct or indirect shoot proliferation from Nicotiana rustica shoot tips. Vine and Jones

(1969) were able to transfer large shoot tips of hop (Humulus) to culture, but shoots only rooted,

and showed a high propensity for callus formation. Haramaki (1971) described the rapid

multiplication of Gloxinia by shoot culture and by 1972 several reports of successful

micropropagation by this method had appeared (Adams, 1972; Haramaki and Murashige, 1972).

Koblitz et al. (1983b) subculture micropropagated shoots of Cinchona ledgeriana and

C. succirubra at weekly intervals and obtained 20,000 shoots from a single apical meristem

within one year. Bajaj et al. (1988) observed 2200 plantlets of Thymus vulgaris from a single

shoot grown in vitro for 5 months. Saxena et al. (1998) reported an average of 3-5 fold

multiplication in Psoralea corylifolia when axillary shoots were allowed to continue in primary

cultures for 8 weeks. In contrast, (Rout et al., 1999) demonstrated a significant improvement in

shoot multiplication rate by sub-culturing Plumbago zeylanica at 4 week intervals. Park et al.

(2009) established an efficient and improved protocol for plant regeneration and

micropropagation from leaf cultures of Rehmannia glutinosa. The regenerated shoots obtained

25

from leaf cultures on solid MS medium contained different concentrations of TDZ. The

highest number of shoots per explants (2.1) and shoot growth (1.2cm) was obtained on MS

medium containing 1mg/l TDZ. The addition of auxins in MS medium containing 1mg/l TDZ

substantially improved the shoot regeneration of R. glutinosa and at the optimal concentration

of 0.1mg/l NAA was the most suitable auxin for the highest number of shoots per explants

(3.8cm) and shoot growth (1.5cm).

Explants

To find out the best suitable explants for micropropagation the response of the explants

taken from different positions of the branches of the field mother plant were assessed in the best

medium found effective in producing the maximum number of multiple shoots. From the field

grown mother plant, the shoot tips were considered for micropropagation and it was observed

that the axillary buds. The survival of explants depends on their rate of microbial contamination

and of explant browning, which pertain not only to the explants used for culture initiation but

also to the physiological stage of mother plants and to the season when explants are collected

(Werner and Boe, 1980; Webster and Jones, 1989; Marin et al., 1993; Yepes and Aldwinckle,

1994; Modgil et al., 1999; Dobránszki et al., 2000a).

Pattnaik et al. (1996) reported that shoot buds were the best for micropropagation, as the

pre existing meristem will easily develop into shoots. The explants were collected from the shoot

tip with an axillary bud and the size varied from 5cm to 20cm (Ohyama, 1970; Ivanicka, 1987;

Tewary and Rao, 1990; Raghunath et al., 1992; Yamanouchi et al., 1999). Salehi and Khosh-

26

Khui (1997) reported that the explants length and diameter played significant roles in

proliferation and shoot growth of miniature roses (Rosa chinensis var. Minima Rehd hybrids),

`Little Buckaroo', `Baby Masquerado' and `Sourati'. They also reported that the best rates of

shoot growth and proliferation were obtained in explants with the greatest length (9.0-10.0cm)

and diameter (3.0-3.5cm). Upadhyay et al., (1989) reported a propagation profile for Picrorhiza

kurroa and observed that the shoot multiplication rate gradually improved as the number of

subcultures increased. They proposed that the effect reflected a time dependant adaptation of the

explants to in vitro conditions, which was essentially completed during the first few subcultures.

Wang and Ma (1978) reported that shoot tip between 5-50mm and shoot meristems between

10-20mm diameters produced only a single shoot. Larger explants (5-50mm diameter) formed

multiple shoots. Mandal et al., (2000) used various explants for regeneration of Dendranthema

grandiflora and regenerated new plants from mutated tissues.

Jones et al. (1990) used phyllode explants for in vitro propagation of Acacia bivenosa,

A. holosericea, A. saligna, A. sclerosperma, and A. salicina. A. salicina alone produced

12 shoots per explants. Shoots from 2 week old and 7 month old seedlings were employed by

Galiana et al., (1991) in A. mangium in which juvenile explants resulted in enhanced axillary

buds. Cotyledonary nodes, epicotyls and shoot tip explants of A. nilotica, A. tortilis and A.

sinuata (Dawan et al., 1992; Nangia and Singh 1996) were used to induce multiple shoots. Of

these three explants, cotyledonary node produced maximum number of shoots per explant. Other

explants were not suitable for in vitro propagation. Reddy et al., (2001) obtained rooting of

microshoots from nodal explants on IBA (4.4µM) exhibiting 100% rooting and showed 90%

field survival. Giridhar et al., (2004) induced somatic embryogenesis from leaf cultures of

27

D. hamiltonii and 70% of the rooted plantlets on IBA on transfer to field survived. Kausal et al.

(2005) studied the effect of explants size and position on tissue browning intensity and survival

of explants. They concluded that axillary buds ranging from 0.2 to 0.6cm in size showed less

browning intensity and higher survival (60%) compared to larger (0.6-1 and 1-2cm) explants;

however, only a few buds survived. When actively growing buds were used as explants,

establishment was successful independent of their position on the mother plant (distal or basal)

but when dormant buds were used as explants, distal buds were most suitable as the source of

explants.

Surface sterilization

Surface sterilization is an important step in the preparation of explants to avoid bacterial

and fungal contamination during tissue culture techniques. Therefore optimization of the

conditions to avoid this contamination is a prerequisite for in vitro culturing. Different chemicals

are used for surface sterilization process e.g. Mercuric Chloride, Sodium hypochlorite and

Calcium hypochlorite etc. Moutia and Dookun (1999) used 2.7% solution of sodium

hypochlorite and 0.1% mercuric chloride for elimination of bacterial contamination in sugarcane

crop and recorded 46.8% and 56.3% clean cultures respectively. The surface sterilization process

with antibiotic proved very useful and greatly reduced the bacterial contamination. Saini et al.

(2004) carried out the surface sterilization in sugarcane with 0.1% (w/v) HgCl2 solution along

with a few drops of teepol for 5 minutes.

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Several surface sterilants were used during the micropropagation process with varying

duration of treatments. Surface sterilants like calcium hypochlorite (Ohyama, 1970;

Chattopadhyay et al., 1990), alcohol (Enomoto, 1987; Srinivasa et al., 2000), mercuric chloride

(Zaman et al., 1993; Pavan et al., 1999) and sodium hypochlorite (Tewary et al., 1995;

Yamanouchi et al., 1999) were used either alone or in combination used chlorine alone to treat

the explants. A successful tissue culture protocol starts with effective explants sterilization

(Dodds and Roberts, 1985). Contamination with microorganisms is considered to be the single

most important reason for losses during in vitro culture of plants. Such microorganisms include

viruses, bacteria, yeast, fungi, etc (Omamor et al., 2007). These microbes compete adversely

with plant tissue cultures for nutrients. The presence of these microbes usually result in increased

culture mortality but can also result in variable growth, tissue necrosis, reduced shoot

proliferation and reduced rooting (Kane, 2003).

Explants contamination is a function of several plant and environmental related factors such

as plant species, age, explants source and prevailing weather condition. Despite the best timing

and selection efforts it is almost impossible to eliminate contamination from in vitro grown

plants. In fact according to Boxus and Terzi, 1987, 1988; Leifert et al., 1990 losses due to

contamination in vitro average between 3 and 15% at every subculture in the majority of

commercial and scientific plant tissue culture laboratories, the majority of which is caused by

fungal, yeast and bacterial contaminants (Leifert et al., 1994). Webster and Jones (1989, 1991)

compared the efficacy of sodium hypochlorite solution (15%) or a solution of mercuric chloride

in sodium hypochlorite (150mg HgCl2 in 12% Domestos) and found them equally effective in

surface sterilization of four cold hardy dwarfing apple rootstocks (‘B.9’, ‘Ottawa 3’, ‘P.2’ and

29

‘P.22’). NaOCl (10%) alone for 10 minutes in the case of ‘M9’ (Grant and Hammatt, 1999) or

10% NaOCl together with 2-3 drops Tween-20 for 20 minutes in the case of ‘MM 111’ was also

effective.

Surface sterilization with carbendazim (0.1%) for 8 minutes to normal sterilization

procedure, fungal contamination could be controlled up to 98%. Control of fungal contamination

by using carbendazim has been reported earlier by Patel et al., 1997 in Momordica dioica and

Reddy et al., 1998 in Gymnema sylvestre. 0.1% solution HgCl2 was sufficient in establishing the

explants in culture medium. The efficacy of HgCl2 solution as surface sterilant in in-vitro

cultures of Andrographis paniculata had been reported by Martin, 2004; Purkayastha et al.,

2008; Kataky and Handique, 2010b.

Chengalrayan et al., (2001) used 10% clorox for 20 minutes to get the explants of sugarcane

crop surface sterilized. Niaz and Quraishi (2002) got the surface sterilization of sugarcane crop

at 20% clorox for 20 minute while treatment with other sterilizing agents are reported as Mamun

et al. (2004) carried out surface sterilization of explants from field grown sugarcane plants with

0.1% HgCl2 for 8 minutes. Similarly Baksha et al., (2002) also carried out surface sterilization

of sugarcane with 0.1% HgCl2. Webster et al., (2003) suggested that prudent selection of

explants from the healthy parent plants coupled with an effective surface sterilization method

should be the goal in avoiding culture contamination. Gantait et al., (2008) sterilized the

Anthurium shoot tips using antifungal solution cetrimide for 5 minutes followed by NaOCl and

30

0.1% HgCl2. Later, Jahan et al., (2009) effectively used 70% (v/v) ethanol for 1 minute, 1.5%

NaOCl for 8 minutes as disinfectant and added few drops Tween-20 as surfactant.

Nutrient media

Culture media contain macro elements, microelements, iron (Fe-ETDA), vitamins, other

organic components, plant growth regulators and sucrose, agar. The composition of the culture

medium depends upon the plant species, the explants, and the aim of the experiments. In general,

certain standard media are used for most plants, but some modifications may be required to

achieve genotype specific and stage dependent optimizations, by manipulating the concentrations

of growth regulators, or by the addition of specific components to the culture medium.

Commercially available readymade powdered medium or stock solutions can be used for the

preparation of culture media. A range of culture media of different formulations, and plant

growth regulators are supplied. Murashige and Skoog MS medium are used most extensively

(Murashige and Skoog, 1962).

Although more than 50 different devised media formulations have been used for the

in vitro culture of tissues of various plant species (Gamborg et al., 1976; Huang and Murashige,

1977) the formulation described by Murashige and Skoog (1962) MS medium is most commonly

used, often with relatively minor changes (Uduebo, 1971; Koblitz et al., 1983a, b; Jha and Sen,

1985; Ravishankar and Venkataraman, 1988; Wakhlu et al., 1990; Zhou et al., 1994; Chand

et al., 1997; Saxena et al., 1998; Rout et al., 1999). Basically, a nutrient medium consists of all

the essential major and minor plant nutrient elements, vitamins, plant growth regulators and a

31

carbohydrate as carbon source with other organic substances as optional additives. The

composition of different nutrient media and the nature, source and use of the ingredients have

been discussed in detail by Murashige (1974) and Torres (1989).

The macronutrients N, P, K, Ca, Mg, and S are required in all types of plant cultures, but

the optimal concentration of each may vary with plant species. Although plant cells in culture

may grow on nitrates alone, the pH of the medium (5.8) is usually more stable and better results

are obtained when the medium contains both nitrate and ammonium ions sources of nitrogen.

The essential micronutrients required in trace amounts for whole plants and for tissue in culture

include Fe, Mn, Zn, B, Cu, Cl, Mo, and Ni. Iron is usually added in a chelated form to avoid

precipitation and facilitate absorption. Co, I, and Na are also included in some media although

their requirement and physiological roles have not been established. The carbohydrate

requirement in culture media is usually met by the incorporation of 2-3% sucrose or less

frequently by glucose. Other carbohydrates including lactose, maltose, galactose, and starch have

been used only rarely. In addition to their role as a carbon source, the carbohydrates act as

osmotica and help maintain an osmotic potential in the culture medium that is conducive to cell

and tissue growth. Whole plants synthesize all of the vitamins; such as, biocatalysts, required for

normal growth and development, but specific vitamins, including thiamine (B1), nicotinic acid

(B3), pyridoxine (B6), and myo-inositol are usually required additions to tissue and cell cultures.

Although, cultured cells are normally capable of synthesizing all of their required amino acids,

addition of L-glutamine, L-cysteine, L-proline, and L-tyrosine or an amino acid mixture like

caesin hydrolysate to the medium may increase cell growth. Agar is most commonly used for

preparing semisolid or solid culture media, but other gelling agents occasionally used including

32

gelatin, agarose, alginate, and gelrite. The two major developments which made shoot culture

feasible were the development of improved media for plant tissue culture (Murashige and Skoog,

1962) and the discovery of the cytokinins as a class of plant growth regulators (Miller, 1961b;

Skoog et al., 1965), with an ability to release lateral buds from dormancy (Wickson and

Thimann, 1958; Sachs and Thimann, 1964).

A rapid rate of propagation depends on the sub-culturing of proliferating shoot cultures.

In the case of prolonged cultures, the nutrients in the medium are gradually exhausted and at the

same time, the relative humidity in the vessels decreases leading to the drying of the culture

medium. Sub-culturing also decreases the effects of competition of the developing shoots for

nutrients. An important biological technique of cloning large number of single cells of higher

plants was, however, developed in 1960 by Bergmann. He filtered the suspension cultures of

Nicotiana tabacum and Phaseolus vulgaris and obtained a population containing about 90% free

cells. These were incorporated into a 1mm layer of solidified medium containing 0.8% agar. In

this experiment some of the single cells divided and formed visible colonies. Percentage of bud

breaks and shoot multiplication was generally higher in MS medium compared to several other

media.

The micropropagation of rose was also found to be influenced by agar concentration

(Ghashghaie et al., 1991), ethylene concentration (Kevers et al., 1992), growth room and vessel

humidity (Sallanon and Maziere, 1992) and different types of gelling agents (Podwyszynska and

Olszewski, 1995). Among the different explants used, the second node was found to be the best

33

for shoot multiplication on MS medium supplemented with 1.0mg/l BAP (Ara et al., 1997).

Davies (1980) reported that the standard MS media induced the best rate of shoot proliferation in

different rose cultivars. Inclusion of BAP (1.0-10.0mg/l) in the culture medium was essential for

bud break and shoots multiplication of Rosa hybrida cultivars (Hasegawa, 1980; Wulster and

Sacalis, 1980). Bressan et al., (1982) reported the maximum promotive effects with BAP

(1.0mg/l) compared to N6; IAA neither enhanced nor repressed shoot multiplication regardless

of the BAP concentration. The presence of cytokines in the culture medium helped in the year

round multiplication of shoots in hybrid roses (Rout et al., 1990).

The nutrient medium is important for successful tissue culture; information is still scanty

on the various media used for Gerbera explants. MS medium was successfully used by many

workers for callus formation as well as shoot regeneration (Pierik et al., 1982; Le et al., 1999;

Modh et al., 2002; Aswath and Wazneen 2004; Kumar and Kanwar 2005, 2006). Parthasarathy

et al. (1996), Verma and Anand (2006) used and N6 (Chu, 1978) media for culture establishment

in Gerbera.

Agar as a gelling agent

Agar is one of the consumables used in large quantities in plant tissue cultures especially

where solid media are required. It is widely used as a gelling agent in most nutrient media

(Pierik, 1987). Since agar binds water and adsorbs compounds from the media, it is thought that

it may limit uptake of growth regulators. It has remained the most frequently used gelling agent

in culture media employed for microbes as well as plants. The properties of agar that make it the

34

gelling agent of choice are stability, high clarity and resistance to metabolism during culture

(Jain and Babbar, 2002).

The seaweeds of the genus Gracilaria and Gelidiella harvested from wild populations

provide the bulk of the world's agar (Santelices and Doty, 1989; Nene and Sheila, 1994). Agar

extracted from species of Gracilaria have shown that the agar content and gel strength

may vary with the stage of the life cycle. The gelling properties depend on the content of

3,6-anhydro-L-galactose and the number and position of the sulfate groups in the

polysaccharide chain (Duckworth and Yaphe, 1971; Lahaye and Rochas, 1991).

The choices of gelling agent can greatly influence the growth of tissue cultured plants.

Several papers have reported that the choice of gelling agent is an important consideration for

micropropagation process (Debergh, 1983; Ebrahim and Ibrahim, 2000). Agar has been widely

used because of its convenient gelling properties and stability during tissue culture. Agar is

insoluble in cold water but soluble in boiling water. A hot aqueous 1.5% agar solution is clear

but when cooled below 40ºC, a firm resilient gel is formed. Originally, gelling agents were

produced by allowing them to freeze and thaw outdoors during the winter months (Foreman and

Whyte, 1997). White (1934) used agar as a gelling agent in the medium. Since, its introduction,

agar could not be satisfactorily replaced and remains the gelling agent of choice for culture

media employed for higher plants as well as microorganisms. However, agar contains some

inhibitors, which result in pre mature abortion of embryoids in anther culture of tobacco (Jain

et al., 1997).

35

Anagnostakis (1974) reported that agar might contain some compounds that influence

differentiation and development of cultured tissue. This was confirmed by Ichimura and Oda

(1995) who found active thermo stable substances in water extracts of agar that promoted shoot

regeneration in tomato. In the well defined media that are used for in vitro culture of plants, agar

is a major source of unknown variation (Scholten and Pierik, 1998). In several studies,

differences have been exported in plant response to agar brands or types attributed to limited

diffusion of medium components and water (Romberger and Tabor, 1971; Stolz, 1971), to

impurities (Kohlenbach and Wernicke, 1978; Nairn et al., 1995) and to differences in gel

strength (Debergh, 1983). Although agar is the most widely used solidifying agent for plant

tissue culture media, it is relatively expensive and it has been reported that sometimes can have

cytotoxic effects (Kohlenbach and Wernicke, 1978).

Agar, a commonly used gelling agent for plant micropropagation, is the most

expensive ingredient of the medium. The brand and concentration of agars were shown to

affect the length of Rosa hybrida and Syringa vulgaris shoots, fresh weight of Gerbera

jamesonii culture (Pierik, 1991) and the multiplication of Cydonia oblonga (Gulsen and

Domanoglu, 1991). The influence of agar brand on cultures is probably caused by

different impurities (Pierik, 1991), rigidity of gel (Stolz, 1971), varying availability of

water, ions and cytokinins (Bornman and Vogelman,1984; Orlikowska and Olszewski, 1993).

36

Plant Growth Regulators

Plant hormones play a crucial role in controlling the way in which plants grow and

develop. They regulate the speed of growth of the individual parts and integrate these parts to

produce the plants. Both auxins and cytokinins are synergistically required to induce cell division

and growth in plant tissue cultures. Shoot branching depends on the initiation and activity of

axillary meristems, which are hormonally controlled mainly by cytokinins; however, they act in

interaction with auxins even though the auxin effect is indirect (Ward and Leyser, 2004). In

angiosperms the meristematic dome in the shoot tip is not autonomous for auxin and it is not the

source of auxin in the plant (Smith and Murashige, 1970; Shabde and Murashige, 1977). Auxin

is probably synthesized by the second pair of youngest leaf primordia. Accordingly, for the

successful culture of meristem explants (without any leaf primordia) of Coleus blumei, Daucus

carota, Nicotiana tabacum, N. glauca, Tropaeolum majus (Smith and Murashige, 1970) the

presence of exogenous auxin was essential.

The plant growth regulators, auxins and cytokinins, are of particular significance in

in-vitro culture. Cytokinins are concerned with the modification of apical dominance (Razdan,

1993). A high cytokinin concentration promotes axillary shoot formation by decreasing apical

dominance (Dodds and Roberts, 1985; Chawla, 2002). Although cytokinin induces the growth of

buds, auxin is required in the culture media. Most cultures require a combination of these two

compounds for growth regulation (Dodds and Roberts, 1985). Usually the manipulation and

variation of auxin and cytokinin levels can successfully alter growth behaviour in plant cultures

(Dixon and Gonzales, 1994). For axillary shoot formation, a low auxin concentration, together

with a high cytokinin concentration is required (Razdan, 1993).

37

Sripichit et al., (1987) also favoured the role of BAP over Kn in inducing shoot formation

in Capscum. Agrawal et al., (1989) found the combinations of IAA and BAP to be the best for

shoot bud induction in comparison with other auxins (IBA, NAA, 2,4-D) and Kn combinations

tested. Combinations of IAA have been shown to induce bud and shoot formation at different

extents in chilli pepper explants from different cultivars (Alibert, 1990; Ochoa-Alejo and García-

Bautista, 1990; Berljak, 1999; Steinitz et al., 1999).

CULTURE CONDITIONSTemperature

Temperature has a great effect on shoot multiplication. There is a little difference in the

number of shoots per explants at constant temperatures of 11, 16 and 21°C, although

multiplication rate is slightly better at 21°C. At temperatures above 21°C, the rate of shoot

multiplication is reduced. For somatic embryogenesis, embryo culture and optimal shoot or root

formation and proliferation, the range 22-28°C is the best temperature, as summarized for

instance, Kaur et al., (2006) used 22°C throughout optimization of medium for embryo culture.

However, San and Hatic (2006) used 25°C for somatic embryogenesis and (Fernandez et al.,

2000) used 26°C for modulation and morphogenic study on embryonic axes via embryo culture

technique. Long et al. (1995) used 25°C for somatic embryogenesis and adventitious shoots in

Juglans nigra L.

Light

Light is a major factor of the culture environment and has been shown to have an effect

on organized development in vitro. Light requirements for differentiation involve a combination

of several components, including intensity, daily light period and quality. The properties of light

38

affect tissue cultures, and influences their growth and development. Reduction of light at the

base of a shoot can provide an environment conducive to the accumulation of photosensitive

auxin or co-factors.

Light intensity plays an important role in satisfactory shoot growth. One to 3000lux

(40-92µmol m-2s-1) is reported to be sufficient for embryo culture, axillary bud culture and

shoot proliferation in walnut (Capellades et al. 1990; Jay-Allemand et al., 1992; Long et al.,

1995; Kaur et al., 2006; Roschke and Pijut, 2006; Paygamzedeh, 2008) however, opined that

cultured plantlets could resemble greenhouse grown plants if these were cultured at a higher light

intensity (80µmol m-2s-1) than that normally available inside the culture vessels (25µmol m-2s-1).

Usually a 16-18hrs photoperiod from Sylvania Grolux type white fluorescent lamps or cool

white fluorescent lamps is provided in culture conditions (Long et al., 1995; Heloir et al., 1996;

Scaltsoyiannes et al., 1997; Sanchez-Zamora et al., 2006; Roschke and Paula, 2006). In general,

in walnut tissue culture, 16-18hrs illumination resulted in a normal growth, multiplication and

germination rate. Light is essential for morphogenetic processes like shoot and root initiation and

somatic embryogenesis. Both quality and intensity of light as well as the photoperiod are very

critical to the success of certain culture experiments (Murashige, 1977). Exposure to light for 12-

16hrs per day under 35-112µmol m-2s-1 provided by cool, white fluorescent lamp is usually

recommended. Blue light promotes shoot formation while, red light induces rooting in many

species (Murashige, 1977). The temperature usually employed in the culture incubation room is

approximately 25°C. Tropical species usually require higher temperature (27-30°C; Tisserart,

1981).

39

The quality of light also influences shoot induction. Gabryszewska and Rudnicki (1997)

developed a micropropagation protocol for Ficus benjamina by using shoot meristems; shoot

numbers increased on MS medium supplemented with 15mg/l 2ip by red light treatment; and

root initiation occurred in all light treatments (white, blue, green and red). However, the rooting

and number of roots per shoot were highest in red light on the medium having 0.5mg/l IAA.

Palai et al., (1998) developed an efficient protocol for mass cloning of Gerbera cv. Fredaisy in

vitro by manipulating growth regulators and culture conditions. Leaves were used as the explant

source for callus culture. The cultures inoculated in the light exhibited a higher rate of shoot bud

differentiation than those inoculated in the dark. In vitro propagation of gerbera was studied by

culturing 3 leaf bud explants on a medium containing 0-4mg/dm3 BAP and 0-1mg/dm3 IAA.

Highest multiplication rates (9.5-11.2) were obtained on 1mg/dm3 BAP, irrespective of the IAA

concentration used (Barbosa et al., 1994). A high frequency of shoot organogenesis and plant

establishment protocol was developed for ex vitro leaf derived callus with 0.4mg/dm3 BAP and

4mg/dm3 NAA (Aswath and Choudhary, 2002a).

Zanandrea et al., (2006) evaluated chlorophyll fluorescence in ‘M.9’. During the

multiplication phase a 16hrs photoperiod was applied and in vitro shoots were irradiated with

15µmol m-2s-1 for 55 or 90 days and additional higher light intensity (15µmol m-2s-1) was added

to plants for 55 days at 2hrs per day before determining fluorescence. They concluded that

in vitro shoots had a functional photosynthetic apparatus but the efficiency of the radiant energy

40

use was low presumably due to low irradiance since the supply of additional light increased the

photosynthetic activity.

Bejoy et al., (2008) reported the effectiveness of continuous dark period over alternative

light and dark period in callus induction of Anthurium. Callus induction frequency was quite

high in dark regime, while the explants kept in light didn't develop callus and turned brown

rapidly. In accordance with the above finding, Atak and Celik (2009) also preferred the role of

continuous dark phase of one month for callus initiation. A favourable effect of controlled

culture condition during Anthurium organogenesis in vitro (pH 5.8), with a temperature range of

25±2°C, 16hrs photoperiod, 60% RH and 1500-3000lux light intensity was studied (Mahanta and

Paswan, 2001; Gantait et al,, 2008; Jahan et al., 2009). For PLB induction and regeneration, a

specific condition at 25±2°C with 12hrs photoperiod at 25-30µmol m-2s-1 proved to be

favourable according to the report of Yu et al., (2009).

In vitro cultures of apple during shoot multiplication were provided with a 16hrs

photoperiod from cool or warm white fluorescent lamps (400-700nm). Several authors applied

continuous illumination (Noiton et al., 1992; Schaefer et al., 2002).The applied light intensity

varied between 38 and 105µmol m−2s−1. Marin et al., (1993) compared the effect of 8-16hrs

photoperiods on the shoot multiplication of columnar apple rootstocks (‘Tuscan’, ‘Trajan’,

‘Telemon’, and ‘Greensleeves’) and concluded that shoot production was significantly higher

under an 8hrs photoperiod. Morel and Martin (1952) for the first time recovered virus free

Dahlia plants from infected individuals by excising and culturing their shoot tips in vitro. Ball

41

(1946) was the first person to produce rooted shoots from cultured shoot apices. His explants

consisted of an apical meristem and 2-3 leaf primordia. There was no shoot multiplication but

plantlets of nasturtium (Tropaeolum majus) and white lupine (Lupinus alba) were transferred to

soil and grown successfully.

Somani et al., (1989) obtained shoots from corm explants of Gloriosa superba on

MS+3mg/l Kn. The multiple shoots formed microcormlets at the base of each shoot. Sivakumar

and Krishnamurthy (2000) reported as many as 35 shoots on average from shoot tip explants on

basal medium consisting of MS salts and 2iP+2.32µM Kn. Hassan and Roy (2005) developed a

protocol for propagation of G. superba using terminal shoot tips and stem nodes. Shoots rooted

on half strength the MS+1.0mg/l IBA or 0.5mg/l IAA. Ninety per cent plants survived in the

field.

Roy et al., (1994) established a protocol for propagation of Rauvolfia serpentina using

shoot tips and lateral buds from field grown plants. Rooting was achieved on

½ strength MS+1.0mg/l IBA+1.0mg/l IAA medium. Ninety five per cent of the plantlets

survived on transfer to field. Ahmad et al., (2002) established plantlet regeneration system from

shoot and nodal explants of field grown plants and from calli of leaf and internode explants of

in vitro grown shoots. Shekhawat and Kataria (2005) obtained 3-5 shoots per node by axillary

bud proliferation on MS medium+10μM BAP+0.5μM IAA. A promising in vitro propagation of

R. serpentina was developed using shoot tips on MS medium supplemented with 4.0mg/l

BAP+0.5mg/l NAA which gave the highest percentage of response with 7 or 8 multiple shoots

42

per culture (Baksha et al., 2007). Sharma and Chandel (1992) reported storage of nodal cultures

of R. serpentina at reduced temperature. After 15 months of storage, the cultures maintained at

15°C were viable, showing 70% survivability in field.

Atta-Alla and VanStaden (1997) succeeded to propagate Yucca aloifolia by using shoot

tip explants and the maximum number shoot production (6.6) was obtained from a single shoot

tip on MS medium supplemented with 4.5μM TDZ and 1.1μM NAA. The proliferated shoots

readily rooted on half strength MS medium containing 2.5-4.9μM IBA and 1% charcoal. The

rooted plants were successfully established in soil.

Benjamin et al., (1987) reported that BAP at high concentration (1-5ppm) stimulated the

development of the axillary meristems and shoot tips of Atropa belladona. Lal et al., (1988)

reported a rapid proliferation rate in Picrorhiza kurroa using Kn at 1.0-5.0mg/l. Further, Barna

and Wakhlu (1988) indicated that the production of multiple shoots was higher in Plantago ovate

on a medium having 4-6mM Kn along with 0.05mM NAA. Similarly, it was observed that

cytokinin was required in optimal quantity for shoot proliferation in many genotypes but

inclusion of low concentration of auxins along with cytokinin triggered the rate of shoot

proliferation (Jha and Jha, 1989; Tsay et al., 1989; Roja et al., 1990; Rout and Das, 1997a, b;

Sharma and Singh, 1997; Shasany et al., 1998).

43

Martin et al., (2003) succeeded in direct shoot bud regeneration from lamina explants of

Anthuriumandr aeanum on MS medium fortified with 1.11μM BA, 1.14μM IAA and 0.46μM

Kn. Furthermore, the regenerated shoots were rooted on ½ strength MS medium supplemented

with 0.54μM NAA and 0.93μM Kn. Nearly 300 plantlets of each cultivar were transferred to soil

with 95% survival rate (Joseph et al., 2003). Beena et al., (2003) established a protocol for

in vitro propagation of Ceropegia candelabrum through axillary bud multiplication by using

BAP (8.88µM) in combination with 2.46µM IBA. Shoots were rooted on half strength MS

medium supplemented with IBA with a maximum of seven roots per shoot on 0.49µM IBA.

Direct shoot induction from leaf explants of Exacum sp. had been reported by using MS

medium supplemented with BAP and NAA and the shoots were subculture on modified MS

medium supplemented with PVP, 2iP, NAA, GA3 and 0.06g/l AC to simultaneously promote

both elongation and rooting (Unda et al., 2007). Baksha et al., (2006) reported that high

frequency of Curcuma longa multiple shoot regeneration was achieved from shoot apices

explants on MS medium supplemented with 2mg/l BAP. Elongated shoot were rooted well in

half strength MS medium with 0.5mg/l auxin.

In vitro rooting

Rooting of shoots derived from in vitro cultures is a prerequisite to make possible their

physiological establishment in soil. NAA or IBA has been the most commonly used auxin for

promoting rooting of in vitro regenerated shoots of pomegranate. Robbins and Maneval (1924)

enabled them to improve root growth, but the first successful report of continuously growing

44

cultures of tomato root tips was made by White in 1934. Initially, White used a medium

containing inorganic salts, yeast extract and sucrose, but later yeast extract was replaced by three

B-vitamins, namely pyridoxine, thiamine and nicotinic acid (White, 1937). On this synthetic

medium, this has proved to be one of the basic media for a variety of cell and tissue cultures. Lal

et al., (1988) examined the effectiveness of various auxins on rooting of Picrorhiza microshoots

and found that NAA at 1.0mg/l was superior to IBA or IAA. Rooting in 9% of the cultured

shoots of Saussurea lappa was achieved within 25-30 days on media containing 0.5-1.0mM

NAA (Arora and Bhojwani, 1989). Many workers reported that roots were easily induced from

excised mature microshoots on MS medium supplemented with low concentrations of auxins

(IAA or IBA or NAA) in the range of 0.1-0.5mg/l with the reduction in the level (2-2.5%) of

sucrose (Hasegawa, 1979). Davies (1980) achieved 100% rooting in several cultivars of rose by

placing them on MS medium supplemented with 40g/l sucrose devoid of growth regulators. Faria

and Illg (1995) obtained 100% rooting in the excised shoots of Zingiber spectabile in liquid or

gelrite gelled medium containing 5mM IAA or NAA. Rout et al., (1999) reported induction of

rooting in microshoots of Plumbago zeylanica on ½ strength MS medium supplemented with

0.25mg/l IBA with 2% sucrose. Leslie and McGranahan (1992) reported that the highest rooting

frequency (75%) occurred on microshoots placed on ½ strength MS containing 2.5µM IBA for

7 days in darkness. Adventitious roots began to emerge within 7 days and elongated when

microshoots were transferred to the light. Although roots were also initiated on microshoots

cultured on media containing 4.9 or 24.6µM IBA, only the 2.5µM IBA rooted plantlets were

successfully acclimatized ex vitro. In vitro and ex vitro rooting of micropropagated Juglans

species has been successful using IBA with or without NAA.

45

Roest and Bokelmann (1975) successfully induced roots in the adventitious shoots of

Chrysanthemum in the liquid MS medium containing 1.0mg/l IAA. In general, shoots and roots

developed on a single medium containing 4.4μM BAP and 5.7μM IAA. Rooting was achieved in

90% cultures of ‘Deep Pink’ rooted with about 2.0Klux (Kilo-lux) of light, whereas higher light

intensities (3.0Klux) gave a lower rooting percentage (Roberts et al., 1992; Rout et al., 1996).

The rooted plants were successfully established in the soil (Roberts and Smith, 1990; Rout et al.,

1996). Sritongkum (1995) found optimal condition for root induction in micro cuttings of

S. rebaudiana (derived from culture of shoot tip) was MS medium supplemented with 0.01mg/l

NAA and MS medium without plant growth regulator.

In vitro culture is one of the key tools of plant biotechnology that exploits the totipotency

nature of plant cell or tissue and organs. Plant tissue culture is the science of growing plant cells,

tissues or organs isolated from the mother plant, on defined media under aseptic and controlled

environment. In fact, tissue culture techniques have played a major role in the development of

plant genetic engineering as well as for studying the regulation of growth and organized

development through examination of structural, physiological, biochemical and molecular bases

underlying developmental processes. Current developments in tissue culture technology indicate

that transcription factors are efficient new molecular tools for plant metabolic engineering to

increase the production of valuable compounds (Gantet and Memelink, 2002). Tissue culture has

become a popular method for vegetative propagation of plants. The most significant advantage

offered by this aseptic method of clonal propagation, popularly called 'micropropagation', over

the conventional methods is that in a relatively short time and space a large number of plants can

be produced starting from a single individual.

46

The lateral sprouts of 8-10cm length are shifted to pro-trays containing equal parts of

coco peat and vermiculite and after sufficient watering left in a shade net (70% shade) at 80-90%

humidity. High humidity is achieved by intermittent misting. Sprouts are usually maintained in

the pro-trays for a period of 15-20 days and then shifted to polythene bags of size 6’x 4’ and

thickness of 120 gauges for secondary hardening. At this stage, the plants are maintained at 50%

shade and 40-50% humidity. Watering is done on alternate days and the plants are ready for field

planting in 30-45 days.

Once the plantlets are ready for shifting outside the laboratory, they are carefully

acclimatized to adapt to the green house and later to least protected field conditions. During

hardening, the plantlets undergo physiological adaptation to changing external factors like water,

temperature, relative humidity and nutrient supply. The plantlets from culture vessels/bottles are

moved from the laboratory to a room at ambient temperature and kept open for 4-6 days. Later

they are shifted to green house for primary hardening where they are first gently washed free of

agar medium. This is important as sucrose in agar encourages microorganisms 8cm shoots with

3-4 ramified roots are planted in individual micro pots in a protray. In places where weather is

conducive (24-26°C temperature and more than 80% humidity), the plantlets are hardened for

4-6 weeks in mini-sand beds. During this period, 90-95% humidity is maintained for the initial

6-8 days under diffused light. The humidity is slowly reduced to 70%, light intensity raised to

normal and temperatures brought to 26°C by the end of 6 weeks.

47

Structures used for primary hardening vary with the climatic conditions. These can be

highly sophisticated with UV-stabilized poly-sheet covering, multiple misting options, thermal

shade net and auto-monitoring of light intensity, temperature and humidity. On the other hand,

the structures can be simple with polycarbonate roofing, shade net on all sides with fogger

facilities. Temperature, RH and light intensities are monitored manually using thermometer,

hygrometer and lux meter, respectively. Planting media for primary hardening range from sieved

sand augmented with nutrition’s to mixtures of coco peat and soil rite with fine sand in equal

proportions NPK is provided in liquid form on weekly basis.

Acclimatize them; this step refers to the final phase of most micropropagation

procedures, where the plants need to adapt to new environmental conditions of relative humidity,

lighting and temperature, among other variables. The intrinsic quality of in vitro seedlings is

considered to be one of the most important factors that determine the success during the

transition to ex vitro conditions. The main factors to overcome are those associated to the

excessive loss of water by transpiration, and to their poorly developed photosynthetic system.

Generally, most of the in vitro plant losses occur when they are moved to new environmental

conditions (Roca, 1991). Consequently there in vitro propagation success lies in being able to

acclimatize the plants to new environmental conditions. During this phase, there occurs a gradual

return to the autotrophic functioning of the plantlets, as well as the recovery of the species

typical morphological and physiological characteristics. Likewise, in this phase the seedlings

often suffer from a type of stress that is provoked by changes in humidity, temperature or

lighting conditions which permit the plants to adapt to of their natural habitats conditions.

48

Davies (1980) reported that a mixture of coarse perlite, peat and loam (2:2:1; v/v/v) was

good for rooting of rose shoots. He achieved 15-85% rooting depending upon the cultivar. Scotti

Campos and Pais (1990) reported that about 83-100% of plantlets of dwarf rose cultivar

Rosamini established well in soil within 45 days of transfer to soil. The survival rate was higher

than that reported in many other rose species (Khosh-Khui and Sink, 1982a, b; Dubois et al.,

1988).Satheesh and Bhavanandan (1988) reported that when micropropagated plants of

Plumbago rosea were transferred to pots containing a1:1 soil and sand mixture under greenhouse

conditions, about 60% of the plants survived. A high survival (96%) was recorded when plantlets

of Pinellia ternata were transplanted into a 1:2:1 mixture of vermiculite: loam soil: peat moss

(Tsay et al., 1989). Jha and Sen (1985) reported that prior to transfer to soil, all of the rooted

plantlets of Bowiea volubilis were maintained for 4-6 weeks in MS salts with 0.5% sucrose and

incubated at 240-300ºC for 4 weeks for hardening. After 4 weeks, the plantlets were transferred

to soil and showed 80% survival.

Mao et al. (1995) reported that rooted plantlets of Clerodendrum colebrookianum

required a minimum of 2 weeks in a mist tunnel, after which they could be transferred

successfully into greenhouse conditions. Approximately 60% of the rooted plants of Centella

asiatica survived in pots containing a 1:1:1 mixture of soil, sand, and well rotted cow dung

(Patra et al., 1998). The plants were supplied with MS inorganic salts twice a week before

transfer to the greenhouse (Palai et al., 1997). Saxena et al., (1997) reported that rooted plantlets

of Psoralea corylifolia were successfully transferred to a 1:1 mixture of soil and sand. About

95% of the regenerated plants survived in the greenhouse. Rout et al., (1999) reported that about

49

95% of micropropagated plantlets of Plumbago zeylanica were established in the greenhouse

within 2-3 weeks of transfer fewer than 85% relative humidity.

The number of shoots increased with subsequent subcultures on the fresh culture

medium. Micro shoots were rooted in moist Sphagnum moss and vermiculite (3:1 ratio), and

90% microshoots survived and grown in the greenhouse. Several researchers have reported on

clonal propagation of Spathiphyllum (Fonnesbech and Fonnesbech, 1979; Orlikowska et al.,

1995; Wated et al., 1997). In photoautotrophically cultured Nicotiana tabacum plantlets, growth

of plants was significantly impaired during acclimatization to ex vitro conditions as compared to

plants cultured in vitro on medium with saccharose, however, photosynthetic parameters were

not affected (Kadlecek et al., 1997). Mahanta and Paswan (2001) successfully transferred in vitro

Anthurium plantlets in the plastic pots containing the growing medium of soilrite perlite at the

ratio of 10:1 and reported 60% survival rate after four weeks of transfer. Effect of vermicompost

and sand mixture (1:3 v/v) in ex vitro establishment under green house followed by net house

condition was reported by Martin et al., (2003), where 95% survival rate was achieved.

Maximum survival rate of 98% was reported by Han and Goo (2003) in cultivar Atlanta using a

combination of vermiculite and per liter (1:1 v/v) as growth substrate. Vargas et al., (2004) and

Anbazhagan et al., (2010) studied the acclimatization of cultivar Rubrun using soil and organic

humus (1:1 v/v). In their study micropropagated plantlets ensured 80% success rate within four

weeks.

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SDS-PAGE

Sodium Dodecyl Sulphate Polyacrylamide Gel Electrophoresis (SDS-PAGE)

is widely used due to its validity and simplicity for describing genetic structure of crop

germplasm. SDS-PAGE is a method used for separating proteins according to their size. Sodium

dodecyl sulfate (SDS) is an anionic detergent that binds to most proteins in constant weight ratio.

Proteins heated in presence of SDS denature into their primary polypeptides and gain an overall

identical negative charge density. These polypeptides, migrating in an electric field towards the

positive anode, can be then separated in a porous gel according to their size with smaller proteins

migrating faster than the larger ones (Hames, 1998).

Electrophoretic techniques are used on large scales in protein and enzyme analysis to

identify and characterize the genotype differences in Araceae plant species and varieties. Matta

et al,. (1981) studied Vicia legumin structure by polyacrylamide gel electrophoresis. Many

authors used the electrophoretic tool to characterize the differences and similarities between

plants. Among these authors are Abdel-Tawab et al., (1982), Giannasi and Crawford (1986),

Gamal El Din et al. (1988), Eweda (1989), Vries (1996); Kamel and Hassan (2001). Recently the

effectiveness of a phenol based extraction method has been shown to give a good protein yield

with different recalcitrant plant tissues, such as olive leaves, tomato tissues, avocado and banana

mesocarp as well as orange albedo and flavedo (Hurkman and Tanaka, 1986; Wang et al., 2003;

Rose et al., 2004; Saravanan and Rose, 2004).

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Protein and secondary metabolites of leaves from variety of plants have been examined

and were found to be promising as markers, such as dehydrin (Lopez et al., 2001; Jiang and

Huang, 2002; Lopez et al., 2003;), superoxide dismutase (Zang and Komatsu, 2007), ASR

(ABA-water stress ripening-induced) protein (Riccardi et al., 1998), actin depolymerizing factor

(Salekdeh et al., 2002). The phenol based method was used to perform a proteome analysis of

grape berry clusters (Vincent et al., 2006), grape berry skin during ripening (Deytieux et al.,

2007) and grape berry cell wall (Negri et al., 2008). For proteome analysis of grapevine leaves,

Castro et al., (2005) used a lysis buffer containing urea, thiourea,3-[(3-Cholamidopropyl)

dimethylammonio] 1-propanesulfonate (CHAPS) and Sodium dodecyl sulfate (SDS) followed by

TCA/acetone precipitation and Vincent et al., (2007) adapted TCA/acetone based method,

according to Damerval et al., (1986), to study proteins from grapevine shoot tips (including the

apex, the stem, four leaves, and tendrils).

Random amplified polymorphic DNA (RAPD)

DNA molecular marker in essence detects nucleotides sequence variation at a particular

location in the genome. The genetic variation/diversity must be found between the parents of the

chosen cross for the marker to be informative among their offspring and to allow its pattern of

inheritance to be analyzed. DNA markers can generate fingerprints, which are distinctive

patterns of DNA fragments resolved by agrose electrophoresis and detected by staining or

labeling.

52

RAPD is one of the most frequently applied molecular techniques (Welsh and

McClelland, 1990; Williams et al., 1990). The technique generates polymorphic band patterns,

produced by polymerase chain reaction (PCR) using arbitrary DNA sequence primers (Xena de

Enrech, 2000). RAPD markers are polymorphic DNA separated by gel electrophoresis after PCR

using short random oligonucleotide primers. It has been particularly used for genetic and

molecular studies, as it is a simple and rapid method for determining genetic diversity and

similarity in various organisms. RAPD also has the advantage that no prior knowledge of the

genome under research is necessary (Fischer et al., 2000; Klinbunga et al., 2000). It has been

concluded that although AFLP analysis is superior in terms of efficiency, RAPDs may still be

used as reliable markers in small low tech laboratories (Kjolner et al., 2004).

The RAPD-PCR method can be applied to detect genetic diversity and similarity in

numerous organisms using the various primers (Bernardi and Talley, 2000). For all of these

reasons the RAPD assay has been used to construct phylogenetic trees for resolving taxonomic

problems in many organisms. It has been concluded that although AFLP analysis is superior in

terms of efficiency, RAPDs may still be used as reliable markers in small low tech laboratories

(Kjolner et al., 2004). On the other hand, Sun et al., (2000) reported that the RAPD and AFLP

techniques are powerful DNA fingerprinting methods for classification of Artemia species and

strains.

The main advantages of RAPD include simplicity, rapidity and small quantities of DNA

required for this technique (Lin et al., 1994). RAPD markers are characterized by a slightly

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lower repeatability than other marker types, but when the relevant procedures are observed, the

repeatability of RAPD analyses may be maintained at a high level (Penner et al., 1993). The use

of RAPD as molecular markers for taxonomic and systematic analyses of plants (Bartish et al.,

2000), as well as in plant breeding and the study of genetic relationships, has considerably

increased (Ranade et al., 2001). Recently, RAPD has been used for the estimation of genetic

diversity in various endangered plant species (Wang et al., 2005; Lu et al., 2006; Liu et al.,

2007; Zheng et al., 2008).

In the case of investigation of polymorphism in closely related strains, the highest

possible complexity of the patterns obtained by RAPD-PCR is required to assure revealing of

limited polymorphisms. Most parameters (reaction components concentration, additives,

different polymerases, and thermal profiles) affecting RAPD-PCR should be examined, in an

effort to increase pattern complexity (Diakou and Dovas, 2001). Fraga et al., (2002) analyzed the

effect of changing concentrations of the primer, template DNA and Taq DNA polymerase with

the goal of determining their optimum concentration for the standardization of the RAPD

technique for genetic studies of Trichomonas vaginalis.

Weeden et al., (1992) investigated the reproducibility and reliability of RAPD markers using

two genetically defined systems. They found that template DNA of high purity was crucial for

reproducible results and the concentration of template DNA could be varied from 3 to 30 ng without

affecting the RAPD pattern. Segregation analysis indicated that RAPD markers scored in

segregating pea and apple (Mains sp.) populations reflected true genetic variation. However, they

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also found that mistakes in scoring were evident, apparently attributable to contamination of

template DNA or faintness of the amplified product. They concluded that the RAPD technique, or

modifications of it, should be very useful for genetic mapping, gene tagging and possibly

cultivar identification.

RAPD mediated DNA fingerprinting has been extensively used for detecting polymorphism

among in vitro micropropagated medicinal plants such as Populus deltoides (Rani et al., 1995),

Picea glauca (Isabel et al., 1996), Panax notoginseng (Shoyama et al., 1997), ZIngiber

officinales (Rout et al., 1998), Piper longum (Philip et al., 1999), Anigozanthos viridis (Turner

et al., 2001), Tylophora indica (Jayanthi and Mandal, 2001), Plumbago zeylanica (Rout and Das,

2002), Curcuma longa (Salvi et al., 2002), Syzgium travancorium (Anand, 2003) and

Chlorophytum arundinaceum (Lattoo et al., 2006) was developed in a relatively short time.

RAPD has been proven to be a suitable molecular technique to detect the variation that is

induced or occurs during in vitro regeneration of plant species (Shu et al., 2003). RAPD is

becoming a widely employed method in the detection of genetic diversity because it has the

advantage of being technically simple, quick to perform and requires only small amounts of

DNA (Ceasar et al., 2010). Many investigators have reported genetic stability of several

micropropagated plants, viz., Picea mariana (Isabel et al., 1993), Pinus thunbergii (Goto et al.,

1998), chestnut rootstock hybrid (Carvalho et al., 2004), Prunus dulcis (Martins et al., 2004),

turmeric etc., using RAPD.