J Appl Phycol (2007) 19:15–25 DOI 10.1007/s10811-006-9104-0
O R I G I NA L A RT I C L E
Callus induction and thallus regeneration from callus of phycocolloid yielding seaweeds from the Indian coast G. Rajakrishna Kumar · C. R. K. Reddy · Bhavanath Jha
Received: 6 December 2005 / Revised and Accepted: 24 May 2006 / Published online: 10 November 2006 C Springer Science + Business Media B.V. 2006
Abstract The tissue culture of phycocolloid yielding seaweeds included preparation of axenic explants, callus induction, subculture of excised callus and regeneration of plantlets from pigmented callus in the laboratory. Treatment of algal material with 0.1–0.5% detergent for 10 min and 1–2% betadine for 1–5 min and 3–5% antibiotic treatment for 48–72 h successively enabled viable axenic explants to be obtained as high as 60% for Gracilaria corticata, Sargassum tenerrimum and Turbinaria conoides and 10% for Hypnea musciformis. Callus induction was more conspicuous in T. conoides than in the other three species investigated. Of the irradiances investigated, 30 μmol photons m−2 s−1 produced calluses in as many as 40% explants in G. corticata and T. conoides and 10% in H. musciformis and S. tenerrimum. The explants cultured at 5 and 70 μmol photons m−2 s−1 did not produce any callus in all the species studied except for H. musciformis in which 10% explants developed callus at 5 μmol photons m−2 s−1 . Most of the species investigated showed uniseriate filamentous type of growths and buds from cut ends and from all over the surface of explants. Nevertheless, T. conoides had three types of callus developments, namely (1) uniseriate filamentous type of outgrowths Dedicated to the memory of Late Dr. Rangarajan. G. R. Kumar · C. R. K. Reddy () · B. Jha Marine Algae and Marine Environment Discipline, Central Salt and Marine Chemicals Research Institute, Bhavnagar 364002, India e-mails:
[email protected]/
[email protected]
from the centre of the cut end of explant, (2) bubbly type of callus and (3) club-shaped callus clumps. The subculture of T. conoides callus embedded in 0.4% agar produced two types of filamentous growth, namely filiform (with elongated cells) and moniliform filaments (with round cells) in the 2 months period after inoculation. Further, friable callus with loose cells was also found associated with excised callus. The moniliform filaments showed prolific growth of micro-colonies resembling to somatic embryo-like growth which, in liquid cultures, differentiated and developed into propagules with deformed shoots and distinct rhizoids. The shoots of these propagules remained stunted with abnormal leaf stalks without forming triangular shaped leaves as the parental plant and rhizoids had prolific growth in the laboratory cultures. The excised callus of G. corticata continued to grow when transferred to liquid cultures and showed differentiation of new shoots within 10 days. The shoots grew to a maximum length of 5–6 cm in the 2 months period in aerated cultures in the laboratory. Keywords Callus induction . Explant culture . Gracilaria . Hypnea . Sargassum . Thallus regeneration from callus . Turbinaria . Uniseriate pigmented filamentous callus Introduction Among the marine living resources the macroalgae (seaweeds) are increasingly viewed as a major potential Springer
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chemical resource, particularly for chemicals of industrial, pharmaceutical and nutraceutical importance (Carte, 1996; Rorrer et al., 1997, 1998; Rorrer, 2000; Schnitzler et al., 2001). Recently, bioprocess technology for the production of high value chemicals from cell and tissue cultures of different macroalgae has been developed using specially designed photo-bioreactors (Huang & Rorrer, 2002; Rorrer & Cheney, 2004). The main advantage of photo-bioreactor cultivation of cell and tissue cultures of macroalgae is the enablement of continuous, steady and defined production of high yields of quality product, thereby circumventing the barriers of seasonality. The optimization of culture conditions for each cell system can also be undertaken with ease and precision for deriving maximum benefit from photo-bioreactor grown cell cultures (Rorrer & Cheney, 2004). Further, the downstream process used for recovery of products from cell culture is more environmental friendly as compared to conventional process that utilizes the whole plants as a source of raw material for extraction. In contrast, the development of traditional macroalgal cultivation technology is aimed at the industrial scale production of biomass. This is labour intensive and requires huge cultivable sea area for farming of seaweeds. The crop yields also are inconsistent and are subjected to several critical factors such as quality of germplasm used for seeding, grazing, pathogens, mariculture practice, seasonal variation and natural calamities like tsunami and cyclones. The success of bioprocess technology for production of valuable compounds from macroalgae largely depends on the development of suitable in vitro tissue culture systems for bioreactor cultivation (Rorrer et al., 1998; Rorrer & Cheney, 2004). Although macroalgal tissue culture is underdeveloped relative to that of land plants, there are more than 40 species of seaweeds from which successful callus formation and subsequent plant regeneration have been accomplished (AguirreLipperheide et al., 1995; Rajakrishna Kumar, 2002). The initial objectives of developing tissue culture techniques for seaweeds has been either for understanding the fundamental aspects of callus formation, morphogenesis, role of plant growth regulators in morphogenesis, role of carbon sources on callus development and growth (Lawlor et al., 1989; Yokoya & Handro, 1996; Yokoya & Handro, 2002; Yokoya et al., 2004), recent studies have successfully employed these techniques for clonal propagation and maintenance of seed stock of economically important seaweeds for mariculture Springer
(Dawes & Koch, 1991; Dawes et al., 1993; Rajakrishna Kumar, 2002; Reddy et al., 2003; Rajakrishna Kumar et al., 2004). The present study reports preparation of axenic explants, callus induction, subculture and its morphogenesis in various red and brown algae namely Gracilaria corticata (J. Agardh) J. Agardh, Hypnea musciformis (Wulfen) Lamouroux, Sargassum tenerrimum J. Agardh and Turbinaria conoides (J. Agardh) K¨utzing. The techniques described in the present study are simple, reproducible and can be carried out with ease and success.
Materials and methods Collection of materials and preparation of axenic explants Gracilaria corticata, Hypnea musciformis, Sargassum tenerrimum from Diu (20◦ 42.73 N, 70◦ 55.48 E) and Veraval (20◦ 54 N, 70◦ 22 E) and Turbinaria conoides from Porbandar (21◦ 38 N, 69◦ 37 E) and Okha along the Saurashtra coast (west coast of India). After cleaning with seawater in the field, all the plants were wrapped in moistened tissue towels and brought in a cool icebox to the laboratory. Healthy plants, preferably free of any visible epiphytes and with few branches, were selected for tissue culture. However, in the case of Turbinaria, rhizoids were used for tissue culture as they were found to be relatively clean. Epiphytes and other microscopic contaminants were removed by manual brushing under a stereoscopic microscope. Then, unialgal cultures for each species were established by following the methods described by Rajakrishna Kumar et al. (2004). The axenic explants from each species in unialgal culture were prepared by exposing to different concentrations of surface sterilants such as detergent, iodine (Reddy et al., 2003) and broad spectrum antibiotic mixture (Polne-Fuller & Gibor, 1984). Selected healthy vegetative fragments of 2–3 cm length in five numbers from each thallus were treated with 0.1, 0.5, and 1% of liquid domestic detergent (Charmy green, Lion. Co., Tokyo, Japan) in filtered (Whatman GF/C) autoclaved seawater for 5, 10 and 20 min, with 0.5, 1, 2 and 5% of betadine (povidone iodine with 0.5% w/v available iodine) in filtered autoclaved seawater for 1, 2.5, 5 and 10 min and with 2, 3, 4 and 5% of broad spectrum
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antibiotic mixture in 100 mL of Provasoli Enriched Seawater (PES) medium (Provasoli 1968) for 24, 48, 72 and 96 h in batch cultures at 20–22 ◦ C under white fluorescence tube lights at 25–35 μmol photons m−2 s−1 with 12:12 light and dark photoperiod. The treatments that facilitated viable explants were regarded as sub lethal concentration of respective chemicals. The explants in subsequent experiment were simultaneously exposed to the sub lethal concentrations of detergent, disinfectant (betadine) and antibiotics to ascertain the cumulative effect of all three treatments on the viability of explants. The treatment that provided viable explants was finally used for testing its efficacy in making axenic explants by growing on ZoBell agar plates (Opprenheimer & ZoBell, 1952) for at least 2 weeks in a bacterial incubator. In order to achieve high level of axenicity and to avoid cross contamination between explants during antibiotic treatment, each explant was incubated separately in individual test tubes. Explant culture and callus induction Following the sterilization treatment, the fragments were thoroughly rinsed with autoclaved seawater to remove sterilants and then excised into 4–5 mm length explants. Each explant was then wiped gently with sterile filter paper to remove excessive moisture and mucilage from the cut ends prior to transferring to the explant culture medium. The explants 10–12 numbers were then inoculated (some vertically and some horizontally) into each Petri dish (90 mm × 15 mm) with 20 mL of solidified PES for red seaweeds or modified PES with 0.4% KI (PESI) for brown seaweeds with 1.5% Bacto agar (Difco, USA) and grown at 20–22 ◦ C under cool white fluorescent tube lights at 5, 30 and 70 μmol photons m−2 s−1 with 12:12 light and dark photoperiod. After 2–3 weeks, callus induction rate was calculated by counting the number of explants with callus from the total number of explants cultured. To ensure sustainable callus growth, the explants bearing callus were transferred regularly to fresh medium at 30-day intervals. Subculture of excised callus After 2–3 months of explant culture, callus outgrowth from each species was excised from the explant and subcultured separately in both liquid as well as solidified PES or PESI medium as described above. During
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subculture, the callus was transferred to fresh medium once every 2 months whereas the liquid medium was replenished every week till the germlings with sizable growth arises from callus. All culture dishes were maintained at 30–50 μmol photons m−2 s−1 .
Results The surface sterilization treatments, i.e. detergent and betadine tested individually or combined for obtaining axenic explants did not show detrimental effect on the viability of the explants from all the species except Hypnea musciformis (Table 1). The H. musciformis was found to be highly sensitive to betadine treatment and the explants exposed either to 0.5% betadine for 1 min alone or simultaneously to detergent and antibiotics bleached in subsequent cultures. Consequently, the explants were directly subjected to 1– 2% antibiotics treatment without detergent and betadine that yielded as low as 10–15% healthy explants. Prolonged exposure of explants from all species to 2% betadine for more than 5 min and 5% antibiotics for more than 72 h showed patches of damaged surface of thallus and explants when used from such material perished gradually on agar plates over a period of a week (data not shown). Generally, treatment of plant material with 0.1–0.5% detergent for 10 min and 1–2% betadine for 1–5 min and 3–5% antibiotics treatment for 48 h enabled viable and axenic explants to be obtained as high as 60% for G. corticata, S. tenerrimum and T. conoides and 10% for H. musciformis (Table 1). This treatment consistently rendered axenic explants in all the species investigated in this study without affecting the potential of callus development. A majority of the cultured explants showed uniseriate branched filamentous type of outgrowths from both cut ends, mostly from the cortical and medullary regions (Figs. 1–7). The callus induction in red seaweeds was comparatively rapid and observed within 2 weeks compared to the brown seaweeds which showed callus induction after 2 weeks. In some explants, especially from red seaweeds, filamentous type outgrowths were also observed over the whole intact surface of explants (Fig. 2). Occasionally, both filamentous type outgrowths and bud formation were also noticed from the cut surface of T. conoides explants. The filamentous outgrowths continued to grow in all species and over a 1–2 months period became prominent and spread Springer
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Treated materials were grown in static cultures in PES (for red) or PESI (for brown) medium for 8–10 days at 20–22 ◦ C under white fluorescence tube lights at 25–35 μmol photons m−2 s−1 with 12:12 light and dark photoperiod b Average of 20 explants a
60 10 60 60 48 48 48 48 3 1–2 5 5 1 0 5 5 1 0 2 2 0.1 0 0.1 0.1 Gracilaria corticata Hypnea musciformis Sargassum tenerrimum Turbinaria conoides
10 0 10 10
Exposure time (h) Concentration (%) Exposure time (min) Concentration (%) Exposure time (min) Concentration (%) Species
Detergenta
Betadinea
Antibiotic mixturea
Viable axenic explants (%)b
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Table 1 Optimised concentration and duration of detergent, betadine and antibiotics that produced highest percentage of viable axenic explants in different red and brown seaweeds
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all over the cut surface, forming an intact whitish caplike structure at cut end of explants, especially in T. conoides (Figs. 4–5). The cells in filaments were elongated (H. musciformis and T. conoides), broad and rectangular (G. corticata) and highly vacuolated with discoid plastids distributed all over the cell. The callus induction rate varied with species and irradiances. Callus induction was more conspicuous in T. conoides. Of the irradiances investigated, 30 μmol photons m−2 s−1 produced calluses in as many as 40% of the explants in G. corticata and T. conoides and 10% in H. musciformis and S. tenerrimum. The explants cultured at 5 and 70 μmol photons m−2 s−1 did not produce any callus in all the species studied except H. musciformis in which 10% explants developed callus when grown at 5 μmol photons m−2 s−1 (Table 2). The present study essentially reports differentiation of plantlets from callus of G. corticata and T. conoides. The callus obtained from H. musciformis and S. tenerrimum (Table 2) was minimal in terms of callus induction and growth and no further studies were possible (Figs. 1, 3). Furthermore, shoot regeneration (without forming callus) was most frequent and common in H. musciformis (Table 2). Though G. corticata developed uniseriate filamentous type of outgrowths from cut ends and from all over the surface of the explants (Fig. 2), three types of callus development were observed from Turbinaria conoides, namely (1) uniseriate filamentous type outgrowths from the centre of cut end of explant (Fig. 4), (2) bubbly type of callus (Fig. 5), and (3) shoots with or without branches sometimes developing callus clumps at the apices (Fig. 6). Nevertheless, more than 60% explants of T. conoides produced direct shoots from rhizoidal explants (Fig. 7). The callus in G. corticata was small in size and remained filamentous throughout the study without forming tufts at cut ends. The subculture of G. corticata excised callus showed shoot development within 10 days when cultured in liquid PES medium (Figs. 8–10) and the same did not survive for more than 2 months when cultured on solid medium (1.5% agar). The plants regenerated from callus filaments were similar to original plants in morphology and attained the size of 5–6 cm length in 2 months in aerated cultures in the laboratory (Fig. 10). Although the growth of subcultured calluses of T. conoides did not cease on 1.5% agar, it showed higher proliferations when grown as embedded culture in 0.4% agar (Fig. 11). There were two types of filamen-
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Fig. 1–7 Callus induction and their morphology in different red and brown seaweeds. Scale bars = 1 mm. (1.) Filamentous callus from the cut surface explant of H. musciformis grown on solidified PES medium at 20–22 ◦ C and 5 μmol photons m−2 s−1 after 30 days culture. (2.) Protruding out filamentous callus from the periphery of both cut and outer surface of the explant of G. corticata grown on solidified PES medium at 20–22 ◦ C and 30 μmol photons m−2 s−1 after 30 days culture. (3.) Forty-five days old explant of S. tennerrimum with bubbly type of callus development grown on solidified PESI medium at 20–22 ◦ C and 30 μmol photons m−2 s−1 . (4.) Rhizoidal explant of T. conoides with
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filamentous callus radiating out from the cut end after 75 days culture. (5.) Rhizoidal explant of T. conoides with bubbly type pigmented filamentous callus grown on solidified PESI medium at 20–22 ◦ C and 30 μmol photons m−2 s−1 after 90 days in culture. (6.) Rhizoidal explant of T. conoides with club callus clumps on both cut ends after 90 days culture on solid PESI medium at 20–22 ◦ C and 30 μmol photons m−2 s−1 . (7.) Rhizoidal explant of T. conoides with callus and shoot on cut ends after 90 days culture on solid PESI medium at 20–22 ◦ C and 30 μmol photons m−2 s−1
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J Appl Phycol (2007) 19:15–25 Table 2 The effect of irradiances on callus induction in red and brown seaweedsa Species
Explants (nos.)
Irradiance (μmol photons m−2 s−1 )
Callus induction (%)
Gracilaria corticata
50 50 50 50 50 50 50 50 50 50 50 50
5 30 70 5 30 70 5 30 70 5 30 70
NC 40 (60)b NC 10 (90) 10 (90) NC NC 10 NC NC 40 (60) NC
Hypnea musciformis
Sargassum tenerrimum
Turbinaria conoides
NC: no callus induction and explants bleached a All the explants were cultured in solidified PES (for red seaweeds) or PESI (for brown seaweeds) medium at 20–22 ◦ C with 12:12 light:dark photoperiod b Values in parenthesis indicate bud induction
tous growth observed from excised callus, i.e. filiform filaments (with elongated cells, Fig. 12) and moniliform filaments (with round cells) in the 2 months period after inoculation. Further, there were also friable-like calluses with loose cells associated with original excised callus culture (Fig. 13). Occasionally, the moniliform filaments had swollen terminal cells. Some of the moniliform filaments showed prolific growth of round cells forming a clump (Fig. 14) resembling somatic embryo-like growth. Culture of such filaments in liquid medium provided bud-like growth (Figs. 15, 16) that eventually in 5 months culture gave rise to deformed plants with abnormal long stalks without triangular shaped leaves and distinct rhizoidal system (Fig. 17). These propagules were found to propagate through fragmentation in culture flasks.
Discussion Preparation of viable axenic explants from seaweeds is considered as an essential step in achieving success in seaweed tissue culture. Simple treatment of explants successively with detergent, chemical disinfectant (betadine) and broad-spectrum antibiotics supplemented with a fungicidal agent (nystatin) allowed the production of a relatively high frequency of viable axenic material that was sufficient enough for initiating tissue culture from all the species, except from Hypnea musciformis which bleached. Yokoya et al. (2003) used Springer
simple washing of explants in sterile seawater with 0.5% sodium hypochlorite and 0.02% detergent for 10 min to obtain axenic explants in H. musciformis. The effectiveness of surface sterilants (detergent and iodine) used in this study in reducing microbiological contamination from explants has also been established (Rajakrishna Kumar et al., 2004). Callus development in multicellular macroalgae has been related to their thallus organization and differentiation (Aguirre-Lipperheide et al., 1995). The earlier studies have often used the term “callus-like formations” to distinguish callus of macroalgae (Garcia-Reina et al., 1991; Yokoya et al., 1993) from that of higher plants where the callus (disorganized growth of cells) develops from differentiated tissues as a result of wounding (Yeoman, 1987). Despite the differences in cellular organization among the red pseudo-parenchymatous type and brown (parenchymatous type) seaweeds, both groups have produced common callus clumps and/or uniseriate, pigmented and branched filamentous outgrowths from both cortical and medullary tissues. Yokoya et al. (2003) also reported apical filamentous callus formation on lateral branches in Hypnea musciformis. The majority of the calluses formed had arisen from both cortical and medullary cells indicating their high regeneration potentials than outer pigmented epidermal cells (Reddy et al., 2003). Light intensity plays an important role in the callus induction in red and brown seaweeds. In the present study, the callus induction confined to explants
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Fig. 8–10 Development of thallus from pigmented callus of G. corticata in liquid PES medium at 20–22 ◦ C and 50 μmol photons m−2 s−1 . Scale bars = 50 μm for Fig. 8, 1 cm for Fig. 9 and 2 cm for Fig. 10. (8.) Excised callus of. G. corticata grown in liquid culture after 10 days. (9.) Young buds of G. corticata regenerated from pigmented callus after 45 days culture. (10.) Fully grown tissue cultured plants after 60 days culture
cultured only at 30 μmol photons m−2 s−1 , although low photon irradiances (5 μmol photons m−2 s−1 ) are reported to have stimulatory effect on callus induction in K. alvarezii and G. acerosa (Reddy et al., 2003;
Rajakrishna Kumar et al., 2004). The differences in callus induction rate have also been reported to vary with species, irradiances used during explant culture and with endogenous substances present in Springer
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Fig. 11–17 Patterns of callus regeneration from callus of T. conoides. Scale bars = 1 mm for Fig. 11, 100 μm for Fig. 12, 75 μm for Fig. 13, 200 μm for Fig. 14, 5 mm for Fig. 15, 2 cm for Fig. 16, 2 cm for Fig. 17. (11) Subculture of excised callus of T. conoides after 45 days culture in 0.4% agar of PESI medium at 20–22 ◦ C and 50 μmol photons m−2 s−1 . (12) Filiformis pigmented callus from subcultured callus in 0.4% agar of PESI medium at 20–22 ◦ C and 50 μmol photons m−2 s−1 after 70 days. (13.) Callus clump with loose cells from subcultured callus in 0.4% agar of PESI medium at 20–22 ◦ C and 50 μmol photons m−2 s−1 after 70 days. (14.) Filamentous callus bearing micro-colonies (somatic embryo-like structures Springer
shown by arrow) of cells from subcultured callus in 0.4% agar of PESI medium at 20–22 ◦ C and 50 μmol photons m−2 s−1 after 3 months. (15.) Plant lets (30 days old) with deformed morphology regenerated from somatic embryo-like structures in aerated cultures of PESI medium at 20–22 ◦ C and 50 μmol photons m−2 s−1 . (16.) Plants (50 days old) with distinct rhizoids and deformed leaves (with arrow) cultured in aerated cultures of PESI medium at 20–22 ◦ C and 50 μmol photons m−2 s−1 . (17) Five months old T. conoides plants with deformed morphology developed from somatic embryo-like structures in aerated of PESI medium at 20–22 ◦ C and 50 μmol photons m−2 s−1
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explants (Yokoya et al., 2003). Studies with Sargassum confusum C. Agardh reported filamentous regenerated cells under dark conditions at 10–15 ◦ C and callus cell clumps at 20 ◦ C under higher irradiances from the stem explants (Kirihara et al., 1997). Although, information on the effect of spectral distribution on the growth of algae in vitro is limited and inconsistent (Lawlor et al., 1989), the maximum callus induction observed at 30 μmol photons m−2 s−1 in the present study could be due to use of optimal light intensity. Yokoya et al. (2004) also used similar irradiances for tissue culture of Gracilaria tenuistipitata Chang et Xia and G. perplexa Byrne et Zuccarello. The profuse filamentous outgrowths occasionally observed from intact apical explants and from external surface of explants in this study could be a response to injury that would have caused while cleaning the explants for obtaining sterilized explants. Callus formation in macroalgae, in general, appears to be a collective response to both wounding and change in external physical environment (explant culture on enriched solid medium), although Sargassum confusum (Kirihara et al., 1997) and Gracilaria chilensis (Collantes et al., 2004) explants produced callus in liquid cultures. The other interesting observation found was bud formation directly from a majority of explants in T. conoides. Bud formation in Gelidiella explants has been found to be associated with agar concentration in the culture medium (Rajakrishna Kumar et al., 2004). Collantes et al. (2004) also observed bud formation from explants of Gracilaria chilensis Bird, McLachlan and Oliveira in liquid cultures. The bud formation directly from explants has been explained as one kind of vegetative proliferation in which, when fronds are excised, lateral initial cells below the cut replace the apical function and therefore, recover their meristematic potential (Kling & Boddart, 1987; Collantes et al., 2004). In contrast to higher plants, the success of subculture of excised callus on solid medium independent of explant is not consistent (Aguirre-Lipperheide et al., 1995). Most calli have been reported to cease growth upon excision from the explants in Grateloupia doryphora (Robaina et al., 1990) and Gracilaria verrucosa (Hudson) Papenfuss (Kaczyna & Megnet, 1993). The failure of subculture of excised callus in Laurencia sp has been attributed to lack of ‘critical size’ of callus that is essential for producing shoots (Robaina et al., 1992). The bleaching of excised callus in subculture
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of both G. corticata and H. musciformis in the present study on solid media could be due to either small size of callus, i.e. smaller than critical size, or callus having cells with poor potential to regenerate and differentiate. On the contrary, the callus tufts excised from T. conoides explants were subcultured successfully in soft agar (0.4% agar). The success in subculturing enabled the production of tiny somatic embryo-like colonies on pigmented filamentous cells with friable callus clumps occasionally. Efforts are underway to grow this type of cells for exploring the possibility of establishing cell suspension cultures from brown seaweeds. The regeneration of somatic embryos first into rhizoidal system was high with most forming stunted shoots with abnormal long stalks without forming triangular leaves in aerated cultures in the laboratory. In the nature, rhizoids of Turbinaria are found almost throughout the year with multiplying high in early winter (November) and then plants emerge from the rhizoids under favourable environmental conditions (November–March). The inability of germlings to grow further into complete plants in laboratory culture needs to be investigated. This is the first study to report callus induction and regeneration from rhizoidal explants of the brown alga T. conoides. It is apparent from these findings that callus with robust growth and abilities to sustain for extended periods with explants will also have greater chances of survival in subculture after excision from the explant. To circumvent these uncertainties, subculture of callus is usually carried out in liquid cultures (Huang & Fujita, 1997; Kirihara et al., 1997). The twin advantages of subculturing in liquid cultures are first that it induces rapid growth and secondly the morphogenesis in callus. In algae which have simple cellular organization and are less evolved organisms, the transfer of excised filamentous tissue to liquid cultures ensures rapid differentiation in most species without the necessity for exogenous supply of growth regulators, unlike higher plants. In conclusion, the plants with firm and thick thallus produced prominent callus tufts with abilities to proliferate in subcultures using both solid and liquid medium. The methods described in the present study collectively assist in pursuing tissue culture research for other seaweeds while proving useful in clonal propagation of desired alga for field cultivation. Further, the callus obtained for these species provides an ideal option for maintenance and storage of germplasm as seed bank for cultivation. Springer
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Acknowledgements The authors are grateful to Dr. P.K. Ghosh, Director, CSMCRI for support and encouragement. This work was supported by the Department of Biotechnology (BT/PR 1721/AAQ/03/89/99) and partly by Council of Scientific and Industrial Research (CSIR), New Delhi, India. The first author likes to thank CSIR for the award of Senior Research Fellowship.
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