Leor Korzena,
Indra Neel Pulidindib,
Alvaro Israelc,
Avigdor Abelsona and
Aharon Gedanken*bd
aDepartment of Zoology, Tel Aviv University, Ramat Aviv, 69978, Israel
bDepartment of Chemistry, Bar-Ilan University, Ramat-Gan 52900, Israel. E-mail: gedanken@mail.biu.ac.il; Fax: +972-3-7384053; Tel: +972-3-5318315
cIsrael Oceanographic and Limnological Research Ltd, The National Institute of Oceanography, P.O. Box 8030, Tel Shikmona 31080, Haifa, Israel
dNational Cheng Kung University, Department of Materials Science and Engineering, Tainan 70101, Taiwan
First published on 1st July 2015
Macro algal seaweeds are a promising feedstock for biofuels production. Yet, their relatively low fermentable carbohydrate content and the inefficient methods used for their conversion hamper their utilization. The optimized production of Ulva rigida co-cultured with fed-fish in an offshore mariculture (fish cages) system is reported. Enhanced production of biomass with elevated content of desired carbohydrates is achieved. The farmed biomass was further converted to bioethanol by a one-step sonication assisted SSF process. An ethanol yield of 16 wt% (based on the dry weight of algae) is obtained.
In addition, advanced conversion techniques should be developed, and sugar yields need to be improved.8–10
Integrated Multi-Trophic Aquaculture (IMTA) is a suitable alternative to the commonly practiced algal mono-culture. This strategy could provide a sustainable solution for the feedstock requirement of bioethanol production. IMTA systems are based on the concept of ecological sustainability. In aquaculture this concept refers to the reuse and recycling of internal feedback within a culture system. This minimizes the input and the output wastes of resources, such as nutrients, water and energy in effluent water.11–13 A common practice among land-based aquaculture operations during the last few decades is the integration of seaweed farming and aquaculture operations where seaweeds are cultured in the effluent water of abalone, prawns, oysters, clams or fish.11,14 Thus, when integrated with fed aquaculture (finfish), extractive organisms (seaweeds and suspension feeders) may turn waste into productive resources there by intensively reducing the impact of derived waste on the local ecosystem.11,13,15
The benefits of integrating seaweed cultivation with fed mariculture in order to recapture waste nutrients are well documented.11,13,15–17 A study conducted at a Mediterranean offshore mariculture farm, has shown intensified growth rates, and elevated cellular contents of fermentable sugars in the marine algae Ulva rigida cultured downstream to fish culture net pens.
Moreover, by the exposure of the cultured biomass to low ambient nutrient levels, a further enhancement of the fermentable sugar fraction was achieved.18 Nevertheless, most of the studies related to the integrated culture of fish and algae in marine open-water systems have focused mainly on the environmental and economic effects. Little attention has been paid, however, to the use of such systems for the production of marine biomass as a feedstock for bioethanol generation.
In this study, we co-cultured the seaweed Ulva rigida with fed-fish culture (Sparus aurata) in an offshore fish cage aquaculture complex. The culture system was focused on obtaining high yields of biomass with high content of fermentable sugars. The carbohydrate rich cultured biomass was then processed by the optimized conversion method so as to achieve optimal yields of bioethanol. Compared with the values in Table 1, 0.16 g of ethanol from 1 g of dry macro algae is the highest yield reported so far.
Macro algae | Hydrolysis conditions & sugar yield (g g−1 dry algae) | Fermentation conditions & ethanol yield (g g−1 sugar) | Reference |
---|---|---|---|
Ulva fasciata | Enzymatic (cellulase 22119) hydrolysis; 36 h; 45 °C; 0.207 | S. cerevisiae; 28 °C; 24 h; 120 rpm; 0.45 | 22 |
Ulva rigida | Enzymatic (cellulase, amylase) hydrolysis; 37 °C; 3 h; sonication; 0.196 | SSF process; cellulase, amylase, S. cerevisiae; 37 °C; 3 h; sonication; 0.33 | 25 |
G. tenuistipitata; R. riparium; G. salicornia; U. intestinalis; | HCl (0.1 to 1 M); 95 °C; 15 h; 0.539, 0.0233, 0.014, 0.0503 respectively | S. cerevisiae TISTR no. 5339; 30 °C; 18 h; 120 rpm; 4.17 × 10−3; 0.86 × 10−4; 0.31 × 10−4; 0.74 × 10−4 respectively | 26 |
Ulva fasciata | Cellulase produced from Cladosporium sphaerospermum was used for hydrolysis; 24 h; 40 °C; 0.112 | S. cerevisiae MTCC no. 180, 12 h, 28 °C, 120 rpm; 0.47 | 27 |
Ulva meridionalis | 2 mM phosphotungstic acid, HPA, 160 °C, microwave irradiation; synergistic effect between HPA and microwave irradiation; 0.336 neutral sugars | — | 28 |
The green macro alga Ulva (Chlorophyceae) is a common marine algae abundantly found in eutrophicated coastal waters. This marine alga could be considered a potential energy crop due to its high growth rates and relatively high carbohydrate content. Most of the studies related to the conversion of marine algae, namely Ulva species, to bioethanol used pretreatment processes prior to the enzymatic hydrolysis of the biomass.18–21 Pretreatment processes are often accompanied with several disadvantages: thermochemical treatments with dilute acids are energy consuming and also generate toxic residues such as hydroxymethyl furfural (HMF) and other furfurals due to the harsh pretreatment conditions.22–24 Korzen et al., reported recently on a one-step sonication assisted simultaneous saccharification and fermentation (SSF) process devoid of pretreatment for the production of bioethanol from Ulva rigida. However, the ethanol yield achieved was still lower (6.2 wt%) than the potentially available fermentable fraction.25 Recent strategies developed for the conversion of marine macro algae to bioethanol are summarized in Table 1.
Culture manipulation trials took place consecutively after the two week grow out culture phase. Algal culture cages housed with cultured U. rigida (n = 3) were moved from the 15 m downstream culture site and were repositioned at the low nutrient control site for 5 more days. So as to monitor the growth and biochemical content of the seaweeds during the culture manipulation, two days after the repositioning of the culture nets, the nets were taken out of water, seaweeds were gently drained with surplus water and weighed (on board), the seaweeds were promptly brought back into the nets and back into the water. 10 g of sampled seaweed from each net (n = 6, control nets were sampled as well) were rinsed with distilled water and stored frozen (−20 °C) for subsequent tissue analysis. At the end of each culture experiment the seaweeds were processed as specified above.
SGR = [ln(Wt/W0)]/t × 100 |
10% constitution of the broth comprised of: 2 g algae in 20 mL solution – (10 mL H2O and 10 mL buffer), 100 μL glucoamylase from A. niger (≥300 U mL−1), 40 μL α-amylase (≥250 units per mL); 0.1 g cellulase (≥0.3 units per mg solid) and 0.5 g yeast (commercial Baker's yeast, Saccharomyces cerevisiae); 15% constitution of the broth comprised of: 3 g algae in 20 mL solution – (10 mL H2O and 10 mL buffer), 150 μL glucoamylase, 60 μL α-amylase; 0.15 g cellulase and 0.5 g yeast; 20% constitution of the broth comprised of: 4 g algae in 20 mL solution – (10 mL H2O and 10 mL buffer), 200 μL glucoamylase, 80 μL α-amylase; 0.2 g cellulase and 0.5 g yeast.
The SSF process was carried out under mild sonication at 37 °C for 2–4 h in a bath sonicator (MRC Clean-01 Ultrasonic cleaner, 40 kHz ultrasound frequency and 120 W ultrasonic power).25
Low loading, enz/2: 3 g algae in 20 mL solution – (10 mL H2O and 10 mL buffer), 75 μL glucoamylase, 30 μL α-amylase, 0.075 g cellulase and 0.5 g yeast.
Middle loading, enz*1: 3 g in 20 mL solution – (10 mL H2O and 10 mL buffer), 150 μL glucoamylase, 60 μL α-amylase; 0.15 g cellulase and 0.5 g yeast.
High loading, enz*2: 3 g in 20 mL solution – (10 mL H2O and 10 mL buffer) 300 μL glucoamylase; 120 μL α-amylase; 0.3 g cellulase and 0.5 g yeast;
The specific growth rate of Ulva rigida grown downstream from the fish cages showed significantly higher (27 times for the September trial (Fig. 2(a)) and 41 times for the November trial (Fig. 2(b))) specific growth rates (SGR) than those grown at the control station upstream from the cages. The availability of inorganic nutrients has been identified as the most important factor controlling the growth and productivity of seaweeds.25,31–33
The achieved daily growth rates are similar to the maximal specific growth rates reported by studies with Ulva species integrated in land-based multi-trophic aquaculture using tank cultivation,34,35 and are comparable to the values found for mass cultivation of Ulva.6,36 The decline in the growth of U. rigida between the September and the November experiments might be the result of intrinsic seasonality effect on the growth of this species. The mean seawater temperature measured during the September experiment was 29.2 °C and dropped to 24.4 °C during the November experiment, other parameters such as irradiance, although not measured during the study, had likely also changed during the different culture seasons. The effect of seasonality on growth and biochemical composition has been studied and observed in different species of algae.37–39 However, in order to fully understand the effects of seasonality on the annual production yields of seaweed biomass further long-term experiments should be carried out.
During both culture trials (September, Fig. 2(c) and November, Fig. 2(d)), the starch content was significantly higher (31.5% of DM, September trial) at the control station than downstream to the cages (24% of DM, September trial, p < 0.01) (Fig. 2(c)). This is in an inverse proportion to the ambient seawater nutrient concentrations In addition, after two days of culture manipulation at the low nutrient site, the starch contents bounced up and levelled with the values of the control site. These results are in line with previous studies on reserve carbohydrates in seaweeds. High nutrient concentrations were found to alter the proximate composition in seaweeds and caused a shift to lower levels of carbohydrates such as starch.39,40 Moreover, it has been shown that nitrogen-deficient green algae accumulate carbon mainly as starch reserves, which could be further used by respiration during growth and reproduction.41,42
Since both high growth rates and high concentrations of desired carbohydrates are crucial parameters for an economically viable biomass for bioethanol production, then two steps must be combined: first a nutrient-rich step for high biomass production and, second, a nutrient-limited phase for the carbohydrate/starch accumulating phase.
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Fig. 3 Efficiency of ultrasonication process for the SSF process with high solid (algae) content (10–20 wt%) (replicate no. n = 3; error bars indicate standard deviation, SD). |
Low loading, enz/2: (3 g algae in 20 mL solution – 10 mL H2O and 10 mL buffer, 75 μL glucoamylase, 30 α-amylase, 0.075 g cellulase and 0.5 g yeast).
Middle loading, enz*1: (3 g in 20 mL solution – 10 mL H2O and 10 mL buffer, 150 μL glucoamylase, 60 α-amylase; 0.15 g cellulase and 0.5 g yeast).
High loading, enz*2: (3 g in 20 mL solution – 10 mL H2O and 10 mL buffer) 300 μL glucoamylase; 120 α-amylase; 0.3 g cellulase and 0.5 g yeast;
Near 2-fold increase in ethanol yield is observed by increasing the enzyme loading by 4 times as depicted in Fig. 4. In view of the cost of the enzymes, the optimum loading of enzymes is the middle loading, enz*1, which could yield 11 wt% ethanol upon sonication for 4 h.
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Fig. 4 Effect of enzyme loading on the ethanol yield in the SSF process with a solid consistency of 15 wt% (replicate no. n = 3; error bars indicate standard deviation, SD). |
The 1H NMR spectrum of the aliquot of sample from the fermentation (SSF) broth under optimal reaction conditions (15 wt% solid consistency, high carbohydrate Ulva rigida, 1 wt% enzyme loading, 4 h sonication) is depicted in Fig. 6.
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Fig. 6 1H NMR spectrum of aliquot of sample collected from the fermentation (SSF) broth under optimal reaction conditions. |
The presence of 3H (t, 1.18 ppm) and 2H (q, 3.64 ppm) indicate the formation of ethanol. The signal at 1.9 ppm is due to the sodium acetate buffer present in the broth. The signal at 8.5 ppm (1H, s) is characteristic of the internal standard, HCOONa. The amount of ethanol estimated based on the relative integral values of the internal standard and ethanol peaks is 16 wt%. From 31.5 wt% starch in the high carbohydrate algae, the expected glucose amount upon complete hydrolysis is 35 wt% (ref. 43) which corresponds to a theoretical ethanol yield value of 17.8 wt%.10 Under the optimized process conditions, 16 wt% ethanol could be obtained from the high carbohydrate algae which corresponds to a process efficiency of 89%. This value is relatively higher than the process efficiency of 65% reported previously.25
The formation of ethanol from Ulva rigida in the SSF process is further confirmed using 13C NMR as represented in Fig. 7. The signals at 17.5 and 58.1 ppm are typical of ethanol. The presence of secondary metabolite, glycerol is also evident from the signals at 63.2 and 72.7 ppm. The signal at 23.8 ppm is due to CH3COONa buffer used in the SSF process. No fermentable sugars are detected in the region of 60–100 ppm typical of glucose indicating the effectiveness of the SSF process.
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Fig. 7 13C NMR spectrum of aliquot of sample collected from the fermentation (SSF) broth under optimal reaction conditions. |
This journal is © The Royal Society of Chemistry 2015 |