Optimization of protoplast yields from the red algaeGracilaria dura (C. Agardh) J. Agardh and G. verrucosa(Huds.) Papenfuss

Vishal Gupta & Manoj Kumar & Puja Kumari &C. R. K. Reddy & Bhavanath Jha

Received: 4 February 2010 /Revised and accepted: 17 August 2010 /Published online: 7 September 2010# Springer Science+Business Media B.V. 2010

Abstract This study reports on the optimization ofprotoplast yield from two important tropical agarophytesGracilaria dura and Gracilaria verrucosa using differentcell-wall-degrading enzymes obtained from commercialsources. The conditions for achieving the highest protoplastyield was investigated by optimizing key parameters suchas enzyme combinations and their concentrations, durationof enzyme treatment, enzyme pH, mannitol concentration,and temperature. The significance of each key parameterwas also further validated using the statistical centralcomposite design. The enzyme composition with 4%cellulase Onozuka R-10, 2% macerozyme R-10, 0.5%pectolyase, and 100 U agarase, 0.4 M mannitol in seawater(30‰) adjusted to pH 7.5 produced the highest protoplastyields of 3.7±0.7×106 cells g−1 fresh wt for G. dura and1.2±0.78×106 cells g−1 fresh wt for G. verrucosa whenincubated at 25°C for 4–6 h duration. The young growingtips maximally released the protoplasts having a size of 7–15 μm in G. dura and 15–25 μm in G. verrucosa, mostlyfrom epidermal and upper cortical regions. A few large-sizeprotoplasts of 25–35 μm, presumably from cortical region,were also observed in G. verrucosa.

Keywords Agarophytes . Commercial enzymes .

Gracilaria . Protoplast production . Rhodophyta

Introduction

Among the red algae, the genus Gracilaria has considerableindustrial importance as an agarophyte and is the principalsource of raw material to the agar industry worldwide(Zemke-White and Ohno 1999; Smit 2004). These agarindustries consume 72,300 dry tons of agarophytes annuallyand produce approximately 9,600 tons agar with a valueof US$173 million. Of this, Gracilaria alone accounts forabout 80% of the world’s agar market with a value ofUS$ 138 million (Bixler and Porse 2010). The increasingdemand for agar worldwide, coupled with short supplies ofagarophytes from wild stocks, has led to the developmentof viable field cultivation methods for their large-scalecultivation in the sea (Critchley 1993). Following the successin large-scale cultivation, cellular biotechnology techniquesare also being applied to improve the cultivated germplasmof this important resource (see reviews of Reddy et al.2008a, b, 2010). However, these techniques have largelyremained underdeveloped and are thus in their nascent stage.The high structural complexity and diversity in cell wallcomposition (Duckworth and Yaphe 1971; Rochas andLahaye 1989) have rendered the agarophytes in particularrecalcitrant to enzymatic digestion of the cell walls and havethus become an impediment for realizing the potentialsoffered by the application of biotechnology tools andtechniques for seaweeds in general. The skeletal and matrixpolysaccharides of cell walls of red seaweeds mainly consistof cellulose and agar. Therefore, the mixture of enzymeshaving cellulase and agarase are invariably required to digestthe cell wall components of agarophytic red algal species forpreparing protoplasts.

To date, protoplast isolation has been accomplished for48 red algal species, including agarophytes, belonging to 13genera (Reddy et al. 2010). In the genus Gracilaria, with

This paper was presented at the 7th Asia Pacific Congress on AlgalBiotechnology, New Delhi, 2009.

V. Gupta :M. Kumar : P. Kumari :C. R. K. Reddy (*) :B. JhaDiscipline of Marine Biotechnology and Ecology,Central Salt and Marine Chemicals Research Institute,Council of Scientific and Industrial Research (CSIR),Bhavnagar 364021, Indiae-mail: crk@csmcri.org

J Appl Phycol (2011) 23:209–218DOI 10.1007/s10811-010-9579-6

more than 150 species across the world, isolation ofprotoplasts has been reported only for 11 species (Reddyet al. 2010). The first report of on the production ofprotoplasts from Gracilaria species using a mixture ofcommercial cell wall lytic enzymes was of Cheney et al.(1986). Subsequent studies have used enzyme mixturescontaining various polysaccharide-degrading enzymes suchas cellulases and pectinases, along with agarase (Cheney et al.1986; Chou and Lu 1989; Bjork et al. 1990; Cheney 1990;Hong and Wang 1993; Mollet et al. 1995; Araki et al. 1998).However, so far, protoplast regeneration into whole plantshas been reported only for Gracilaria chilensis (Cheney1990), Gracilaria asiatica (Hong and Wang 1993), andGracilaria changii (Yeong et al. 2008). The only study thathas successfully accomplished interspecific protoplast fusionand regeneration of putative somatic hybrids from Gracilariatikvahiae and G. chilensis is that of Cheney (1990).

Gracilaria dura (C. Agardh) J. Agardh has beenreported to yield as much as 32–35% agar on a dry wtbasis with gel strength of 263–600 gcm−2 (Marinho-Soriano and Bourret 2005). Recently, Meena et al. (2007)reported preparation of high-quality agarose-form G. durawith suitability for biotechnology and molecular biologyapplications. The feasibility of field cultivation of this algausing both vegetative fragments (Dr. K. Eswaran, personalcommunication) and carpospores has also been successfullydemonstrated (Mantri et al. 2009). Rath and Adhikary(2004) and Christiaen et al. (1987) have examined thescope for industrial production of agar from Gracilariaverrucosa (Huds.) Papenfuss.

The present study describes the optimization of protoplastyield by standardizing the key parameters of protoplastisolation such as enzyme composition, enzyme pH, mannitolconcentration, incubation temperature, and duration for thetwo important agarophytes G. dura and G. verrucosa usingdifferent cell wall lytic enzymes obtained from commercialsources. These parameters were further validated statisticallyusing central composite design (CCD).

Materials and methods

Gracilaria dura and G. verrucosa were collected fromCSMCRI experimental seaweed farm at Mandapam (9° 16′ N,79° 8′ S), Tamil Nadu, southeast coast of India, and weretransported to the main laboratory at Bhavnagar under coolconditions in thermos flasks with ice. Selected thalli werecleaned manually several times in autoclaved seawater (ASW).The seawater used in this study for preparation of both culturemedium and enzyme solution was filtered using GF/C filters(47 mm diameter, Whatman, UK). The cleaned thalli wereestablished as unialgal cultures by growing in enriched seawatermedium (Provasoli 1968) supplemented with GeO2

(10 mg L−1) for a week in aerated flat-bottom round flasksunder cool white fluorescent lights at 15 μmol photons m−2 s−1

with a 12:12 h light/dark photoperiod. The medium wasreplenished every alternate day during the acclimatizationperiod to ensure exponential growth of thalli.

Protoplast isolation and purification

The commercially available cell wall lytic enzymes cellulaseOnozuka R-10, macerozyme R-10 (Yakult Co. Ltd., Japan),agarase, papain, and pectolyase (Sigma-Aldrich, USA) wereused for preparing the protoplasts. These enzymes were firstassessed individually for their ability in digesting the cellwalls, and those found effective were mixed in variousconcentrations and combinations to identify the right enzymemix for producing large numbers of protoplasts (Table 1). Theinitial experiments consisted of enzymes dissolved in60% seawater (30‰ seawater diluted with Milli-Q water)containing 25 mM MES adjusted to pH 6.0. Subsequentenzyme mixtures (A, B, C, D, E, and F) with appropriateosmoticum concentrations were made in autoclaved seawaterand then centrifuged at 10,000×g for 20 min at 4°C toremove the insoluble materials. The clear supernatant wasdispensed in cryo-vials as 5-mL aliquots and stored at −40°C.

Protoplast isolation and purification were carried outfollowing the protocol of Reddy et al. (2006). The growingapical tips from each species were excised and chopped intosmaller tissues (≤1 mm thin). The chopped tissues werethen rinsed several times with ASW supplemented with 2%NaCl to remove debris. The tissues thus cleaned weretreated with different enzyme mixtures as shown in Table 1and incubated on rotary shaker (50 rpm) for 6–8 h in dark at25°C. During the incubation period, periodic microscopicobservations were made to ascertain protoplast release. Theprotoplast yields were estimated by counting the cells usinga hemocytometer under an inverted microscope. The effectof pre-plasmolysis on the protoplast yields was also studiedby treating the chopped tissue in hyperosmotic ASW (1 Mmannitol, 450 mM NaCl, 100 mM MgCl2·6H2O, 20 mMKCl, and 40 mM BaCl2 as sulfate chelator) for 30 min priorto the commencement of enzymatic digestion.

Optimization of protoplast isolation parameters

The enzyme pH, incubation temperature, incubation duration,and osmoticum concentration were optimized for both algaeusing enzyme mix F. The ideal concentration of mannitol wasdetermined by the estimation of protoplast yield in the enzymemix F supplemented with various concentrations of mannitol(0.4, 0.6, 0.8, and 1.0 M). Similarly, optimal pH of enzymemix was determined at pH 5 and 6 (with 25 mMMES buffer)and 7.0, 7.5, and 8.0 (with 50 mM Tris–Cl buffer). Foroptimizing the incubation temperature, protoplast isolation

210 J Appl Phycol (2011) 23:209–218

was carried out at different temperatures (15°C, 20°C, 25°C,and 30°C). The protoplasts yield in each set of experiment wasquantified as described earlier.

Statistical analysis

Each experiment was conducted in triplicate. All the data werereported as mean ± SD. Analysis of variance (ANOVA) wasperformed for the comparison of the results under differentconditions. Any statistical property p≤0.05 was considered assignificant.

Multivariate optimization

All statistical calculations made for the optimisation processwere performed using the Unscrambler version 9.8 (CAMO,USA). The multivariate optimization of experiments wascarried out in two steps. The initial screening design served toidentify the variables having the highest influence on theyields of protoplasts. Additionally, it also enabled monitoringof the effect of interactions between the variables. Thescreening design used was a two-level full-factorial design,requiring 36 experiments (32 experimental points and fouradditional center points), which were conducted in random-ized order to avoid interferences from systematic errors. Thefactors which were significant at p<0.05 were considered tohave a significant effect on protoplast yields. CCD wasthereafter applied to determine the optimum level of thesignificant factors identified by full-factorial design. A totalof 34 run experiments were generated by Unscrambler 9.8which includes five center samples. The mean value ofprotoplast yields was taken as response. A multipleregression analysis was applied to the data thus obtained.Statistical analysis of the resulted model for the optimumconditions of variables was evaluated by ANOVA.

Results

The optimization of different physicochemical conditions(enzyme pH, osmoticum concentration, incubation tempera-

ture, and time) as well as enzyme combinations in the presentstudy had led to the development of an enzyme compositionfor obtaining abundant yields of viable protoplasts from G.dura and G. verrucosa. Individual enzyme treatmentsemployed in the present study did not yield any protoplasts.Nevertheless, the individual enzymes 4% cellulase onozukaR-10, 2% macerozyme R-10, and 100 U agarase did showsoftening of the tissues by the digestion of the components ofrigid cell wall. The protoplast yields for enzyme mixes A, B,C, and D were in the range of 102 to 104 protoplasts g−1 FWfrom both algae (Table 2). In contrast, enzyme mix E yielded7.75±0.7×105 protoplasts g−1 FW from G. dura and 1.2±0.7×104 protoplasts g−1 FW from G. verrucosa. Addition of0.5% pectolyase to enzyme mix E (termed as enzyme mix F)further enhanced the protoplast yield almost by fivefold in G.dura (3.7±0.7×106 protoplasts g−1 FW) and 1.8-fold in G.verrucosa (2.15±0.35×104 protoplasts g−1 FW). The pre-plasmolysis of the tissue before their direct incubation withenzyme mixture F substantially improved the protoplastyield in G. verrucosa from 2.15±0.35×104 to 1.2±0.78×106

protoplasts g−1 FW, while no appreciable effect on protoplastyield was observed in G. dura.

The optimization of pH has revealed a linear increase inprotoplast yield with increasing pH up to pH 7.5 and adecline thereafter. In G. dura, the protoplast yield at pH 6.0was 1.2±0.46×106 protoplasts g−1 FW and increased to3.25±0.63×106 protoplasts g−1 FW (about 2.5 times) for

Commercial enzymes Composition of enzyme mixtures

A B C D E F

Cellulase Onozuka R-10 (%) 3 3 2 5 4 4

Macerozyme R-10 (%) 2 2 1 2.5 2 2

Agarase (U/mL) 20 20 125 125 100 100

Abalone acetone powder (%) 2 5 – – – –

Pectolyase (%) – – – – – 0.5

Pre-plasmolysis – – – + + +

pH 6.0 6.0 6.0 6.0 7.5 7.5

Table 1 Combinations andconcentrations of enzymemixtures for isolation ofprotoplasts from G. dura and G.verrucosa

+ present, – Absent

Table 2 Protoplast yield data of G. dura and G. verrucosa obtainedfor different enzyme mixtures

Enzyme mixture Yield (protoplasts g−1 fresh wt.)

G. dura G. verrucosa

A 9.8±0.8×101 9.4±0.3×101

B 9.6±0.2×101 9.6±0.1×101

C 1.6±0.2×103 0.8±0.1×103

D 1.2±0.35×104 1.1±0.2×104

E 7.75±0.7×105 1.2±0.7×104

F 3.7±0.7×106 2.15±0.35×104

J Appl Phycol (2011) 23:209–218 211

pH 7.5 (Fig. 1a). The protoplast yield in G. verrucosa forpH 6.0 was 0.8±0.12×106 and showed a 1.5-fold increase(1.2±0.42×106 protoplasts g−1 FW) for pH 7.5 (Fig. 1a).

Protoplast release increased steadily with increasingtemperature and reached the optimum at 25°C with a yieldof 2.89±0.15×106 protoplast g−1 FW in G. dura and 1.8±0.12×106 protoplast g−1 FW in G. verrucosa as compared to1.75±0.12×106 and 1.5±0.32×106 protoplasts g−1 FWobtained at 20°C, respectively. Further increase in tempera-ture above 30°C greatly affected the protoplast yields(Fig. 1b).

Amannitol concentration of 0.4Mwas found to be optimaland gave yields as high as 3.2±0.2×106 protoplasts g−1 FWfor G. dura and 1.6±0.4×106 protoplasts g−1 FW for G.verrucosa (p≤0.05). The protoplast yield for both species didnot vary significantly from that of 0.4 M when mannitolconcentration was increased to 0.6 M (Fig. 1c). Furtherincrease of mannitol concentration to more than 0.8 Mreduced the protoplast yield nearly to half and to one fourthof that obtained at 0.4 M for G. dura and G. verrucosa,respectively.

An incubation period of 6 h significantly enhanced theyield and produced as high as 3.7±0.7×106 protoplasts g−1

FW for G. dura and 1.2±0.78×106 protoplasts g−1 FW forG. verrucosa (p≤0.05). The yield for 4 h was 1.8±0.8×103

protoplasts g−1 FW in the former and 2.4±0.2×103

protoplasts g−1 FW in the latter. The corresponding yieldsfor 2 h were 1±0.2×102 protoplasts g−1 FW and 1.2±0.7×102 protoplasts g−1 FW, respectively. Further extension ofincubation period to more than12 h resulted in bleaching oftissue and poor protoplast yields (Fig. 1d).

It is evident from this study that enzyme mix Fconsisting of 4% (w/v) cellulase onozuka R-10, 2% (w/v)macerozyme R-10, 100 U (v/v) agarase, 0.5% (w/v)pectolyase, and 0.4 M mannitol in natural seawater(adjusted to pH 7.5) incubated at 25°C for 6 h in the darkproduced maximum yield of 3.7±0.7×106 protoplasts g−1

FW for G. dura and 1.2±0.78×106 protoplasts g−1 FW forG. verrucosa.

The full-factorial design revealed a significant positiveeffect of pH and incubation period while mannitol concen-trations and temperature were found to have a negativeeffect. The interaction of pH and mannitol concentrationwith temperature showed a significant positive effect onprotoplast yields (Fig. 2a). For the design of optimization,the ranges of parameters with significant positive effectswere narrowed down to avoid high variability. The CCDanalysis resulted in optimized conditions of pH andincubation period as pH 7.5 and more than 6 h. As aresult, the response surface analysis based on multipleregressions and analysis of variance and quadratic surfacemodel was computed for the optimization of protoplast

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Fig 1 Effect of pH (a),temperature (b), mannitolconcentration (c), andincubation period (d) onprotoplast yield of G. dura(white square) and G. verrucosa(black square). All experimentswere conducted using enzymemix-F. Different letters abovethe bars indicate significantdifferences at probabilityp≤0.05 according to two-wayANOVA

212 J Appl Phycol (2011) 23:209–218

yields (Fig. 2b). This plot clearly showed that pH in therange of pH 7–8 and incubation period of 4–6 h is optimumfor obtaining the maximum protoplast yields. The statisticalsignificance of the second-order quadratic model wasevaluated by ANOVA (Table 3). The model p value of0.05 (desired p≤0.05) indicated that model terms weresignificant. The CCD analysis further supported thatmannitol concentration is significant.

Freshly isolated protoplasts were spherical in shape withdiameter ranging from 7 to 15 μm in G. dura and 15–25 μm in G. verrucosa and with a chloroplast at one pole.A few large-size protoplasts (25–35 μm), presumably fromcortical regions, were also released in G. verrucosa.

Discussion

The present study demonstrates a successful method forisolation of a large number of protoplasts from two important

tropical agarophytes G. dura and G. verrucosa using acombination of cell wall lytic enzymes obtained fromcommercial sources. The optimization of parameters wasstatistically validated for the first time using one-factor-at-a-time design and finally resulted in optimized model usingCCD. One-factor-at-a-time design is useful in selecting thebest experimental treatment. Further, CCD improved theoptimization method by reducing the number of tests. Tothe best of our knowledge, this is the first report of theapplication of statistical methods to optimize the productionof protoplasts.

The highest protoplast yield of 1.2±0.78×106 proto-plasts g−1 FW obtained for G. verrucosa in this study iscomparable with that of 105–107 protoplasts g−1 FWreported by Bjork et al. (1990). Also, Mollet et al. (1995)reported a similar yield of 8–10×106 protoplasts g−1 FWfrom G. verrucosa using a combination of cellulaseonozuka R-10 and crude agarase prepared from bacterialsources (Pseudomonas atlantica). Nevertheless, Araki et al.(1998) reported relatively higher yield of 1.03×108 proto-plasts g−1 FW using an enzyme mix containing agaraseprepared from Vibrio sp. in the laboratory. The interspecificvariations in the cell wall compositions of differentGracilaria species also plays an important role and partlyaccounts for varying protoplast yields among the species(see review by Reddy et al. 2010) and thus warrants studiesaimed at optimization of enzyme compositions and otherconditions. In contrast, for green seaweeds, especiallymembers of Ulvales with relatively homogenous cell wallcompositions, single-enzyme preparations were sufficientenough to produce large number of protoplasts (Reddy et

a

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Fig 2 Multivariate optimizationof protoplast isolationparameters for optimumprotoplast yields. Results ofscreening design afterfull-factorial experiment (a),response surface plot ofoptimized parameter with theinteraction of pH and incubationperiod (b)

Table 3 ANOVA table of the central composite design model foroptimized parameters

Source SS df MS F value p value

Model 20.47 14 1.46 3.10 0.021

Error 6.59 14 0.47

Adjusted total 27.07 28 0.96

Multiple correlation 0.87

R2 0.75

SS sum of squares, df degree of freedom, MS mean square

J Appl Phycol (2011) 23:209–218 213

al. 2006). The variations in protoplast yields from Gracilar-iales, especially with enzymatic methods, have beenattributed partly to physiological and biochemical as wellas seasonal changes of the material being investigated(Cheney et al. 1986; Kloareg et al. 1989; Chen and Shih2000). Moreover, the activity of crude enzymes also variesfrom batch to batch, leading to inconsistent protoplast yieldfrom time to time.

The cell wall composition of the genus Gracilariaconsists of an outer fibrillar layer of cellulose and a smallnon-cellulosic portion of xylose and mannose (Bellanger etal. 1990) and an inner amorphous matrix consisting ofsulfated polygalactan units (agar and agarose). The outer-most mucilaginous cuticle layer also hinders the penetrationand function of cell-wall-degrading enzymes. Further, thevariability in composition of polygalactans due to seasonaland physiological conditions of the thalli requires thepreparation of more specific enzyme mixtures includingthe incorporation of crude extracts from phycophages andmarine invertebrates prepared in the laboratory for proto-plast release from anatomically more complex seaweedssuch as Gracilaria (Table 4).

Although Yeong et al. (2008) showed release ofprotoplasts from G. changii with individual cell wall lyticenzyme, this could be due to lower degree of complexity inthe polygalactans units present in the matrix region of cellwall. In the present study, no individual enzyme was foundto be adequate for protoplast release, but the optimizedcombination of these enzymes (enzyme mix F) under definedphysical–chemical conditions led to optimal protoplastrelease.

The incubation of tissue in hyperosmotic solution leads toloosening of the cell wall from the plasma membrane,facilitating efficient penetration of cell wall enzymes leadingto efficient hydrolysis and giving rise to higher yields ofprotoplasts. One-molar mannitol has been used in pre-plasmolysis for the improvement of protoplast yields fromG. asciatica and G. changii (Hong and Wang 1993; Yeong etal. 2008). The pre-incubation of tissue of Laminariasaccharina and Laminaria digitata in hypertonic solutioncontaining salts of sodium, magnesium, and calcium chelator(EGTA) has also been shown to improve the protoplast yield(Butler et al. 1989). The pre-plasmolysis treatment in thepresent study has increased the protoplast yield from 2.15±0.35×104 to 1.2±0.78×106 protoplasts g−1 FW in G.verrucosa, but no effect of pre-plasmolysis was recordedfor G. dura. The overall higher yield of protoplasts from G.dura compared to G. verrucosa could be due to the lesssulfated residues of polygalactans in its matrix region of cellwall.

Mannitol in the concentration of 0.8 M has been reportedas suitable osmotic stabilizer for the protoplasts in G.changii, G. verrucosa, and G. asciatica (Yeong et al. 2008;

Araki et al. 1998; Hong and Wang 1993) and 1.0 M for G.tikvahiae and Gracilaria lemaneiformis (Cheney et al.1986). On the other hand, Bjork et al. (1990) showedoptimal protoplast yield in an enzyme mix with 0.2–0.4 Mmannitol when exponentially growing plants were used.We, in the present study, also found 0.4 M mannitol asoptimal for both species. While no significant differencewas observed at 0.6 M mannitol compared to 0.4 M, theyield sharply declined at 0.8 and 1.0 M mannitol.

Previous studies on Gracilaria have shown that enzymepH 5.8–6.5 is optimal for protoplast release, while in thepresent study the enzyme mix at pH 7.5 resulted in 2.5-foldincrease in the yield from G. dura and 1.5-fold from G.verrucosa over the yields obtained for pH 6.0. The low yieldsin G. verrucosa with the same enzyme could presumably bedue to high sulfate content in their cell-wall-suppressing theenzyme activity particularly of agarase which is crucial forisolating protoplasts from red algae. Meena et al. (2007)while studying the agars from different tropical agarophytesreported sulfate as much as 3.3% in G. dura and more than5% for other agarophytes used for food-grade agar.

The incubation temperature of 20–25°C has beenreported as optimal for protoplast release from differentspecies of Gracilaria (see review by Reddy et al. 2010). Inthe present study, we also have had the maximum yields at25°C for both the species. Increased incubation temperature(>25°C) in the present study decreased the protoplast yielddue to bleaching of tissue resulting from degradation of thephycobilisomes. This bleaching of tissue (or chlorosis) isyet another major hurdle to be overcome for successfulprotoplast isolation and culture from Gracilaria spp.

Generally, the reported incubation periods for producingprotoplasts from Rhodophyta are highly variable and rangefrom 1 to 48 h depending upon the complexity of cell wallsof test species and types of enzyme compositions employed.Enzyme incubation periods of 1–3 h for Palmaria palmata(Liu et al. 1992; Le Gall et al. 2004), 2 h for Porphyrayezoensis (Liu et al. 2004), 18 h for Grateloupia sparsa andGrateloupia filicina (Chen and Chiang 1995), and 48 h forKappaphycus alvarezii (Salvador and Serrano 2005) havebeen used for successful protoplast production. The incuba-tion period of 2 to 4 h has been optimized for differentGracilaria spp. (Cheney et al. 1986; Bjork et al. 1990; Yanand Wang 1993; Araki et al. 1998; Yeong et al. 2008) withthe only exception being G. verrucosa (Mollet et al. 1995)for which 12-h incubation resulted in optimal yield ofprotoplasts. Therefore, the incubation period of 6 h asoptimized in the present study is in accordance with earlierstudies.

The apical tips from exponentially growing thalli haveshown the highest yield of protoplasts in both species.Cheney et al. (1986) also reported increased yield ofprotoplasts from young sporelings compared to mature

214 J Appl Phycol (2011) 23:209–218

Table 4 Enzyme mixtures used for protoplast isolation from different species of Gracilaria

Species Enzyme composition Yield (g−1 f wt.) Status Reference

G. asiatica Pretreatment NA PR Yan and Wang (1993)SW: DW 60:40

Mannitol 1 M

Treatment

Crude Agarase

Cellulase 4%

Sea snail enzyme 1%

SW:DW 60:40

Mannitol 0.8 M

CaCl2 5 mM

pH 6.2–6.5

G. changii Pretreatment 5–8×104 PR Yeong et al. (2008)SW:DW 60:40

Mannitol 1.0 M

Treatment

Cellulase 2%

Macerozyme 1%

Agarase 10 U/mL

MES 50

pH 6.0

G. chilensis NA NA NA PR Cheney (1990)

G. chorda Cellulase 3% 0.2–20×105 PI Chou and Lu (1989)Macerozyme 3%

Pectolyase 0.5%

Abalone acetone powder 10%

Mannitol 0.8 M

CaCl2 5 mM

pH 5.8–6.0

G. gigas Cellulase 3% 0.2–20×105 PI Chou and Lu (1989)Macerozyme 3%

Pectolyase 0.5%

Abalone acetone powder 10%

Mannitol 0.8 M

CaCl2 5 mM

pH 5.8–6.0

G. lemaneiformis Cellulase R-10 3% 3–10 × 105 CW Cheney et al. (1986)Macerozyme 3%

Agarase 1%

Pectolyase 0.5%

Mannitol 1 M

CaCl2 5 mM

SW:DW 60:40

MES 50 mM

pH 6.0

G. lemaneiformis Cellulase 2% 105×107 PI Bjork et al. (1990)Agarase 0.01%

Mannitol 0.4 M

pH 7.0

G. salicornia Cellulase 3% 0.2–20×105 PI Chou and Lu (1989)Macerozyme 3%

Pectolyase 0.5%

Abalone powder 10%

Mannitol 0.8 M

J Appl Phycol (2011) 23:209–218 215

Table 4 (continued)

Species Enzyme composition Yield (g−1 f wt.) Status Reference

CaCl2 5 mM

pH 5.8–6.0

G. sordida Cellulase 2% 105×107 PI Bjork et al. (1990)Agarase 0.01%

Mannitol 0.4 M

pH 7.0

G. tenuistipitata Cellulase 3% 0.2–20×105 PI Chou and Lu (1989);Macerozyme 3%

Pectolyase 0.5%

Abalone powder 10%

Mannitol 0.8 M

CaCl2 5 mM

pH 5.8–6.0

G. tikvahiae Cellulase R-10 3% 3–10×105 PI Cheney et al. (1986);Cheney (1990)Macerozyme 3%

Agarase 1%

Pectolyase 0.5%

Mannitol 1 M

CaCl2 5 mM

SW:DW 60:40

MES 50 mM

pH 6.0

G. verrucosa Cellulase 2% 105–107 PI Bjork et al. (1990)Agarase 0.01%

Mannitol 0.4 M

pH 7.0

G. verrucosa Pretreatment 8–10×106 PI&CW Mollet et al. (1995)CaCl2.2H2O 10 mM

Mannitol 0.4 M

Tris MES 10 mM

pH 6.0

Treatment

Agarase 25U

CaCl2. 2H2O 10 mM

Cellulase 1.25%

Mannitol 0.8 mM

Tris Mes 10 mM

pH 6.0

G. verrucosa Pretreatment 1.03×108 PI Araki et al. (1998)Papain 5%

Mannitol 0.7 M

MES 20 mM

pH 7.5

Enzyme Treatment

Cellulase R-10 4%

Macerozyme 2%

Agarase 4 U

Mannitol 0.7 M

MES 20 mM

pH 6.5

CW cell wall formation, PI protoplast isolation, PR plant regeneration

216 J Appl Phycol (2011) 23:209–218

thalli of G. tikvahiae and G. lemaneiformis. Bjork et al.(1990) also concluded that growth rate and cultureconditions strongly influence the protoplast yield indifferent Gracilaria spp., and Mollet et al. (1995) found agradual decline in the protoplasts yield due to prolongedculturing of algae in the laboratory.

The two Gracilaria species investigated in the presentstudy for protoplast isolation have been reported to containquality agar with high gel strength compared to otherGracilaria species from the Indian coast (Meena et al.2008). G. dura is considered to be a superior agarophytewith high gel strength of 250±8.16 g cm−2 and yield of 27±0.81% of native agar (Meena et al. 2007) while G. verrucosahas about 50% lesser gel strength (160–180 g cm−2) than G.dura and yields in the range of 27–30% (Rath and Adhikary2004). With the optimization of protoplast production fromthese two Gracilaria species, future work will focus onregeneration and protoplast fusion for developing strainswith improved traits such as higher growth rate andimproved agar yield and gel strength.

Acknowledgement The financial support received from theDepartmentof Science and Technology (DST), New Delhi (SR/SO/PS-53/2005), isgratefully acknowledged. The first author (VG) also expresses his gratitudeto DST, for the financial support and the second (MK) and third authors(PK) to the CSIR, NewDelhi, for awarding the Senior and Junior ResearchFellowships, respectively.

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