Pretreatment and process optimization of spent seaweed biomass (SSB) for bioethanol production using yeast (Saccharomyces cerevisiae)

Renewable Energy 153 (2020) 456e471

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Pretreatment and process optimization of spent seaweed biomass (SSB) for bioethanol production using yeast (Saccharomyces cerevisiae) M.P. Sudhakar a, *, K. Arunkumar b, K. Perumal c a

Marine Biotechnology Division, National Institute of Ocean Technology, Ministry of Earth Sciences (Government of India), Chennai, 600100, Tamilnadu, India b Department of Plant Science, Central University of Kerala, Periye, Kasaragod, Kerala, 671 314, India c Biodyne Research Institute, Annai Lea Community College, The School of Biodynamic Farming, Inba Seva Sangam, Sevapur, Kadavur T.K, Karur Dt, Tamilnadu, India

a r t i c l e i n f o

a b s t r a c t

Article history: Received 23 July 2019 Received in revised form 20 January 2020 Accepted 8 February 2020 Available online 11 February 2020

The study aimed to utilize the industrial spent seaweed biomass (SSB) for effective ethanol production using yeast as a fermenting microorganism. Pretreatment of SSB was optimized using different acids. The highest percentage of spent biomass was obtained from G. corticata (12.53 ± 2.66% DW). The proximate, ultimate and biochemical constituents of spent biomass were calculated. The total sugar (440 ± 40 mg/g DW), reducing sugar (129.85 ± 10.23 mg/g DW) and protein (11.08 ± 0.11 mg/g DW) content of SSB were analysed. Pretreatment was optimized using three different acids. The effect of different pH (4.5, 5.0, 5.5 and 6.0) and temperature (30 and 35
C) on ethanol production using baker’s and MTCC yeast was studied. At 35
C, the maximum (4.85% w/w) ethanol production was achieved in a fermentation process maintained at pH 4.5 and 5.0 at 24 h and 72 h, respectively. Substrate fermented with MTCC yeast recorded the maximum production of ethanol (4.98% w/w) at pH 4.5 within 48 h. The fermentation process was scaled up to 300 mL for ethanol production, and achieved 3.75% w/w ethanol (72 h, pH 5.5). To conclude, in future SSB would be a potential renewable novel substrate for bioethanol production when compared to other lignocellulosic substrates. © 2020 Elsevier Ltd. All rights reserved.

Keywords: Spent seaweed biomass Acid pretreatment Saccharomyces cerevisiae Bioethanol

1. Introduction As world energy demand continues to rise and fossil fuel resources are getting depleted, seaweeds are receiving increasing attention as an attractive renewable source for producing fuels and chemicals [1]. Growing demand for fuel (3.0 million barrels per day in 2040) to meet the technological advancement and decrease in natural oil resource due to overexploitation urged researchers to find an alternate way for fuel generation by renewable and green energy production technology methods [2e4]. Algae (micro- and macroalgae) emerged as potential renewable biomass that has enormous photosynthetic efficiency, generate and stored sugars in

Abbreviations: SSB, Spent seaweed biomass; YPD, Yeast potato dextrose; MTCC, Microbial Type Culture Collection; DW, Dry weight; HMF, hydroxymethylfurfural. * Corresponding author. Marine Biotechnology Division, National Institute of Ocean Technology, Ministry of Earth Sciences, Government of India, VelacherryTambaram Main Road, Narayanapuram, Pallikaranai, Chennai, 600 100, Tamil Nadu, India. E-mail address: [email protected] (M.P. Sudhakar). https://doi.org/10.1016/j.renene.2020.02.032 0960-1481/© 2020 Elsevier Ltd. All rights reserved.

it [5]. Macroalgae (also called seaweeds) are classified into three types: brown algae (Phaeophyceae), red algae (Rhodophyceae), and green algae (Chlorophyceae). Each type has its own food storage mechanism and generates polysaccharides through photosynthetic light harvest and nutrients available in the sea [6]. Red seaweeds mainly contain agar polysaccharides and some cellulose. The breakdown of agar into linear polymer gives agarose which is comprised of galactose subunits [7]. Seaweeds are abundant throughout the world, especially in countries with the more covering area of coastlines such as Japan, Philippines, Malaysia, Singapore, Thailand, most of the European countries, the United States and Australia. The macroalgae production during the year 2008 was 15.5 million tonnes fresh weight and 93% of it had good commercial value [8,9]. The total annual production value is estimated at almost US$ 6 billion, of which food products for human consumption represent US$ 5 billion [10]. Seaweeds are found abundantly in the south-east and west coastal areas of India (08.04e37.06 N and 68.07e97.25 E) with a coastline of about 7500 km comprising of 271 genera and 1153 species. Nearly, 7 lakh tonne of standing stock (wet weight) of seaweed is

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found along different coastal states of India and Tamil Nadu alone contributing around 2 lakh tonnes (wet weight). Seaweeds are rich in bioactive compounds like pigments, polyphenols and sulphated polysaccharides (agar, carrageenan and alginates) which are being utilized by various industries. Around 30e40% of the coastline is covered with drift seaweeds in Tamilnadu, India [11]. These drifted seaweeds remain unexplored. Biomass of marine origin has an advantage over terrestrial plant biomass for its application as feedstock for fuels [12], by converting diverse carbohydrates of seaweed biomass into liquid biofuels (e.g., bioethanol, biobutanol, biomethanol) through metabolic engineering [1]. The spent biomass obtained after extraction of pigment and phycocolloids still contains polysaccharides which are found to be a rich source for biofuel production [13e15]. These spent biomasses of seaweeds can be effectively utilized for bioethanol production through fermentation, the process to meet the demand for ethanol production and address energy shortage [16]. There are limited investigations carried out by researchers on pigment extraction followed by utilization of spent biomass for ethanol production from red seaweeds [17e21]. Conversion of spent seaweed biomass into biofuel production is still a major challenge. Ethanol production from seaweeds reported was shown in Table 1. The global production of ethanol using seaweeds is increasing tremendously. The analytical studies towards global ethanol production show that most of the ethanol is produced by the fermentation process which contributes 97% while only 3% of ethanol is produced via catalytic hydration of ethylene, a petroleum by-product [22]. The techno-economic analysis of macroalgae for biorefinery approach was analysed in detail [23], in which feedstock price, yield, moisture content of seaweeds, solids and enzyme loading were considered. Consider, feedstock price is $ 100/MT, then the minimum ethanol selling price was observed to be in the range of $ 3.6e8.5/gal and this will again reduce if the feedstock price reduced to $ 50/MT [23]. The production of synthetic ethanol is economically less attractive as compared to fermentation in the United States due to the high cost of ethylene and abundance of agricultural feedstocks for ethanol production. World bioethanol production reached 4.5 billion gallons in the year 2000, then increased to 22.7 billion gallons in the year 2012 [24]. The production of synthetic ethanol is growing in Middle East countries, especially Iran [22]. The largest synthetic ethanol plants in Germany and Scotland can produce 4.4 million gallons per year. There are several multinational companies which produce synthetic ethanol such as Sasol (Europe and South Africa), SADAF (Saudi Arabia), Shell (UK and Netherlands), BP (UK) and Equistar (US) [25,26]. The utilization of a low-value by-product from waste seaweeds for bioethanol production will pave the way towards the development of an economical biorefinery with zero-waste technology [27].

457

Hence, the present work is aimed at the extraction of sugars from selected red seaweeds and its conversion to bioethanol production through the fermentation process using yeast. 2. Materials and methods 2.1. Seaweed collection and processing Red seaweeds such as Gracilaria corticata, Gracilaria crassa and Gracilaria edulis (Fig. 1) were collected from different places such as Mandapam (9
160 57.2500 N, 79
120 5.9100 E) and Thondi (9
460 3500 N, 79
00 2800 E), Tamilnadu, India, respectively. The collected seaweeds were washed thoroughly in seawater followed by tap water to remove salts and silt. The processed seaweeds were stored at 4
C for further use. Red seaweeds contain valuable pigments such as phycoerythrin (water-soluble pigment protein) which were removed by cold distilled water (below 10
C) extraction method using mixer grinder [6,28,29]. The ground slurry was filtered and squeezed through a muslin cloth (200 mesh size). The obtained liquid contains phycoerythrin, and rest of the biomass was called as spent seaweed biomass (SSB), which is rich in polysaccharides and other pigments such as chlorophyll and carotenoids. The obtained SSB was dried under room temperature (35 ± 2
C) for further use as a biofuel substrate. This research work was carried out at Shri A.M.M. Murugappa Chettiar Research Centre (MCRC), Chennai, India. 2.2. Proximate, ultimate and biochemical analysis of SSB Analysis such as moisture [30], ash [31], volatile matter [32], total organic carbon, C, H, N, S, O [33], total sugar [34], reducing sugar [35], total protein [36,37] and trace elements [38] from three species of Gracilaria were analysed and recorded. 2.3. Extraction of sugars from SSB Acids such as sulphuric acid (H2SO4), hydrochloric acid (HCl) [39e41] and phosphoric acid [42] taken in different concentrations such as 1, 2, 3, 4 and 5% (v/v) were used for the hydrolysis of dry red seaweed, Gracilaria corticata, to yield maximum amount of reducing sugars. In this experiment, 4% (w/v) of dry seaweed in 10 mL of respective acids was taken in a test tube and autoclaved at 121
C for 15 min. After the hydrolysate was cooled to room temperature (35 ± 2
C), the hydrolysate was squeezed out using muslin cloth (500 mesh size) and was analysed for reducing sugar yield using DNS method [35]. The best acid that yielded maximum reducing sugar was taken for further studies. To optimize the substrate and acid concentration to get the maximum amount of reducing sugar, different concentrations of

Table 1 Ethanol yield reported in different red seaweeds. Seaweeds

Biomass type

Microorganism

Country

Ethanol yield

Reference

Gracilaria verrucosa Gracilaria salicornia Kappaphycus alvarezii Kappaphycus alvarezii (cottonii) Gelidium amansii Gelidium elegans Gelidium amansii Gracilaria chorda Gracilaria tenuistipitata Kappaphycus alvarezii Eucheuma cottonii Red macroalgae

Residual agar pulp Whole biomass Spent biomass Whole biomass Whole biomass Whole biomass Whole biomass Whole biomass Whole biomass Whole biomass Seaweed solid wastes Whole biomass

Saccharomyces cerevisiae HAU strain E. coli KO11 Saccharomyces cerevisiae (NCIM 3523) Commercial brewer’s yeast Brettanomyces custersii KCTC 18154P Saccharomyces cerevisiae IAM 4178 Commercial Saccharomyces cerevisiae Commercial Saccharomyces cerevisiae Commercial Saccharomyces cerevisiae Saccharomyces cerevisiae CBS1782 S. cerevisiae (YSC2, type II) E. coli KO11

India China India Indonesian South Korea Japan Vietnam Vietnam Vietnam Brazil Malaysia Republic of Korea

14.89 g/L 7.9% 2.06% 1.7 g/L 0.13e0.38 g/g 55.0 g/L 0.83 g/L 0.5 g/L 0.6 g/L 38e64.3 g/L 0.27 g/g 1.4 g/L

[19] [61] [14] [39] [65] [66] [16] [16] [16] [20] [27] [71]

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Fig. 1. List of red seaweeds collected from Mandapam and Thondi. (a) Gracilaria corticata, (b) Gracilaria crassa, (c) Gracilaria edulis.. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

sulphuric acid such as 0.1, 0.5 and 1% (v/v) were used. Different concentrations of Gracilaria corticata spent biomass (2, 4, 6, 8 and 10% w/v) were transferred to 10 mL of different concentrations of sulphuric acid. Then the mixture was autoclaved at 121
C for 15 min. The hydrolysate was analysed for reducing sugar using the DNS method [35]. 2.4. Analysis of furfural in the hydrolysate of SSB The hydrolysate was prepared as per the procedure mentioned in section 2.3. The hydrolysate obtained was analysed for furfural. Furfural formed in the hydrolysate was detected using UVevisible spectrophotometer absorbance at 284 nm [43]. Standard furfural was prepared at different concentrations such as 1, 2, 3, 4 and 5% (v/ v) and the absorbance was read at 284 nm and plotted a graph along with hydrolysate samples. 2.5. Preparation of ethanol production medium Pretreatment of the spent biomass of Gracilaria sp. was hydrolysed using mild sulphuric acid (1% v/v). Spent seaweed biomass (4% w/v) was used for hydrolysis. The mixture was sterilized at 121
C for 15 min [16]. After cooling to bearable heat, the hydrolysate was filtered through a muslin cloth and the pH was adjusted to 6e6.5 and stored overnight at 4
C in a separating flask for unwanted solids to settle at the bottom. Then if needed centrifuged the hydrolysate to remove the few unwanted solids. After removal of unwanted solids in the hydrolysate, it was added with supplementary nutrients (g/L) such as urea (1), sodium hydrogen phosphate (0.5), potassium dihydrogen phosphate (2.5), magnesium sulphate (1), ammonium sulphate (1), ferrous sulphate (0.001), and yeast extract (0.5) according to Ge et al. [12]. Zinc sulphate (10 mg/ L) was also added to production media as a stimulant for ADH (adenosine dehydrogenase) production. ADH supports the sugar consumption and ethanol production during fermentation by yeast [44] and helps in the regeneration of NADþ by alcohol dehydrogenase while fermentation, which requires zinc as an essential cofactor for its activity [45]. After pretreatment and the addition of supplementary nutrients, 70 mL of sterile hydrolysate (triplicate) was used for ethanol production. All the experiments were performed in triplicates. 2.6. Preparation of yeast inoculum Commercially available active dry yeast (Baker’s yeast) was purchased from Shri Kamakshi products, Chennai, India and MTCC yeast 180 (Saccharomyces cerevisiae) was procured from IMTECH, Chandigarh, India. Yeast peptone and dextrose (YPD) medium (HiMedia) was prepared for 100 mL and sterilized at 121
C for

15 min About100 mg of dry yeast pellets was added to the sterilized YPD liquid medium at the light warm condition to trigger the growth of yeast and incubated at 30e33
C for 24 h. The grown yeast was subcultured in petriplates containing potato dextrose agar (PDA) by streak plate method and incubated at 30e33
C in for 48 h. The well-grown yeast culture was stored at 4
C for further studies. The yeast culture was subcultured every 30-day interval in PDA media. For batch culture, 12e15 h grown culture (log phase) of 10% (v/v) yeast inoculum was added to the seaweed extract with supplementary nutrients, and the temperature was maintained at 30 and 35
C for ethanol production in different pH in all the experiments. 2.7. Optimization of fermentation process In this experiment, ethanol production was optimized using different pH (4.5, 5.0, 5.5 and 6.0) and temperatures (30 and 35
C). The experiment was done in batch fermentation [46]. Batch fermentation details are explained in Table 2. 2.7.1. Optimization of pH Different pH ranges, 4.5, 5.0, 5.5 and 6.0, were tested for optimization of ethanol production. The yeasts such as baker’s yeast (Batches 1 and 2) and MTCC yeast (Batches 3 and 4) were used in this experiment for ethanol production. The fermentation process was carried out in 100 mL vials with the working volume of 70 mL. The pretreated hydrolysate (filtered and sterilized) was used for ethanol production after adjusting the pH to 4.5, 5.0, 5.5 and 6.0. The supplementary nutrients were added after filtration and pH adjustment, but before sterilization at 121
C for 15 min. Before inoculation of yeast into the vials, the sterilized hydrolysate was again filtered using muslin cloth (500 mesh size) under sterile condition after which 70 mL of fermentation medium was filled in 100 mL vials with 10% (v/v) of yeast inoculum (baker’s yeast and MTCC yeast). The vials were then sealed using rubber cork and crimps and incubated at 30
C (Batches 1 and 3) and 35
C (Batch 2) in an incubator cum shaker at 100 rpm. About 2 mL of fermented samples were withdrawn every 24 h interval up to 144 h from vials using a sterile syringe. The yeast growth was monitored using UVevisible spectrophotometer recording absorbance at 600 nm [47]. Reducing sugar and ethanol yield was calculated for every 24 h interval up to 144 h. The CO2 produced during the process was removed while a taking sample for analysis using a sterile syringe needle. 2.7.2. Optimization of temperature Two different temperature studies were performed at 30
C (Batches 1 and 3) and 35
C (Batch 2) to standardize the ethanol production at pH 4.5, 5.0, 5.5 and 6.0 in 100 rpm using baker’s yeast

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459

Table 2 Experimental set up for the ethanol optimization. Batch No.

Substrate

Hydrolysate volume (mL)

Yeast

pH

Temperature (oC)

1 2 3 4 5 6

SSB SSB SSB SSB YPD and D-Galactose (Standard) YPD and D-Galactose (Standard)

70 70 70 300 70 70

Baker’s Baker’s MTCC MTCC Baker’s MTCC

4.5, 5.0, 5.5, 6.0 4.5, 5.0, 5.5, 6.0 4.5, 5.0, 5.5, 6.0 5.5 5.5 5.5

30 35 30 30 30 30

(10% v/v) and MTCC yeast (10% v/v). Ethanol fermentation was carried out in 100 mL glass vials with a working volume of 70 mL. Fermented samples (2 mL) were withdrawn from vials using sterile syringe at every 24 h interval. The yeast growth (OD at 600 nm), reducing sugar and ethanol yield was calculated for every 24 h interval up to 144 h. The CO2 produced during the process was removed while the taking sample for analysis using a sterile syringe needle. The processing of fermentation medium preparation was followed as mentioned in section 2.5. All the experiments were performed in triplicates. 2.7.3. Scale-up production of ethanol The spent biomass of Gracilaria corticata (4% w/v) was used for this study. In this study, 500 mL glass vials containing 300 mL of hydrolysates were fermented with 10% (v/v) MTCC yeast (Batch 4) and incubated at 30
C under shaking condition (100 rpm). Yeast growth, reducing sugar and ethanol yield were calculated every 24 h period up to 6 days. In this experiment, MTCC yeast was used based on the previous batch 3 experimental results. 2.8. Ethanol production using standard substrate (YPD and DGalactose) In this experiment, to determine the efficiency of baker’s yeast (Batch 5) and MTCC yeast (Batch 6), a standard substrate such as YPD (contains dextrose 4% w/v) and D-Galactose (4% w/v) were used for ethanol production. The pH of the standard substrate was adjusted to 5.5 using 1 N HCl. The original pH values of YPD and DGalactose media were 6.6 and 6.0, respectively. The composition of YPD was yeast extract (1% w/v), peptone (2% w/v) and dextrose (4% w/v) and D-galactose (4% w/v) medium supplemented with nutrients as mentioned in section 3.7.4. Fermentation of standard sugars (YPD and D-Galactose) was carried out using baker’s yeast and MTCC yeast. The temperature at 30
C and 100 rpm was maintained throughout the experiment. The experiment was conducted in a 100 mL vial with 70 mL working volume Yeast growth, reducing sugar and ethanol yield were monitored and calculated for every 24 h interval. 2.9. Estimation of ethanol yield Ethanol yield was calculated by retrieving 2 mL of sample and centrifuging to remove the yeast cells. The resultant supernatant containing ethanol was analysed using Gas chromatography [48] (Chemito GC-7610) (Table 3). For every 24 h, fermented samples (1 mL) were withdrawn and analysed for ethanol yield. The standard ethanol (99.99% purity) was used for calibration. Standard ethanol graph was plotted for each batch and the ethanol yield of each batch was calculated based on the standard graph. Formulae for ethanol calculation are as follows [27,41,49]. Ethanol concentration ¼ [ethanol produced (g)/volume of reaction mixture (L)]

Table 3 Gas chromatography specifications for ethanol analysis. Parameters

Specifications

Column Oven temperature (oC) Injector temperature (oC) Detector temperature (oC) Carrier gas H2 Air Sample volume

10% SE 30 100 130 200 N2 (1 bar pressure) 2.0 bar pressure 1.2 bar pressure 1 mL

Ethanol yield ¼ [ethanol produced (g)/weight of substrate (g)] Specific ethanol yield ¼ [ethanol produced (g)/sugar consumed (g)] Ethanol yield (g/g) ¼ [ethanol concentration (g/L)/initial concentration of sugars (g/L)]

2.9.1. Statistical analysis All the experiments were conducted in triplicates and the data were analysed statistically using software SPSS 14 and one-way ANOVA was performed to interpret the result with standard deviation using Tukey’s B method. 3. Results and discussion 3.1. Physico-chemical analysis of Gracilaria sp. After extraction of phycoerythrin from seaweeds, the SSB obtained was characterized for its physicochemical properties. The highest percentage of spent biomass was obtained from G. corticata (12.53 ± 2.66%) followed by G. edulis (10.03 ± 2.28%). The least amount of spent biomass was obtained in G. crassa (3.8 ± 0.86%). The shade dried spent seaweed biomass had a moisture content of 14 ± 1% and the proximate, ultimate and biochemical constituents of spent biomass are presented in Table 4. The ash content (14.86 ± 1.47% DW), volatile matter (70.5 ± 1.04% DW) and total organic carbon (25.08 ± 4.15% DW) content of spent biomass was recorded in G. corticata SSB. The ash content was found less in this study when compared to Baghel et al. [50] and reported 43.18 ± 1.15% DW in Gracilaria crassa. The difference in ash content in this study was due to moisture content and the nature of seaweeds and also reported the moisture content of 7.46 ± 0.3% DW in G. crassa by Baghel et al. [50]. The higher ash content in seaweeds was reported to contain microelements important for human and animal nutrition. Wong and Cheung [51] reported ash content in Hypnea charoides and Hypnea japonica that ranged from 21.3% to 22.8% DW. In Gracilaria conforvoides and Gracilaria foliifera, the ash content was found to be 31% and 38%, respectively. In some cases, the ash figures may be slightly low owing to the presence of volatile inorganic salts

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Table 4 Proximate, ultimate, total organic matters, biochemicals and trace elements analysis of SSB of G. corticata. Parameters

Quantity pH

7.0 ± 0.2

Proximate (% DW)

Moisture Ash Volatile matter

14 ± 1 14.86 ± 1.47 70.5 ± 1.04

Total organic matters ((% DW)

Total organic carbon Total carbon Inorganic carbon

25.08 ± 4.15 25.13 ± 4.17 0.05 ± 0.04

Ultimate (% DW)

Carbon Hydrogen Nitrogen Sulphur Oxygen

32.77 ± 0.05 5.64 ± 0.91 1.33 ± 0.35 1.86 ± 0.05 43.56 ± 1.25

Biochemicals (mg/g DW)

Total sugar Reducing sugar Total protein

440 ± 40 129.85 ± 10.23 11.08 ± 0.11

Trace elements (mg/Kg FW)

Arsenic Cadmium Copper Lead Mercury Zinc

BDL (DL:0.1) BDL (DL:0.5) 0.57 BDL (DL:0.5) BDL (DL:0.1) 4.26

Values are expressed as the mean ± SD (n ¼ 3). BDL e Below Detection Limit; DL e Detection Limit.

reported by Ross [52]. Bae et al. [33] reported ash (10.44% DW) and volatile matter (69.66% DW) composition of Porphyra sp., and in this study, the volatile matter in spent biomass also corroborates with Porphyra sp. Norziah and Ching [53] reported ash content of 22.7% and total protein of 6.9% on a wet weight basis in Gracilaria changgi. In this study, the protein content of spent biomass of Gracilaria sp. was calculated as 11.08% DW. The C, H, N, S and O contents of spent biomass were found to be 32.77 ± 0.05%, 5.64 ± 0.91%, 1.33 ± 0.35%, 1.86 ± 0.05% and43.56 ± 1.25% DW, respectively. The total sugar (440 ± 40 mg/g DW), reducing sugar (129.85 ± 10.23 mg/g DW) and protein (11.08 ± 0.11 mg/g DW) content of spent seaweed biomass was recorded (Table 4). The ultimate analysis such as C, H, N, S and O was found low in Gracilaria corticata spent biomass when compared to the study by Ross [52] that reported Nitrogen (3.8% and 2.6%) and Sulphur (2.8% and 4.5%) from Gracilaria conforvoides and Gracilaria foliifera, respectively. Bae et al. [33] reported C, H, N, S and O of Porphyra sp. were found to be 40.6%, 4.65%, 6.13%, 1.26% and 47.4%, respectively, on ash free basis and C and N % found less in this study may be due to seasonality and biomass nature. Oxygen, H and S were recorded high in this study when compared to Bae et al. [33] on ash-free basis. Baghel et al. [50] reported the C, H, N and S contents of G. crassa were 31.75 ± 1.62, 5.18 ± 0.64, 0.83 ± 0.11 and 1.56 ± 0.01%, respectively, on a DW basis. The sugar present in the spent biomass upon pigment extraction undergoes some chemical changes due to heat on over grinding, buffer interference and seasonal variation. 3.2. Characterization of SSB using FT-Raman spectroscopy After acid hydrolysis, the spent hydrolysate was analysed through FT-Raman spectroscopy and determined for its sugar properties. FT-Raman spectra for agar, D-Galactose and spent biomass hydrolysate of G. corticata are presented in Figs. 2e4 respectively. The FT-Raman spectrum for standard commercial grade agar recorded strong bands at 740.8 cm1 and 771.9 cm1 that were assigned to the skeletal bending of the galactose ring. In

addition, strong bands at 843.7 cm1 and 891.5 cm1 were mainly associated with the CH vibration coupled with CeOH mode in methyl 3,6-anhydro- D-galactose residues with an a-configuration and CeH bending at the anomeric carbon in b-galactose residues, respectively. The weak band at 937.5 cm1 indicated the presence of 3,6-anhydo-D-galactose. The most prominent band at 1080.4 cm1 in the FT-Raman spectrum of agar has several vibrational modes, namely the CeO and CeC stretchings and the CeCeO, CeOeC and CeOeH deformations. The FT-Raman spectrum of acid treated hydrolysate of red seaweed spent biomass recorded bands at 748 and 781 cm1 and standard D-Galactose at 764 cm1 representing skeletal bending of the galactose ring (Table 5). The bands at 888.21 cm1 and 892.5 cm1 of D-galactose and RS indicates the CeH bending in galactose, respectively. The bands at 956 cm1 and 927 cm1 of D-galactose and RS indicates the presence of 3,6anhydo-D-galactose, respectively. Other weak centered peaks at 1067.5 and 1073.4 cm1 of D-galactose and RS, respectively, contribute to several stretching of CeO and CeC and deformations of CeCeO, CeOeC and CeOeH. The bands at 1413, 1402.9 and 1400 cm1 in agar, D-Galactose and RS, respectively, represent C6 vibration of galactopyranose rings. Some weak bands at 1150 and 1370 cm1 correspond to SO2 4 in agar and D-Galactose samples, but not observed in acid treated RS sample. The available sulphate in the RS sample may be precipitated while adjusting pH with sodium hydroxide. The spectra of the graph revealed that the functional group of standard substrate matches with seaweed hydrolysate and also corroborates with the findings of Pereira et al. [54] confirming the presence of galactose units (bands 740 and 770 cm1). The presence of galactose in the hydrolysate favours ethanol production. Matsuhiro [55] and Pereira et al. [54] also reported 837 cm1 band associated with the CH vibration attached with CeOH related modes in methyl 3,6-anhydro-D-galactose residues with a-configuration. Bands at 890 and 930 cm1 correspond to b-galactose and 3,6-anhydro-D-galactose residues, respectively. The most prominent band at 1079 cm1 in FT-Raman contributes several vibrational modes, especially CeO and CeC stretchings and to the
mez-Ordo
n ~ ez and CeCeO and CeOeH deformations [54]. Go
rez [56] also reported the FT-IR spectrum for agar that Rupe corroborated with the present study with respect to different wave numbers such as 740, 771.9, 891.5, 937.5 and 1252.5 cm1 observed in the FT-Raman spectrum. 3.3. Pretreatment of SSB using different acids Among the 3 different acids (sulphuric acid, hydrochloric acid and phosphoric acid) were used for pretreatment of SSB. Sulphuric acid at 1% (v/v) was recorded for the maximum yield of reducing sugars (0.64 ± 0.03 g/g dry weight) than 1% HCl (v/v) (0.52 ± 0.06 g/ g dry weight) and 1% H3PO4 (v/v) (0.15 ± 0.02 g/g dry weight) (Table 6). Increase in concentration of all the 3 acid percentages such as 2, 3, 4 and 5% (v/v) for hydrolysis favourably increased reducing sugar content. At 4% (v/v) acid concentration there was no significant difference in sugar concentration in the H2SO4 (0.76 ± 0.14 g/g) and HCl (0.79 ± 0.16 g/g) treated at 121
C for 15 min. Increase in the concentration of reducing sugars was observed while using 4% acid, meanwhile, the HMF also increased due to acids at higher concentration and heating. The effect of acid hydrolysis on different concentrations of SSB was analysed; an experiment was conducted with different concentrations of acid and substrate. In that experiment, 1% sulphuric acid and 4% spent biomass were found effective for hydrolysis to produce monosugars at 121
C for 15 min. According to Kim et al. [40], H2SO4 found to be effective for hydrolysis and it is widely utilized in the industry for hydrolysis process. Longer reaction time

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461

Fig. 2. FT- Raman spectrum of agar standard.

Fig. 3. FT- Raman spectrum of D-Galactose standard.

for hydrolysis was ineffective in producing polysaccharides, taking into account energy and cost, and might lead to the acceleration of sugar degradation [57]. After 30 min, there were no significant differences in the amount of reducing sugars compared to the 15 min samples reported by Choi et al. [57]. Hence, in this study also the same parameters were followed for hydrolysis of SSB. Meinita et al. [39] reported the highest galactose yield (23.87 ± 0.73 g/L) during H2SO4 hydrolysis and was obtained at a K. alvarezii concentration, H2SO4 concentration, temperature and reaction time of 10%, 0.2 M, 130
C and 15 min, respectively. However, the highest formation of galactose(12.72 ± 1.91 g/L) in HCl hydrolysis was obtained by using 10% K. alvarezii, 0.2 M HCl, 130
C and 15 min. Sunwoo et al. [58] reported the monosaccharide content of 29 g/L in 8% sun dried Gracilaria verrucosa using 70 mM H2SO4 for 90 min at 121
C. The acid hydrolysis differs for different biomass, and the concentration, temperature and time will vary for sugar extraction. Kim et al. [40] reported that 0.1 N H2SO4 hydrolysis yielded (7.47 g/

L) more reducing sugars than 0.1 N HCl (6.99 g/L) in Gracilaria verrucosa. The effect of these degraded compounds on cell growth and biofuel production, the presence of by-products are known to decrease the bioenergy production. Some previous studies have examined the inhibition effect of by-products during acid hydrolysis. Alves et al. [59] reported that 1 g/L of HMF might be sufficient to inhibit cell growth and ethanol production by S. cerevisiae. Larsson et al. [60] reported that acetic acid, formic acid, and levulinic acid concentrations below 100 mM/L increased the ethanol yield, whereas high concentrations decreased ethanol productivity. 3.4. Furfurals estimation in the hydrolysate The hydrolysis of spent seaweed biomass produced not only reducing sugars, but also HMF and furfurals. These furfurals hinder the growth of yeast during the fermentation process. The absorbance of the standard furfurals was compared with seaweed

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Fig. 4. FT-Raman spectrum of SSB hydrolysate of G. corticata.

Table 5 FT-Raman spectra of standard substrates and spent biomass. Wave number (cm1) Substrate

1

2

3

4

5

6

7

8

9

10

Agar D-Galactose RS

740.8 703.3 748.5

771.9 764.3 781.6

843.7 828.9 827.6

891.5 888.21 892.5

937.5 956.3 927.7

1080.4 1067.5 1073.4

1252.5 1247.0 1248.0

1286.6 1307.02 1280.0

1413.0 1402.9 e

1472.2 1487.8 e

RSe Red seaweed spent biomass hydrolysate.

Table 6 Pretreatment of G. corticata (4% DW) using various concentration of different acids. Acids

Concentration of acid (%) 1

2

3

4

5

0.67 ± 0.02b 0.65 ± 0.02b 0.28 ± 0.11a

0.65 ± 0.02ab 0.75 ± 0.08b 0.36 ± 0.12a

0.76 ± 0.14a 0.79 ± 0.16a 0.35 ± 0.05b

0.76 ± 0.15b 0.62 ± 0.03ab 0.31 ± 0.06a

Reducing sugar (g/g) H2SO4 HCl H3PO4

0.64 ± 0.03b 0.52 ± 0.06b 0.15 ± 0.02a

Values are expressed as the mean ± SD (n ¼ 3). Values in the same column having the same letter are not significant (P < 0.05).

hydrolysate absorbance read at 284 nm. The furfural content recorded in the H2SO4, HCl and H3PO4 hydrolysate at all concentration of acids was calculated based on the absorbance of standard furfural and recorded to be  1% furfural in all the hydrolysate (Fig. 5a). Low reducing sugar may be due to the decomposition of galactose to other chemicals such as HMF or levulinic acid. The level of HMF will increase gradually in the hydrolysate while an increase in acid concentration. HMF will hinder the growth of yeast and lowers ethanol production. The seaweed species, reaction time, and acid concentration affect the amount of reducing sugars produced (Fig. 5b). 3.5. Optimization of spent seaweed biomass hydrolysis (SSB) using H2SO4 In order to extract simple sugars from SSB, pretreatment and

hydrolysis were carried out using different concentrations of sulphuric acid (0.1, 0.5 and 1% v/v). Among the various concentrations, sulphuric acid (1% v/v) hydrolysis yielded the highest amount of reducing sugar (0.74 g/g dry weight). Increase in concentration of substrate, hydrolysis with 1% acid produced less reducing sugar (Table 7). The acid concentration at 1% with 4% spent biomass ratio was used throughout the fermentation study. The concentration of reducing sugar in spent seaweed substrate varied in the samples based on the period of collection and type of seaweed species. 3.6. Ethanol production using Baker’s yeast 3.6.1. Ethanol production using SSB hydrolysate incubated at 30
C at different pH (Batch 1) In this study, ethanol was produced using spent hydrolysate (treated with 1% H2SO4) fermented with baker’s yeast and incubated at 30
C adjusted with different pH (4.5, 5.0, 5.5 and 6.0). The

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0.14 ± 0.07, 0.24 ± 0.03 and 0.11 ± 0.03 at pH 4.5, 5.0, 5.5 and 6.0, respectively. The growth of yeast was recorded at every 24-h interval. An increase in the growth of yeast was recorded at 24 h. After 24 h there were decreases in the growth of yeast at all the pH such as 4.5, 5.0, 5.5 and 6.0. After 48 h, a steady increase in growth was again observed at all the pH (Fig. 6e). The reducing sugar gradually decreased at all the four pH tested. On the 6th day, the reducing sugar was calculated at all the four different pH (4.5, 5.0, 5.5 and 6.0) and was recorded 3.23 ± 0.22 g/L, 2.70 ± 0.32 g/L, 3.06 ± 0.14 g/L and 2.48 ± 0.22 g/L, respectively (Fig. 6aed). The ethanol produced by baker’s yeast was calculated at every 24-h interval and found that the highest production of ethanol (1.78 ± 0.21 g/L) at pH 6.0 was recorded at 96 h followed by pH 4.5 (1.76 ± 0.12 g/L) (Fig. 6a and d). Whereas at pH 5.0 and 5.5, the ethanol yield was found to be 1.61 ± 0.10 g/L and 1.65 ± 0.04 g/L at 144 h and 96 h, respectively (Fig. 6b and c). The amount of reducing sugar consumed by baker’s yeast at pH 6.0 was 5.12 g/L on the 4th day and 8.21 g/L on 6th day from initial reducing sugar (10.69 ± 0.87 g/L) (Fig. 6d).

Fig. 5. (a) Furfural and (b) reducing sugar contents in the acid hydrolysate of SSB.

Table 7 Optimization of sulphuric acid and SSB substrate used for hydrolysis. SSB (%)

2% 4% 6% 8% 10%

Reducing sugar (g/g) 0.1% acid

0.5% acid

0.08 ± 0.02c 0.05 ± 0.01bc 0.03 ± 0.00ab 0.01 ± 0.00a 0.004 ± 0.002a

0.40 0.64 0.37 0.26 0.19

± ± ± ± ±

0.03b 0.07c 0.03b 0.01a 0.01a

1% acid 0.51 0.74 0.52 0.29 0.29

± ± ± ± ±

0.01b 0.01c 0.02b 0.07a 0.02a

Values are expressed as the mean ± SD (n ¼ 3). Values in the same column having the same letter are not significant (P < 0.05).

initial reducing sugar of the hydrolysate was 10.72 ± 0.40 g/L, 10.90 ± 0.38 g/L, 10.44 ± 0.63 g/L and 10.69 ± 0.87 g/L at pH 4.5, 5.0, 5.5 and 6.0, respectively. The yeast growth was taken at 600 nm using a spectrophotometer. The initial growth of pre-cultured yeast (15-h old culture) was 1.16 (OD value). The initial OD (0 h) of yeast after inoculation in the fermentation broth was 0.16 ± 0.04,

3.6.2. Ethanol production using SSB hydrolysate incubated at 35
C at different pH (Batch 2) In this experiment, ethanol was produced using baker’s yeast at 35
C. The growth of yeast at 600 nm was 2.32 (OD) at 15 h. The initial growth (OD at 600 nm) of baker’s yeast was 1.15 ± 0.17, 1.50 ± 0.65, 1.38 ± 0.34 and 1.43 ± 0.59, respectively at pH 4.5, 5.0, 5.5 and 6.0 (Fig. 7e). There is a steady increase in growth from 0 h to 24 h and then stable growth was observed throughout the experiment up to 96 h. The reducing sugar was calculated every 24-h interval. The initial reducing sugar was 4.55 ± 0.26 g/L, 4.02 ± 0.05 g/L, 4.36 ± 0.43 and 4.17 ± 0.28 g/L at pH 4.5, 5.0, 5.5 and 6.0, respectively (Fig. 7aed). Gradually the reducing sugar decreased at 24 h but increased at 48 h till 96 h. This may be due to the production of substrate degrading enzymes by the yeast during the fermentation process. This was observed in the pH study for fermentation. The ethanol production observed maximum at pH 4.5 in 24 h was 1.94 ± 0.18 g/L (Fig. 7a) followed by pH 6.0, 5.5 and 5.0 calculated as 1.85 ± 0.33, 1.64 ± 0.10 and 1.52 g/L, respectively in 24 h (Fig. 7bed). The maximum ethanol was produced within 72 h at pH 5.0 (1.94 ± 0.09 g/L) and pH 5.5 (1.82 ± 0.14 g/L). But at pH 6.0, the maximum ethanol was 1.87 g/L in 96 h (Fig. 7d). The amount of sugar consumed by the baker’s yeast was 1.78, 0.46, 0.19 and 0.38 g/ L, respectively, at the pH 4.5, 5.0, 5.5 and 6.0 (Fig. 7aed). Wang et al. [61] reported ethanol yield of 8 g/100 g DW by 2stage hydrolysis of Gracilaria salicornia. The ethanol yield was 61% less in this study due to one-step hydrolysis and the yeast used for fermentation. On the other hand, a high concentration of strong acid pretreatment converts the glucose molecule to hydroxymethylfurfural (HMF) which inhibits the fermentation process. Further steam pretreatment produces furfural, HMF, and soluble phenolic compounds which can inhibit the ethanol fermentation. The presence of 3,6-anhydrogalactose in the galactan hydrolysis, a notable amount of 5-hydroxymethylfurfural (yield 0.7e21.8%) was formed during the decomposition in a soluble state [62]. In this study, the maximum amount of reducing sugars released while using 1% sulphuric acid and the yield is low when compared to other studies due to the formation of inhibitor compounds during pretreatment [12]. Utilization of glucose for ethanol production is well known, whereas another monosaccharide such as galactose from Gracilaria sp. for ethanol production has not been fully explored yet. This is because glucose is directly consumed by yeast through the glycolytic pathway while galactose needs intermediate pathway prior to the glycolytic pathway for conversion [63].

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Fig. 6. (aee). Growth, reducing sugar and ethanol yield by yeast (Baker’s) using Gracilaria corticata spent biomass incubated at 30
C in different pH (Batch 1). (a) pH 4.5, (b) pH 5.0, (c) pH 5.5, (d) pH 6.0 and (e) Growth of yeast.

3.7. Ethanol production using MTCC Yeast 180 3.7.1. Ethanol production using SSB hydrolysate incubated at 30
C at different pH (Batch 3) In this batch Saccharomyces cerevisiae (MTCC yeast 180) was used for ethanol production. The spent biomass of Gracilaria corticata was used for hydrolysis using 1% H2SO4. The hydrolysate was fermented for 6 days, and at every 24-h interval, reducing sugar and ethanol yield was calculated.

The growth of yeast was monitored periodically at 600 nm. The OD of the 15-h grown MTCC yeast was 2.20. The initial (0 h) OD of MTCC yeast was found to be 0.18 ± 0.06, 0.52 ± 0.26, 0.46 ± 0.20 and 0.52 ± 0.19 at the pH 4.5, 5.0, 5.5 and 6.0, respectively (Fig. 8e). Increase in growth of MTCC yeast at 24 h was observed, after which yeast growth was stable throughout the experiment. There is no much increase in growth was observed. The reducing sugar content in the hydrolysate gradually decreased from 11.91 ± 1.29 g/L, 11.91 ± 0.58 g/L, 12.37 ± 0.39 g/L

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Fig. 7. (aee). Growth, reducing sugar and ethanol yield by yeast (Baker’s) using Gracilaria corticata spent biomass incubated at 35
C in different pH (Batch 2). (a) pH 4.5, (b) pH 5.0, (c) pH 5.5, (d) pH 6.0 and (e) Growth of yeast.

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Fig. 8. (aee). Growth, reducing sugar and ethanol yield by yeast (MTCC) using Gracilaria corticata spent biomass incubated at 30
C in different pH (Batch 3). (a) pH 4.5, (b) pH 5.0, (c) pH 5.5, (d) pH 6.0 and (e) Growth of yeast.

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and 11.35 ± 0.33 g/L to 7.30 ± 0.55 g/L, 7.72 ± 0.82 g/L, 6.93 ± 0.54 g/ L and 8.01 ± 0.88 g/L, respectively, in 24 h period (Fig. 8aed). Later, on the consumption of sugar by the yeast is found less for the remaining days. The ethanol yield was calculated and the highest was 1.99 ± 0.07 g/L and 1.97 ± 0.09 g/L at 48 h and 72 h at the pH 4.5 and 6.0 (Fig. 8a and d). Whereas, the ethanol yield was found to be 1.93 ± 0.07 g/L and 1.92 ± 0.15 g/L for the pH 5.0 and 5.5 at 144 h and 48 h, respectively (Fig. 8b and c). Slow consumption of sugar was observed at pH 5.0 (Fig. 8b). In order to produce a higher concentration of ethanol, sugars in the hydrolysate should be increased. An increase in the concentration of seaweeds in the acid treatment is not feasible for fermentable sugars production. In addition, dilute-acid pretreatment appears to be ineffective for hydrolysis of seaweed glucans for biofuel production. Concentrating the hydrolysate through evaporation and repeated hydrolysis of spent seaweeds with the first hydrolysate will favour the fermentation process to yield a high concentration of ethanol. Other than acid hydrolysis an alternate method will be necessary to produce a high concentration of fermentable sugars from red spent seaweeds such as enzymatic or both acids cum enzymatic treatment may favour the fermentation process for ethanol production. Moreover, 3,6-anhydrogalactose contained in the red spent seaweeds hydrolysate was not converted to ethanol [64e66]. In order to get maximum yeast growth and ethanol production, the pH of the medium plays a vital role. Kim et al. [67] patented ethanol production from galactose and glucose extracted from G. amansii using S. cerevisiae and reported that S. cerevisiae did not consume all the mixed sugars even after 48 h of fermentation. It consumes glucose first then galactose for metabolism and tolerates only 20% of sugar in the medium composition. 3.7.2. Scale-up production of ethanol from SSB hydrolysate using yeast (MTCC) incubated at 30
C at pH 5.5 (Batch 4) In this study, ethanol production was scaled up to 300 mL cycle. The growth (OD at 600 nm) of yeast was 2.36 (15 h grown yeast). The yeast growth (OD) after inoculation at 0 h was 0.79 ± 0.08 (Fig. 9a). Gradually, the growth of yeast was recorded in the seaweed extract throughout the experiment. The reducing sugar was 9.42 ± 1.05 g/L initially in the hydrolysate and gradually decreased to 4.04 ± 0.09 at 144 h at pH 5.5 (Fig. 9b). The sugar consumption was 5.38 g/L for 6 days. The ethanol yield (1.50 ± 0.08 g/L) and sugar consumed (1.58 g/L) were recorded at 72 h (Fig. 9b). 3.7.3. Ethanol production using standard substrate (YPD and DGalactose) and Baker’s yeast (Batch 5) A Standard substrate such as YPD (Yeast extract, Peptone and Dextrose) and D-galactose was used for ethanol production. The growth was monitored at 600 nm as OD. The initial yeast inoculum OD was 0.9 and 0.92 for YPD BY and Gal BY, respectively. The initial reducing sugar was 0.87 ± 0.01 and 0.79 ± 0.01, respectively at 0 h. The growth increased to 2.45 ± 0.01 (YPD BY) and 2.24 ± 0.12 (Gal BY) in 24 h and 2.50 ± 0.03 (YPD BY) and 2.42 ± 0.02 (Gal BY) in 48 h (Fig. 10e). The initial sugar concentration was 40.20 ± 0.16 g/L used for fermentation (Fig. 10a and c). The reducing sugar concentration was analysed every 24-h interval and was found to be 0.25 ± 0.10 g/ L and 1.37 ± 1.22 g/L for YPD and D-galactose, respectively. The ethanol concentration was analysed every 24-h interval. The ethanol yield was 17.44 ± 1.03 g/L (YPD BY) and 15.66 ± 1.84 g/L (Gal BY) at 24 h. The ethanol yield increased to 18.42 ± 0.96 g/L at 48 h in YPD BY medium (Fig. 10a). The ethanol yield decreased to 15.49 ± 1.93 g/L in Gal BY medium at 48 h (Fig. 10c). The sugar consumption was found maximum in both the medium at 30
C. These results showed that baker’s yeast ferments both simple sugars such as dextrose and galactose effectively.

Fig. 9. (a&b). Growth, reducing sugar and ethanol yield by yeast (MTCC) using Gracilaria corticata spent biomass (300 mL) incubated at 30
C (Batch 4). (a) Growth of yeast and (b) Fermentation process at pH 5.5.

3.7.4. Ethanol production using standard substrate (YPD and DGalactose) and yeast (MTCC) (Batch 6) In this batch, standard substrate such as YPD and D-galactose (Gal) was used. The inoculum OD of MTCC yeast in YPD and Dgalactose (Gal) was 2.14 and 2.16, respectively. The initial growth of MTCC yeast after inoculation was 0.92 ± 0.01 and 0.91 ± 0.02, respectively, for YPD MTCC and Gal MTCC in 0 h (Fig. 10e). The yeast growth (OD) increased to 1.94 ± 0.53 and 1.95 ± 0.63, respectively, for YPD and Gal medium in 24 h and 48 h increase in OD was observed as 2.24 ± 0.35 and 2.08 ± 0.60 for YPD and Gal medium respectively. The initial reducing sugar concentration was 38.83 ± 0.58 g/L and 39.26 ± 0.94 g/L in YPD and Gal medium. There was a decrease in reducing sugar concentration observed in 24 h interval and was found to be 0.10 ± 0.05 g/L and 3.26 ± 1.03 g/L in YPD and Gal medium, respectively (Fig. 10b and d). The ethanol yield was calculated as 18.29 ± 1.06 g/L and 15.61 ± 0.78 g/L in 24 h and 48 h, respectively, in YPD medium. The Gal medium ethanol yield was 15.15 ± 0.81 g/L and 17.48 ± 1.08 g/L at 24 h and 48 h interval, respectively (Fig. 10b and d). S. cerevisiae supports only zinc-dependant alcohol dehydrogenase (ADH) gene [68]. The ADH enzyme of S. cerevisiae has zinc

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Fig. 10. (aee). Growth, reducing sugar and ethanol yield by yeast (Baker’s and MTCC) using YPD and Galactose incubated at 30
C (Batch 5 & 6). (a) YPD BY, (b) YPD MTCC yeast, (c) Gal BY, (d) Gal MTCC yeast and (e) Growth of yeast.

binding and induces the activity of ADH enzyme. The dissociation of half (2 zinc atoms/monomer) of the total zinc content of the enzyme is associated with the full inhibition of its activity. Hence, zinc sulphate was used in the spent seaweed extract or YPD or D-

galactose media might have to augment the enzyme alcohol dehydrogenase activity of S. cerevisiae [44,69]. In this study, zinc sulphate (10 mg/L) was added to the fermentation broth whereas Taloria et al. [44] reported that 20 mM zinc sulphate supports the

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ethanol production up to 8% (v/v) in YEPD media. But, in this study, the ethanol yield was achieved maximum of 0.25% (v/v) that may be due to inhibitors present in the fermentation medium (spent seaweed extract). Whereas, in YPD and galactose medium, MTCC yeast inoculated without zinc and the yield of ethanol was 2.32 ± 0.13% (v/v) and 2.22 ± 0.14% (v/v), respectively. The ethanol yield of MTCC yeast 180 was comparable to the randomly UV mutated MTCC yeast strain no. 3786 used by Taloria et al. [44]. The increase in ethanol yield achieved by UV mutation, concentration of zinc sulphate and without any HMF in the production medium (i.e. YPD medium) has been reported by Taloria et al. [44]. Ethanol yield depends on the yeast metabolism of galactose and the enzymes of the Leloir pathway converts galactose to most useful glucose-6-phosphate. This pathway is essentially required as galactose itself cannot be used for glycolysis directly; the yeast, Saccharomyces cerevisiae, requires five enzymes to catalyze this conversion: a galactosemutarotase, a galactokinase, a galactose-1phosphate uridyltransferase, a UDP-galactose-4-epimerase, and a phosphoglucomutase. In yeast, the genes encoding these above enzymes are tightly controlled at the stage of transcription and are only transcribed under specific sets of conditions [70]. 3.8. Ethanol yield and analysis Ethanol yield was calculated at every 24 h interval through gas chromatography. Ethanol yield was expressed in g/L. The ethanol yield was calculated based on the substrate concentration used. 3.8.1. Baker’s yeast and ethanol yield In batch 1, the highest ethanol (4.5% w/w) yield was recorded at pH 6.0 in 96 h and at other pH such as 4.5, 5.0 and 5.5, the ethanol yield was recorded as 4.0, 3.8 and 4.1% (w/w), respectively, in 96 h. However, there was an increase in ethanol yield recorded after 96 h at pH 4.5 and 5.0, but ethanol yield was saturated at pH 5.5. In batch 2, the ethanol yield (4.85% w/w) was recorded high at pH 4.5 in 24 h followed by pH 5.0 (4.85% w/w), 6.0 (4.63% w/w) and 5.5 (4.55% w/w) at 72 h, 24 h and 72 h, respectively. There was no increase in ethanol yield at all pH after 72 h. In batch 5, the standard substrate such as YPD and galactose fermented using Baker’s yeast showed the highest ethanol yield (46.05% w/w) at 48 h in YPD followed by galactose (39.15% w/w) at 24 h in pH 5.5.

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of Gracilaria sp. Overall, the maximum ethanol yield (4.98% w/w) was achieved using yeast procured from MTCC than baker’s yeast (4.5% w/w). Furfurals present in the hydrolysate inhibited the growth of yeast and ethanol production in the present study. Further removal of furfurals from the hydrolysate will yield higher ethanol production. The following ideas are being proposed based on the outcome of this research 1. Naturally grown seaweeds are not sufficient to produce ethanol for throughout the year, instead, commercial cultivation of these seaweeds paves the way to generate revenue and energy in future. 2. Though in this preliminary study the ethanol production from spent biomass of seaweed is less, the utilization aspects of this spent biomass from seaweed industry can be a target for the integrated biofuel production. 3. Further detailed research is needed to expand the commercialization aspect and yield improvement. 4. The cost of production of ethanol will be reduced while using acid þ heat treatment method of sugar extraction used in the present study. Enzymes usage for sugar extraction will be an alternate method; however cost of production will become higher than expected.

Declaration of author’s contributions Dr M.P. Sudhakar contributes to the whole experimental lab work, seaweed collection, sample analysis and writing manuscript with interpretation. Dr K. Arunkumar contributes to seaweed sample collection and identification. Dr K. Perumal contributes to experimental design and manuscript writing. Declaration of competing interest The authors declare that no conflicts, informed consent, human or animal rights applicable. Acknowledgements

3.8.2. Yeast (MTCC) and ethanol yield In batch 3, the ethanol yield was calculated highest in pH 4.5 (4.98% w/w) at 48 h followed by pH 6.0, 5.0 and 5.5 as 4.93%, 4.83% and 4.80% (w/w) at 72 h, 144 h and 48 h, respectively. In this batch, at pH 5.0, the gradual increase in ethanol yield was observed from 24 h to 144 h (4.40%e4.83% w/w). In batch 4, the scaled-up substrate showed maximum ethanol yield (3.75% w/w) at 72 h in pH 5.5, whereas in 24 h, the ethanol yield was 3.65% w/w and gradually increased afterwards. In batch 6, the standard substrate such as YPD and galactose fermented using yeast (MTCC) showed maximum ethanol yield 45.72% w/w and 43.7% w/w in 24 h and 48 h, respectively, at pH 5.5. Based on the above fermentation batches, yeast (MTCC) recorded maximum ethanol production than baker’s yeast incubated at 30
C with pH 4.5 by using the spent seaweed biomass hydrolysate. 4. Conclusion The spent seaweed biomass (SSB) obtained after extraction of phycobiliproteins was characterized, studied its properties and utilized for production of ethanol using Saccharomyces cerevisiae. The method was developed for pretreatment process (using 1% H2SO4 showed better sugar yield) and ethanol production using SSB

Authors thank Shri A.M.M. Murugappa Chettiar Research Centre for providing the laboratory facilities, instruments handling etc. This project (Ref. No.DST/TSG/AF/2010/12) was financially supported by Department of Science and Technology, New Delhi, India. Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi.org/10.1016/j.renene.2020.02.032. References [1] N. Wei, J. Quarterman, Y. Jin, Marine macroalgae: an untapped resource for producing fuels and chemicals, Trends Biotechnol. 31 (2013) 70e77. [2] International Energy Outlook, 2014. https://www.eia.gov/pressroom/ presentations/sieminski_09222014_columbia.pdf. [3] H.M. Kim, C.H. Oh, H.J. Bae, Comparison of red microalgae (Porphyridium cruentum) culture conditions for bioethanol production, Bioresour. Technol. 233 (2017) 44e50. [4] P. Vo Hoang Nhat, H.H. Ngo, W.S. Guo, S.W. Chang, D.D. Nguyen, P.D. Nguyen, X.T. Bui, X.B. Zhang, J.B. Guo, Can algae-based technologies be an affordable green process for biofuel production and wastewater remediation? Bioresour. Technol. 256 (2018) 491e501. [5] C. Herrmann, J. FitzGerald, R. O’Shea, A. Xia, P. O’Kiely, J.D. Murphy, Ensiling of seaweed for a seaweed biofuel industry, Bioresour. Technol. 196 (2015)

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M.P. Sudhakar et al. / Renewable Energy 153 (2020) 456e471

301e313. [6] M.P. Sudhakar, A. Jagatheesan, C. Poonam, K. Perumal, K. Arunkumar, Biosaccharification and ethanol production from spent seaweed biomass using marine bacteria and yeast, Renew. Energy 105 (2017) 133e139. [7] Q. Fu, H. Zhang, H. Chen, Q. Liao, A. Xia, Y. Huang, X. Zhu, A. Reungsang, Z. Liu, Hydrothermal hydrolysis pretreatment of microalgae slurries in a continuous reactor under subcritical conditions for largeescale application, Bioresour. Technol. 266 (2018) 306e314. [8] FAO, Fishery Statistical Collections Global Aquaculture Production, 2010. http://www.fao.org/fishery/statistics/global-aquaculture-production/en. (Accessed 24 September 2010). [9] S. Kraan, Mass-cultivation of carbohydrate rich macroalgae, a possible solution for sustainable biofuel production, Mitig. Adapt. Strategies Glob. Change 18 (1) (2013) 27e46. [10] B. Kılınç, S. Cirik, G. Turan, H. Tekogul, E. Koru, Seaweeds for Food and Industrial Applications, InTech open access, 2013, pp. 735e748, https://doi.org/ 10.5772/53172. [11] P.V. SubbaRao, V.A. Mantri, Indian seaweed resources and sustainable utilization: scenario at the dawn of a new century, Curr. Sci. 91 (2006) 164e174. [12] L. Ge, P. Wang, H. Mou, Study on saccharification techniques of seaweed wastes for the transformation of ethanol, Renew. Energy 36 (2011) 84e89. [13] S.J. Horn, I.M. Aasen, K. Østgaard, Ethanol production from seaweed extract, J. Ind. Microbiol. Biotechnol. 25 (2000) 249e254. [14] Y. Khambhaty, K. Mody, M.R. Gandhi, S. Thampy, P. Maiti, H. Brahmbhatt, K. Eswaran, P.K. Ghosh, Kappaphycus alvarezii as a source of bioethanol, Bioresour. Technol. 103 (2012) 180e185. [15] Y. Khambhaty, D. Upadhyay, Y. Kriplani, N. Joshi, K. Mody, M.R. Gandhi, Bioethanol from macroalgal biomass: utilization of marine yeast for production of the same, Bioenergy Res. 6 (2013) 188e195. [16] M.D.N. Meinita, B. Marhaeni, T. Winanto, G. Jeong, M.N.A. Khan, Y. Hong, Comparison of agarophytes (Gelidium, Gracilaria, and Gracilariopsis) as potential resources for bioethanol production, J. Appl. Phycol. 25 (2013) 1957e1961. [17] M. Kumar, P. Kumari, V. Gupta, C.R.K. Reddy, B. Jha, Biochemical responses of red alga Gracilaria corticata (Gracilariales, Rhodophyta) to salinity induced oxidative stress, J. Exp. Mar. Biol. Ecol. 391 (2010) 27e34. [18] C.S. Goh, K.T. Lee, A visionary and conceptual macroalgae-based third generation bioethanol (TGB) biorefinery in Sabah, Malaysia as an underlay for renewable and sustainable development, Renew. Sustain. Energy Rev. 14 (2010) 842e848. [19] J.M. Adams, T.A. Toop, J.M. Gallagher, I.S. Donnison, Seasonal variation in Laminaria digitata and its impact on biochemical conversion routes to biofuels, Bioresour. Technol. 102 (2011) 9976e9984. [20] P.I. Hargreaves, C.A. Barcelos, A.C. Augusto da Costa, N. Pereira Jr., Production of ethanol 3G from Kappaphycus alvarezii: evaluation of different process strategies, Bioresour. Technol. 134 (2013) 257e263. [21] D. Rodrigues, A.C. Freitas, L. Pereira, T.A. Rocha-Santos, M.W. Vasconcelos, M. Roriz, L.M. Rodríguez-Alcal
a, A.M. Gomes, A.C. Duarte, Chemical composition of red, brown and green macroalgae from Buarcos Bay in central west coast of Portugal, Food Chem. 183 (2015) 197e207. [22] M.N.A.M. Yusoff, N.W.M. Zulkifli, B.M. Masum, H.H. Masjuki, Feasibility of bioethanol and biobutanol as transportation fuel in spark-ignition engine: a review, RSC Adv. 5 (2015) 100184e100211. [23] N.V.S.N. Murthy Konda, S. Singh, B.A. Simmons, D. Karl-Marcuschamer, An investigation on the economic feasibility of macroalgae as a potential feedstock for biorefineries, Bioenerg. Res. 8 (2015) 1046e1056. [24] L.R. Brown, Food or fuel? in: Linda Starke (Ed.), Full Planet, Empty Plates: the New Geopolitics of Food Scarcity E.P. Institute, W.W. Norton & Company, New York, 2012, pp. 36e44. [25] B. Roozbehani, M. Mirdrikvand, S.I. Moqadam, A.C. Roshan, Chem. Technol. Fuels Oils 49 (2013) 115e124. [26] Sugar Industry News. http://www.sugarinds.com/2011/08/worlds-top-20ethanolproducing.html. (Accessed 4 September 2015). [27] I.S. Tan, K.T. Lee, Enzymatic hydrolysis and fermentation of seaweed solid wastes for bioethanol production: an optimization study, Energy 78 (2014) 53e62. [28] M.P. Sudhakar, A. Jagatheesan, K. Perumal, K. Arunkumar, Methods of phycobiliprotein extraction from Gracilaria crassa and its applications in food colourants, Algal Res. 8 (2015) 115e120. [29] M.P. Sudhakar, R. Merlyn, K. Arunkumar, K. Perumal, Characterization, pretreatment and saccharification of spent seaweed biomass for bioethanol production using baker’s yeast, Biomass Bioenergy 90 (2016) 148e154. [30] J.W. Agger, P.J. Nilsen, V.G.H. Eijsink, S.J. Horn, On the determination of water content in biomass processing, BioEnergy Res. 7 (2014) 442e449. [31] ASTM E1755-01, Standard Test Method for Ash in Biomass, American Society for Testing and Materials (ASTM), USA, 2007. [32] ASTM E872-82, Standard Test Method for Volatile Matter in the Analysis of Particulate Wood Fuels, American Society for Testing and Materials (ASTM), USA, 2006. [33] Y.J. Bae, C. Ryu, J.-K. Jeon, J. Park, D.J. Suh, Y.W. Suh, D. Chang, Y.K. Park, The characteristics of bio-oil produced from the pyrolysis of three marine macroalgae, Bioresour. Technol. 102 (2011) 3512e3520. [34] M. Dubois, K.A. Gilles, J.K. Hamilton, P.A. Rebers, F. Smith, Colorimetric method for determination of sugars and related substances, Anal. Chem. 26 (1956) 350.

[35] G.L. Miller, Use of dinitrosalicylic acid reagent for determination of reducing sugars, Anal. Chem. 31 (1972) 426e428. [36] E. Barbarino, S.O. Lourenco, An evaluation of methods for extraction and quantification of protein from marine macro and micro algae, J. Appl. Phycol. 17 (2005) 447e460. [37] O.H. Lowry, N.J. Rosenbrough, A.L. Farr, R.J. Randall, Protein measurement with the folin phenol reagent, J. Biol. Chem. 193 (1951) 265e275. [38] Aoac 17th edition, Official method 999.11, Determination of Lead, Cadmium, Copper, Iron and Zinc in Foods. Atomic Adsorption Spectrophotometry after Dry Ashing, 2000. [39] M.D.N. Meinita, Y. Hong, G. Jeong, Comparison of sulfuric and hydrochloric acids as catalysts in hydrolysis of Kappaphycus alvarezii (cottonii), Bioproc. Biosyst. Eng. 35 (2012) 123e128. [40] S.W. Kim, C.H. Hong, S.W. Jeon, H.J. Shin, High-yield production of biosugars from Gracilaria verrucosa by acid and enzymatic hydrolysis processes, Bioresour. Technol. 196 (2015) 634e641. [41] C. Park, J.H. Lee, X. Yang, H.Y. Yoo, J.H. Lee, S.K. Lee, S.W. Kim, Enhancement of hydrolysis of Chlorella vulgaris by hydrochloric Acid, Bioproc. Biosyst. Eng. 39 (2016) 1015e1021. [42] M.Z.B. Hussein, M.B.B.A. Rahman, A.H.J. Yahaya, T.Y.Y. Hin, N. Ahmad, Oil palm trunk as a raw material for activated carbon production, J. Porous Mater. 8 (2001) 327e334. [43] A. Martinez, M.E. Rodriguez, S.W. York, J.F. Preston, L.O. Ingram, Use of UV absorbance to monitor furans in dilute acid hydrolysates of biomass, Biotechnol. Prog. 16 (2000) 637e641. [44] D. Taloria, S. Samanta, S. Dasa, C. Pututundaa, Increase in bioethanol production by random UV mutagenesis of S. cerevisiae and by addition of zinc ions in the alcohol production media, APCBEE Procedia 2 (2012) 43e49. [45] G.M. Walker, Fuel alcohol: current production and future challenges, J. Inst. Brew. 117 (2011) 3e22. [46] C. Laluce, J.O. Tognolli, K. Fernanda de Oliveira, C.S. Souza, M.R. Morais, Optimization of temperature, sugar concentration, and inoculum size to maximize ethanol production without significant decrease in yeast cell viability, Appl. Microbiol. Biotechnol. 83 (2009) 627e637. [47] L.W. Bergman, Two-Hybrid systems: methods and protocols, Chapter 2: growth and maintenance of yeast, in: P.N. MacDonald (Ed.), Methods in Molecular Biology, vol. 177, Humana Press Inc., Totowa, NJ, 2001, pp. 9e14, https://doi.org/10.1385/1592592104. [48] Y. Gerchman, A. Schnitzer, R. Gal, N. Mirsky, N. Chinkov, A simple rapid gaschromatography flame-ionization detector (GC-FID) method for the determination of ethanol from fermentation processes, Afr. J. Biotechnol. 11 (2012) 3612e3616. [49] C. Pothiraj, R. Arumugam, M. Gobinath, Sustaining ethanol production from lime pretreated water hyacinth biomass using mono and co-cultures of isolated fungal strains with Pichia stipitis, Bioresour. Bioprocess. 1 (2014) 27. [50] R.S. Baghel, P. Kumari, C.R.K. Reddy, J. Bhavanath, Growth, pigments, and biochemical composition of marine red alga Gracilaria crassa, J. Appl. Phycol. 26 (2014) 2143e2150. [51] K.H. Wong, P.C.K. Cheung, Nutritional evaluation of some subtropical red and green seaweeds Part I e proximate composition, amino acid profiles and some physico-chemical properties, Food Chem. 71 (2000) 475e482. [52] A.G. Ross, Some typical analyses of red seaweeds, J. Sci. Food Agric. 4 (1953) 333e335. [53] M.H. Norziah, C.Y. Ching, Nutritional composition of edible seaweed Gracilaria changgi, Food Chem. 68 (2000) 69e76. [54] L. Pereira, A. Sousa, H. Coelho, A.M. Amado, P.J.A. Ribeiro-Claro, Use of FTIR, FT-Raman and 13C-NMR spectroscopy for identification of some seaweed phycocolloids, Biomol. Eng. 20 (2003) 223e228. [55] B. Matsuhiro, Vibrational spectroscopy of seaweed galactans, Hydrobiologia 326e327 (1996) 481e489.
mez-Ordo
n ~ ez, P. Rupe
rez, FTIR-ATR spectroscopy as a tool for poly[56] E. Go saccharide identification in edible brown and red seaweeds, Food Hydrocolloids 25 (2011) 1514e1520. [57] J.I. Choi, H.J. Kim, J.H. Kim, M.W. Byun, B.S. Chun, D.H. Ahn, Y.J. Hwang, D.J. Kim, G.H. Kim, J.W. Lee, Application of gamma irradiation for the enhanced physiological properties of polysaccharides from seaweeds, Appl. Radiat. Isot. 67 (2009) 1277e1281. [58] I.Y. Sunwoo, C.H. Ra, G.T. Jeong, S.K. Kim, Evaluation of ethanol production and bioadsorption of heavy metals by various red seaweeds, Bioproc. Biosyst. Eng. 39 (2016) 915e923. [59] L.A. Alves, M.G.A. Felipe, J.B. Silva, S.S. Silva, A.M.R. Prata, Pretreatment of sugar cane bagasse hemicellulose hydrolysate for xylitol production by Candida guilliermondii, Appl. Biochem. Biotechnol. 70 (1998) 89e98. [60] S. Larsson, E. Palmqvist, B.H. Hagerdal, C. Tengborg, K. Stenberg, G. Zacchi, N. Nils-Olof, The generation of fermentation inhibitors during dilute acid hydrolysis of softwood, Enzym. Microb. Technol. 24 (1999) 151e159. [61] X. Wang, X. Liu, G. Wang, Two-stage hydrolysis of invasive algal feedstock for ethanol fermentation, J. Integr. Plant Biol. 53 (2011) 246e252. [62] M. Robal, K. Truus, O. Volobujeva, E. Mellikov, R. Tuvikene, Thermal stability of red algal galactans: effect of molecular structure and counterions, Int. J. Biol. Macromol. 104 (2017) 213e223. [63] N.Z. Zakaria, D. Arbain, M.N. Ahmad, M.I.H.M. Dzahir, Galactose consuming microbes for ethanol production from seaweed, Adv. Mater. Res. 925 (2014) 219e222. [64] C. Kim, H.J. Ryu, S.H. Kim, J.J. Yoon, H.S. Kim, Y.J. Kim, Acidity tunable ionic

M.P. Sudhakar et al. / Renewable Energy 153 (2020) 456e471 liquids as catalysts for conversion of agar in to mixed sugars, Bull. Kor. Chem. Soc. 31 (2010a) 511e514. [65] J.H. Park, J.Y. Hong, H.C. Jang, S.G. Oh, S.H. Kim, J.J. Yoon, et al., Use of Gelidium amansii as a promising resource for bioethanol: a practical approach for continuous dilute-acid hydrolysis and fermentation, Bioresour. Technol. 108 (2012) 83e88. [66] M. Yanagisawa, S. Kawai, K. Murata, Strategies for the production of high concentrations of bioethanol from seaweeds: production of high concentrations of bioethanol from seaweeds, Bioengineered 4 (2013) 224e235. [67] G.S. Kim, M.K. Shin, Y.J. Kim, K.K. Oh, J.S. Kim, H.J. Ryu, K.H. Kim, Method of Producing Biofuel Using Sea Algae. US Patent Pub. No.: US 2010/0124774 A1.

471

2010b, pp. 1e15. [68] K.F. Mackenzie, C.F. Eddy, L.O. Ingram, Modulation of alcohol dehydrogenase isoenzyme levels in Zymomonas mobilis by iron and zinc, J. Bacteriol. 171 (1989) 1063e1067. [69] V. Leskovac, S. Trivi, M. Latkovska, State and Accessibility of zinc in yeast alcohol dehydrogenase, Biochem. J. 155 (1976) 155e161. [70] C.A. Sellick, R.N. Campbell, R.J. Reece, Chapter 3: galactose metabolism in yeast-structure and regulation of the Leloir pathway enzymes and the genes encoding them, Int. Rev. Cell Mol. Biol. 269 (2008) 111e150. [71] E.J. Yun, I.-G. Choi, K.H. Kim, Red macroalgae as a sustainable resource for biobased products, Trends Biotechnol. 33 (2015a) 247e249.