Synseed technology—A complete synthesis

Biotechnology Advances 31 (2013) 186–207

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Research review paper

Synseed technology—A complete synthesis Shiwali Sharma a, Anwar Shahzad a,⁎, Jaime A. Teixeira da Silva b a b

Plant Biotechnology Laboratory, Department of Botany, Aligarh Muslim University, Aligarh- 202 002, U.P., India Faculty of Agriculture and Graduate School of Agriculture, Kagawa University, Miki-Cho, Ikenobe, 2393, Kagawa-Ken, 761-0795, Japan

a r t i c l e

i n f o

Article history: Received 29 December 2011 Received in revised form 23 September 2012 Accepted 26 September 2012 Available online 2 October 2012 Keywords: Cryopreservation Dehydration Encapsulation Hydrogel Synthetic seed

a b s t r a c t Progress in biotechnological research over the last two decades has provided greater scope for the improvement of crops, forest trees and other important plant species. Plant propagation using synthetic seeds has opened new vistas in the field of agriculture. Synseed technology is a highly promising tool for the management of transgenic and seedless plant species, polyploid plants with elite traits and plant lines that are difficult to propagate through conventional propagation methods. Delivery of synseeds also alleviates issues like undertaking several passages for scaling up in vitro cultures as well as acclimatization to ex vitro conditions. Optimization of synchronized propagule development followed by automation of the whole process (sorting, harvesting, encapsulation and conversion) can enhance the pace of synseed production. Cryopreservation of encapsulated germplasm has now been increasingly used as an ex vitro conservation tool with the possible minimization of adverse effects of cryoprotectants and post-preservation damages. Through synseed technology, germplasm exchange between countries could be accelerated as a result of reduced plant quarantine requirements because of the aseptic condition of the plant material. © 2012 Elsevier Inc. All rights reserved.

Contents 1. 2.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Types of synseed . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Encapsulated desiccated . . . . . . . . . . . . . . . . . . . . . . 2.2. Encapsulated hydrated . . . . . . . . . . . . . . . . . . . . . . . 3. Hydrogel encapsulation techniques . . . . . . . . . . . . . . . . . . . . 3.1. Single layered synseed . . . . . . . . . . . . . . . . . . . . . . . 3.2. Double-layered synseed . . . . . . . . . . . . . . . . . . . . . . 3.3. Hollow beads . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Propagules used for synseed production . . . . . . . . . . . . . . . . . . 4.1. Bipolar propagules . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Unipolar propagules . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1. Nodes with apical or axillary buds and microshoots . . . . . 4.2.2. Microbulbs, microtubers, rhizomes and corms . . . . . . . . 4.2.3. Meristemoids, cell aggregates and primordia . . . . . . . . 5. Possible modes of synseed utilization . . . . . . . . . . . . . . . . . . . 5.1. In vitro plant production . . . . . . . . . . . . . . . . . . . . . . 5.2. Direct sowing . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3. Short-term germplasm conservation . . . . . . . . . . . . . . . . 5.4. Cryopreservation: an effective approach for long-term germplasm storage . 6. DNA marker technology in synseed experimentation . . . . . . . . . . . . 7. Problems, limitations and future prospects . . . . . . . . . . . . . . . . . Acknowledgment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

⁎ Corresponding author. Tel.: +91 9837061683. E-mail addresses: [email protected], [email protected] (A. Shahzad). 0734-9750/$ – see front matter © 2012 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.biotechadv.2012.09.007

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S. Sharma et al. / Biotechnology Advances 31 (2013) 186–207

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1. Introduction

2. Types of synseed

An active research front has emerged over the last decade with the goal of developing non-zygotic embryogenesis into a commercially useful method of plant propagation. From Haberlandt's postulate (1902) of artificial embryo cultivation to the concept proposed by Murashige (1977), artificial seeds have evolved from a futuristic idea into a real field of experimental research. The term “artificial seed”, which was first coined by Murashige, is now also known by other names including manufactured seed, synthetic seed or synseed. The original definition of an artificial seed, as given by Murashige (1978), was “an encapsulated single somatic embryo”, i.e., a clonal product that could be handled and used as real seed for transport, storage and sowing and that, therefore, would eventually grow either in vitro or ex vitro, into a plantlet (“conversion”). Gray and Purohit (1991) also defined synseed as “a somatic embryo that is engineered for the practical use in commercial plant production”. Thus, synseed production was previously limited to those plants in which somatic embryogenesis had been reported. However, many plant species remain recalcitrant to somatic embryogenesis. However, Bapat et al. (1987) proposed that synseeds could be produced from in vitro derived propagules other than somatic embryos, especially in non-embryogenic species; in Morus indica, for example, they proposed the use of encapsulated axillary buds. Thus, a synseed is referred to as artificially encapsulated somatic embryo, shoot bud or any other meristematic tissue that can be used as functional mimic seed for sowing, possesses the ability to convert into a plant under in vitro or ex vitro conditions, and can be stored (Ara et al., 2000; Capuano et al., 1998). This definition extends the concept of the synthetic seed from its bonds to somatic embryogenesis and links the term to its use (storage, sowing) and product (plantlet). In response to this shortcoming, the possibility of using non-embryogenic vegetative propagules such as shoot tips, nodal segments/axillary buds, protocorm like bodies (PLBs), organogenic or embryogenic callus has been explored as a suitable alternative to somatic embryos (Ahmad and Anis, 2010; Ara et al., 2000; Danso and Ford-Llyod, 2003; Faisal and Anis, 2007; Nhut et al., 2005; Ozudogru et al., 2011; Rai et al., 2008b; Sharma et al., 2009a,b; West and Preece, 2009). To complete this definition, it should be emphasized that the propagule must be able to grow into a plantlet after sowing (Piccioni, 1997). Even though in vitro-derived propagules were used in most synseed studies for encapsulation, it is also possible to encapsulate propagules excised directly from in vivo cultivated mature plants. For example, Pattnaik et al. (1995) successfully encapsulated the dormant vegetative buds of an in vivo-grown three-year old mature mulberry tree. More recently, Banerjee et al. (2012) produced synseed containing young sprouted vegetative microshoots together with a small basal rhizome portion excised from in vivo-grown rhizomes of Curcuma amada which were stored in lightly packed polythene packets. Over the past two decades, extensive progress has been made in synseed technology. Rai et al. (2009) presented a brief overview on synseed technology development in fruit crops only while Ara et al. (2000) and Saiprasad (2001) described the applications, prospects and limitations of sysnseed technology, but both those reviews are either incomplete, or outdated. The present review provides an upto-date, elaborate and refreshing perspective on synseed technology covering as wide a range of plant species as possible. Synseed technology is highly promising for the conservation and mass clonal propagation (Singh et al., 2006) of valuable rare hybrids, elite genotypes, sterile unstable genotypes and genetically engineered plants for which seeds are either not available or that require a mycorrhizal-fungal association for their germination as in the case of orchids. Recently, encapsulation technology has attracted the interest of researchers for germplasm delivery and for various analytical studies (Ara et al., 2000). The possible applications of synseed are summarized in Fig. 1.

Since the formulation of the concept of synseed by Murashige (1977), a number of studies have been undertaken in this area of plant biotechnology. The basic hindrance to synseed technology was the lack of a natural endosperm and protective coatings in somatic embryos that made them inconvenient to store and handle (Redenbaugh et al., 1993). Furthermore, the absence of a quiescent resting phase and the inability of undergoing dehydration limited the utility of somatic embryos as a source of synseed production. Thus, the primary effort in synseed technology was to treat somatic embryos in such a way that they mimicked zygotic embryos during storage and other applications. This was the first major step in the success of synseed technology (Ara et al., 2000). Synseed technology has been extended by several research groups for a variety of plant species including cereals, fruits, vegetables, medicinal plants, forest trees, orchids and other ornamentals (Ara et al., 1999; Germanà et al., 1998; Ipekci and Gozukirmizi, 2003; Janeiro et al., 1997; Rai et al., 2008a,b; Utomo et al., 2008). Based on the literature available to date, synseeds can be separated into two categories: 2.1. Encapsulated desiccated Coated desiccated embryos represent an ideal form of synseed (Pond and Cameron, 2003) for which somatic embryos are first hardened to withstand desiccation before encapsulation. This induces quiescence in the embryos and provides more handling flexibility in large-scale production systems. Thus, the ability of somatic embryos to withstand drying to low moisture content is an important factor for storage and plays a critical role in the developmental transition between maturation and conversion. Such types of synseeds can only be produced in those plants whose somatic embryos are desiccation-tolerant. Desiccation can be achieved either slowly over a period of one or two weeks sequentially using chambers of decreasing relative humidity, or rapidly by unsealing the Petri dishes and leaving them overnight to dry (Ara et al., 2000). The drying rate is one of the critical factors for the efficient survival of somatic embryos. If the embryos are immature, slow drying over one week is optimal, but if there is large number of fully mature embryos, rapid drying in a laminar flow bench is preferable (Senaratna et al., 1990). Desiccation tolerance can also be induced with maturation medium with high osmotic potential induced by either increased levels of permeating osmoticants (e.g., sucrose, mannitol), non-permeating osmoticants (e.g., polyethylene glycol or PEG) or high gel strength media (to limit water availability). While working with ginger synseeds, Sundararaj et al. (2010) found that sucrose-dehydration was more effective than air-dehydration in terms of re-growth ability by providing required nutrients; moreover, rapid moisture loss during air dehydration resulted in poor conversion frequency. For sucrose-dehydration, Sundararaj et al. (2010) transferred the synseeds to liquid nutrient medium containing various concentrations of sucrose for 16 h and kept them in an incubator-shaker for 16 h at 25±2 °C. Synseeds dehydrated resulted in 86% conversion whereas higher concentrations (0.50 M and 0.75 M) resulted in no conversion. Other sub-lethal stresses such as low temperature and nutrient deprivation also have a similar effect on desiccation tolerance (Pond and Cameron, 2003). Properly pretreated embryos remain viable when they are rapidly desiccated to less than 10–15% moisture content. Pretreatment with abscisic acid (ABA) also improves the conversion of somatic embryos both in desiccated and hydrated systems (Nieves et al., 2001; Pond and Cameron, 2003). Nieves et al. (2001) reported the effect of ABA and jasmonic acid (JA) on partial desiccation of encapsulated sugarcane somatic embryos. Before encapsulation, embryogenic callus with somatic embryos were placed on MS medium supplemented with 3.8 μM ABA and/or 4.7 μM JA, as described by Tapia et al. (1999),

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S. Sharma et al. / Biotechnology Advances 31 (2013) 186–207

Synseeds

Propagation

• Large-scale monoculture of rare,endangered, genetically engineered elite genotypes. • Mixed genotype plantation • Uniform genetic constitution of plantlets

Analytical tool

Conservation

• Cost-effective approach for ex situ germplasm conservation • Ensures economy of space, medium, storage period • Protection from environmental disasters • Extra protection to explants against pests and diseases

• Comparative study aid for zygotic embryogeny • Determining the role of endosperm in embryo development and germination • Seed coat formation studies • Study of somaclonal variation

Short to medium-term storage

Long-term storage

Slow-growth conservation

Cryopreservation

Transport

• Easy, cost-effective disease-free germplasm transportation • Direct greenhouse and field delivery • Germplasm exchange between countries without obligations form quarantine department

• Two-step freezing • Simple desiccation • Encapsulation-dehydration • Vitrification • Encapsulation-vitrification

• Maintenance under reduced temperature and/or reduced light intensity • Use of growth retardants such as ABA • Use of minimal growth medium • Use of osmoticum • Reduction in oxygen concentration • Combination of more than one treatment

Fig. 1. Potential uses of synseeds.

and maintained over 5 days at 36 °C. After encapsulation in calcium alginate enriched with artificial endosperm medium and osmotically dehydrated in 0.5 M sucrose over 24 h or non-dehydrated, somatic embryos were dehydrated over 96 h in chambers containing silica gel until the beads reached either 60% or 30% of water content. Water content was assessed according to wet weight and was evaluated until a thermodynamic balance was reached; thereafter the water activity (aw) of somatic embryos was determined according to Timbert et al. (1996). Survival of encapsulated-dehydrated embryos was achieved with ABA rather than with JA. Further, they reported that ABA induced an increase in protein, polyamine, free proline and starch levels in response to desiccation tolerance. Rai et al. (2008a) induced quiescence in somatic embryos of guava (Psidium guajava) using different concentrations of ABA and sucrose. Maturation was possible by transferring earlier stages of non-encapsulated somatic embryos (globular to cotyledonary stages) to full-strength MS medium solidified with 0.8% agar and supplemented with 1 mg l−1 ABA and 10% sucrose for 4 weeks. As the concentration of sucrose (3–9%) or ABA (0.01–1.0 mg l−1) increased in MS medium, the percentage conversion of encapsulated somatic embryos decreased significantly. Encapsulated somatic embryos after storage on MS medium supplemented with 9% sucrose or 1 mg l−1 ABA for different durations (0–60 days) converted to plantlets when transferred to MS medium containing only 3% sucrose. About 20.8% and 37.5% of encapsulated somatic embryos converted after storage on MS medium containing ABA (1 mg l−1) or sucrose (9%) for 60 days. After desiccation, embryos are coated with a protective and nutritive layer that inhibits mechanical damage during handling and provides nourishment during the early stages of conversion (Pond and Cameron, 2003). The coating must be non-toxic, non-aqueous (to

prevent the rehydration of embryos), it should be able to melt at a relatively low temperature (so the embryos do not suffer thermal damage during coating) and must be able to adhere to embryos. Moreover, the rigidity of the coating should be soft enough in order to allow the emergence of shoot and root primordia. The first report on synseed production was published by Kitto and Janick (1982). They produced desiccated synseeds of carrot (Daucus carota) by encapsulating multiple somatic embryos in a water-soluble resin, polyoxyethyelene glycol (Polyox) then desiccating these embryos. Of the various compounds tested for encapsulation of celery (Apium graveolens) embryos, Kitto and Janick (1985) selected polyoxyethylene which is readily soluble in water and dries to form a thin film, does not support the growth of microorganisms and is non-toxic to the embryo. Later on, Janick et al. (1993) extended this technology for encapsulating a PEG-based coating mixture for carrot somatic embryos and embryogenic callus that was allowed to dry for several hours on a Teflon surface in a sterile hood. The dried mixture was then placed onto a culture medium, allowed to rehydrate and then scored for embryo survival. Three years later, Timbert et al. (1996) suggested that the survival percentage of encapsulated carrot somatic embryos was dependent on the type of encapsulating matrix and on the speed of dehydration. Some hydrogels delayed dehydration and preserved the water content of somatic embryos. They found alginate with gellan gum to be the most efficient hydrogel in maintaining high water activity (aw) around somatic embryos. To exert different dehydration speeds, they used a slow dehydration protocol (95–15% relative humidity (RH) into the chamber within 11.5 days). In the absence of any maturing pretreatment, 72.9% of alginate-gellan gum-encapsulated carrot somatic embryos, dehydrated

S. Sharma et al. / Biotechnology Advances 31 (2013) 186–207

to 15% RH and rehydrated in moisture air (90% RH), germinated. Desiccation in alginate-encapsulated and non-encapsulated somatic embryos from two cell lines (P28H9 and P29H17) of oak tree (Quercus robur) was also induced by Prewein and Wilhelm (2003). They compared the effect of desiccation under two treatments: either after 1 week desiccation in a multi-well chamber (partial drying) at 27 °C in the dark at a relative humidity of more than 95% or after drying for 2 h in a laminar flow cabinet. No significant effect of desiccation compared to alginate-encapsulation was observed, either on the germination rate or on the conversion rate. Both cell lines responded poorly with only 8% and 7% conversion rates following rapid-drying. When the two desiccation methods used with oak synseeds of cell line P29H17 were compared, no differences were detected. The conversion of encapsulated somatic embryos was significantly higher with slow desiccation than non-encapsulated controls. This can be attributed to the effect of desiccation rather than the effect of encapsulation in general. 2.2. Encapsulated hydrated Redenbaugh et al. (1984) developed hydrogel encapsulation of individual somatic embryos of alfalfa (Medicago sativa) and patented this technology in 1988. Since then it remains the most studied strategy of synseed production (Ara et al., 2000; Rai et al., 2009). A number of coating agents such as agar, sodium alginate, potassium alginate, sodium pectate, carrageenan, sodium alginate with carboxymethyl cellulose, gelatin, gelrite, guargum, tragacanth gum, etc. have been tested as hydrogels (Ara et al., 2000; Rai et al., 2009). Among all, sodium alginate has been frequently selected because of its moderate viscosity, low spin ability of solution, no toxicity for propagules, quick gellation, low cost and bio-compatibility (Saiprasad, 2001; Swamy et al., 2009). Sodium alginate and calcium salt are reported to be the best combination for encapsulation since these ions are non-damaging, have a low price, are easy to use and result in high embryo-to-plant conversion. The capsule gel potentially serves as a reservoir of nutrients that helps in the survival and speedy growth of embryos (Redenbaugh et al., 1987). 3. Hydrogel encapsulation techniques Since hydrogel encapsulation is the most successful and widely accepted approach to synseed production, the next section presents an overview of hydrogel encapsulation and its modifications. 3.1. Single layered synseed This is the simplest way to achieve hydrogel encapsulation. To produce single layered synseeds, propagules are carefully isolated either from an in vitro or an in vivo source and mixed with a suitable hydrogel such as sodium alginate (0.5–5.0%, w/v). Alginate is a straight chain, hydrophilic, colloidal polyuronic acid primarily composed of hyrdo-βD-mannuronic acid residues with 1–4 linkages (Cameron, 2008; Martinsen et al., 1989). Alginate solution is prepared either in double distilled water (DDW) or in liquid nutrient medium and dropped, along with the propagule, as a bead into a complexion agent such as calcium chloride (CaCl2·2H2O) or calcium nitrate [Ca(NO3)2] solution (30–150 mM). The pH of both the alginate matrix and complexing agent is adjusted to 5.8 prior to sterilization. Generally, for sterilization, both the gel matrix and complexing agent are autoclaved at 1.06 kg cm−2 and 121 °C for 20 min (Kavyashree et al., 2006; Sharma et al., 2009a,b), although Rihan et al. (2011) sterilized the sodium alginate solution by tyndallisation and calcium chloride by autoclaving. The major principle involved in the alginate encapsulation process is the formation of round and firm calcium alginate beads due to an ion exchange process between sodium (Na+) and calcium (Ca++) ions. The permeability, hardness or rigidity of the beads and the subsequent

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success of the encapsulation method is affected by alginate and calcium chloride concentration used and curing time and it may vary with different propagules as well as with the different plant species (Table 1). Hence, the concentrations of these two solutions and complexion time must be optimized for the formation of an ideal bead (Saiprasad, 2001). In most of the reports, 3% (w/v) sodium alginate and 100 mM calcium chloride for 20–30 min has proved to be the best combination for the formation of an ideal synseed (Ahmad and Anis, 2010; Alatar and Faisal, 2012; Hung and Trueman, 2011, 2012a,b; Ozudogru et al., 2011; Sarkar and Naik, 1998; Tabassum et al., 2010). However, in several studies, 3% sodium alginate upon complexion with 75 mM calcium chloride for 20–30 min was the optimum combination for proper hardening of beads viz., Dendrobium, Oncidium and Cattleya orchids (Saiprasad and Polisetty, 2003) and Fragaria × ananassa (strawberry) and Rubus idaeus (raspberry) (Lisek and Orlikowska, 2004). In contrast, for the encapsulation of nodal segments of Pogostemon cablin (Swamy et al., 2009) and Spilanthes acmella (Sharma et al., 2009b) and the microshoots of Zingiber officinale (ginger) (Sundararaj et al., 2010), 4% sodium alginate with 100 mM calcium chloride was optimum. This variation in sodium alginate concentration for synseed formation in different plant species might be due to the variation in commercial source from which the chemicals were purchased, as reported earlier by Ghosh and Sen (1994) and Mandal et al. (2000). In Ocimum spp. (basil), the best results with respect to capsule formation were obtained with Sigma's (Sigma, St. Louis, USA) 4% sodium alginate (with 75 mM calcium chloride) than with the product of CDH (Central Drug House, Mumbai, India) (Mandal et al., 2000). In Spilanthes mauritiana, optimum bead formation was possible with 3% sodium alginate of Loba Chemie (Mumbai, India) and 100 mM calcium chloride but for S. acmella this was possible with the CDH product (4% sodium alginate with 100 mM calcium chloride) (Sharma et al., 2009a,b). At lower concentrations (1–2%), sodium alginate became unsuitable for encapsulation because of a reduction in its gelling ability following exposure to high temperature during autoclaving (Larkin et al., 1998). On the contrary, high concentrations of sodium alginate (5–6%) beads were isodiametric but too hard, causing considerable delay in sprouting of shoots (Ahmad and Anis, 2010; Sharma et al., 2009a,b). To make synseed beads, peristaltic pumps (Blandino et al., 2000) and pipettes are usually employed (Ara et al., 2000; Hung and Trueman, 2012a,b). The solution in which beads are formed is constantly stirred/shaken in order to avoid them sticking to each other and to increase the formation of spherically-shaped beads during polymerization (Mallón et al., 2007). The size of the beads is controlled by varying the inner diameter of the pipette nozzle (Ara et al., 2000). Nishitha et al. (2006) used a sterilized pipette, 10 mm in diameter, for shoot tip encapsulation in Chonemorpha grandiflora while Hung and Trueman (2011, 2012a, 2012b) used a 7 mm diameter Pasteur pipette for the encapsulation of shoot tips and nodal segments of Khaya senegalensis (mahogany) and Corymbia torelliana × C. citriodora (eucalypt). For the encapsulation of gametophyte buds of an endangered moss, Splachnum ampullaceum, Mallón et al. (2007) used a drip because: a) peristaltic pumps delivered the material too quickly to be employed with fragile plant material and since sterile conditions required for in vitro culture are difficult to obtain; b) The use of pipettes is tiring and the precision necessary to form uniform beads (i.e. with a low standard deviation in diameter of 0.2 or 0.3 mm) is unlikely to be reached. These firm beads are washed with sterile DDW to remove traces of adhering chemicals (Fig. 2) and placed immediately either on a nutrient medium or sown in different planting substrates such as wet filter paper, cotton or Soilrite™ moistened with nutrient solution or DDW (Rai et al., 2009). The composition of the gel matrix is an important factor that significantly affects the conversion performance of encapsulated tissue. For effective re-growth and conversion of the encapsulated plant

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Table 1 Optimum conditions for ideal synseed or ca-alginate bead formation in different plant species (2000 onwards, in chronological order and alphabetical order within each year). Proliferation medium of the mother culture before synseed production

Propagule Pretreatment or Propagule size after treatment used for (as required) encapsulation

Actinidia deliciosa (‘Hayward’ kiwifruit)

Macro- and micro-elements as in QL medium, organic compounds as in the 2nd formulation of Walkej (1972)+ sucrose (25 g/l)+ 1 mg/l GA3 +1 mg/l BA

MC (apical and axillary buds)

Ocimum americanum (hoary basil), O. basilicum (sweet basil), O. gratissimum (shrubby basil), O. sanctum (sacred basil)

Explant taken from in vivo Axillary bud source

Morus alba, M. australis, M. bombycis, M. cathyana, M. latifolia and M. nigra (mulberry)

Modified MS medium + 100 mg/l myo-inositol + 30 g/l sucrose + 8 g/l agar + 4.4 μM BA MS medium + 4.5 μM 2,4-D, 1 mg/l D-biotin, 100 mg/l L-glutamine + 100 mg/l malt extract (suspension culture) B5 medium + 4.65 μM Kn + 396.48 μM PG MS medium supplemented with 5 × 10−6 μM NAA + 1 × 10-5 μM Kn

Musa spp. AAB group (banana cv. ‘Rasthali’)

Adhatoda vasica (vasaka)

Allium sativum (garlic)

4 mm

Encapsulating matrix composition

Concentration Polymerization Composition of of CaCl2·2H2O time (min) regrowth medium for or ⁎CaCl2 synseed

References % Conversion (complete plantlet recovery in vitro)

2.5% SA + macro- and microelements of QL medium + 25 g/l sucrose + 1 mg/l GA3 + 1 mg/l BA

100 mM

30

½-strength proliferation medium without PGR

Adriani et al. 56.2–36.1% (2000) (after cold treatment) 31.4– 25.7% (after root induction treatment) 50– 57% (after adding sucrose in sowing and encapsulation matrix)

4% SA + MS medium + 100 mg/l myo-inostiol + 30 g/l + 1.1 μM BA (O. americanum) +2.2 μM BA (O. gratissimum)+4.4 μM BA (O. basilicum and O. sanctum) 4% SA + MS medium + 30 g/l sucrose + 4.4 μM BA

75 mM

NA

MS medium + 30 g/l sucrose + 1.1 μM BA (O. americanum) + 2.2 μM BA (O. gratissimum) + 4.4 μM BA (O. basilicum and O. sanctum)

95–99%

Mandal et al. (2000)

⁎75 mM

30

MS medium + 4.4 μM BA

78-98% (for shoot development)

Pattnaik and Chand (2000)

–

–

MS medium

66%

Ganapathi et al. (2001)

20

B5 medium

66.28%

Anand and Bansal (2002)

30

½-MS 95% medium + 4.4 × 10-2 μM sucrose + 5 × 10–6 μM NAA + 1 × 10-5 μM Kn 100%

NA

i. Cold treatment was given to 1 month old proliferation culture by placing in cold chamber for 1 month of darkness at 4 °C. ii. MCs were pretreated with 15 g/l sucrose and 1 mg/l or 3 mg/l IBA. The cultures were kept in darkness inside the growth chamber on a 100 rpm rotatory shaker for 24 or 48 h. After root induction MCs submitted to root initiation on sterile filter paper moistened with nutrients. iii. 30 g/l sucrose increase in the sowing medium as well as in artificial endosperm. -

NS or axillary bud

5 mm

-

SE

–

½-MS medium + 10 μM Zea + 30 g/l sucrose (maturation medium)

5% SA + MS medium

Shoot bud

1–4 mm

–

Calli

NA

-

1.1% 4% SA + B5 Medium + 4.65 μM Kn + 50 mg/l PG 1.5% SA + half-strength *50 mM MS medium + 5 × 10–6 μM NAA + 1 × 10-5 μM Kn

MS

2-5 mm

30

Kim and Park (2002)

S. Sharma et al. / Biotechnology Advances 31 (2013) 186–207

Plant species (common name)

Proliferation medium of the mother culture before synseed production

Propagule used for encapsulation

Ananas comosus (pineapple)

MS basal medium + MS vitamins + 0.56 mM myo-inositol + 0.06 M sucrose + 9.67 μM NAA + 9.84 μM IBA + 9.29 μM Kn for the induction of multiple shoots

Paulownia elongata (Royal Empress tree)

MS medium + 500 mg/l casein hydrolysate + 10 mg/l TDZ + 3% sucrose ½-MS medium + 6.97 μM Kn

Ipsea malabarica (Malabar daffodil orchid)

Dendrobium, Oncidium and Cattleya (orchids)

Pretreatment or after treatment (as required)

Encapsulating matrix composition

Concentration Polymerization Composition of of CaCl2·2H2O time (min) regrowth or ⁎CaCl2 medium for synseed

Pretreatment of shoots in liquid medium of W basal medium + W vitamins + 0.56 mM myo-inositol + 0.03 M sucrose + 10.8 μM NAA + 39.4 μM IBA and agitated on a gyratory shaker at 100 rpm for 12 h MS basal medium (maturation)

3% SA + MS basal medium + 0.56 mM myo-inositol + 0.06 M sucrose

1.36 g/ 150 ml

3% SA + MS medium

*50 mM

30

Peat + perlite (1:1)

53.3%

Ipekci and Gozukirmizi (2003)

Shoots developed on proliferation medium transferred on ½-MS + 6.97 μM + 6 or 8% sucrose P24 medium + 0.9 μM BA + 3% sucrose (maturation medium)

3% SA + ½-MS medium + 6.97 μM Kn

*0.7%

30

½-MS medium with or without 6.97 μM Kn

100%

Martin (2003)

4% SA + P24 medium + 3% sucrose

50 mM

20

26%

Prewein and Wilhelm (2003)

NA

-

75 mM

30

100% (for all three species)

Saiprasad and Polisetty (2003)

Nodal segments were placed on ½-MS supplemented with 1.0 mg/l IBA for 10 days Pre-conditioning with 10 mg/l mannitol added to proliferation media

3% SA + ¾-MS medium + 0.44 μM BA with 0.54 μM NAA (for Dendrobium), 2.69 μM NAA (for Oncidium) and 5.38 μM (for Cattleya) 3% SA + half-strength MS medium

*75 mM

20

P24 medium + 0.1 μM IBA + 0.9 μM BA + 3% sucrose MS + 0.44 μM BA + 0.54 μM NAA (for Dendrobium) MS + 2.69 μM NAA (for Onicidium) MS + 5.38 μM NAA (for Cattleya) ½-MS medium

85%

Chand and Singh (2004)

3% SA + 4% glucose + water (strawberry) or 3% SA + MS medium (raspberry)

75 mM

30

Proliferation medium

3.7 shoots/bead (after 9 month of storage and 2nd subculture) 5.6 shoots/bead (after 9 month of storage and 2nd subculture)

Lisek and Orlikowska (2004)

10

Whatman filter paper irrigated with ½-MS medium + vitamins + 2% sucrose

47%

Arun Kumar et al. (2005)

30

½-MS medium

100%

Chithra et al. (2005)

SE

1.5– 2.0 mm

Bulbs

NA

Embryogenic culture lines SE were maintained on P24 medium + 0.9 μM BA + 3% sucrose MS medium + 4.44 μM BA PLB

4–8 mm

Dalbergia sissoo (sissoo)

Explants taken from in vivo source

NS

10 mm

Fragaria ananassa (strawberry) and Rubus idaeus (raspberry)

Boxus medium + 2.2 μM BA + 2.46 μM IBA (for strawberry) and MS medium with NH4NO3 and KNO3 reduced by 50% + 3.55 μM BA + 0.49 μM IBA, both supplemented with WPM vitamins Embryogenic calli from dehusked seed induced on MS medium + 30 g/l sucrose + 2 mg/l 2,4-D. Calli were then transferred to MS medium + 30 g/l sucrose + 2 mg/l BA + 0.5 mg/l NAA + 0.5 mg/l MS medium + 0.45 μM 2,4-D (induction medium); liquid ½-MS medium + 0.23 μM 2,4-D

ST

3 mm

SE

NA

Self breaking treatment was given to the prepared synseed, by dipping them in 200 mM KNO3 for 60 min and rising in sterile tap water for 40 min till the bead became swollen

SE

–

Semi-solid ½-MS medium

Oryza sativa (hybrid rice)

Rotula aquatic (takad)

4% SA + MS medium *1.5% (except CaCl2) + 30 g/l sucrose + plant growth regulators (0.5 mg/l NAA, 0.5 mg/l IAA, 2 mg l BA) + antibiotics (1 mg/l bavistin, 1 mg/l streptomycin) + 1.25% AC *50 mM 3% SA + 1/2-MS medium (except CaCl2) + 3% sucrose

% Conversion References (complete plantlet recovery in vitro) Soneji et al. (2002)

MS basal medium, MS vitamins + 0.56 mM myo-inositol + 0.06 M sucrose + 9.67 μM NAA + 9.84 μM IBA and 9.29 μM Kn

191

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S. Sharma et al. / Biotechnology Advances 31 (2013) 186–207

Quercus robur (pedunculate oak)

Propagule size

192

Table 1 (continued) Concentration Polymerization Composition of of CaCl2·2H2O time (min) regrowth medium for or ⁎CaCl2 synseed

References % Conversion (complete plantlet recovery in vitro)

2.5% SA + MS medium

NA

30

86.13% (after 15 days of storage at 8 °C)

Gangopadhyay et al. (2005)

2.5% SA + DCR basal DCR medium + 175 mM salts maltose + 80 μM ABA + 9 g/l Gellan gum was used for maturation. All cultures were placed in the dark at 25 ± 2 °C for 8–14 weeks 3% SA 100 mM (calcium nitrate) – 3% SA + MS + 1.0 mg/l IAA

*100 mM

5

MS medium + 0.022 mM BA + 0.003 mM IAA. For root induction, microshoots were placed in liquid MS medium with Luffa-sponge) + 0.01 mM IBA ½-DCR basal medium

89%

Malabadi and van Staden (2005)

25

25

MS medium

60.6%

*75 mM

30

Perlite moistened with MS medium

50% (without IAA treatment)

Manjkhola et al. (2005) Pinker and Abdel-Rahman (2005)

3–5 mm

–

4% SA

1.036 g/ 150 ml

30–40

MS medium + 1 mg/l BA + 0.5 mg/l IBA

Axillary bud

0.5 cm

-

40

Naik et al., 1999, 2000

NS

3–6 mm

–

*5 × 104 μM 4% SA + LSBM medium + 8.88 μM BA + 2.0 μM TIBA *100 mM 3% SA + MS medium + 3% sucrose and 100 mg/l myo-inositol + 4.44 μM BA + 0.54 μM NAA

Chonemorpha grandiflora (Bengal creeper)

MS medium + 13.3 μM BA + 2.46 μM IBA

ST

NA

–

LSBM medium + 8.88 μM BA + 2 μM TIBA MS medium + 100 mg/l myo-inositol + 3% sucrose + 4.44 μM BA + 0.54 μM NAA (for Source A) MS medium + 3% sucrose and 100 mg/l myo-inositol (for sprouting, source B) MS medium + 0.54 μM NAA (rooting medium) ½-MS medium

88% (after 30 days of storage at 22 ± 2 °C) 48.2%

Hibiscus moscheutos (hardy hibiscus)

DKW medium + 3% sucrose + 1 × 10-7 M TDZ

NS

3–5 mm

Proliferation medium of the mother culture before synseed production

Propagule Pretreatment or Propagule size after treatment used for (as required) encapsulation

Ananus comosus (pineapple)

MS medium + 0.55 mM myo-inositol + 88 mM sucrose + 0.22 mM BA + 0.003 mM IAA

MS

1.5– 2.0 mm

–

Pinus patula (Mexican weeping pine)

DCR basal medium + 120 mM maltose + 2 g/l Gellan gum + 2 μM 2,4-D + 2.5 μM NAA + 1 μM BA

SE

NA

Arnebia euchroma (zicao; arnebia root) Dendranthema × grandiflora (chrysanthemum)

MS medium + 2.5 μM SE IBA + 2.5 μM BA Modified MS salts (mac- NS ro-nutrient content reduced by half) + 2.5 mg/l thiamine HCl + 0.2 mg/l pyridoxine-HCl + 0.2 mg/ l biotin + 100 mg/l + 8 g/ l + 30 g/l sucrose + 0.5 mg/l BA MS medium + 1.0 mg/l SB, PC BA + 0.5 mg/l IBA

5–6 mm

Morus sp. (mulberry variety-S54)

LSBM medium + 8.88 μM BA + 2 μM TIBA

Punica granatum (pomegranate)

Vanilla planifolia (vanilla)

4–5 mm

3% SA + ½-MS (except CaCl2) + 0.49 μM IBA + 11.7 μM silver nitrate Beads were exposed to light 2.75% SA for at least 2 weeks prior to planting in the greenhouse

30

*50 mM

30

*50 μM

30

Medium grade vermiculite placed under intermittent mist (bead planted

Divakaran et al. (2006)

Kavyashree et al. (2006)

75% (spouting only for source A) 81% (sprouting for source B) 21% conversion only from source B 87% rooting (source A) 90% rooting (source B)

Naik and Chand (2006)

95%

Nishitha et al. (2006)

Preece and West (2006)

S. Sharma et al. / Biotechnology Advances 31 (2013) 186–207

Encapsulating matrix composition

Plant species (common name)

Proliferation medium of the mother culture before synseed production

Phyllanthus amarus (bahupatra) Populus tremula × P. tremuloides (hybrid aspen) Hibiscus moscheutos (hardy hibiscus) Acca sellowiana (pineapple guava, feijoa)

Propagule used for encapsulation

MS medium + 0.75 mg/l BA NA

DKW medium + 3% sucrose + 10-7 M TDZ LPm medium + 8 mM Glu + 20 μM 2,4-D + 0.1 mg/l myo-inositol Tylophora indica (antamul) Explant taken from in vivo source Protonema cultured Splachnum ampullaceum on Heller's medium (purple-vased stink + 3% sucrose moss, bryophyte)

OMM

Citrus nobilis × C. deliciosa (Kinnow mandarin)

For somatic embryogenesis, ovules were cultured on MS medium + 9.29 μM Kn + 3% sucrose Coleus forskohlii (makandi) MS medium + 2.22 μM BA

Pogonatherum panicum (golden hair grass)

MS medium + 2 mg/l BA + 0.2 mg/l NAA

Pretreatment or after treatment (as required)

Encapsulating matrix composition

Concentration Polymerization Composition of of CaCl2·2H2O time (min) regrowth or ⁎CaCl2 medium for synseed 1 cm deep and not covered in greenhouse) MS medium

% Conversion References (complete plantlet recovery in vitro)

ST

NA

–

3% SA + MS medium

75 mM

20

ST

0.5– 0.7 cm

–

*1.4%

5

MS medium + 1.3 μM BA + 3% sucrose

100%

NS

4 mm

–

4% SA + MS medium + 1.3 μM BA + 3% sucrose 2.75% SA

50 mM

30

SE

NA

–

1% SA + LPm medium

*50 mM

15

DKW medium + 3% sucrose + 10-7 M TDZ LPm medium

80% (after 1.5 year storage) 70.3% (after 60 days of culture)

NS

3–5 mm

–

3% SA + MS medium

100 mM

30

91%

Gametophyte bud

–

–

1% SA

*100 mM

20 min

MS medium + 2.5 μM BA + 0.5 μM NAA Heller's medium + 3% sucrose + B5 vitamins

NS

3–4 mm

2.5% SA + ½-OMM + 1 mg/l Zea + 50 g l-1 sucrose

*1.1%

35

OMM

SE

NA

Two pretreatment a) sprouting induction (micro-cuttings were placed in 50 ml sealed glass vessel containing 15 ml of 1 mg/l GA3 + 30 g/l sucrose + vessel put on a rotator shaker (100 rpm for 24 h) in growth chamber under darkness at 23 ± 2 °C b) sprouting initiation treatment (after sprouting induction, micro-cuttings were placed on filter paper which placed on vessel containing ½-MS proliferation medium + 0.7% agar, kept in growth chamber at 23 ± 2 °C under darkness for 6 days) For maturation embryos allowed to grow on same medium as used for induction

4% SA + MS medium

75 mM

45

MS medium + 9.29 μM Kn

81.94%

Singh et al. (2007)

ST, NS

–

–

3% SA + MS + 2% sucrose

1.36 g/ 150 ml

NA

92% (shoot multiplication) 75% (complete plantlet recovery)

Swaroopa et al. (2007)

SB

3–5 mm

–

3% SA + MS medium + 1% AC

*2% with 0.3% 20 bavistin

MS medium + 2.22 μM BA + 3% sucrose (for multiplication) MS medium + 3% sucrose (for complete recovery of plantlet) ½-MS medium + 2% sucrose

>90%

Wang et al. (2007)

90%

Singh et al. (2006) Tsvetkov et al. (2006) West et al. (2006) CangahualaInocente et al. (2007)

Faisal and Anis (2007) 100% (after short Mallón et al. to medium-term (2007) storage) 60% (after 2 years of storage) 50% (after 2.5 years of storage) Data was not in Micheli et al. (2007) terms of % conversion (1.8 and 1.9 shoots/ capsule after 30 days of storage at 18 and 4 °C)

193

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S. Sharma et al. / Biotechnology Advances 31 (2013) 186–207

Olea europaea (olive)

Propagule size

194

Table 1 (continued) Propagule Pretreatment or Propagule size after treatment used for (as required) encapsulation

Plant species (common name)

Proliferation medium of the mother culture before synseed production

Psidium guajava (guava)

SE Somatic embryogenesis induced from zygotic embryos on MS medium + 4.52 μM 2, 4-D + 5% sucrose. After 8 days of treatment with 2,4-D explants transferred to PGR-free MS medium +5% sucrose Rai et al. 2007 ST

Psidium guajava (guava)

Encapsulating matrix composition

Concentration Polymerization Composition of of CaCl2·2H2O time (min) regrowth medium for or ⁎CaCl2 synseed

References % Conversion (complete plantlet recovery in vitro)

Globular and cotyledonary stage embryos transferred to MS medium + 1.0 mg/l ABA or 10% sucrose for maturation

2% SA

100 mM

20–30

MS medium + 3% sucrose

91.6%

Rai et al. (2008a)

3–5 mm

–

3% SA

*100 mM

20–30

Liquid MS medium

97.2%

Anderson medium + 7 mg/l 2-iP + 100 mg/l PVP + 100 mg/l ascorbic acid + 10 mg/l citric acid R4 medium

96%

Rai et al. (2008b) Singh (2008)

Rhododendron maddeni (rhododendron)

Anderson medium + 7 mg/l 2-iP + 100 mg/l PVP + 100 mg/l ascorbic acid + 10 mg/l citric acid

ST

NA

–

3% SA + Anderson medium + 3% sucrose

60 mM

30

Spartina alterniflora (smooth cordgrass)

Calli proliferated on modified R4 medium followed by suspension culture in liquid medium modified by adding FeSO4.7H2O, myo-inositol, thiamine HCl, pyridoxine HCl and casein acid hydrolysate –

SE, MP

NA

–

2% SA

*100 mM

30

SE and zygotic embryos

3–4 mm (SEs)

–

*15 g/l (for SEs) * 5.5 g/l (for zygotic embryos)

10 (for SEs) 30 Macro and micronutrients of MS (for zygotic medium embryos)

50 mM

30

MS medium + 0.5 μM TDZ + 0.075% PPM

100%

Lata et al. (2009)

100 mM

5

½-MS medium + 3% sucrose + 10% coconut water ½-MS medium

68% (shoot development)

Nagesh et al. (2009)

>90% (after 3 week of storage at 4 °C) 87.8%

Sharma et al. (2009a)

80%

Siddique and Anis (2009) Srivastava et al. (2009)

Cannabis sativa (marijuana)

MS medium + 0.5 μM TDZ NS

NA

–

Curculigo orchioides (black musali)

MS medium + 1 mg/l BA + 1 mg/l Kn

Shoot bud

5–6 mm

–

4% SA (for SEs) or 3% SA (for zygotic embryos) + MS medium + 0.5 mg/l IAA + 0.5 mg/l NAA + 2 mg/l BA + 30 g/l sucrose 5% SA + MS + 0.5 μM TDZ + 2.5 μM IBA +0.3–0.5% PPM 2.5% SA + MS medium

Spilanthes mauritiana (toothache plant)

MS medium + 1.0 μM BA + 0.5 μM IAA

NS

3–4 mm

–

4% SA + MS medium

100 mM

20–25

Spilanthes acmella (toothache plant)

In vitro seedling grow on ½-MS medium + 0.5 μM GA3 Explants taken from in vivo source NA

NS

3–4 mm

–

4% SA + MS medium

100 mM

20–25

MS medium + 1.0 μM BA + 0.5 μM NAA

NS

NA

–

3% SA + MS medium

75 mM

20

ST, NS

3–5 mm

–

3% SA + 1.5% sucrose

*3%

20–30

½-MS + 5 μM BA + 0.5 μM IAA MS medium

5 mm

–

4% SA + MS medium + 2% sucrose

*100 mM

30

Nothofagus alpina (rauli-beech)

Ocimum basilicum (sweet basil) Cineraria maritima (dusty miller or silver dust)

Pogostemon cablin (patchouli)

MS medium + 2.22 μM BA NS

MS medium + 2.22 μM BA

>90%

Utomo et al. (2008)

Cartes et al. 44% (for SEs) 100% (for zygotic (2009) embryos)

11.76% (after 6 month of storage at 25 ± 2 °C) 91.1%

Sharma et al. (2009b)

Swamy et al. (2009)

S. Sharma et al. / Biotechnology Advances 31 (2013) 186–207

2–3 mm

Proliferation medium of the mother culture before synseed production

Propagule used for encapsulation

Propagule size

Pretreatment or after treatment (as required)

Encapsulating matrix composition

Concentration Polymerization Composition of of CaCl2·2H2O time (min) regrowth or ⁎CaCl2 medium for synseed

Vitex negundo (five-leaved chaste tree) Simmondsia chinensis (jojoba)

MS medium + 5.0 μM BA + 0.5 μM NAA MS medium + 2.0 mg/l BA + 0.5 mg/l NAA

NS

3 mm

–

3% SA + MS medium

*100 mM

30

ST

3–5 mm

–

3% SA

100 mM

20–30

Vitis vinifera (grape) Eclipta alba (yerba de tago)

Das et al. (2002)

SEs

5–7 mm

–

2% SA

*100 mM

40–50

MS medium + 8.8 μM BA

ST, NS

–

–

3% SA

*1.11%

30–45

Vanda coerulea (orchid)

IY medium+20 g/l+TDZ (concentration not mentioned)+NAA (concentration not mentioned ) MS medium + 4.4 μM BA

PLB

3 mm

–

3% SA + IY medium + 20 g/l sucrose

100 mM

30

NA

NS

3–5 mm

–

3% SA + MS medium

100 mM

20–30

100%

½-MS medium + 2% sucrose

SE

NA

2.5% SA + ½-MS medium + 2% sucrose

43.3%

20

Zingiber officinale (ginger) Cucumis sativus (cucumber)

MS medium + 2.5 mg/l BA + 3% sucrose For embryogenesis, hypocotyl pieces cultured on MS medium + 2 μM 2,4-D + 0.5 μBA + 3% sucrose followed by cell suspension in liquid MS medium + 5 μM NAA + 1 μM BA

MSs

3–5 mm

For proper maturation cotyledonary stage SEs transferred to ½-MS medium + 10% sucrose for 2 weeks prior to encapsulation –

MS medium + 4.4 μM BA ½-MS medium + 2% sucrose

4% SA + MS medium

100 mM

30

66%

Cell suspension having SEs

NA

3% SA

*100 mM

30

Citrus sinensis × Poncirius trifoliate (carrizo citrange)

MS salts + vitamin mixture + 30 g/l sucrose + 1 mg/l NAA + 10 mg/l BA

NS

3–4 mm

2.5% SA + ½-MS medium + 50 g/l sucrose

*1.1%

35

MS medium without PGR

100% (sprouting) Only 17% of rooting after encapsulation

Germanà et al. (2011)

Khaya senegalensis (mahogany)

MS medium + 3% sucrose + 4.4 μM BA

ST, NS

4–6 mm

3% SA + MS medium + 4.4 μM BA + 3% sucrose

⁎100 mM

30

½-MS medium + 3% sucrose + 4.4 μM BA

52-98%

Hung and Trueman (2011)

Picrorhiza kurrooa (katuka)

NA

ST, NS

3–5 mm

For synchronization, all the cell passed through 150 μM sieve after every 10 day of subculturing and then incubated on gyratory shaker, shifted to modified suspension media containing CH+ NAA for 2–3 days. Finally maturation was done on MS basal medium. Micro-cuttings were immersed for 3 days in the dark and kept at 23 ± 1 °C in 50 ml glass jars (10 micro-cuttings per jar) containing 15 ml of rooting solution (5 mg/l IBA and 15 g/l sucrose (root induction pretreatment) Excised shoot tips were pre-treated by culturing on agar-solidified 1/2-MS medium containing 245 μM IBA + 2% sucrose for 24 h in darkness –

MS medium + 3% sucrose Neutral gel formed by dissolving phytagel in autoclaved tap water

3% SA + ½-MS medium + 1.5% sucrose

*3%

30

MS basal medium

Mishra et al. (2011)

Flickingeria nodosa (orchid)

Burgeff's N3F basal medium + 2% sucrose

PLBs

NA

–

2% SA + Burgeff's N3F medium + 2% sucrose

100 mM

30

21.43% (after 3 month of storage in a mist environment at 25 ± 2 °C) 95%

92.6% 66.6% (sprouting) and 70% (rooting) 36% 94.3% (after 4 week of storage) 94.9%

Ahmad and Anis (2010) Kumar et al. (2010)

Nirala et al. (2010) Ray and Bhattacharya (2010) Sarmah et al. (2010)

Singh et al. (2010) Singh and Chand (2010)

43.3%

57% (after 10 weeks of storage at 4 °C)

Sundararaj et al. (2010) Tabassum et al. (2010)

S. Sharma et al. / Biotechnology Advances 31 (2013) 186–207

Eclipta alba (yerba de tago) Dalbergia sissoo (sissoo)

MS medium + 2.5 μM Kn + 1.0 μM NAA MS medium (for sprouting) and MS medium + 1 mg/l IBA (for rooting) Quarter strength B5 macrosalts. MS medium + 3% sucrose

% Conversion References (complete plantlet recovery in vitro)

Nagananda et al. (2011) 195

(continued on next page)

196

Table 1 (continued) Plant species (common name)

Proliferation medium of the mother culture before synseed production

Propagule Pretreatment or Propagule size after treatment used for (as required) encapsulation

Encapsulating matrix composition

Concentration Polymerization Composition of of CaCl2·2H2O time (min) regrowth medium for or ⁎CaCl2 synseed

212– 300 μm

–

2% SA

*15 g/l

30

NS

3–5 mm

–

3% SA + WPM

100 mM

30

ST, NS

3–5 mm

⁎100 mM

15–20

NS

3–5 mm

Synseed were treated with 3% SA + MS medium 200 mM KNO3 for 5 min followed by rising with sterile DDW and 1.0 mg/l GA for 2 min (self-breaking treatment) – 3% SA + WPM

100 mM

30

MS medium + 1.5 mg/l Aranda Wan Chark Kuan TDZ ‘Blue’ × Vanda coerulea Grifft. Ex. Lindl. (orchid)

PLBs

4 mm

–

3% SA + ½-MS medium 75 mM

Corymbia torellina × MS medium + 3% C. citriodora (eucalyptus) sucrose + 2.2 μM BA

ST, NS

4–6 mm

Pretreatment with 19.6 μM IBA

3% SA + MS medium + 2.2 μM BA

⁎100 mM

SE MS medium + 2 mg/l BA + 0.2 mg/l NAA + 3 mg/l ABA or 4% sucrose MS medium + 0.5 μM TDZ NS

3–4 mm

–

3–5 mm

–

MS medium + 4.52 μM 2,4-D

SE

NA

-

4% SA + MS medium + 3% sucrose + 1 mg/l BA + 0.2 mg/l NAA 5% SA + MS medium + 0.5 μM TDZ + 2.5 μM IBA + 0.3–0.55 PPM 2.5% SA + 3% sucrose

In vitro raised seedlings

NS

3–4 mm

–

4% SA + MS medium

Rauvolfia tetraphylla (be still tree) Stevia rebaudiana (stevia)

Rauvolfia serpentina (Indian snakeroot or Sarpagandha)

Clitoria ternatea (butterfly pea)

Cannabis sativa (marijuana)

Catharanthus roseus (Madagascar periwinkle) Decalepis hamiltoni i (swallow root)

Liquid MS medium + 2 mg/l Kn + 1 mg/l IBA WPM + 7.5 μM BA + 2.5 μM NAA NA

WPM + 5.0 μM BA + 1.0 μM NAA

97%

Rihan et al. (2011)

90.3%

Alatar and Faisal (2012) Ali et al. (2012)

100%

WPM + 5.0 μM BA + 1.0 μM NAA (for sprouting) and ½-liquid MS medium + 0.5 μM IAA MS medium

80% (after 4 weeks of storage at 4 °C)

Faisal et al. (2012)

96.2%

Gantait et al. (2012)

30

MS medium + 3% sucrose + 2.2 μM BA

62–100%

100 mM

45

92%

50 mM

30

MS medium supplemented with 2 mg/l BA + 0.5 mg/l NAA MS medium + 0.5 μM TDZ

Hung and Trueman (2012a, 2012b) Kumar and Thomas (2012)

100 mM

15

MS medium + 1.34 μM NAA + 1.1 μM BA

84.33%

Maqsood et al. (2012)

100 mM

20–25

MS medium + 5 μM BA + 0.5 μM IAA + 30 μM ADS

77%

Sharma and Shahzad (2012)

20

Lata et al. 60% (after (2012) 24 weeks of storage at 15 °C)

Abbreviations: ABA—abscisic acid, AC—activated charcoal, ADS—adenine sulphate, Anderson medium (Anderson, 1975), B5—Gamborg medium (Gamborg et al., 1968), BA—6-benzyladenine, DCR—Gupta and Durzan basal medium (Gupta and Durzan, 1985), DKW medium—Driver and Kuniyuki medium (Driver and Kuniyuki, 1984), 2,4-D—2, 4-dichlorophenoxyacetic acid, GA3—gibberellic acid, Glu—glutamic acid, Heller's medium—Heller (1953), IBA—indole-3-butyric acid, 2-iP—(Δ2-isopentenyl) adenine, IY medium—Ichihashi and Yamashita's medium (Ichihashi and Yamashita, 1977), Kn—kinetin, LPm medium—von Arnold and Eriksson (1981), LSBM—Linsmaier and Skoog's basal medium (Linsmaier and Skoog, 1965), MC—microcutting, MP—microplantlet, MS—microshoot, MS medium—Murashige and Skoog's medium (Murashige and Skoog, 1962), NAA—α-naphthalene acetic acid, NA—information not available, Nitsch medium—Nitsch (1951), NS—nodal segment, OMM—olive modified medium (Mencuccini et al., 1997), P24 medium—(Teasdale, 1992), PC—protocorm, PG—phloroglucinol, PGR—plant growth regulator, PPM—Plant Preservative Mixture™, PVP—polyvinylpyrrolidone, QL—Quoirin and Lepoivre medium (Quoirin and Lepoivre, 1977), R4 medium—Chaleff and Stolarz (1981), SA—sodium alginate, SB—shoot bud, SE—somatic embryo, ST—shoot tip, TDZ—thidiazuron, TIBA—2,3,5-triidobenzene acid, W medium—White's medium (White, 1943) medium, WPM—woody plant medium (Lloyd and McCown, 1980), Zea—zeatin.

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MS

Brassica oleracea var botrytis (cauliflower)

Burgeff's N3F medium + 2 mg/l AdSO4 + 1 mg/l NAA Perlite + MS medium + 2 mg/l Kn and 2 mg/l NAA WPM + 7.5 μM BA + 2.5 μM NAA MS medium + 1.0 mg/l BA

References % Conversion (complete plantlet recovery in vitro)

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Gelation from “outside to inside” Polyelectrolyte solution (e.g. Na-alginate) + Somatic embryo

Cross-linking solution (CaCl2)

Ca++

Ca++

Ca++

Ca++

Inotropic gel (Ca-alginate)

Fig. 2. Simple bead (Single layered synseed) formation [redrawn from the report of Patel et al. (2000) with permission].

material, the requirements of definite ingredients of the hydrogel matrix (viz., inorganic nutrients, organic nutrients, plant growth regulators (PGRs), carbon sources, etc.) are species-specific. Preparation of a gel matrix in nutrient medium improves the re-growth of encapsulated plant tissue (Ahmad and Anis, 2010; Chand and Singh, 2004; Sundararaj et al., 2010). Various PGRs and growth additives were also added to the gel matrix to further enhance synseed conversion. In Adhatoda vasica (vasaka), a comparison was made between the additive effect of PGRs in gel matrix and conversion media (Anand and Bansal, 2002). They reported that a synseed prepared in Gamborg's medium (B5, Gamborg et al., 1968) supplemented with kinetin (Kn, 4.65 μM) and phloroglucinol (PG, 50.0 mg/l) resulted in highest synseed conversion (66.28%) when inoculated on basal B5 medium. However, such beads showed reduced conversion frequency when planted on B5 medium supplemented with a similar combination of Kn and PG as added to the gel matrix. Similarly, seeds prepared in a B5 matrix exhibited reduced conversion frequency (54.58%) on B5 medium supplemented with Kn (4.65 μM) and PG (50.0 mg/l). Thus, their study revealed that the presence of nutrients and PGRs in the gel matrix was more crucial than in the sowing medium for A. vasica synseed. Tsvetkov et al. (2006) reported that 3% sucrose in the gel matrix had a more pronounced effect on synseed conversion than PGRs in hybrid aspen (Populus tremula × P. tremuloides). Synseeds are reported to be highly susceptible to bacterial, fungal and other microbial infections in the greenhouse (Vij and Kaur, 1994). To reduce microbial contamination, various antimicrobial agents such as bavistin (Pattnaik et al., 1995), vitrofural G-1 (Nieves et al., 2003), plant preservative medium (PPM) (Micheli et al., 2002) can be added to the gel matrix. However, such chemicals impair convertibility which can be successfully alleviated by adding PGRs to the gel matrix. When activated charcoal (AC) is incorporated into synseeds, it also improves the conversion and vigor of the encapsulated propagules of tropical forest trees. AC not only stimulates the diffusion of gases and nutrients, but also helps in breaking down alginate which in turn facilitates enhanced respiration of propagules, thus preventing the loss of vigor that extends the storage period significantly (Saiprasad, 2001). Furthermore, AC absorbs unwanted exudates such as 5-hydroxymethylfurfural (a toxic breakdown product of sucrose formed during autoclaving) and other harmful phenolic products (Wang et al., 2007). In addition, AC retains nutrients within the hydrogel capsule and releases them slowly, thus

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providing an uninterrupted, long-term supply of nutrients to the growing tissue. Inclusion of 1.25% AC also improved the conversion frequency of encapsulated somatic embryos in Oryza sativa (rice) (Arun Kumar et al., 2005). Besides this, natural PGRs like coconut water and tomato juice have also been successfully used for synseed conversion. Being inexpensive, natural additives provide a cheaper substitution for expensive, sometimes synthetic, PGRs (Swamy et al., 2009). Another mechanical problem in encapsulation technology is the hindered emergence of the root or shoot of the encapsulated propagule by the gel capsule, although adopting self-breaking alginate gel bead technology could overcome this problem (Onishi et al., 1994). In this technology, synseeds are pretreated with potassium nitrate (KNO3). This self-breaking synseed technology was adapted to Feijoa sellowiana (goiabeira serrana), O. sativa (hybrid rice) and Stevia rebaudiana (stevia) (Ali et al., 2012; Arun Kumar et al., 2005; Guerra et al., 2001). O. sativa synseeds with a synthetic endosperm were applied a self-breaking treatment by dipping them in 200 mM KNO3 solution for 60 min and rising in sterile tap water for 40 min or more until the beads became swollen (Arun Kumar et al., 2005) while F. sellowiana and S. rebaudiana synseeds were dipped in 100 mM and 200 mM KNO3 solution for 20 and 5 min, respectively (Ali et al., 2012; Guerra et al., 2001). During pretreatment with KNO3, the K + ions replace the Ca2+ ions of the calcium alginate capsule thus allowing the synseeds to soften and open the subsequent conversion to plantlets (Onishi et al., 1994). In F. sellowiana, Guerra et al. (2001) reported 81.2% opening of the capsule vs. 0% in the treatment with water while in O. sativa, Arun Kumar et al. (2005) observed a 47% conversion frequency relative to normal synseeds (without a self-breaking coat). Hydrated synseeds are sticky and difficult to handle on a large scale and dry rapidly in the open air. This problem can be resolved by coating the beads with Elvax 4260 (ethylene vinyl acetate acrylic acid terpolymer; Dupont, USA) (Redenbaugh and Walker, 1990). 3.2. Double-layered synseed To prepare double-layered synseed, a single-layered bead is further coated with a similar concentration of sodium alginate solution and dropped into calcium chloride solution for 30 min then washed with sterile DDW. These double-encapsulated synseeds have all the merits of a single-layered synseed, with the added advantage of double encapsulation for better protection. Pinker and Abdel-Rahman (2005) tested the feasibility of forming a second layer in Dendranthema × grandiflora (chrysanthemum) synseeds. They observed that if the second layer was prepared in MS medium, the infection rate was higher, while the second layer prepared by Ca-alginate dissolved in water or mannitol reduced contamination considerably. Similarly, double-layered synseeds were prepared for Malus pumila (M.26 apple rootstock) using in vitro-derived apical buds (Micheli et al., 2002). 3.3. Hollow beads In conventional single-layered synseeds, propagules are usually located near the surface of beads which does not ensure their complete protection in contrast to a natural seed where the seed coat provides better protection. Hollow beads seem to be a promising tool in the realization of true synseeds by providing complete protection to the propagule. Hollow beads have various advantages over singleand double-layered synseeds, although the process of hollow bead formation is labor-intensive and costlier (Table 2). Considering various potentials of hollow beads over single- and double-layered synseeds, Patel et al. (2000) adopted this approach for the encapsulation of embryogenic calli of D. carota (carrot), as well as calli and shoot tips of Solanum tuberosum (potato). In this technique, propagules were suspended into a solution containing carboxymethyl cellulose (CMC) and calcium chloride and then

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Table 2 Advantages of hollow bead encapsulation [modified from Patel et al. (2000)]. Advantages over natural seeds

Advantages over naked somatic embryos

Advantages over encapsulated somatic embryos

Rapid clonal propagation

Easy handling (storage, sowing)

Economical propagation of plants which cannot be easily propagated via natural seeds (i.e. hybrids, transgenic plants, trees) Economic production of virus-free plants Easy maintenance of genetic resources

Protection from adverse environmental conditions (temperature, dryness)

Closer to natural seeds because of complete protection of somatic embryos in a sterile core Continued development of somatic embryo within the hollow bead

Protection against pathogens Longer shelf life

No early emergence Entrapment and coating realized in one step

dripped into a constantly-stirred sodium alginate solution (Fig. 3). In initial experiments with carrot, it was found that after 14 days of culture, 100% of hollow beads encapsulated in calcium alginate converted in the liquid core, 13% of which protruded out by bursting from the capsule. Embryogenic callus, encapsulated inside hollow beads, induced somatic embryogenesis while callus encapsulated in conventional calcium alginate beads detached from the beads early in development and no somatic embryogenesis occurred in any of the media. With potato, callus developed in 50% of hollow beads, whereas 81% of shoot tips encapsulated in hollow beads sprouted and grew out of the capsules. In contrast to this, in cyclamen (Cyclamen persicum), lower conversion of somatic embryos was reported from hollow beads than from conventional alginate beads (Winkelmann et al., 2004). This approach was not found to be suitable also for encapsulation of moss bud (S. ampullaceum) (Pourjavadi et al., 2006); regeneration was not satisfactory, for the following reasons: a) the strong net developed by the highly viscose alginate impeded a steady exchange of nutrients with the growth medium. That could be avoided by mechanically breaking the beads with tweezers, as is often done. b) Excessive uptake of water by beads during cold storage (5± 1 °C) led to a loss of sphericity, as well as loss of insulation and subsequent damage to the plant material in a relatively short period of time. These occurrences may have been due to the inclusion of CMC in the bead, which has a high capacity to absorb water. This may have led to the opening-up of large pores in the bead matrix wall and to the loss of protection of the plant material. 4. Propagules used for synseed production A synseed acts as an alternative seed for those species in which zygotic seeds are either not produced or not viable. A variety of Gelation from “inside to outside” Crosslinking solution (e.g. CaCl2) + Polymer + somatic embryo Polymer solution (Na-alginate)

Core (sol or gel) + somatic embryo

micropropagules have been successfully utilized for synseed production. These encapsulating units can be categorized into two types, as described next: 4.1. Bipolar propagules A somatic embryo is regarded as a bipolar structure as it possesses shoot and root poles at the same time (Standardi and Piccioni, 1998). Among various propagules, somatic embryos have been found to be the best suited entity for synseed production, and being bipolar in nature (each with a radicle and plumule), they are able to develop roots and shoots in a single step. Considering the advantages of somatic embryos over other propagules, these have been successfully exploited for synseed production in a number of plant species: C. persicum (Jalali et al., 2012; Winkelmann et al., 2004), Arnebia euchroma (Manjkhola et al., 2005), Rotula aquatica (Chithra et al., 2005), P. guajava (Rai et al., 2008a), Spartina alterniflora (Utomo et al., 2008), O. sativa (Roy and Mandal, 2008), Nothofagus alpina (Cartes et al., 2009), Vitis vinifera (Nirala et al., 2010), Catharanthus roseus (Maqsood et al., 2012). Somatic embryos have been utilized in both dried and hydrated state with encapsulation for synseed production. The process used for desiccating somatic embryos has already been described in subsection 2.1. The basic problem in the exploitation of somatic embryos for synseed production is an asynchronous and late maturation of the embryonic pole (Castellanos et al., 2004). Maturation has been hampered in many woody plant species due to precocious conversion, spontaneous repetitive embryogenesis, and embryo dormancy (Teixeira da Silva and Malabadi, in press). Thus, a well-tuned embryogenic system is required to improve plantlet conversion from synseeds. To address this need, various osmoticants have been used to enhance embryo maturation in cork oak (Garcia-Martin et al., 2001, 2005). Embryo weight increment is an indicator of embryo quality and a pre-requisite for successful conversion (Garcia-Martin et al., 2001, 2005). Nowadays, mathematical models have been developed with the aim of monitoring the large-scale uniform production of microplants (Konan et al., 2006). Combining these two pre-requisites, a technique was suggested for monitoring the growth of cork oak somatic embryos inside Petri dishes with a standard system of image capture and a digital system of image analysis which simplified the problem of quantifying and comparing growth in different treatments and culture media (Pintos et al., 2008). This method permitted growth assessment without the risk of contamination and opened up the possibility of automated control of culture growth for scaling-up plant production. The practical use of somatic embryogenesis, in terms of synseeds, has not evolved accordingly. Since embryogenic ability is often found only in a few genotypes within a species or is achieved by only using zygotic embryos or juvenile plant material as initial explants, this limits its use for cloning selected superior varieties (Standardi and Piccioni, 1998). 4.2. Unipolar propagules

Ionotropic gel (Ca-alginate) Fig. 3. Hollow bead formation [redrawn from the report of Patel et al. (2000) with permission].

Plant propagules with only a shoot or root pole are referred as unipolar propagules. A new perspective in synseed technology was initiated with the use of non-embryogenic unipolar plant propagules

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2011), Brassica oleracea var botrytis (Rihan et al., 2011), Carrizo citrange (Germanà et al., 2011), K. senegalensis (Hung and Trueman, 2011), C. torelliana×C. citriodora (Hung and Trueman, 2012a,b) and Decalepis hamiltonii (Sharma and Shahzad, 2012). Gangopadhyay et al. (2009) developed an encapsulation-based antibiotic selection technique for encapsulated microshoots of pineapple var. Queen. In most reports, a 3–5 mm long single nodal segment or shoot tip explant, possessing one or two axillary/apical buds were used for synseed production (Ahmad and Anis, 2010; Germanà et al., 2011; Hung and Trueman, 2012a,b), although Rihan et al. (2011) excised a 0.2-0.3 mm size microshoot for synseed production in B. oleracea var botrytis (cauliflower) while Chand and Singh (2004) dissected 10-mm long nodal segments for synseed preparation in D. sissoo. A highly effective protocol was developed for shoot tip encapsulation in four clones (K61, K522, K584 and K686) of K. senegalensis, a medicinally important and timber-yielding plant (Hung and Trueman, 2011). The capsules were prepared in sodium alginate solution containing full-strength MS medium with 4.4 μM 6-benzyladenine (BA) and 3% sucrose followed by polymerization in calcium chloride. Shoot regrowth from capsules was evaluated on agar-solidified and liquid MS medium with or without 4.4 μM BA. Shoots emerged from 92 to 100% of capsules after 6 weeks of incubation. Only medium containing BA stimulated multiple shoots, which were longer than those from their corresponding agar-solidified or liquid hormone-free media in clones K522 and K686. Liquid medium with BA provided more shoots than agar-solidified medium in the presence of BA for clones K522 and K686. Shoots produced on BA-containing culture medium were sturdy and each excised shoot produced multiple shoots or a plantlet upon transfer to the same medium or an indole-3-butyric acid (IBA)-containing medium, respectively. Thus, culture media supplemented with BA was considered to be optimal for subsequent shoot proliferation from K. senegalensis-encapsulated explants. They

whose potential advantages are easy to understand. It would be possible to exploit the natural ability of the species to produce vegetative propagules or, when nothing else is available, to use easy-to-obtain somatic fragments of the plant directly, to reduce the difficulties and generalize the application of synseed technology to as many genotypes and species as possible. Also, the risks of somaclonal variation would be reduced—relative to somatic embryos—by using unipolar propagules (Standardi and Piccioni, 1998). The scientific literature reports several examples of unipolar propagules used as synseed. Standardi and Piccioni (1998) divided unipolar propagules into three groups according to the type of propagule: 4.2.1. Nodes with apical or axillary buds and microshoots These types of propagules, also referred to as microcuttings, are one of the most frequently considered for synseed production, probably because of the relative ease with which these explants are produced once the micropropagation system has been established (Piccioni and Standardi, 1995). Shoot tips and nodal segments have been successfully exploited for synseed development in a number of plant species as they assure a high degree of genetic stability without encountering somaclonal variations (Piccioni, 1997). Non-embryogenic plant propagules have been widely used in synseed technology for a range of plant species: Morus spp. (Pattnaik and Chand, 2000; Pattnaik et al., 1995), Eucalyptus grandis (Watt et al., 2000), A. vasica (Anand and Bansal, 2002), Dendranthema×grandiflora (Teixeira da Silva, 2003), Dalbergia sissoo (Chand and Singh, 2004), Ananas comosus (Gangopadhyay et al., 2005), C. grandiflora (Nishitha et al., 2006), Punica granatum (Naik and Chand, 2006), Gerbera jamesonii (Taha et al., 2009), Cineraria maritima (Srivastava et al., 2009), Musa sp. (Sandoval-Yugar et al., 2009), Cannabis sativa (Lata et al., 2009), S. mauritiana (Sharma et al., 2009a; Fig. 4), S. acmella (Sharma et al., 2009b), Z. officinale (Sundararaj et al., 2010), Vitex negundo (Ahmad and Anis, 2010), Picrorhiza kurrooa (Mishra et al.,

B

E A

C

D Fig. 4. Different stages of plantlet recovery from encapsulated nodal segments of Spilanthes mauritiana. (A) Encapsulated nodal segments in 3% sodium alginate and 100 mM calcium chloride. (B) Sprouted synseeds after 2 weeks on MS medium supplemented with BA (1.0 μM) and IAA (0.5 μM). (C) Complete plantlets (shoot and root development) after 4 weeks of culture. (D) An acclimatized plantlet. (E) A twig showing flowering.

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performed a similar synseed experiment with shoot tips and nodal segments of four highly proliferating clones (A80, C46, C48 and C79) of C. torelliana × C. citriodora, a eucalypt (Hung and Trueman, 2012a). They found higher growth frequencies (78–100% and 76–100% for encapsulated shoot tips and nodal segments, respectively) and length of regrowing shoots (2.7–5.7 and 2.5–6.0 nodes from encapsulated shoot tips and nodal segments, respectively) on PGR-free half- and full-strength MS media. However, mean number of shoots per capsule was higher (1.4–4.4 shoots/capsule) on full-strength MS media supplemented with 2.2 μM BA with or without 0.3 μM α-naphthalene acetic acid (NAA) relative to PGR-free re-growth media. Thus, the optimal medium for K. senegalensis and Corymbia shoot re-growth depended upon the purpose of the capsules. BA-free sowing media would be preferable when a germplasm collection of a few capsules from many clones was transferred between laboratories and where it would be important to maximize the probability of at least one shoot emerging from each clone. Sowing medium containing BA may be preferable when many capsules of only a few clones are transferred, although rapid shoot proliferation is required from each clone. In contrast to this, maximum shoot development (2–5 shoots/bud) from encapsulated axillary buds of Morus species (mulberry) was possible on MS medium containing 4.4 μM BA (Pattnaik and Chand, 2000). Among six species of Morus (M. alba, M. australis, M. cathyana, M. nigra and M. bombycis), one step regeneration i.e., both shoot and root formation was recorded in M. alba, M. bombycis and M. latifolia although the shoots that were recovered from encapsulated buds of M. australis, M. cathyana and M. nigra failed to root on re-growth media and needed an additional in vitro root induction step. Naik and Chand (2006) reported encapsulation of nodal segments of P. granatum (pomegranate) for germplasm distribution and exchange. Two types of explant source i.e., source A and source B were used for collection of nodal segments followed by hydrogel encapsulation. Source A was established using mature nodal segments while source B was derived from cotyledonary nodes. In both cases, encapsulated nodal segments sprouted best on MS medium supplemented with 100 mg/l myo-inositol, 4.44 μM BA and 0.54 μM NAA. A maximum of 68% (source A) and 81% (source B) sprouting was noticed with this combination of PGRs and 2–3 shoots were recovered from each encapsulated nodal segment. However, a one-step conversion, i.e. simultaneous shoot and root formation, was possible only with encapsulated nodal segments of source B. About 21% of the encapsulated nodal segments cultured on PGR-free full-strength MS medium supplemented with 100 mg/l myo-inositol showed conversion within 40 days which was the maximum among planting media tested while no rooting was noticed with source A on any of the planting media tested. Thus, similar to Morus species, a separate experiment was needed for efficient root induction of capsulederived P. granatum microshoots (Naik and Chand, 2006). Although nodal segments are the most suitable (among various unipolar propagules) for encapsulation studies as they possess a preexisting axillary meristem (mostly two axillary buds/node relative to single apical buds in shoot tip explants), three sets of problems can be highlighted. The first is related to the nature of the organ used, i.e., a vegetative plant segment, devoid of any storage tissues and strongly adapted to in vitro culture conditions. The second is related to the lack of a root apex and the inability of explants to spontaneously form roots. The third is the overall additional cost of the micropropagation system. Among all, in vitro root induction is the major obstacle, especially encountered for recalcitrant woody plant species, although this can now be more easily overcome with CO2-enrichment in aerated vessels (see, for example, Teixeira da Silva et al., 2006). It is assumed that encapsulation inhibits oxygen supply to the propagule and suppresses root induction (Piccioni, 1997). However, in some species, such as C. grandiflora, Coleus forskohlii, C. maritima, Eclipta alba and P. kurrooa, encapsulated nodal segments with apical or axillary buds demonstrated a high adventitious rooting capacity after sowing on PGR-free nutrient

media (Mishra et al., 2011; Nishitha et al., 2006; Ray and Bhattacharya, 2010; Srivastava et al., 2009; Swaroopa et al., 2007). On the other hand, in many other species, an appropriate root induction protocol was suggested to be integrated with the encapsulation protocols. In the literature, the following methods listed next have been tested for root induction of synseed-derived propagules with varying degrees of success. 4.2.1.1. Pretreatment of explants prior to encapsulation. Piccioni (1997) suggested a method of incubating explants in the dark for inducing root primordia followed by the addition of PGRs in the gel matrix for higher conversion from encapsulated beads. Pretreatment of explants with cytokinin(s) and auxin(s) has also been used to enhance the conversion frequency of synseeds i.e., complete plantlet development (with a healthy shoot and root system) (Chand and Singh, 2004; Germanà et al., 2011; Pattnaik et al., 1995; Soneji et al., 2002) (Table 1). Recently, Hung and Trueman (2012a) reported that the conversion of C. torelliana × C. citriodora (four clones viz., A80, C46, C48 and C79) synseed depended almost entirely on IBA pretreatment, particularly for nodal segments which rarely (0–4%) produced roots without IBA. Pretreatment with 19.6 μM IBA consistently resulted in optimal conversion (62–100%) for all four Corymbia clones and two explant types (shoot tips and nodal segments). The use of 4.9 μM IBA afforded a high conversion from encapsulated shoot tips but in three of the four clones, this concentration resulted in a lower conversion (44–56%) from encapsulated nodes than when 19.6 μM IBA was used. Similarly, pretreatment with 245 μM IBA consistently provided one of the highest conversion percentages for each K. senegalensis clone (K61, K522, K584 and K686) (52–100%) (Hung and Trueman, 2011). This auxin level was much higher than 10.8 μM NAA plus 39.4 μM IBA (79.17% conversion) and 4.90 μM (85% conversion) used for A. comosus (Soneji et al., 2002) and D. sissoo (Chand and Singh, 2004), respectively. This was very close to the optimal concentration (260 μM IBA) used for rooting K. senegalensis microshoots (Danthu et al., 2003). The concentrations (4.9–39.4 μM IBA) used for A. comosus, D. sissoo and M. pumila synseeds resulted in similar conversion frequencies (58–100%) (Chand and Singh, 2004; Piccioni, 1997; Soneji et al., 2002). 4.2.1.2. Incorporation of PGRs to the gel matrix. The addition of PGRs to the gel matrix improves the efficiency of synthetic endosperm around the vegetative propagules and thus provides a simpler method for the successful recovery of complete plantlets. Pinker and Abdel-Rahman (2005) emphasized that the addition of 5.71 μM indole-3-acetic acid (IAA) to the gel matrix (prepared in modified MS) resulted in 100% root formation from encapsulated nodal segments of C. grandiflora. Nishitha et al. (2006) suggested the addition of 11.7 μM silver nitrate (AgNO3) along with 0.49 μM IBA to enhance the conversion frequency in synseeds of C. grandiflora. 4.2.1.3. Incorporation of PGRs to the regrowth media. Supplementation of PGRs to the germination medium has been found to eliminate the requirement of an additional in vitro root induction step prior to acclimatization. Ahmad and Anis (2010) found that the addition of 2.5 μM kinetin (Kn) and 1.0 μM NAA to MS basal medium induced a mean of 2.8 roots/shoot from encapsulated nodal segments of V. negundo. Sharma et al. (2009b) reported a maximum of 87.8% conversion frequency on MS medium supplemented with 1.0 μM BA and 0.5 μM NAA for S. acmella synseeds. For Tylophora indica synseeds, optimum conversion (91%) was achieved on MS medium supplemented with 2.5 μM BA and 0.5 μM NAA (Faisal and Anis, 2007). In Ocimum basilicum, maximum conversion frequency (80%) was obtained on half-strength MS medium supplemented with 5.0 μM BA and 0.5 μM IAA (Siddique and Anis, 2009). However, Alatar and Faisal (2012) reported 90.3% complete plantlet recovery when Rauvolfia tetraphylla synseeds were placed on woody plant medium (WPM, Lloyd and McCown, 1980) augmented with 7.5 μM BA and 2.5 μM NAA.

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4.2.1.4. Re-culturing of capsule derived microshoots on rooting media. Gangopadhyay et al. (2005) devised a two-step method to achieve maximum bead conversion in A. comosus (pineapple). In the first step, shoots were retrieved from encapsulated beads and in the second step, these microshoots were rooted in liquid medium (supplemented with 0.01 mM IBA and 0.002 mM Kn) supported with Luffa-sponge. In contrast, Pattnaik and Chand (2000) used half-strength MS medium supplemented with different concentrations of IAA, IBA and indole-3-propionic acid (IPA) for root initiation in capsule-derived microshoots of Morus species (mulberry). In M. australis and M. cathyana, optimum rooting was achieved on half-strength MS medium containing 5.7 μM IAA, 4.9 μM IBA and 5.3 μM IPA while that of M. nigra required only 4.9 μM IBA. In contrast, shoots of M. alba, M. bombycis, M. latifolia and M. nigra rooted best on half-strength MS medium augmented with 4.9 μM IBA. In P. granatum, rooting was possible by re-culturing the capsulederived microshoots on half-strength MS medium supplemented with 50 mg/l myo-inositol, 1.5% sucrose and NAA (0.054–5.37 μM) or IBA (0.049–4.9 μM). Among various treatments, maximum rooting efficiency (87–90%) was obtained with 0.54 μM NAA (Naik and Chand, 2006). In contrast, Singh (2008) reported successful rooting in capsule-derived microshoots of Rhododendron maddeni using liquid Anderson medium (Anderson, 1978) supplemented with 1.0% AC and 0.20 mg/l IBA. In contrast, Swamy et al. (2009) reported rooting on PGR-free half-strength MS basal medium for microshoots retrieved from encapsulated beads of P. cablin. Srivastava et al. (2009) reported healthy root formation in capsule-derived microshoots of C. maritima following a 2-week transfer to half-strength MS medium containing 1.0 mg/l NAA. 4.2.2. Microbulbs, microtubers, rhizomes and corms These explants are generally naturally ready to convert, since they are natural propagules that have been differentiated by evolution for vegetative multiplication of individuals and often also include storage tissues. Encapsulation of microbulbs, rhizomes and protocorms has been reported for a number of plant species (Datta et al., 1999; Martin, 2003; Saiprasad and Polisetty, 2003; Standardi and Piccioni, 1998). In Ipsea malabarica (Malabar daffodil orchid), encapsulated bulbs cultured either on PGR-free half-strength MS or 6.97 μM Kn-supplemented medium facilitated 100% conversion (Martin, 2003). Although conversion was independent from culture medium, the number of shoots developed varied. A maximum of 7.2 shoots/ capsule formed on half-strength MS medium supplemented with 6.97 μM Kn while 4.6 shoots/capsule were induced on PGR-free half-strength MS medium. Moreover, the growth of shoots on Kn-enriched medium was faster than that on medium devoid of Kn. Bekheet (2006) established a synseed protocol for the conservation of garlic by encapsulating in vitro-proliferated bulblets of garlic. Maximum shoot recovery (6.30), shoot length (6.00 cm) as well as shoot fresh mass (3.80 g) were obtained on MS medium containing 2 mg/l BA and 2 mg/l NAA. Synseed production in orchids is of special significance in view of the tiny and non-endospermic seeds they produce. Corrie and Tandon (1993) used protocorms for synseed production in Cymbidium giganteum and induced healthy plantlets upon transfer either to nutrient medium or directly to sterile sand and soil. The conversion frequency was comparable under both in vitro (100%) and in vivo (88% in sand, 64% in sand and soil mixture) conditions. These observations have made it feasible to transplant aseptically-grown protocorms directly to soil, and to cut the cost of raising in vitro plantlets and their subsequent acclimatization. Encapsulation of PLBs is well documented in many orchids such as C. giganteum, Dendrobium wardianum, Dendrobium densiflorum, Phaius tonkervillae, Oncidium, Cattleya and Spathoglottis plicata (Ara et al., 2000; Saiprasad and Polisetty, 2003; Vij et al., 2001). Saiprasad and Polisetty (2003) optimized the most suitable stage of PLBs for encapsulation in three

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orchid genera (Dendrobium, Oncidium and Cattleya). They found that fractionated PLBs, 13–15 days after culture, at the leaf primordia stage were most suitable for encapsulation. An encapsulation matrix prepared with MS medium (¾-strength) supplemented with 0.44 μM BA and 0.54 μM NAA gave 100% conversion of encapsulated PLBs when cultured on MS medium supplemented with 0.44 μM BA and 0.54 μM NAA (Dendrobium) or MS medium supplemented with 2.69 μM NAA (Oncidium) or MS medium supplemented with 5.38 μM NAA (Cattleya). Sarmah et al. (2010) produced synseeds in Vanda coerulea, an endangered monopodial orchid by encapsulating PLBs regenerated from the leaf base. A maximum of 94.9% conversion frequency was noticed when encapsulated PLBs were inoculated immediately on IY medium (Ichihashi and Yamashita, 1977). Nagananda et al. (2011) encapsulated the PLBs of Flickingeria nodosa and achieved 95% conversion on Burgeff's medium (Withner, 1955) supplemented with 2% sucrose, 2 mg/l adenine sulphate (AdSO4) and 1 mg/l IAA, after 3 months' storage at 4 °C. Recently, Gantait et al. (2012) reported alginate-encapsulation of Aranda × Vanda PLBs. Individual PLBs (4 mm long) were best encapsulated with 3% sodium alginate and 75 mM calcium chloride. The highest percentage conversion (96.4%) was obtained on PGR-free half strength-MS medium. 4.2.3. Meristemoids, cell aggregates and primordia These propagules are more complex to examine since they are not homogenous groups like microcuttings. In fact, differentiating propagules might eventually develop into one of the previous groups, i.e., evolve into buds or shoots (Yoshida, 1996), or form a natural propagule, such as a corm (Sharma et al., 1992; Tandon et al., 1994). The characteristic feature of the reports concerning this type of propagule is that the explants are encapsulated before the end of the differentiation phase, for example encapsulation of proliferating callus. Proliferating embryogenic callus clumps are also known to regenerate easily and show great potential for genetic transformation and can therefore be used for synseed production with promising results. Kim and Park (2002) reported a single-layer encapsulation of dehydrated embryogenic garlic callus using alginate gel. Encapsulated callus with a water loss of less than 50% showed 93% conversion frequency on half-strength MS medium. The microshoots that regenerated from encapsulated calli were much longer than those recovered from naked calli. Patel et al. (2000) successfully encapsulated the embryogenic callus of D. carota and S. tuberosum using a hollow bead approach. 5. Possible modes of synseed utilization Synseed technology has unraveled new vistas in the field of plant biotechnology. Synseeds can be employed in different ways for the management of plant germplasm and some modes of their utilization are described next. 5.1. In vitro plant production Encapsulation provides a convenient and reliable means of producing plants in vitro. Synseeds can be efficiently planted in vitro either on semi-solid culture medium or planting substrate (perlite, vermiculite, vermicompost, soilrite, soil, sand, gravel) for conversion into complete plantlets (Mandal et al., 2000; Pinker and AbdelRahman, 2005). Generally, nutrient-rich media are more effective than nutrient-deficient substrates for successful recovery of plantlets (Mandal et al., 2000), but in Dendranthema × grandiflora, synseeds germinated more efficiently on perlite moistened with MS medium than on solidified nutrient-rich medium (Pinker and Abdel-Rahman, 2005). The PGR requirement in nutrient medium varies from plant to plant. For instance, encapsulated axillary buds of Ocimum spp. exhibited maximum conversion (95–99%) on MS basal medium supplemented with 1.1 μM BA (O. americanum), 2.2 μM BA (O. gratissimum) or

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4.4 μM BA (O. basilicum and O. sanctum) (Mandal et al., 2000). However, for encapsulated embryos of O. sativa, MS medium supplemented with 1 mg/l BA, 1 mg/l Kn and 0.5 mg/l NAA was critical for synseed conversion (maximum = 87.5%) (Roy and Mandal, 2008). In contrast, encapsulated nodal segments of D. sissoo (Chand and Singh, 2004) and Z. officinale (Sundararaj et al., 2010) showed maximum conversion response on PGR-free half- and full-strength MS media, while Kavyashree et al. (2006) reported multiple shoot formation from synseeds of mulberry var. S54 on Linsmaier and Skoog (LS, Linsmaier and Skoog, 1965) medium augmented with 8.88 μM BA and 2 μM 2, 3, 5-tri iodo benzoic acid (TIBA). Gelling agent is also an important factor for in vitro synseed conversion. Generally, agar is used as a gelling agent for synseed conversion medium (Cameron, 2008). In contrast, Singh (2008) reported better conversion of synseed on phytagel compared to agar.

5.2. Direct sowing Sowing synseeds ex vitro provides a commercially important and cost-effective technique for direct recovery of plantlets. This technology will greatly impact the large-scale production of commercially important plant species at a reduced cost. Although a large number of plants can be produced in tissue culture through embryogenesis and/or direct or indirect organogenesis, in some cases, delivery is cumbersome. Direct sowing of synseeds to soil or other substrates helps to avoid the acclimatization procedure required for tissue culture-raised plantlets. Synseeds are also an ideal delivery system enabling ease and flexibility of handling and transport of propagules compared to voluminous parcels of seedlings or tissue-cultured plantlets. Conversion of synseeds on nutrient-free substrate is a prerequisite for sowing under non-sterile conditions. Mandal et al. (2000) suggested that the successful conversion of synseeds into plantlets on a simple planting substrate such as sand, soil, soilrite or vermi-compost is necessary for their use in commercial-scale propagation. Despite this, successful conversion of encapsulated tissues on various planting substrates has been reported only for a few plant species either in a controlled culture room environment or greenhouse conditions (Chand and Singh, 2004; Faisal and Anis, 2007; Kavyashree et al., 2006; Lata et al., 2009; Sharma et al., 2009b; Singh et al., 2006). Moderate conversion (40%) of encapsulated somatic embryos was found on sterile sand (Roy and Mandal, 2008). The major limiting factor reducing conversion is the low-nutrient availability. Therefore, it is necessary to build up a nutrient reservoir for the encapsulated plant tissue, either endogenously or exogenously. Singh et al. (2006), Faisal and Anis (2007) and Sharma et al. (2009b) supplied half- or quarter-strength MS nutrients to soilrite for effective in vivo conversion of synseeds and reported 62, 43 and 63% conversion in Phyllanthus amarus, T. indica and S. acmella, respectively. Similarly, Chand and Singh (2004) reported 45% conversion on peat moss moistened with half-strength MS medium, after 30 days of ex vitro sowing of Dalbergia sissoo synseeds. Kavyashree et al. (2006) exogenously supplied half-strength LS nutrients in horticultural grade soilrite mix (peat: perlite: vermiculite, 1:1:1 (v/v)) for ex vitro conversion (45.5%) of mulberry synseeds with healthy shoots and roots. After sowing for 30 days, they reported a maximum of 11.6 shoots/synseed with 7.7 roots/synseed on this planting substrate. The feasibility of encapsulated cauliflower microshoots in commercial substrates (compost, vermiculite, perlite and sand) irrigated with different solution mixtures including sterilized DDW, MS medium with or without different concentrations of Kn and NAA was evaluated by Rihan et al. (2011). The use of MS medium supplemented with 2 mg/l Kn and 2 mg/l NAA gave an optimal response with both perlite and compost. This study showed the high growth capacity of cauliflower synseed in commercial substrate which is considered to be a promising step for their direct use in vivo.

During direct sowing, contamination by microorganisms is one of the major hurdles for the commercialization of encapsulation technology. Nutrients, especially organic, released by the beads are responsible for severe contamination (Abdel-Rahman, 2003; Nhut et al., 2005). The addition of fungicide (carbendazim or bavistin and benomyl, 50–100 mg/l) to the alginate matrix has been suggested for M. indica (Bapat and Rao, 1993) to counter this problem to a certain extent. Germanà et al. (2007) added another fungicide, 100 mg/l thiophanate-methyl® to the gel matrix of Citrus reticulata synseeds and reported 96.7 and 100% viability on filter paper and perlite, respectively. However, Lata et al. (2009) used PPM in both the gel matrix and planting substrate used for C. sativa. They reported 100% conversion on potting mixture of fertilome with coco natural growth medium (1:1), moistened with full-strength MS medium supplemented with 3% sucrose and 0.5% PPM. Occasionally, fungicides can reduce the viability of encapsulated propagules, thus to restrict contamination without affecting an explant's viability while the use of double-layered synseeds was suggested by Sharma et al. (1994) and Pinker and Abdel-Rahman (2005) for Z. officinale and Dendranthema × grandiflora, respectively. When the second layer was formed by Ca-alginate with water or 0.2 M mannitol, contamination was reduced considerably and only 6.7% of the beads were contaminated (Pinker and Abdel-Rahman, 2005). These findings were similar to those of Kinoshita and Saito (1992), who also used two-layered beads (with the second layer containing water) for encapsulation of axillary buds from Betula papyrifera (white birch) cultures but observed a low level of rooting. Leakage of sucrose from beads to the substrate stimulated the proliferation of microorganisms and inhibited rooting. Rinsing away the sucrose by spraying distilled water onto the perlite enhanced rooting to 78% (Kinoshita and Saito, 1992). However, Nhut et al. (2005) developed a much easier method in which Cymbidium synseeds were coated with a solution of a fungicide, chitosan. Preece and West (2006) studied suitable conditions for ex vitro synseed conversion in Hibiscus moscheutos cultivars, ‘Lord Baltimore’ and ‘Southern Belle’. After pre-treating with light providing a photon flux of approximately 40 μmol m −2 s −1 for a 16-h photoperiod at 25 °C for 5 weeks, they planted synseeds in vermiculite and provided intermittent mist during the course of incubation, resulting in 100% germination. Rooting was best in vermiculite if the encapsulated nodal segments were planted 1 cm deep and without any covering as covering might block the penetration of light. Ex vitro conversion of synseeds has been accomplished in sterile (autoclaved) potting media (Chand and Singh, 2004; Faisal and Anis, 2007; Kavyashree et al., 2006; Lata et al., 2009; Rihan et al., 2011; Sharma et al., 2009b; Singh et al., 2006). Ex vitro conversion avoids the use of an in vitro culture passage, but practical application of this method is restricted by the high cost of sterilizing potting media. Both problems could be circumvented by pre-converting the synseeds in moistened Petri dishes prior to transferring to non-sterile potting media as reported in K. senegalensis (Hung and Trueman, 2011) and Corymbia spp. (Hung and Trueman, 2012a). For pre-conversion of synseeds, shoot tips and nodal segments were pre-treated with IBA (for induction of root primordia), and maintained under light (16-h photoperiod at 100 μmol m − 2 s − 1) in moistened Petri dishes for 4 weeks at 25 °C. Explants were pre-treated by culturing them on semi-solidified MS medium supplemented with IBA (245 μM for K. senegalensis and 19.6 μM for Corymbia spp.) and 2% sucrose for 24 h in darkness. For both plant species, organic compost was found to be most suitable for direct conversion of synseeds under non-aseptic conditions. For K. senegalensis, 42–86% conversion was noticed for encapsulated shoot tips while in Corymbia spp. 46–90% conversion from shoot tips and 32– 50% from nodal segments was recorded 4 weeks after sowing onto organic compost. Pre-conversion may enhance plantlet development in non-sterile soils because of two factors. Firstly, microbial contamination may be minimized due to a reduction in nutrient content of the synseed matrix during pre-conversion and secondly, the shoots may be pre-

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adapted to ex vitro conditions because of pre-acclimatization during the pre-conversion period. Pre-conversion provides a simple and effective technique for synseed distribution and direct transfer to nursery conditions.

availability can be a limiting factor for the conversion ability of Actinidia deliciosa (kiwi-fruit) synseeds.

5.3. Short-term germplasm conservation

Biochemical and physiological mechanisms of low temperature tolerance in higher plants have been intensively studied, leading to a comprehension of the mechanisms required for plant adaptation to low temperatures. The attempt to conserve the plant propagules under ultra low temperature for a long duration is termed cryopreservation (Walter et al., 2006). Cryopreservation of biological tissues is useful only if intracellular ice crystal formation is avoided as it causes irreversible damage to cell membranes, thus destroying their semi-permeability. To date, various strategies have been employed for cryopreservation of different plant tissues i.e., the two-step freezing, simple dessication, encapsulation-dehydration, vitrification and encapsulation-vitrification. The encapsulation-dehydration strategy originally developed by Fabre and Dereuddre (1990) for Solanum shoot tips is more convenient than others. It also avoids the use of a costly programmable freezer and high concentrations of harmful cryoprotectants (Reinhoud et al., 2000). This strategy is based on a successive osmotic and evaporative dehydration of plant cells which allows gradual extraction of water from encapsulated propagules in sucrose-rich medium. Sucrose concentration in the beads is gradually increased by additional air-drying or desiccation in a laminar air flow cabinet to reach the saturation point which results in a glass transition during cooling to −196 °C in LN, thus preventing ice crystal formation (a cause of lethal damage to living cells) during exposure to ultra low temperature (Engelmann and Takagi, 2000). Encapsulation-dehydration has been used for cryopreservation of shoot tips in different genera of hardwood species including Malus, Pyrus and Prunus. It is noteworthy that for 50% of the species cryopreserved through this procedure, a survival rate of 80% or more was achieved (Lambardi and De Carlo, 2003). Encapsulation-dehydration and vitrification are simple and inexpensive techniques and maintain genetic stability while encapsulationvitrification is a hybrid of these two techniques that minimizes any potential injury from vitrification (Moges et al., 2004) and offers various advantages over encapsulation-dehydration in terms of greater recovery (Wang et al., 2004). Embryogenic callus of Dioscorea bulbifera (Yin and Hong, 2010) and PLBs of Dendrobium candidum (Yin and Hong, 2009) were successfully cryopreserved using encapsulation-vitrification.

Synseed technology also acts as a tool for germplasm exchange between countries. For this purpose, synseed storage is a critical factor that determines their successful conversion after transportation abroad. Therefore, appropriate storage conditions and a definite storage period are prerequisites to maintain synseed viability during transportation that leads to successful commercialization of synseed technology. Generally, 4 °C has been found to be most suitable for synseed storage (Faisal and Anis, 2007; Ikhlaq et al., 2010; Kavyashree et al., 2006; Pintos et al., 2008; Saiprasad and Polisetty, 2003; Sharma et al., 2009a,b; Singh et al., 2007; Tabassum et al., 2010). Gangopadhyay et al. (2005) stored encapsulated microshoots of A. comosus in different racks of a refrigerator over a range of temperatures (4, 8, 12 and 16 °C) for 60 days. Beads stored at 8 °C showed maximum conversion frequency on MS medium supplemented with 0.022 mM BA and 0.003 mM IAA. Few investigations revealed the requirement of higher temperature (25 °C) rather than low temperature for amenable storage of synseeds in certain tropical and sub-tropical crops. Sundararaj et al. (2010) observed 100% re-growth of Z. officinale synseeds incubated at 25 °C while no re-growth was observed for synseeds stored at 4 °C in the dark. Similarly, encapsulated microshoots of C. maritima (Srivastava et al., 2009) and P. kurrooa (Mishra et al., 2011) were successfully stored up to 6 and 3 months, respectively at 25 ± 2 °C in moist conditions. A few previous studies on preservation of encapsulated explants of eucalypt and mahogany have identified storage medium, irradiance or storage temperature as important factors in maintaining their regenerative potential. Encapsulated axillary buds of E. grandis were stored successfully in water for 6 months at 10 °C and 4 μmol m-2 s−1 irradiance but could only be stored for 2 months at 28 °C and 200 μmol m −2 s −1 irradiance (Watt et al., 2000). Encapsulated shoot tips and cotyledonary nodes of Cedrela fissilis were stored for 3–6 months in nutrient and sugar-free media at 25 °C and 20–25 μmol m -2 s -1 irradiance, with higher regrowth capacity of encapsulated shoot tips on 0.4% agar-solidified medium than on 1% agar-solidified medium (Nunes et al., 2003). For Cedrela odorata, 0–20, 0–40 and 30-80% of encapsulated shoot tips survived 12-month storage on 1% agar-solidified media at 20, 25 °C (both at 35 μmol m −2 s−1 irradiance) or 12 °C (in darkness), respectively (Maruyama et al., 1997). Eucalypt and mahogany species occur naturally in the tropics and subtropics and the temperatures used for storage of their encapsulated explants parallel temperatures typically used (10–25 °C) for maintaining in vitro cultures of tropical and subtropical species (Trueman, 2006; Withers, 1991). Encapsulated shoot tips of K. senegalensis survived longer at 25 °C than at 4 °C, with 73–88% viability after 8 weeks (Hung and Trueman, 2011). They noticed enhanced regrowth frequency of encapsulated K. senegalensis explants up to 12 months when the storage conditions were 14 °C and darkness. Under these conditions, encapsulated Corymbia and Khaya explants were preserved most effectively on half-strength MS medium supplemented with 1% sucrose which provided very high frequencies of shoot regrowth (92–100% for Corymbia and 71–98% for Khaya) and excellent shoot development after 12 months' storage (Hung and Trueman, 2012b). Ray and Bhattacharya (2010) optimized the best storage environment for E. alba synseed by changing in vitro physicochemical conditions. They extended the duration of storage up to 12 weeks by decreasing sucrose concentration in an alginate matrix from 3 to 1 or 2%. Adriani et al. (2000) also analyzed the pronounced effect of sucrose on re-growth ability of synseed and suggested that sucrose

5.4. Cryopreservation: an effective approach for long-term germplasm storage

6. DNA marker technology in synseed experimentation Although synseeds have been widely utilized for micropropagation and conservation of various medicinal plant species (Anand and Bansal, 2002; Ara et al., 2000; Ikhlaq et al., 2010; Lata et al., 2009; Nor Asmah et al., 2011; Nyende et al., 2003; Ray and Bhattacharya, 2010; Singh et al., 2006), the genetic stability of synseed-derived plantlets has been less evaluated. In contrast, the assessment of genetic variability of cryopreserved materials via encapsulation/dehydration or vitrification methods has attracted more attention (Bekheet et al., 2007; Hirai and Sakai, 2000; Scocchi et al., 2004). The increasing utilization of synseed for germplasm conservation and propagation necessitates the assessment of genetic stability of conserved propagules following their conservation (Dehmer, 2005). In recent years, systematic sampling of germplasm and analysis of their molecular status through the use of DNA marker technology has become common practice. Among different DNA polymorphism detection techniques, random amplified polymorphic DNA (RAPD) and inter simple sequence repeats (ISSR) analysis has widely been used to study clonal integrity, detect genetic and somaclonal variations (Agnihotri et al., 2009; Borner, 2006; Jokipii et al., 2004; Mandal et al., 2007) because of their simple genotyping, easy handling, cost effectiveness, and wide availability. Bekheet (2006), Srivastava et al. (2009) and Mishra et al. (2011) used RAPD

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analysis to ascertain the genetic fidelity of synseed/capsule-derived plantlets of Allium sativum (garlic), C. maritima and P. kurrooa, respectively. Srivastava et al. (2009) used 20 RAPD markers for this purpose. Of 20 primers tested, 14 produced amplification products, and a total of 69 bands with an average of 4.93 bands/primer were observed. Of these 69 scorable bands, only 20% of bands were polymorphic. Cluster analysis of the RAPD profiles revealed an average similarity coefficient of 0.944 thus confirming the molecular stability of C. maritima plants derived from encapsulated microshoots following 6 months of storage. Mishra et al. (2011) used 45 RAPD markers to ascertain the genetic fidelity of P. kurrooa plants growing after storage in encapsulated form. Of 45 primers tested, 14 produced scorable amplified products. A total of 68 bands were observed, 7.35% of which were polymorphic. They reaveled an average coefficient of 0.966 thus confirming genetic stability of plants derived from encapsulated microshoots following 3 months of storage. However, Gangopadhyay et al. (2005), Alatar and Faisal (2012) and Faisal et al. (2012) used RAPD and ISSR to assess the genetic fidelity of A. comosus (pineapple), R. tetraphylla and Rauvolfia serpentina plantlets, respectively growing after storage in encapsulated form. Gangopadhyay et al. (2005) used 10 RAPD and 3 ISSR primers and observed identical profiles in all the samples of A. comosus. Alatar and Faisal (2012) used 20 RAPD and 5 ISSR primers for R. tetraphylla. Out of 20 primers, 17 yielded clear and reproducible bands. The number of bands varied from 1 to 13. All the ISSR primers tested gave clear and scorable bands. The number of bands for each primer ranged from 11 to 15. In contrast to these studies, Tabassum et al. (2010) employed amplified fragment length polymorphism (AFLP) analysis to detect the level of variations among synseed derived regenerants of Cucumis sativus (cucumber). For this study, they used five EcoRI + MSeI primer combinations. Using five primer combinations, a total of 109 bands were scored (an average of 21.8 primers or 35.77%). The number of bands scored with each primer ranged between 13 and 28. The polymorphism ranged between 7.14% with primer E-AA + M-CTC to 15.38% with primer E-AG + M-CTC. The percentage polymorphism obtained was almost negligible, depiciting genetic integrity among regenerants.

non-sterile conditions, so as to make this technique feasible and practical for farmers and producers, is still a limitation (Jung et al., 2004). Several factors are involved in this process such as poor survival, which might be attributed to the lack of nutrients and oxygen supply, but primarily the protection of synseed beads from attack by microorganisms (Bapat and Rao, 1993; Nhut et al., 2005). Further refinement of existing protocols and correct formulation of the gel matrix would definitely improve ex vitro germination and enhance synseed germination, although this is species-specific and ideal conditions need to be established on a case-by-case basis. Loss of tissue viability after short- to long-term storage and occurrence of somaclonal variations are other frequently encountered limitations of synseed technology (Rai et al., 2009). One of the future applications of synseeds would be in germplasm conservation through cryopreservation, thus further experimentation is needed by using either hydrated calcium alginatebased or desiccated polyoxyethylene glycol-based encapsulation (Ara et al., 2000). Similar to all micropropagation techniques, conventional alginate encapsulation is a very labor-intensive process. Each explant is handled multiple times, including excision, cutting to the appropriate size, coating with sodium alginate, dipping in calcium chloride, washing in water and finally placement in a vessel for use or storage. In this situation, bulk encapsulation has evolved as an efficient technique with reduced labor (West and Preece, 2009). However, there are several problematic issues related to bulk alginate encapsulation including exposure of explants as a result of matrix shrinkage and reduced shoot and root growth due to high sodium alginate concentration (West and Preece, 2009). These problems need to be resolved in future research for proper utilization of this technique in long-term conservation practices. Aside from the cost of developing encapsulating units, additional investments are necessary to develop methods and machinery for handling synseed, both during production and sowing. Although little progress has been made to demonstrate the feasibility of synseeds, their successful implementation can only be accomplished on a small scale while commercial use is still more of a concept than a reality.

7. Problems, limitations and future prospects

Anwar Shahzad gratefully acknowledges the financial support provided by the Council of Science and Technology, Uttar Pradesh (Project no. CST/D3836), UGC (Project no. 39-369/2010) and DST-FIST Programme 2005 (Project no. SR/FST/LSI-085/2005). Shiwali Sharma is thankful to UGC, for the award of BSR Fellowship in Science (1st April 2010) for providing research assistance.

Synseed technology has drawn tremendous attention in recent years because of its wide application in germplasm conservation and exchange between countries. Despite various achievements, several major problems still need to be resolved for its commercialization. The first requirement for the practical utility of synseed technology is the large-scale production of high quality viable micropropagules in a cost-effective manner which remains a major limiting factor (Ara et al., 2000). Although somatic embryos are the best suited entity for synseed development, several bottlenecks such as loss of embryogenic potential with aging cultures, asynchronous development, precocious germination, structural anomalies and lack of desiccation tolerance restrict their utilization (Ara et al., 2000). Thus, a judicious coupling of synseed technology with mathematical modeling would allow automated encapsulation and regulation of plant development to take place (Pintos et al., 2008). This would tremendously increase the efficiency of encapsulation and generate homogenous high quality synseeds. Alternatively, the use of non-embryogenic propagules for synseed production holds great promise to those species which are recalcitrant to somatic embryogenesis. For most woody plant species, a single rooting step is still the major obstacle for non-embryogenic encapsulated beads and requires further research to improve their conversion ability (Chand and Singh, 2004; Hung and Trueman, 2012a; Naik and Chand, 2006; Piccioni, 1997; Soneji et al., 2002). Moreover, ex vitro or direct sowing of synseeds to soil under

Acknowledgment

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