Two-stage Hydrolysis of Invasive Algal Feedstock for Ethanol Fermentation

Authors

  • Xin Wang,

    1. Department of Oceanography, University of Hawaii at Manoa, HI 96822, USA
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  • Xianhua Liu,

    1. School of Environmental Science and Engineering, Tianjin University, Tianjin 300071, China
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  • Guangyi Wang

    Corresponding author
    1. Department of Oceanography, University of Hawaii at Manoa, HI 96822, USA
    2. School of Environment and Energy, Peking University Shenzhen Graduate School, Shenzhen 518055, China
      Corresponding author
      Tel: +1 808 956 3744, +86 755 2661 1617; Fax: +1 808 956 9225, +86 0755 2603 5227; E-mail: guangyi@hawaii.edu gywang@pkusz.edu.cn
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Corresponding author
Tel: +1 808 956 3744, +86 755 2661 1617; Fax: +1 808 956 9225, +86 0755 2603 5227; E-mail: guangyi@hawaii.edugywang@pkusz.edu.cn

Abstract

The overall goal of this work was to develop a saccharification method for the production of third generation biofuel (i.e. bioethanol) using feedstock of the invasive marine macroalga Gracilaria salicornia. Under optimum conditions (120 °C and 2% sulfuric acid for 30 min), dilute acid hydrolysis of the homogenized invasive plants yielded a low concentration of glucose (4.1 mM or 4.3 g glucose/kg fresh algal biomass). However, two-stage hydrolysis of the homogenates (combination of dilute acid hydrolysis with enzymatic hydrolysis) produced 13.8 g of glucose from one kilogram of fresh algal feedstock. Batch fermentation analysis produced 79.1 g EtOH from one kilogram of dried invasive algal feedstock using the ethanologenic strain Escherichia coli KO11. Furthermore, ethanol production kinetics indicated that the invasive algal feedstock contained different types of sugar, including C5-sugar. This study represents the first report on third generation biofuel production from invasive macroalgae, suggesting that there is great potential for the production of renewable energy using marine invasive biomass.

Introduction

Like plants, many algal species have rigid cellulose-based cell walls and accumulate starch as their main carbohydrate storage compounds and cell wall structure, which contains an astonishingly diverse range of simple and complex carbohydrates (Goh and Lee 2010; Guerriero et al. 2010). Some of marine algal species contain up to 70% of polysaccharides, i.e., cell wall polysaccharides (cellulose, hemicelluloses, xylan, and mannan), intercellular polysaccharides (sulfated glucuronoxylorhamnan, algine, agar, and carrageenin), and storage polysaccharides (amino pectin, laminaran and floridean starch) (Okuda et al. 2008). Both intercellular and cell wall polysaccharides can be converted into fermentable sugars. The majority of algal polysaccharides are potential biochemical feedstock and can be fermented to produce ethanol. Additionally, algal feedstocks have several advantages over other types of feedstocks. These include high area productivity, no competition with conventional agriculture for land, utilization of different water sources (e.g., seawater, blackish water, saline water, and wastewater), recycling of carbon dioxide, and compatibility with integrated production of fuels and co-products within biorefineries. Hence, algal feedstocks are considered one of the most promising non-food feedstocks for biofuels (Wijffels and Barbosa 2010). Previous studies of algal biofuel production have largely focused on microalgae (Mata et al. 2010). Mannitol extracted from the brown seaweed Laminaria hyperborean has been used as a substrate for ethanol production by Zymobacter palmae with a yield of 0.38 g ethanol per gram mannital (Horn et al. 2000). A conceivable biorefinery production process of third generation ethanol using the seaweeds Euchema spp. has been proposed (Goh and Lee 2010). However, detailed studies of macroalgal feedstock hydrolysis for ethanol fermentation are rare. Particularly, production of third generation biofuel from invasive macroaglae has not been reported.

Ethanol was successfully implemented as the chief component of automotive fuel in Brazil nearly three decades ago. Currently, ethanol is commonly used in transportation fuel, with blends up to 15% with gasoline and constitutes 99% of the total biofuel consumption in the USA. Regardless of its low energy content in comparison with gasoline, ethanol has many advantages as an alternative energy, including reduction of pollutant emissions and production from environmental wastes (Rao et al. 2007). Corn (Zea mays) grain and other cereals like sorghum (Sorghum bicolor) constitute the primary feedstock for ethanol production in the USA and other countries (Schmer et al., 2008). However, these feedstocks are major food sources and ethanol production from these grains can seriously impact food prices and the economic viability of ethanol production. Thus, ethanol from nonfood feedstock is a key factor for the economic viability of its production. Among various nonfood feedstocks, algae (macro- and microalgae) have a higher photosynthetic efficiency and have long been considered as one of the most promising non-food feedstock for biofuels (i.e., ethanol and diesel) production (Hossain et al. 2008; Wijffels and Barbosa, 2010).

Marine macroalgae contribute significantly to global primary production and play critical roles in the stability and function of marine ecosystems (Inderjit et al. 2006; Williams 2007). However, macroalgae can also become invasive in a newly introduced environment and have profound adverse ecological impacts including the alteration of ecosystem structure, reduction of indigenous biodiversity, and economic losses (Smith et al. 2002; Smith et al. 2004; Pandolfi et al. 2005; Goreau et al. 2008). There are about two dozen alien algal species in Hawaii with half of them reported as invasive (O’Doherty and Sherwood 2007). Management of invasive algae in Kihei (Maui) alone costs the state of Hawaii $20 million annually (Smith et al. 2004). Invasive algae are also found in other coastal areas of the state, including Waikiki and Kaneohe Bay (Oahu) and southern Molokai (Goreau et al. 2008). Therefore, the statewide economic impact of invasive algae is much greater than $20 million annually. During blooming time, some of these species (e.g., Gracilaria salicornia) are washed up on beaches in large quantities. The large piles of decomposing biomass draw complaints from residents and drive tourists away (Squair et al. 2003). The invasive species G. salicornia is commonly found in Hawaii and has been reported to kill corals in Waikiki and Kaneohe (Smith et al. 2004; Goreau et al. 2008). Currently, physical removal has been the only reported efficient method for management of invasive algae. Over twenty tons of biomass for the invasive species G. salicornia were removed from the area of the War Memorial Natatorium in Waikiki (Oahu) at a clean-up event (Smith et al. 2004). For these reasons the management of large quantities of invasive algae biomass presents a great environmental challenge.

In this study, we took advantage of the high content of cellulosic biomass of the invasive species G. salicornia and demonstrated the possibility of producing ethanol from the invasive algal feedstocks. It represents the first report of ethanol production from invasive macroalgae.

Results

Optimizing dilute acid hydrolysis of algal biomass

Dilute acids break down the cellulose and hemicellulose polymers in cellulosic biomass to release individual sugars, which can be fermented into ethanol (Lee et al. 1999; Lenihan et al. 2010). Kinetic studies on the dilute acid hydrolysis of various cellulosic materials indicated that the hydrolysis kinetic parameters are strongly dependent on the substrate and acid concentration, temperature, and reaction time (Malester et al. 1988; Lenihan et al. 2010). However, macroalgal feedstocks have rarely been investigated for saccharification using dilute acid hydrolysis. To optimize the conditions for the hydrolysis, the homogenates resulting from the invasive algal plants were heated to 100°C and cooled down to room temperature. The agar fraction was separated from cellulosic one. Individual fractions were used to optimize their conditions for saccharification using dilute acid hydrolysis. Of the four tested sulfuric acid concentrations, dilute acid hydrolysis using 2.5% sulfuric acid yielded the highest glucose concentration from the cellulosic fraction and hydrolysis with 2% sulfuric acid gave the best results from the agar fraction (Figure 1). At their optimal sulfuric acid conditions, the hydrolysis at 125°C gave the highest glucose concentration from both the agar and cellulosic fractions of algal biomass (Figure 2). To reduce the cost of separating agar from cellulose fraction, the homogenates of the invasive algal plants was hydrolyzed at the level of 2% sulfuric acid and 120 °C for 30 min. Glucose concentration in the resulting hydrolysates contained approximately 4.1 mM (i.e., 4.3 g glucose/kg fresh algal biomass) and failed to yield detectable ethanol in the batched fermentation (data not shown).

Figure 1.

Effect of sulfuric acid on the hydrolysis of the invasive algal feedstock.

Figure 2.

Effect of temperature on the hydrolysis of the invasive algal feedstock.

Two-stage hydrolysis of invasive algal biomass

To improve the saccharification efficiency of invasive algal plants, the homogenate containing both agar and cellulose components was subjected to dilute acid hydrolysis containing 2% sulfuric acid at 120 °C for 30 min. The resulting hydrolysates were hydrolyzed using cellulase as described previously. Glucose concentration in the hydrolysates resulting from the two-stage hydrolysis reached 15.1 mM at 4-h incubation and continued to rise up to 16.6 mM after 26-h incubation (Figure 3). Overall, after 4-h incubation, further enzymatic hydrolysis did not significantly increase glucose concentration. Therefore, 4-hour incubation was used for enzymatic hydrolysis in this study.

Figure 3.

Enzymatic hydrolysis of the invasive algal feedstock.

Ethanol production from algal hydrolysates

The ethanologenic strand E. coli KO11 (Jarboe et al. 2007) was used in batch fermentation for the production of ethanol with the algal hydrolysate as the sole carbon source. In media containing the algal hydrolysate, 13.8 mM glucose was completely consumed within 18 h, with little variation in all three replicate fermentation experiments (Figure 4). The ethanol concentration reached approximately 28 mM when the glucose was used up. However, ethanol concentration continued to increase up to 34 mM with further incubation. The continued increase in ethanol concentration clearly indicated that other non-glucose sugars occurred in the algal hydrolysate and were fermented by the E. coli KO11 cells into ethanol (Suppl. Figure 1). Ethanol concentration reached the maximum (34 mM) at 30-h incubation, which indicates that extended fermentation did not increase ethanol concentration in the fermentor (Figure 4). The described process using the invasive algal plants as feedstock, two-stage hydrolysis, and E. coli KO11 as the ethanologenic strains in fermentation was able to achieve 79.1 g EtOH per kg of dried invasive algal plants. In the positive control, 30 mM of glucose supplemented in Neidhardt MOPS defined minimal media as carbon source was completely consumed at 18-h fermentation and yielded the highest ethanol concentration of 44 mM (data not shown).

Figure 4.

Ethanol production from hydrolysate derived from invasive algal feedstock using the ethanologenic E. coli strain KO11.

Discussion

Algal feedstocks are one of the most promising non-food feedstocks for biofuels (Wijffels and Barbosa 2010). In this manuscript, we developed a two-stage hydrolysis method for the saccharification of invasive marine algal plants and use E. coli KO11 as the ethanogenic strain to successfully ferment invasive algal monosaccharide into ethanol. This is the first report on the production of ethanol from marine invasive species. It provides useful techniques to produce biofuel from marine biomass, particularly from those that are an environmental hazard. Given the fast-growing nature of the invasive algae, the resulting methods and technology can be used to produce biofuel from marine biomass collected in Hawaii or any other coastal region of the world's oceans. This study contributes to the pursuit of viable renewable energy production from marine bioresources and also methods to reduce CO2 emissions.

First generation biofuels refer to fuels derived from sources like corn starch, sugar cane or sugar beet, and animal and vegetable fats (Scott et al. 2010). The major concern related to first generation biofuels is that they compete for arable lands with food crops (Hill et al. 2006; Stephenson et al. 2008). Considering the requirements of freshwater and fertilizers, it is difficult to produce those fuels in a sustainable manner. Second generation biofuels are derived from cellulosic biomass feedstock using advanced biotechnological processes. Compared with first generation biofuels, they hold greater promise because of their more favorable greenhouse gas emissions, lack of competition with food production, and lower land usage (Lynd et al. 1991; Li et al. 2007; Hossain et al. 2008; Hu et al. 2008; Gouveia and Oliveira 2009). However, the production of second generation biofuels faces several challenges: relatively high production costs; technological demands for enzyme, pretreatment, and fermentation; and the essential development of a whole new infrastructure for harvesting, transporting, storing, and refining biomass (Berndes et al. 2009). Algae fuels are considered third generation biofuel (Sheehan 2009). Although algae fuels are superior to the other two types of biofuels in several aspects, developing a technological process to produce third generation biofuel is technologically more demanding than those for other types of biofuels. Here, we demonstrate that combining dilute acid hydrolysis with enzymatic hydrolysis can provide efficient saccharification of biomass of the invasive algae Gracilaria salicornia. Use of engineered E. coli KO11 enabled us to successfully ferment monosacharides derived from G. salicornia into ethanol (Figure 4).

Pretreatment of feedstocks, especially those containing cellulose and hemicelluloses, and subsequent hydrolysis of these polysaccharides into monosaccharides are key steps in the production of second and third generation biofuels (Rubin 2008). Dilute sulfuric acid method was used for biomass hydrolysis as early as 1819 when Braconnot retrieved fermentable sugar from linen (Braconnot 1819). The sulfuric acid was able to break the hydrogen bond among the cellulose polymers, making them available for sulfuric acid and allowing hydrolyzing glycosidic bonds to release sugars from cellulose and hemicelluloses (Xiang et al. 2003; Binder and Raines 2009). This dilute acid hydrolysis is still commonly used in today's industrial processes (Mosier et al. 2005). However, dilute acid hydrolysis only yielded a low amount of monosaccharides from the invasive algal plants (Figures 1 and 2). The enzymatic hydrolysis released much more monosaccharides from algal feedstocks than dilute acid hydrolysis alone (Figure 3). To ferment monosaccharides derived from invasive algae, the genetically engineered E. coli strain KO11, which is capable of fermenting a variety of sugars (C6 and C5-sugars) (Ingram et al. 1987; Ingram et al. 1999), was used in batch fermentation. After the glucose was completely consumed, the continuous production of ethanol from the invasive algae hydrolysate confirmed the metabolic ability of KO11 to ferment other C5-sugars (Figure 4, Suppl. Fig. 1). Furthermore, ethanol yield of this study was lower than those previously reported using extracts of the brown seaweed Laminaria hyperborean (Horn et al. 2000) and the microalga Chlamydomonas reinhardtii biomass (Nguyen et al. 2009). The low ethanol yield may be ascribed to following reasons. First, macroalgal feedstocks have been reported to contain large amount of polysaccharides that are made up from monomers such as α(1–3)D-galactose-4-sulphate, β(1,4)-3,6-anhydro-D-galactose, β(1–3)-D-glucopyranose, D-mannitol, and mannose (Horn et al. 2000; Lin et al. 2000; Tong et al., 2007; Goh et al. 2010). Those sugars are not readily fermented by E. coli KO11. Second, ethanologenic microbial strains used in the other studies are Zymobacter palmae (Horn et al. 2000) and Saccharonmyces cerevisiae (Nguyen et al. 2008), which have better capacities to ferment the previously mentioned sugars into ethanol than E. coli KO11. Finally, hydrolysate of the invasive feedstocks may contain inhibitory chemicals of ethanol fermentation by E. coli KO11. Clearly, chemical analysis of biomass of the invasive species G. salicornia will be an interesting follow-up project. Metabollically engineered E. coli KO11 with capabilities of fermenting more monosaccharides derived from G. salicornia will surely increase the overall ethanol yield from invasive algal hydrolysate.

With increased demands for renewable biofuels, the development of technological processes for the production of second and third generation biofuels will be one of the greatest challenges facing our society. Production of third generation biofuel from invasive algal biomass will surely contribute to our global efforts to reduce global warming and dependence on conventional fossil fuels. To that end, this study provides the first report on third generation biofuel production from marine environmental hazards.

Materials and Methods

Algae specimen collection

Algal plants of the invasive species Gracilaria salicornia were collected on a reef in the subtidal zone near Waikiki Beach, Oahu Island. Specimens were directly transferred to ziplock plastic bags. Latex gloves were worn during collection. The samples were transported to the laboratory within 1 h and processed immediately for homogenization.

Dilute acid hydrolysis of algal biomass

Algal plants were rinsed briefly with sterile water and dried with paper towels. The washed algal plants were homogenized in a homogenizer. Separation of agar and cellulosic fibrin used the method described by Guerin and Bird (1987). The resulting agar and cellulosic fibrin fractions were individually hydrolyzed using the dilute acid hydrolysis (Nguyen et al. 1999). To optimize acid concentration, each part of the two fractions was mixed with sulfuric acid with final concentrations of 0.5%, 1%, 2.5%, and 5% (v/v) and hydrolyzed at room temperature for 30 min. To optimize hydrolysis temperature, agar and cellulosic fractions containing 2% and 2.5% of sulfuric acid, respectively, were autoclaved at different temperatures (105, 115, 125 and 128°C) for 30 min. The resulting hydrolysates were centrifuged at 6,000 ×g for 10 min and the resulting supernatant containing mono-sugars was collected and neutralized with NaOH to pH 7. Glucose content of the hydrolysates was determined using Glucose Assay Kit (Sigma) and then used to choose the optimal hydrolysis condition. All treatments were done in three replicates.

Enzymatic hydrolysis of algae biomass

For enzymatic hydrolysis, the homogenate of the invasive algal plants containing 2% sulfuric acid was hydrolyzed at 120 °C for 30 min. The resulting hydrolysates was neutralized to pH 5 with NaOH and mixed with NaAc/HAc buffer (0.05 M, pH5) at the volume ratio of 1:1. One thousand of the buffered hydrolysates were incubated with 5g of cellulase (MP Biomedicals, LLC) at 40°C. During the incubation, glucose concentration was monitored using HPLC (Shimadzu). At the end of incubation, the enzyme mixture was centrifuged at 14,000 ×g for 10 min. The resulting suppernant containing mono-sugars was filtered through Whatman filter paper, 2.0-μm and 0.2-μm polycarbonate filters sequentially. The glucose concentration of the resulting filtrates was measured using a high-performance liquid chromatography (HPLC) (Shimadzu) equipped with a BioRad Aminex HPX-87H Ion Exclusion column (300 mm×7.8 mm) and a refractive index detector (RID 10A, Shimadzu) (Chavez-Servin et al. 2004).

Fermentation of algal hydrolysate

Neidhardt MOPS defined minimal broth with supplements was used for anaerobic batch fermentation following the method described by Mathews et al. (2010). In brief, the batch fermentation was carried out in a four-part assembly 500 ml capacity spinner flask (Bellco Blass), with a total volume of 400 ml of minimal Neidhardt MOPS defined broth with algal hydrolysate as the carbon source (Mathews et al. 2010). The 400 ml of fermentation broth was made from 300 ml of algal hydrolysate, 74.4 mL of Neidhardt MOPS defined minimal broth (containing 10× MOPS, 100× K2HPO4 (0.132 M), 40× ACGU solution, 20× supplement EZ, 1 000× chloramphenicol (50 mg/mL)) and 25.6 ml sterile water. The media inside the spinner flask and the space in the attached gas trap was sparged with nitrogen gas for 20 min to remove oxygen. All fermentations were done at 30 °C using a circulating water bath.

For the fermentative production of ethanol, a Escherichia coli KO11 strain (Jarboe et al. 2007) was inoculated into 100 mL of LB broth supplemented with 2% xylose and incubated overnight at 37 °C. On the day of fermentation, 5 ml of the resulting bacterial culture was reinoculated into 100 mL LB broth supplemented with 2% xylose in the morning and incubated at 37 °C till OD600 reached 1.00. Bacterial cells were then collected by centrifuging at 5 000 ×g for 10 minutes and washed twice with sterile distilled water. Finally, the washed bacterial cells were suspended in 25 mL of fermentation media and injected into the fermentation system.

Ethanol production

For ethanol and other analysis, 1 mL of fermentative broth was drawn from the fermentation system at time points of 0, 2, 4, 6, 9, 12, 15, 17, 20, 23, 26, 30, 34, 42, 45, and 50 h. The fermentative broth was centrifuged at 16 000 ×g for 1 min, and the supernatant was filtered through a 0.22-μm of polycarbonate membrane. The filtrate was used to determine glucose and ethanol concentrations using HPLC analysis and glucose (Mathews et al. 2010). Triple experiments were conducted for fermentation analysis. Neidhardt MOPS defined minimal broth supplemented with 30 mM of glucose was used as a positive control.

(Co-Editor: Hai-Chun Jing)

Acknowledgements

This work is funded by the University of Hawaii Sea Grant (NA09OAR4170060). The views expressed herein are those of the authors and do not necessarily reflect the views of NOAA or any of its subagencies.

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