Morpho-physiological disorders in in vitro culture of plants

Scientia Horticulturae 108 (2006) 105–120 www.elsevier.com/locate/scihorti

Review

Morpho-physiological disorders in in vitro culture of plants B.N. Hazarika * Biotechnology Laboratory, Division of Horticulture, ICAR Research Complex, Umiam 793 103, Meghalaya, India Received 9 September 2004; received in revised form 31 December 2005; accepted 31 January 2006

Abstract The special conditions during in vitro culture results in the formation of plantlets of abnormal morphology, anatomy and physiology. Tissue culture conditions that promote rapid growth and multiplication of shoots often results in the formation of structurally and physiologically abnormal plants. They are often characterized by poor photosynthetic efficiency, malfunctioning of stomata and a marked decrease in epicuticular wax. Qualitatively also, the waxes present on the surface of the leaves of in vitro cultured plants may vary. The conditions under which most laboratories done tissue culture is high relative humidity and low light, no supplemental CO2, high sucrose and nutrient containing medium may contribute to a phenotype that cannot survive the environmental conditions when directly placed in a greenhouse or field. Understanding these abnormalities is a prerequisite to develop efficient transplantation protocols. The present review summaries the major abnormalities in in vitro culture of plants and also highlight the current and developing methods that are satisfactory for acclimatization of in vitro cultured plantlets. # 2006 Elsevier B.V. All rights reserved. Keywords: Photosynthetic efficiency; Tissue culture; Stomatal function; Epicuticular wax; Anatomy; Culture-induced phenotype; Acclimatization

Contents 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

11.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Photosynthetic efficiency . . . . . . . . . . . . . . . . . . . . . . Stomatal functioning . . . . . . . . . . . . . . . . . . . . . . . . . The cuticle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anatomy. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hyperhydricity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Development of volatiles in vessel headspace . . . . . . . . The persistent leaves . . . . . . . . . . . . . . . . . . . . . . . . . The new leaves . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Strategies for acclimatization. . . . . . . . . . . . . . . . . . . . 10.1. In vitro acclimatization . . . . . . . . . . . . . . . . . . . 10.1.1. Abiotic approaches to acclimatization . . . 10.1.2. Biological approaches to acclimatization . 10.2. Ex vitro acclimatization . . . . . . . . . . . . . . . . . . 10.2.1. Environment control . . . . . . . . . . . . . . . 10.2.2. Antitranspirants . . . . . . . . . . . . . . . . . . 10.2.3. Ex vitro rooting and hardening . . . . . . . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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* Present address: Department of Horticulture, Assam Agricultural University, Jorhat 785 013, Assam, India. E-mail address: [email protected]. 0304-4238/$ – see front matter # 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.scienta.2006.01.038

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1. Introduction In vitro propagation has been extensively used for the rapid multiplication of many plant species. But the ultimate success of in vitro propagation on commercial scale depends on the ability to transfer plants out of culture on a large scale, at a low cost and with high survival rates. In vitro cultured plants are generally susceptible to transplantation shocks leading to high mortality during final stage of micropropagation. Plantlets or shoots that have grown in vitro have been continuously exposed to a unique microenvironment that has been selected to provide minimal stress and optimum conditions for plant multiplication. Plantlets developed within the culture vessels under low level of light, aseptic conditions, on a medium containing ample sugar and nutrients to allow for heterotrophic growth and in an atmosphere with high relative humidity. Due to these conditions, in vitro plantlets can develop certain characteristic features that are inconsistent with the development under greenhouse or field conditions. The heterotrophic mode of nutrition and poor mechanism to control water loss render micropropagated plants vulnerable to the transplantation shocks when directly placed in a greenhouse of field. Although some aspects of culture-induced phenotypes are known (Brainerd et al., 1981; Debergh and Maene, 1984; Donnelly et al., 1984; Donnelly and Vidaver, 1984c; Sutter, 1985; Fabbri et al., 1986; Dhawan and Bhojwani, 1987; Grout, 1988; Ziv and Ariel, 1992; Hazarika et al., 2001b, 2002b; Lamhanedi et al., 2003) but the transplantation stage continues to be a major bottleneck in the micropropagation of many plants. Understanding the physiological and morphological characteristics of tissue culture plants and the changes they undergo during the hardening process should facilitate the development of efficient transplantation protocols. This article discuss the major abnormalities of in vitro cultured plants that accounts for the fragility of cultured plants and reviews the methods used to harden the plants for transplantation to soil.

altogether The low RubPcase activity may be due to presence of sucrose during the development of leaves (Grout and Aston, 1977a; Wetzstein and Sommer, 1982; Donnelly and Vidaver, 1984a). The photosynthetic apparatus of regenerating cauliflower meristem culture is not sufficiently active to produce a net positive carbon balance in vitro. The chloroplasts have light stimulated electron transport comparable to control material but lower level of chlorophyll and RubPcase activity as shown in Fig. 1 resulting in correspondingly low carbon assimilation. This photosynthetic system does not develop further at transplanting as the in vitro foliage deteriorates rapidly, contributing little to net carbon uptake (Grout and Donkin, 1987). Studying cauliflower plantlets growing in vitro, Grout and Aston (1978) measured CO2 uptake using radiolabelled carbon and gas exchange using an infrared gas analyzer. They found negligible carbon dioxide uptake in the light while the plantlets were in vitro. A negative CO2 balance persisted up to 2 weeks after the plantlets had been transferred to soil in the greenhouse. The regenerated plantlets also had lower chlorophyll content than 4-week-old seedlings in a greenhouse (Grout and Aston, 1977b). Similar results were obtained with strawberry plantlets grown in vitro (Grout and Millam, 1985). After transplanting the strawberry to the greenhouse, most of the persistent leaves deteriorated rapidly and those that remained on the plants showed no increase in carbon fixation, indicating lack of development of photosynthetic competency. Birch plantlets regenerated in vitro had approximately one-

2. Photosynthetic efficiency High sucrose and salt containing media, low light level and the carbon dioxide concentration in the air in the culture vessel are some of the important limiting factors among various physical microenvironmental factors which influence photosynthesis of in vitro cultured plants (Fujiwara and Kozai, 1995; Jeong et al., 1995). For in vitro growth, a continuous supply of exogenous sucrose is required (2–3%) as a carbon source (Hazarika et al., 2000b, 2004; Hazarika, 2003a, 2003b; Wainwright and Scrace, 1989). High sucrose and salt containing media often employed for raising cultures and poor light conditions seems to restrict photosynthetic efficiency of leafy shoots. Although such plantlets may appear normal, they are unlikely to be actively photosynthesizing. This is because of the exogenous supply of sucrose, which does not necessitate the normal development of photosynthetic apparatus. Therefore in vitro cultured plants are either poor in chlorophyll content or the enzymes responsible for photosynthesis i.e. ribulose bisphosphate carboxylase (RubPcase) are inactive or absence

Fig. 1. Aspects of the photosynthetic system of cauliflower meristem cultures in vitro, compared to seedlings and plantlets established in soil (cultures 4 weeks after initiation, seedlings 2 weeks post-germination, transplants 4 weeks after transplanting—de novo foliage only) (Grout and Donkin, 1987).

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third the photosynthetic rate (expressed as mmol O2 mg 1 h 1 chlorophyll measured by a Clark-type oxygen monitor) compared to greenhouse-grown seedlings (Smith et al., 1986). The photosynthetic rate of cultured shoots declined over a 4-week passage in vitro. There was no change in net photosynthesis in shoot culture leaves over light levels ranging from 200 to 1200 mmol m 2 s 1. Greenhouse-grown leaves, serving as controls, showed almost a two-fold increase in net photosynthesis under the same light levels. Donnelly and Vidaver (1984b) measured a very low level of CO2 uptake in red raspberry leaves using an infracted gas analyzer. They found that the amount of CO2 uptake in cultured red raspberry plants was approximately 2.6 mg dm 2 h 1 CO2 compared with 10.5 mg dm 2 h 1 CO2 in field controls. During a 1-month period after removal from culture, CO2 uptake in persistent leaves remained the same or was in slight negative balance regardless of the light level under which the plantlets were grown. In Leucoena leucocephala, the leaves from cultured shoots and plantlets exhibited total lack of starch grains normally found in mesophyll cell of field-grown plants, thereby suggesting the photosynthetic inefficiency of the plants in cultures (Dhawan and Bhojwani, 1987). Studies by Kozai et al. (1987a) on nine different species of ornamental plants showed that the level of CO2 in the culture vessels decreased from a range of 3000–9000 ppm in the dark period to less than 90 ppm in the light period indicating that the plants had photosynthetic competency during the light period. The low level of CO2 in the vessel during the light period limited photosynthesis of the shoots and resulted in a net negative balance of CO2 uptake per day. The plantlets compensated for this by using the sucrose supplied in the medium. Increasing both CO2 and light level resulted in striking increases in growth of several plant species including orchids, carnation and statice both in culture tubes and in an acclimatization chamber. Differences in photosynthetic rates of various species in vitro may be attributed to inherent species-specific differences as well as to the methods used to measure photosynthesis. Grout (1988) postulated that cultured plantlets can be divided into two discrete groups of photosynthetic response. In one group represented by strawberry and cauliflower, leaves formed in vitro never developed photosynthetic capability if they grew while the plant was on medium containing sucrose. After transfer to medium lacking sucrose, these leaves functioned as storage organs supplying carbon, stored as starch, to the new developing leaves. The persistent leaves were not photosynthetically competent and eventually declined. The second group included species such as Dieffenbachia and had persistent leaves that eventually adapted to autotrophic conditions, with concomitantly increasing rates of photosynthesis. The addition of sugars to the medium had a positive influence on the formation of biomass and leaf area of tobacco plantlets (Ticha et al., 1998). Similar results have been obtained for various species (e.g. Cournac et al., 1991; Galzy and Compan, 1992; Paul and Stitt, 1993; Kovtun and Daie, 1995), and they could simply reflect that the additional carbohydrate pool was used for biomass formation. Also the accumulation of chlorophyll and the photosynthetic capacity were positively

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affected by sugar feeding. This corresponds to the results of some (e.g. Galzy and Compan, 1992; Paul and Stitt, 1993; Kovtun and Daie, 1995), but not all (e.g. Capellades et al., 1991; Hdider and Desjardins, 1994) groups working on the topic. The positive effect of sugar feeding on photosynthesis is not consistent with the hypothesis that excess sugars cause the down regulation of photosynthesis (Koch, 1996). A down regulation was observed when sugars were fed to suspensioncultured cells (Schafer et al., 1992) or tobacco leaves (Krapp et al., 1991), or when the balance between the production and the consumption of carbohydrates was disturbed (Koch, 1996), Paul and Stitt (1993) argued that the lack of down regulation in photomixotrophically grown tobacco plantlets might be due to the low light regime which results in low photosynthetic rates and a source limitation of growth. The light saturated photosynthetic rate of 3% high light plantlets was about 50% lower than in field-grown tobacco plants. This observation indicates that the source limitation is indeed stronger in plantlets grown in vitro than in field-grown plants. On the other hand the positive effect of sugar feeding on chlorophyll content was more pronounced under high light conditions than under low light conditions, although the source limitation should be considerably smaller in high light. This indicates that source limitation might not be the decisive factor. Kovtun and Daie (1995) observed that exogenous sugars accelerate leaf development and sink-to-source transition in Beta vulgaris plantlets grown in vitro. They suggest that this effect prevents the down regulation of photosynthesis and the problem is not a source but rather a sink limitation and the enhanced development of photomixotrophically grown plantlets increases the capacity to use carbohydrates. Ticha et al. (1998) speculated that sugar feeding prevented the occurrence of photoinhibition. The photosynthetic capacity was considerably higher in the presence than in the absence of sugars and this might have increased the capacity to use the absorbed light. However, this effect was partly counteracted by the high chlorophyll content of 3% high light plantlets which increased the absorption of light energy. The adequate carbohydrate supply in 3% high light plantlets might enable them to maintain manifold protective and repair mechanisms, thus reducing the risk of photoinhibition. Part of this effect could be due to the enhanced growth rate which helps to replace impaired photosystems and to consume a surplus of assimilates. The accumulation of xanthophyll and zeaxanthin are regarded as plant responses to excess light energy (Bjorkman, 1981). Zeaxanthin and antheraxanthin could be detected in 3% high light and 0% high light plantlets, indicating that high light represented excess light energy (Table 1). The high pool size of xanthophyll cycle pigments in 0% high light plantlets shows that these plantlets experienced a sustained light stress during leaf development. Concerning the production of plantlets in vitro, Ticha et al. (1998) reported that photomixotrophic conditions yield better results than photoautotrophic conditions. However, they are in the opinion that net photosynthesis rate (NPR) and growth of chlorophyllous explants can be increased by increasing carbon dioxide concentration and by high photosynthetic photon flux

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Table 1 The contents of photosynthetic pigments in plantlets after 35 days cultured under given conditions besides the totals for xanthophylls cycle pigments (V + A + Z) and carotenoids (total Car) and also the chlorophyll a/b ratio, the percentage of different carotenoids in the total carotenoid pool and the deepoxidation state {DEPS,(Z + 0.5A)/(V + A + Z)} are shown Pigment 1

(V + A + Z)/Chl (mol mol ) Total Car/Chl (mol mol 1) Chl a/b (mol mol 1)

3% HL

3% LL

0% HL

0% LL

0.041 0.173 2.62

0.048 0.224 2.70

0.106 0.448 1.28

0.045 0.220 2.69

Carotenoid pattern (% of total carotenoid) Violaxanthin 2.8 Antheraxanthin 2.8 Zeaxanthin 18.3 Neoxanthin 16.5 Lutein 34.9 b-Carotene 23.9 DEPS 0.83

16.9 2.1 2.8 14.1 33.1 30.3 0.18

2.7 9.6 12.0 9.6 36.4 29.9 0.69

14.5 1.4 5.1 15.2 29.7 34.1 0.29

The data are means of three HPLC runs from three plantlets (10 leaf discs) (Ticha et al., 1998). HL = high light (200 mmol m 2 s 1); LL = low light (60 mmol m 2 s 1).

(PPF) conditions in the growth vessel/chambers. A large number of reports are published on supplementation of photosynthesis by CO2 enrichment and high PPF condition during in vitro culture of various plant species including avocado (Prusky et al., 1996), cacao (Figueira et al., 1991), carnation (Kozai, 1988), raspberry (Desjardins et al., 1988), carrot, citrus, kale, lettuce, radish and tomato (Tisserat et al., 1997). The CO2 concentration inside the culture during the photoperiod can be increased by increasing the number of air exchange of vessel per hour (Kozai and Seikimoto, 1988) and/or by increasing the CO2 concentration inside the culture room (Kozai and Iwanami, 1988). Employing an air diffusive filter on the vessel increases the number of air exchanges of the vessel. Moreover, the CO2 concentration inside large culture vessels would be increased and easily controlled with a forced ventilation system and, if necessary, CO2 enrichment in the incoming air. 3. Stomatal functioning Stomatal structure and impaired stomatal functioning have been considered as factors contributing to excessive loss of water by cultured plants. SEM studies indicated that stomatal structure in some species of cultured plants differed markedly from that in greenhouse or field-grown plants. Stomata had raised, rounded guard cells compared to normal elliptical,

sunken guard cell in a variety of species including sweet gum (Wetzstein and Sommer, 1983; Lee et al., 1988), red raspberry (Donnelly and Vidaver, 1984b), apple (Blanke and Belecher, 1989), rose (Capellades et al., 1990). SEM studies in Citrus leaves (Hazarika et al., 2002b) indicated that stomata with kidney shaped guard cell (Fig. 2a) was observed in greenhouse leaves while crescent shaped and rounded guard cell (Fig. 2b and c) were observed in in vitro leaves. When expressed as an index per number of epidermal cells, there was no significant difference in stomatal frequency among in vitro, acclimatized and greenhouse-grown plants (Conner and Conner, 1984). But Zaid and Hughes (1995) reported that the stomatal frequency of greenhouse green leaves of date palm was significantly higher than the control plantlets. However polyethylene glycol (PEG) treatment did not increase the number of stomata of in vitro plantlets of date palm. In Liquidamber styraciflua, Vacciniuum carymbasum and Nicotiana tabacam stomatal density decreased after transplantation (Wetzstein and Sommer, 1983; Ticha et al., 1999). After a short period of acclimatization, stomatal density on adaxial and abaxial leaf epidermis of N. tabacum plants was not changed, but later the total number of stomata per leaf were more than double in ex vitro plants due to enormous leaf area growth after transferred to ex vitro conditions (Pospisilova et al., 1999). On the other hand in Prunus serotina and Rhododendron spp. plants stomatal density increased and stomata pore length decreased after transplantation (Drew et al., 1992) leaves from in vitro grown Prunus cerasus, Vaccinium corymbosum or N. tabacum plantlets had ring shaped stomata, but in leaves of ex vitro transferred plants, stomata were elliptical (Marin et al., 1988; Ticha et al., 1999). Guard cells of in vitro grown Rosa hybrida plantlets contained numerous ribosomes and mitochondria, starch rich plastids and relatively large vacuoles indicating that they may exhibit metabolic activity similar to normal guard cells (Sallanon et al., 1991). Many researchers have reported that stomata of cultured plants show a characteristic inability to close when first removed from culture. Stomata on excised leaves of micropropagated apple plants did not close when treated with ABA, CO2 or mannitol immediately after removal from culture whereas stomata on plants that had been acclimatized responded as expected by closing immediately when treated (Brainerd and Fuchigami, 1982). Stomata on epidermal strips of chrysanthemum also did not close when incubated in darkness or in the presence of ABA (Wardle and Short, 1983). Likely in vitro cultured sweet gum leaves did not respond to applications of ABA, whereas stomata on leaves of

Fig. 2. Scanning electron micrograph of stomata from normal greenhouse leaves (a) and in vitro leaves (b and c).

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greenhouse-grown and acclimatized plants closed in response to this treatment (Wetzstein and Sommer, 1983). Marin et al. (1988) reported that up to 80% of stomata of excised leaves of P. cerasus closed when removed from culture and placed in 45% relative humidity. They concluded from a histochemical study in which they studied the cellulose, pectin, callose, cutin and birefringence patterns produced by the cellulose microfibrils in epidermal cells that a non-functional state existed in stomata in in vitro leaves. These stomata, however were able to revert to a functional state to some degree relatively rapidly after removal from culture. In addition, Sutter (1988) noted that there appeared to be a district difference in response of stomata on excised compared with those intact leaves and among different species. Seventy-five percent of stomata on detached in vitro leaves of P. cerasus closed after 15 min (Marin et al., 1988). Stomata of apple shoot remained open after removal from culture, but up to 78% of stomata of cherry and sweet gum plants closed after 1 h of exposure to ambient condition on a laboratory bench (Sutter, 1988). One cannot generalize that stomata of all micropropagated plants are unable to close in response to chemical treatment, darkness or an increased water vapour pressure gradient. Sutter and Langhans (1982) stated that stomata on excised leaves of cultured cabbage plants close after the leaves were allowed to wilt for 5 min. Shackel et al. (1990) reported that stomata of intact apple shoots do have the ability to close in an atmosphere of 90% relative humidity. They showed that water loss of micropropagated plants immediately after removal from culture, measured in a specially adapted cuvette attached to a steady-state porometer decreased over a period of 24 h to steady continuous rate. This steady rate, which was maintained for as long as 3 days was indicative of a low rate of cuticular transpiration combined with transpiration from any open stomata. The size of stomatal aperture is controlled by the relative volume of the guard cells depending on the turgor. Guard cell turgor in turn depends on vascular osmotic and water potential. The failure of stomata to close could be due to either abnormal cell wall properties or improper protoplast function (Zeiger, 1983). In carnation the failure of the guard cells to contract in hypertonic solutions resulted from defects in the cell wall and correlated with abnormal orientation of cellulose microfibrils (Ziv et al., 1987). However, dark treatment did not induce stomatal closure and vacuolar

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volumes remained unchanged. K content in guard cells did not vary significantly and a very low concentration of Ca2 ions was found. However, after ex vitro acclimatization, stomatal sensitivity to the dark was developed. Simultaneously, the light induced opening of stomata and K influx into guard cells were observed and calcium amount was 10 times higher than in guard cell of in vitro grown plantlets (Sallanon et al., 1991). 4. The cuticle The cuticle is a superficial non-cellular layer secreted by the epidermis and composed of a cutin matrix together with embedded and surface waxes that covers above ground tissues of plants. The primary function of the cuticle is to limit transpirational water loss. Water permeability through the cuticle is influenced primarily by the structure and amount of cuticular and epicuticular waxes (Martin and Juniper, 1970). Scant deposition of protective epicuticular wax on the surface of the leaves of the in vitro grown plants, has been regarded as one of the most important factor responsible for excessive loss of water, leading to poor transplantation success (Fuchigami et al., 1981; Hazarika et al., 2000a, 2000c, 2001a, 2002a; Wetzstein and Sommer, 1982). Fig. 3 illustrates the epicuticular wax cover of citrus leaves from greenhouse (a) and in vitro culture (b) (Hazarika et al., 2002b). Measurement of the amount of epicuticular wax on leaves of cultured plants revealed that the lack of crystalline structure was correlated with significantly less epicuticular wax compared to that on greenhouse-grown plants. Epicuticular wax on cauliflower and cabbage leaves in vitro was only 25% that of plants grown in the greenhouse (Sutter and Langhans, 1982). The relationship between the amount of wax formed in vitro and that formed on greenhouse-grown plants was not consistent or predictable in foliage plants with naturally glossy surfaces (Sutter, 1985). Sutter concluded that SEM alone was not a reliable method for determination of amount of wax on leaf surfaces but that gravimetric measurements were necessary as well. Consequently conclusions about the amount of epicuticular was inferred from SEM micrographs without gravimetric corroboration may be suspect. The chemical nature of wax deposited on the surface of the leaves under in vitro conditions is also known to differ from that

Fig. 3. Scanning electron micrograph illustrating epicuticular wax cover on the leaf surface of greenhouse (a) and in vitro leaves (b).

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formed under natural conditions allowing excessive diffusion of water from in vitro formed leaves (Sutter, 1984). Epicuticular wax on leaves of cultured cabbage plants differed chemically from that on greenhouse-grown plants (Sutter, 1984). Wax on leaves formed in vitro had a higher proportion of esters and polar compounds and significantly less long chain hydrocarbons than that on greenhouse-grown leaves. Since polar compounds are less hydrophobic and afford greater water permeability than long chain hydrocarbons, it was probable that the chemical composition of the wax also contributed to water loss. Leaves of cabbage plants grown in vitro never became as glaucous as new leaves that formed after removal from culture. Newly formed leaves developed increasingly grater amount of wax and more complex crystalline structure over time (Sutter and Langhans, 1982). Similar results were noted in carnation (Sutter and Langhans, 1979), cauliflower (Grout and Aston, 1977a), and cabbage (Sutter, 1988). The scanning electron microscope studies in Leucaena leucocephala have revealed a definite increase in the amount of epicuticular wax deposited on the leaves following the transfer of plants out of culture. The micropropagated plants attained wax density comparable to that of field-grown plants, within 6–7 weeks of transplantation (Dhawan and Bhojwani, 1987). These observations corroborate well with observations on the rate of water loss from leaves at different stages of micropropagation and hardening. The decline in the rate of water loss coincided with the increase in the amount of wax deposited on the leaves. However, the efficient water economy of the nature-nurtured plants, i.e. naturally grown plants could not be matched by the transplanted plants even after 5 months. This may perhaps be due to the difference in the chemical nature of the wax deposited; hydrophobic wax, typical of the leaves of plants grown in vitro (Sutter, 1984), being more abundant on the transplants than the hydrophilic wax predominantly found in the plants growing in the field. However, this hypothetical assertion needs to be confirmed by proper chemical analysis. Differences in the rate of water loss by leaves at different stages of micropropagation have also been reported in Malus domestica (Brainerd and Fuchigami, 1981), Prunus insititia (Brainerd et al., 1981), Brassica oleracea (Sutter and Langhans, 1982) and Solanum laciniatum (Conner and Conner, 1984). The reduced amount of epicuticular wax was directly correlated with substantially increased water loss in cultured shoots. Transpiration rates were significantly higher in leaves of cultured plants lacking epicuticular wax compared with rtes in greenhouse-grown plants (Wardle et al., 1983; Sutter and Langhans, 1982). Increased water loss directly related to the presence of epicuticular wax was shown when greenhousegrown cabbage leaves lost significantly more water after epicuticular wax was removed by chloroform (Sutter and Langhans, 1982). Sutter (1985) noted however that the quantity of epicuticular wax alone was not a good predictor of survival of micropropagated plantlets in the greenhouse during acclimatization. 5. Anatomy The poor mesophyll differentiation and weak vasculature of the leaves formed in vitro render the plants highly susceptible

to transplantation shock. Leaves of plants grown in vitro were thinner and had a characteristically poorly developed palisade layer with significant amount of mesophyll air space compared to greenhouse-grown plants. Both micropropagated cauliflower (Grout and Aston, 1978) and sweet gum (Wetzstein and Sommer, 1982) plantlets failed to develop a clearly defined palisade layer in vitro. Leaves on micropropagated plum shoot had only one layer of palisade cells rather than the usual two to three layers and greater air space in mesophyll tissue compared with leaves on greenhouse or field-grown plants (Brainerd et al., 1981). Such dissimilarities in leaf anatomy of in vivo and in vitro grown plants were also observed in Liquidamber styuraciflua (Wetzstein and Sommer, 1982) and Rubus idaeus (Donnelly and Vidaver, 1984a). Stems of red raspberry plantlets grown in vitro were most slender and had considerably less collenchyma and sclerenchyma supportive tissue than plants that were grown in the field. Roots on in vitro plantlets were slender, covered with root hairs and had much less periderm than the field-grown red raspberries (Donnelly et al., 1985). The connection between roots and shoots was shown to be incomplete in cauliflower plants resulting in insufficient water transfer between the roots and shoots (Grout and Aston, 1977a). However, xylem appeared continuous and was functional between the roots and shoot of in vitro rooted Prunus creases (Marin et al., 1988). The changes in leaf anatomy that occurred during acclimatization were noted most in leaves that developed after the plants were removed from culture. Persistent leaves of strawberry plants became thicker due to enlargement of the palisade cells (Fabbri et al., 1986), but there was no change in numbers of layers of palisade cells or in amount of mesophyll air space. During acclimatization leaves present as primordial leaves in vitro assumed intermediate characteristics between leaves grown in vitro and greenhouse or field leaves. Only new leaves that formed completely after removal from culture resembled greenhouse-grown leaves (Wetzstein and Sommer, 1982; Donnelly et al., 1985). Hydathodes on leaves of micropropagated blackberry (Donnelly et al., 1987) and strawberry (Donnelly and Skelton, 1987) were open, whereas those on leaves of greenhouse-grown plants were closed or had smaller apertures. Since the hydathodes appeared to be open and exhibited guttation, it was hypothesized that they might contribute to adaxial water loss. 6. Hyperhydricity Hyperhydricity (also formerly known as vitrification) is a morphological and physiological disorder of plants vegetatively propagated in vitro. Hyperhydric plants are so named because they have a glassy appearance. Their stems and leaves are often thick, rigid and easily breakable (Fig. 4). They are characterized by decreased protein and chlorophyll content (Phan and Letouze, 1983; Frank et al., 2004), low phenolics (Perry et al., 1999), an increased water content (Bottcher et al., 1988) and an altered ion composition (Kevers and Gaspar, 1986). In many cases, hyperhydric leaves do not have palisade tissues, instead they have only a spongy and largely vacuolated mesophyll with large intercellular spaces (Vieitez et al., 1985).

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Fig. 4. Various stages of vitrification of carnation shoots during in vitro culture.

Many workers demonstrated that hyperhydricity is dependent on water availability, micronutrient content and hormonal imbalance in the media (Doneso, 1987; Kataeva et al., 1991). However, microenvironment of the culture vessel and ethylene composition inside the culture vessel were found to have a prolonged effect on hyperhydricity. Histochemical studies of guard cells from hyperhydric leaves revealed lower levels of lignins, cellulose, pectins and cutin, as well as deposits of callose (Kevers and Gaspar, 1985; Marin et al., 1988; Ziv and Ariel, 1988). The size of the stomatal apparatus is controlled by the relative volume of the guard cells depending on the turgor. Guard cells’ turgor, in turn, depends on vacuolar osmotic and water potential. The failure of stomata to close could be due to either cell wall properties or improper protoplast function (Zeiger, 1983). Guard cell walls have a regulating function in stomatal movement, their thickening, conformation and composition as well as the orientation of the microfibrils, contribute to their functioning (Palevitz, 1981). Structural defects in guard cells are associated with the development of hyperhydric leaves, which lack or have lost cell wall constituents (Gaspar et al., 1987). Microscopic studies indicate that the pore surrounded by the guard cells in hyperhydric leaves is more rounded in contrast to the elliptical pore in normal leaves (Ziv et al., 1987). Ziv et al. (1987) showed that in carnation even though stomata pores in hyperhydric leaves did not close in the dark, in ABA or in a hypertonic solution, the protoplasts in the guard cells responded to such signals. They concluded that the defect lies in the wall of the guard cells. Microscopic studies indicated that the pore surrounded by the guard cues in hyperhydric leaves is more rounded in

contrast to the elliptical pore in normal leaves (Ziv et al., 1987). In addition to an abnormal shape of the stomatal pore, the cell walls adjacent to the pore are torn and damaged. This could have resulted from failure of the process of cell division during cell plate formation as well as deficiency in pectic substances and cellulose during secondary wall formation (Ziv and Ariel, 1992). The walls are supported by microfibrils, oriented in parallel to the longitudinal axis of the pore and are distributed anisotropically. This special orientation controls stomatal movement, in response to changes in the guard cells’ turgor pressure. Faulty orientation and callose presence affected the stomatal apparatus movement in hyperhydric leaves. The availability of Ca2+ and other ions, known to affect callose formation (Kauss, 1987) may have affected cell wall thickening, microfibril orientation and callose formation in the walls. Callose instead of cellulose deposition under such conditions may have caused stomatal malfunction. Hyperhydricity and stomatal deformation are affected by changes in cell wall structure, depending on the developmental state of the leaf. A close relation between hyperhydricity and guard cell wall deformation justifies the use of stomatal function as a precised parameter to define hyperhydricity. Olmos and Hellin (1998) studied the anatomy of normal and hyperhydric leaves of Dianthus caryophyllus plantlets regenerated from leaves using SEM and transmission electron microscopy. Hyperhydric leaves had large vacuolated mesophyll cells, showing hypertrophy of cells and preventing the formation of larger intercellular spaces; also the lack of cuticular wax and chloroplasts prevented the formation of abundant plastoglobuli. Pectins are a highly heterogeneous

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group of polymers that control cell adhesion, cell wall architecture and cell wall mechanical strength. Pectins control cell wall porosity and cell ionic status and are implicated in intercellular space development. The degree off etherification of pectins is controlled by the activity of cell wall pectin methyl esterases; their different actions can affect the properties of the cell wall, which have been considered important with respect to controlling the development of hyperhydricity (Saher et al., 2005). They reported that the pectate fraction was significantly increased in hyperhydric leaves of carnation while soluble pectin and protiopectins were significantly lower. Prunus avium shoots evolving towards a hyperhydric state produced higher amounts of ethylene, polymines and praline which are substances considered as stress markers (Frank et al., 2004). Piqueras et al. (2002) studied the changes in polyamine levels and patterns were studied in response to hyperhydricity in micropropagated carnation plants. Hyperhydric carnation leaves showed high peroxidase activity, low lignification and high malondialdehyde content, suggesting oxidative damage. The most predominant fraction of polyamine corresponded to free polymine in hyperhydric leaves as well as in control nonhyperhydric leaves. Regarding individual amines, hyperhydricity brought about an almost complex depletion of free 1,3-diaminopropane, a rise in conjugated form of the amine and a great reduction in bound spermidine in relation to non-hyperhydric leaves. A very high percentage up to 80% of reverted shoots was obtained by maintaining hyperhydric carnation shoots in a culture chamber with bottom cooling device that lowered the relative humidity inside the culture jars. Reversion of hyperhydricity was associated with changes in PA patterns. Thus, compared with both non-hyperhydric and hyperhydric leaves, reverted plants showed a drastic reduction in free PA, and a major increase in conjugated diamines (especially important in the case of cadaverine, Cad). The PA profile in non-hyperhydric and hyperhydric plants could indicate stress condition and a more suitable physiological situation in reverted plants. An integrated optimization of salt, antioxidant and growth regulator composition and potassium-phosphate buffer-mediated pH stabilization of culture media resulted in an effective tissue culture system that prevented the undesirable hyperhydricity of Ramonda myconi tissue cultures (Toth et al., 2004). Shetty et al. (1996) developed a Pseudomonas spp.-mediated approach to control hyperhydricity in oregano. This bacterium-induced prevention of hyperhydricity helped the establishment of clonal plants in the greenhouse without excessive acclimatization. The prevention of hyperhydricity and specifically linked to mucoid Pseudomonas spp. and was characterized by high chlorophyll and reduced water content in oregano shoots. The Pseudomonas spp.mediated hyperhydricity reduction in oregano is partially due to its extracellular polysaccharide. 7. Development of volatiles in vessel headspace In tissue culture closed vessels are used with the purpose of avoiding contamination. Sometimes, these may cause abnormal plant growth due to gas accumulation such as ethylene in tissue culture vessel. Ethylene, a plant growth regulator produce by

tissue, callus and plantlets is known to influence in vitro morphogenesis and well known to induce chlorophyll break down leading to senescence and leaf abscission. Excessive ethylene gas accumulation in closed vessel may inhibit plant growth and establishment of appropriate plantlets to use for repropagation and reduce the availability of oxygen for respiration (Adkins, 1992). Ethylene also lowers hairy root formation and L-DOPA (L3,4-dihydroxyphenylalanine) production (Sung and Huang, 2000). There have been several reports in which excessive ethylene accumulation at later stages of in vitro culture might be one of the major factors inducing hyperhydricity. To overcome this problem reducing relative humidity by providing good gas exchange through culture vessel (Buddendorf-Joosten and Woltering, 1995), increasing the agar concentration in the media (Debergh and Harbaoui, 1981), use of ethylene absorbent like charcoal (Mensuali et al., 1993), KMnO4 (Dimasi-Theriou et al., 1993), or alginate STS capsule (Sarker et al., 2002), utilization of large culture vessels (Jackson et al., 1991), forced ventilation (Zobayed et al., 2001) are some of the remedies that has been in practice. Among the chemicals AgNO3 has been widely and in most cases the most successfully used one. AgNO3 was also used in order to reduce the occurrence of hyperhydricity in tissue culture of sunflower (Mayor et al., 2003). Vessels that allow ventilation to the internal atmosphere have been designed in such a way to make the internal conditions more similar to those found outside for controlling hyperhydricity. Ventilation modified the anatomical characteristics of micropropagated carnation of shoots and leaves, and pushed the culture-induced phenotype closer to that of ex vitro acclimatized plants. 8. The persistent leaves Leaves that developed in culture were retained after transplantation for a week to several months prior to senescing (Grout and Millam, 1985). Persistence depends on the plant species and the degree of environmental stress ex vitro. These persistent leaves increased in size slightly mainly due to cell elongation (Fabbri et al., 1986). In most cases stomatal function has been equated with closure (Brainerd et al., 1981; Sutter, 1988; Wetzstein and Sommer, 1982). The role of persistent leaves is a very important issue. Photosynthetic capacity appears to vary with plant species in culture and may determine the ex vitro contribution of persistent leaves. Cultured plants are divisible in photosynthetic non-competent and competent species. In the non-competent group cultured cauliflower and strawberry leaves that develop in culture deteriorated rapidly after transplantation (Grout and Aston, 1978; Grout and Millam, 1985). These leaves contributed only those nutrients which could be restored by the transplant. Such leaves have been referred to as storage organ or pseudocotyledonary tissues (Wardle et al., 1983). Non-competence in strawberry has been attributed to irreversibly reduced levels of Rub‘P’case activity in leaves developed in the presence of sucrose. Strawberry plantlets defoliated in the absence of sucrose in the medium were competent (Grout and Price, 1987). Dieffenbachia (Dieffenbachia picta), as well as potato and chrysanthemum

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were photosynthetically competent in vitro. They achieved a positive carbon balance in culture and continued to contribute photosynthetically after transplantation. Leaves of competent species did not deteriorate rapidly after transplantation (Grout and Donkin, 1987). Persistent leaves of Asian white birch (Smith et al., 1986) and red raspberry (Donnelly and Vidaver, 1984a) seem to fall in to competent group. Red raspberry plantlets photosynthesized at a low level after transplantation. However, persistent leaves shifted to become both net respires and sinks for photo-assimilates formed in the new leaves by 1 month ex vitro (Donnelly and Vidaver, 1984a). 9. The new leaves The phenotypes of new leaves formed ex vitro varies with the species, the culture and transplant environments and the age of the transplant. New leaves of cauliflower (a noncompetent species) that formed the second week after transplantation apparently exhibited greenhouse control levels of CO2 uptake (Grout and Aston, 1977b). However, new leaves of red raspberry (a competent species) were transitional in the sense that weekly flushes of new leaves became progressively larger, eventually with control type anatomy, functional stomata and improved CO2 uptake capability (Donnelly et al., 1985). The number of transitional leaves produced by a transplant may depend on the number of immature leaf buds formed in culture. The degree of transition of these leaves and how closely they resemble those of control plants is probably a reflection of the stage of development of leaf primordia when the plantlets was transferred from culture and the conflicting stresses imposed on leaf development by both the culture environment and the new ambient environment (Donnelly and Vidaver, 1984a). It is likely too that the retention of any culture type organ on the transplant influences the physiological status of the rest of the plant (Donnelly et al., 1985). 10. Strategies for acclimatization Acclimatization of a micropropagated plants to a greenhouse or a field environment is essential because there is difference in micropropagation environment and the greenhouse or field environment. Successful acclimatization procedures provide optimal conditions for higher survival, subsequent growth and establishment of micropropagated plants. The physiological and anatomical characteristics of micropropagated plantlets necessitate that they be gradually acclimatized to the environment (greenhouse or field). Techniques that are more satisfactorily address the changes required for successful acclimatization require lower relative humidity, higher light level, autotrophic growth and a septic environment that are characteristic of the greenhouse or field. Although specific details of acclimatization may differ, certain generalizations can be noted.

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10.1.1. Abiotic approaches to acclimatization Three major strategies have emerged that focus on substantially changing the in vitro environment, especially in the later stages of propagation in order to modify the cultureinduced phenotype with improved storage capability, photosynthetic competence or water relations, and thus facilitate transplantation ex vitro. The first strategy assumes that larger persistent leaves, packed with greater amount of storage compounds would contribute more after transplantation. Increasing the concentration of sugar in the medium might maximize the nutrient function of persistent leaves (Grout and Millam, 1985; Desjardins et al., 1987). To some extent this strategy has been discounted for higher evapotranspiration losses in transplants however it seems to hold promise for some plants (Maene and Debergh, 1985). Wainwright and Scrace (1989) found that maximum values for shoot height, fresh and dry weight of Potentilla fruticosa and Ficus lyrata were obtained in vivo when previously conditioned with 2% or 4% sucrose. Sucrose concentration of 40 g l 1 prior to transferring watercress microcuttings to in vivo conditions was shown to maximize the dry weight of established plantlets (Wainwright and Marsh, 1986). The objective is to modify the CIP away from the characteristic hydrophytic-type anatomy and promote epicuticular and cuticular wax development, stomatal function and possibly overcome other deficiencies. Capellades et al. (1991) found that size and number of starch granules of Rosa cultivated in vitro increase with the sucrose level in the culture medium. Studying acclimatization of Asiatic hybrid lily under stress conditions after propagation through tissue culture, Mishra and Dutta (2001) reported that liquid medium having 9% sucrose and other phytohormones was found suitable for growth of bulblets in the isolated unrooted shoots. Due to the high concentration of sucrose the size of the bulblets increased from 0.5 cm in diameter to approximately 1–1.5 cm within 2 months of inoculation. Mehta et al. (2000) reported that increased in sucrose concentration from 2 to 4%. In the medium increases caulogenic response in tamarind plantlets from 34 to 48% in explants. Further increase of sucrose to 6% induced browning of media which was detrimental for growth of the shoots. Hazarika et al. (2000b) reported that in vitro preconditioning of citrus microshoots with sucrose concentration of 3% was found optimum for subsequent ex vitro survival (Fig. 5) and growth. There was a linear increase of biochemical constituents on addition of sucrose to the medium.

10.1. In vitro acclimatization The process of acclimatization can begin while the plantlets are still in vitro.

Fig. 5. Ex vitro survival percent of citrus microshoots as influenced by in vitro preconditioning with varying level of sucrose (Hazarika, 2003a).

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The second strategy assumes that autotrophic cultures will have persistent leaves that live longer and would be more photosynthetically productive ex vitro (Grout and Millam, 1985). The objective is to modify the culture induce phenotypes towards autotrophy in culture. To do this, oxygen concentration can be reduced in the culture environment which depresses the photorespiration rate (Shimada et al., 1988). Alternatively, the sugar is reduced or completely eliminated from the medium (Kozai, 1988) while the photosynthetic photon flux (PPF) and CO2 concentration are increased. In the system a gas permeable clear plastic film is used as a vessel closure (Kozai et al., 1988). This plastic film improves gas exchange in culture, CO2 enrichment or O2 reduction, increases the light penetration to the container content and decreases the relative humidity of the vessel. Such a system has the added advantages that microbial contamination is less of a problem when sugar is eliminated from the medium (Fujiwara et al., 1988). Kozai (1988) reported that the growth of plantlets in vitro is often greater under photoautotrophic conditions than under heterotrophic conditions, provided that the in vitro environment is properly controlled for promoting photosynthesis. He also reported that use of plant growth regulating substances, vitamins and other organic substances can be minimized because some of these will be produced endogenously in sufficient quantities by plantlets growing photoautotrophically. Short et al. (1987) found that growth by photoautotrophy could be stimulated by culturing chrysanthemum meristems on sucrose-free medium. Plantlets culture under this regime exhibited a comparable rate of photosynthesis to those found in seedlings. These procedures therefore can be used to facilitate the successful transfer of tissue culture-derived plants to soil conditions and thereby obviate the need for any hardening regime. The photoautotrophic tissue culture method makes it possible to use a large culture vessel without risk of increasing the loss of plantlets due to contamination. Use of a large vessel facilitates the automation, robotization and automatic environmental control. The third strategy assumes that plants developed under lower relative humidity will have fewer transpiration and translocation problems ex vitro and persistent leaves look like normal leaves. Lowering the relative humidity in vitro has been done experimentally with varying results. A range of methods has been used including the use of desiccant, by coating the medium with oily materials (Ziv et al., 1983; Short et al., 1987), by opening culture containers into a low relative humidity atmosphere (Brainerd and Fuchigami, 1981), adjusting the culture closure or using a special closure that facilitates water loss (Fari et al., 1987) or by cooling container bottoms (Vanderschaeghe and Debergh, 1988). In all cases these treatments were found to promote the surface wax development. Increasing the sugar or agar concentration or adding osmotic agents such as polyethylene glycol to the medium will also lower the relative humidity and in some cases serve the same purpose of desiccant (Leshem, 1983). Generally the relative humidity could not be lowered to loss than 80–85% without culture injury (Short et al., 1987). A relative humidity of 85% decreased the multiplication rate of carnation but

increased the number of glaucous leaves, the pigment and protein content, decreased the percentage water content and improved ex vitro survival (Ziv et al., 1983). At 80% relative humidity, growth rates of cauliflower and chrysanthemum were similar to those of controls grown under 100% relative humidity but ex vitro transplantation was greatly facilitated by functional stomata and greater epicuticular was deposition in plantlets rooted at the lower relative humidity (Short et al., 1987) By decreasing the relative humidity in culture containers, both transpiration and translocation systems are presumably improved in culture plants (Debergh, 1986, 1988a, 1988b) with associated improvement in mineral ion uptake through the transpiration stream and other benefits (Debergh, 1988a, 1988b). The obvious disadvantage of more extreme relative humidity reduction was to decrease the multiplication rate (Short et al., 1987; Ziv et al., 1983), posing and obvious dilemma (Ziv et al., 1983), Ziv (1986) recommended that relative humidity reduction should be considered in vitro, even at the expense of reduced propagation rates. As the propagation rate of cauliflower and chrysanthemum was not apparently compromised at 80%, relative humidity reduction was followed by the elimination of sucrose which successfully promoted autotrophy in Stage III cultures of both these plants (Short et al., 1987). This resulted in comparable photosynthetic rates for plantlets rooted at 80% relative humidity in vitro and seedling plants and underlined the viability of this approach. A number of innovative culture vessels have been developed to facilitate the establishment and growth of plantlets under reduced humidity in vitro. Tanaka et al. (1988) developed a disposable fluorocarbon polymer film culture vessel which is gas permeable and it may therefore facilitate a reduction in the relative humidity normally found in the culture flask. A preliminary evaluation of containers by Smith et al. (1990b) indicated that they result in the in vitro acclimatization of chrysanthemum plantlets which can be successfully transferred from the culture vessel to soil conditions without wilting. It is well known that plant growth retardants results in the production of more durable, compact plants with stronger shoots and roots. The inclusion, therefore, of the anti-gibberellin, paclobutrazol in the rooting medium has been studied as a methods of inducing in vitro hardening of cultured chrysanthemum plantlets. The culture of shoot meristem inoculated with sorbarods in liquid medium supplemented with 5 and 10 mM of paclobutrazol and incubated under high humidity (98%) resulted in the production of plants with reduced growth, increased deposition of surface wax and reduced wilting on transfer to field conditions (Ritchie et al., 1991; Smith et al., 1991). In addition, the precocious developments of trichomes on chrysanthemum have been reported by Ritchie et al. (1991). Hazarika et al. (2000a, 2001a) also reported that preconditioning microshoots with paclobutrazol influence higher ex vitro survival (Fig. 6) by intensifying internode length, thickening roots and reducing leaf dehydration by regulating stomatal function and increasing epicuticular wax per unit area of leaf of in vitro cultured citrus plantlets. Inclusion of paclobutrazol in the growth medium produced stomata with minimum apertures unlike normal stomata possibly due to a general reduction in cell expansion caused

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Fig. 6. Ex vitro survival percent of citrus microshoots as influenced by in vitro preconditioning with varying level of paclobutrazol (Hazarika et al., 2001a).

by anti-gibberrelin activity (Hazarika et al., 2002b). Paclobutrazol inhibits kaurene oxidase and thus blocks the oxidative reactions from ent-kaurene to ent-kaurenoic acid in the pathway leading to gibberrelic acid (Graebe, 1987). The strategies reviewed indicated that it is possible to acclimatize plantlets during in vitro culture by exposure to reduced humidity or paclobutrazol. 10.1.2. Biological approaches to acclimatization The micropropagation does not take into consideration the existence of mutualistic symbiosis of mycorrhiza and other associated plant growth promoting fungi and rhizobacteria. The media used are devoid of symbiotic propagules and therefore the plantlets obtained from these systems are not associated with any symbiotic fungi or bacteria in vitro, but often develop a negative association with a pathogenic organism when they are field planted. However, an early inoculation of these plants with an appropriate symbiotic organism promises to improve plants’ survival and performance. Steps in tissue culture where microorganism are applied potentially to give beneficial/fruitful results are

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symmetrically shown in Fig. 7. In recent years the role of endomycorrhiza in the outplanting performance of seedlings/ plantlets raised through tissue culture has been exploited (Bojan et al., 1995). Prior inoculation with symbiotic fungi in vitro would appear to offer solutions relating to hyperhydricity in micropropagated plantlets, as demonstrated for chrysanthemum (Nhut et al., 2003). As the fungi responsible for mycorrhiza are lost during micropropagation several workers have tried to discover effective methods of reinfection for better survival during acclimatization. Three major principles can be listed that are likely to be determined the extent of benefit to a plant as a result of the mycorrhizal symbiosis. Firstly that, in general, plants benefit primarily due to an increased supply of phosphorus. Secondly, some plant species are more dependent than others and therefore responsive species tend to have coarser roots and fewer root hairs. And thirdly isolates of the mycorrhizal fungi differ in their ability to benefit plants and both plant species and environmental conditions can modify this. The persistence of latent microbial contaminants in micropropagated plantlets has been suggested in many reports to increase vigour of the micropropagules and also the hardiness of micropropagated plantlets without showing any disease symptoms (Khan, 2003). The microbes may stimulate the growth by altering the phytohormone level in the host plants, secreting growth promoting substances and/or increasing photosynthetic and nutrient absorption efficiencies. Legumes and plants of a few other families develop a symbiotic association with specific nitrogen-fixing bacteria. The bacteria can grow in nodules on the roots of the infected plants where they utilize translocated carbohydrates. In return, the plant is able to mobilize some of the nitrogenous compounds produced by the bacteria through reduction of atmospheric nitrogen and consequently plant and bacteria usually exhibit a greatly improved growth rate. It can therefore be advantageous to inoculate the roots of micropropagated plantlets with suspension at acclimatization stage for better survival and growth. Dhawan and Bhojwani (1987) found that although micropropagated Leucena leucocephala plantlets took 3 weeks longer to develop root nodules than comparable seedlings, 80% of the plantlets become nodulated if they were inoculated with Rhizobium during acclimatization. 10.2. Ex vitro acclimatization

Fig. 7. Steps in biological hardening of in vitro cultured plants (Sahay and Verma, 2000).

10.2.1. Environment control Acclimatization of micropropagated plants to a greenhouse or a field environment is essential. Successful acclimatization provides optimal conditions for higher survival, subsequent growth and establishment of micropropagated plants. Traditionally, the acclimatization environment is adjusted to accommodate transplants from culture, generally weaning them towards ambient relative humidities and light levels. Novel approaches to ex vitro acclimatization include CO2 enrichment (Lakso et al., 1986) or with supplementary lighting (Desjardins et al., 1987). This reduces the ex vitro acclimatization interval in humidity chambers or in the greenhouse but fails

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to eliminate the requirements for habituation to low humidity. The most sophisticated ex vitro acclimatization procedures utilize the ‘‘acclimatization units’’ (Maene and Debergh, 1985; Kozai et al., 1987), an apparatus which immerged from the engineering science of climate controlled greenhouses. The microcomputer controlled acclimatization unit can determine the relative humidity, temperature, light intensity, CO2 concentration, air flow rate and even the temperature of the nutrient solution and has the potential to control almost every other feature of the environment. In the beginning changes are made in small increments, which are later increased. Special emphasis is placed on minimizing water stress in the early stages ex vitro. It is not surprising that in such a unit both transplant survival and growth rate are significantly increased. Plantlets acclimatized usually transplanted into big size pots, polybags or other containers with soil or suitable potting mixtures or directly to the field for further growth. It is necessary to be especially careful to water micropropagated plants adequately during the first few days. Dormancy requirements or rest period must be satisfied for each crop to allow successful propagation and growth. 10.2.2. Antitranspirants Spraying of coating of plantlets with impermeable materials (50% aqueous glycerol, low melting point paraffin or petroleum grease, dissolved in diethyl ether), growth retardants (paclobutrazol) and antitranspirants (abscisic acid) has been advocated as a method of reducing water loss from newly planted material. Antitranspirants have not proven useful in promoting ex vitro survival of performance; phytotoxicity and interference with photosynthesis were both cited as possible reasons (Sutter and Hutzell, 1984). Exogenous application of ABA to cell cultures can induce rapid hardening of cells to a significant level (Chen and Gusta, 1983). Application of ABA to whole plants has

consistently shown less dramatic hardening responses compared to cells (Gusta et al., 1982). Inadequate uptake, rapid metabolism and microbial degradation are suggested as possible reasons for the minor hardiness in whole plants following ABA application. ABA treatments along were not able to harden plantlets to the extent attained under low temperature acclimation conditions, suggesting that factors other than or additional to ABA are involved in hardening. 10.2.3. Ex vitro rooting and hardening Although it is a common practice to root microshoots in vitro, this may result in problems. When cauliflower shoots were rooted in vitro, the transition zone between the root and shoot was abnormal (Grout and Aston, 1977a). The vascular connections were poorly formed and narrow when observed at the time the plantlets were removed from culture. This restricted water uptake from the root into the shoot. After acclimatization, the vascular connections were more substantial, but water uptake remained less than in seedlings. Roots growing in agar often lack root hairs and may die shortly after transplanting, resulting in plantlets the case to grow (Debergh and Maene, 1981). Many commercial laboratories do not root microcuttings in vitro, because it is labor intensive and expensive. The process of rooting in vitro has been estimated to account for approximately 35–75% of the cost of micropropagation (Debergh and Maene, 1981). However, depending on the species and facilities, rootless microshoots may die in large numbers when removed from culture, whereas those with roots may have a higher survival rate. If losses of unrooted microcuttings are low, then rooting and acclimatization can be done simultaneously. This can substantially reduce costs of micropropagation and add efficiency to production schemes. Sharma et al. (1999) reported that acclimatization and hardening in tea micropropagation could

Fig. 8. Direct rooting and acclimatization of citrus microshoots in carriers.

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be accomplished as a one-step procedure with a short period of time before transplanting. Optimization of time of harvesting of microshoots, shot size, soil pH (4.0–6.4), plant growth regulator treatment (IBA 500 mg 1 1, 30 min), CO2 enrichment and light (15 mol m 2 s 1) conditions is specially designed hardening chambers made a significant impact on the percent of success for hardening in tea micropropagation. Das et al. (1990) reported that direct rooting of tea shoots was achieved by dipping the cut ends in IBA (50 mg 1, 30 min) and subsequently planting these in soil:peat moss 1:1 mixture. Shoots which were directly rooted in soil showed higher percent survival in the field than those rooted under in vitro conditions. An approach combining the advantages of in vitro and ex vitro has been successful for apples (Zimmerman and Fordham, 1985). They exposed microshoots to a sterile medium of IBA or NAA together with sucrose for 3–7 days during which root initials were formed. When combining rooting and acclimatization, the same environmental consideration apply as when acclimatizing rooted plantlets. Particular attention must be paid to humidity, light and temperature. Hazarika et al. (1996) reported direct rooting and acclimatization using 2 cm long microshoots of Aegle marmelos after treating them with IBA at 10 ppm for 2 min. Simultaneous ex vitro rooting and acclimatization was achieved using microshoots in carriers (Hazarika et al., 1995, 1999; Parthasarathy et al., 1999). The reported that among different carriers soilrite layered over farm yard manure showed better rooting (80–91%) (Fig. 8) and high ex vitro survival (90–97%) which was important over conventional rooting in agar-based medium. 11. Conclusion This review of morpho-physiology in vitro cultured plants indicates that the culture-induced phenotypes reflect epigenetic variations and their anatomy, physiology and biochemistry are atypical. In vitro plants are very delicate owing to high humidity in the culture flask, low light intensity and hetero or mixotrophic mode of nutrition. As a result, they lack the protective mechanisms like waxy cuticle, stomatal regulation, poor development of photosynthetic apparatus which make them vulnerable to transplant shock when exposed to the ambient conditions ex vitro. Therefore, understanding the physiology of plant cultured in vitro and the changes they undergo during the hardening process should facilitate the development of a successful transplantation protocol. The strategies reviewed indicated that it is possible to acclimatize plantlets during in vitro as well as in vivo by various methods and thereby facilitating the successful transfer of in vitro cultured plantlets to soil. Acknowledgements The author is grateful to the Department of Biotechnology, Ministry of Science and Technology, Government of India, New Delhi, for financial support. Dr. V.A. Parthasarathy, Dr. V. Nagarju and A. Bora are gratefully acknowledged for their support and constant encouragement.

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