Saccharification of microalgae biomass obtained from wastewater treatment by enzymatic hydrolysis. Effect of alkaline-peroxide pretreatment

1 Saccharification of microalgae biomass obtained from wastewater treatment by enzymatic hydrolysis. Effect of alkaline-peroxide pretreatment. Judit Martín Juáreza, Ana Lorenzo Hernandoa, Raúl Muñoz Torrea, Saúl Blanco Lanzab1, Silvia Bolado Rodrígueza* a Department of Chemical Engineering and Environmental Technology, University of Valladolid, Calle Doctor Mergelina s/n, 47011, Valladolid, Spain b The Institute of the Environment, La Serna, 58, 24007 León, Spain. Authors e-mail addresses: [email protected], [email protected], [email protected], [email protected], [email protected] *Corresponding author Telephone number: +34 983 423 958 E-mail address: [email protected]s 1Permanent address: Department of Biodiversity and Environmental Management, University of Leon, 24071 Leon, Spain. *Manuscript Click here to view linked References 2 ABSTRACT An enzymatic method for the carbohydrate hydrolysis of different microalgae biomass cultivated in domestic (DWB)† and pig manure (PMWB) wastewaters, at different storage conditions (fresh, freeze-dried and reconstituted), was evaluated. The DWB provided sugars yields between 40 and 63%, although low xylose yields (< 23.5%). Approximately 2% of this biomass was converted to byproducts as succinic, acetic and formic acids. For PMWB, a high fraction of the sugars (up to 87%) was extracted, but mainly converted into acetic, butyric and formic acids, which was attributed to the bacterial action. In addition, the performance of an alkaline-peroxide pretreatment, conducted for 1 hour, 50ºC and H2O2 concentrations from 1 to 7.5% (w/w), was essayed. The hydrolysis of pretreated microalgae supported a wide range of sugars extraction for DWB (55-90%), and 100% for PMWB. Nevertheless, a large fraction of these sugars (~30% for DWB and 100% for PMWB) was transformed to byproducts. Highlights Tested biomass showed different behaviours depending on the algae/bacteria ratio. Enzymatic hydrolysis of DWB yielded high glucose and low xylose extraction. Sugars from PMWB were completely released by enzymatic hydrolysis but oxidized. Acetic, formic and succinic acids were the main byproducts from released sugars. Pretreatment enhanced enzymatic hydrolysis performance for almost all biomass tested. † Abbreviations: DWB, domestic wastewater biomass; PMWB, pig manure microalgae biomass; HRT, hydraulic retention time; SRT, sludge retention time; CO2, carbon dioxide; CH4, methane. 3 Keywords: Enzymatic hydrolysis; Glucose; Xylose; Wastewater; Alkaline-peroxide pretreatment 1. Introduction World human population and industrial activity have exponentially increased during last decades, with a concomitant raise in global energy demand. This growth has been traditionally based on fossil fuels, whose side effects have turned this dependence environmentally unsustainable (Chisti, 2007). New renewable fuel sources and biorefinery approaches for designing cost-effective and “green” processes are expected to create more efficient and sustainable economies (Daroch et al., 2013). During the past decade, microalgae have experimented a continuous and positive development due to their wide range of practical applications: wastewater treatment, nitrogen and phosphorous recovery, biogas upgrading, production of biofuels, biofertilizers, animal and fish feed, etc. Despite Oswald and co-workers were pioneers in introducing the microalgae biorefinery concept in the 60’s, the combination and optimization of processes for the valorisation of microalgae biomass obtained from wastewaters treatment remains a challenge nowadays (Acién et al, 2014). Microalgae biomass is mainly composed of proteins (6% - 52%), lipids (5% - 23%) and carbohydrates (7% - 23%) (Tijani et al., 2015). This content may vary within microalgae strains and is highly dependent on cultivation conditions, especially under nutrients-deprivation scenarios. Among them, carbohydrates are one of the preferred feedstock for obtaining a variety of biofuels. Carbohydrates are mainly present in microalgae cell wall as cellulose and hemicellulose, and/or inside the cell as starch. Cell walls are mainly composed of biopolymers such as sporopollenin or algaenan, which 4 confer the cell a high rigidity and resistance to chemical attack (González-Hernández et al., 2012) and are characteristic of microalgae strains like Scenedesmus (Miranda et al., 2012). In order to make available the valuable compounds present inside microalgae cells, pretreatments are often needed in order to disrupt cell walls. Microalgae pretreatment allows for an efficient release of the carbohydrate content, enhancing saccharification and sugars bioavailability to maximize biofuels production (Hernández et al., 2015). Due to the lack of lignin, microalgae-based biofuels are expected to be cheaper compared to second-generation biofuels (Chen et al., 2013), but most of the literature references use pure cultures of microalgae grown on synthetic media, which would turn microalgae biofuel production prohibitive from an economic point of view (Lam and Lee, 2015). For instance, Miranda et al., (2012) evaluated the performance of several chemical and mechanical pretreatments for cell disruption and sugar extraction of wet and dried Scenedesmus obliquus biomass. H2SO4 hydrolysis was selected and optimized (120ºC, 2N sulfuric acid, 50 g biomass/L, one single step), and a synergistic effect between microalgae drying and sugar extraction for the acid pretreatment was reported. This study also confirmed the key role of cell disruption on the efficiency of sugar extraction from Scenedesmus. Harun and Danquah, (2011a) and (2011b) assessed the efficiency of pretreatments such as acid hydrolysis and ultrasound followed by enzymatic hydrolysis with cellulose on Chlorococcum humicola for bioethanol production. Despite no values of released sugars or byproducts were provided after acid hydrolysis, the authors obtained a maximum released glucose yield of 68.2% with 10g/L of biomass concentration after enzymatic hydrolysis at 40ºC and pH 4.5. Furthermore, it is also desirable to develop pretreatment methods with chemicals and 5 effluents streams that have a lower impact on the environment. Some works have been published studying the use of green solvents, as supercritical fluids and ionic liquids, (Silveira et al., 2015) for pure culture algae pretreatment. For example, Schultz-Jensen et al., (2013) applied ozonolysis to increase the digestibility of Chaetomorpha linum macroalgae, reporting 75% of xylan recovery; and Zhou et al., (2012) obtained 0.65g of released sugars/g algae applying [Emim]Cl and 7 wt% HCl at 105°C for 3 h to Chlorella sp. biomass (73.58% of initial sugars). Similarly, Ometto et al., (2014) evaluated the energy consumption and impact of four pretreatments (enzymatic treatment, thermal, thermal hydrolysis and ultrasound) on the preferential release of the biochemical fractions of axenic Scenedesmus obliquus, Chlorella sorokiniana and Arthrospira maxima strains. The authors concluded that enzymatic hydrolysis was the best method for carbohydrate release and the only one with a positive energy balance due to the mild operational conditions needed. Based on the benefits and popularization of microalgae-based wastewater treatment, there is a recent interest on developing strategies for the valorisation of this residual microalgae biomass. This biomass often contains significant concentrations of heterotrophic and nitrifying bacterial due to the high concentration of organic matter and ammonium present in domestic or livestock wastewaters, which could have some effect on the pretreatments results. Nevertheless, only some authors mentioned this bacteria contribution, like Alzate et al., (2012) working in biogas production or Castro et al., (2015) who considered necessary to apply sterilization process (autoclaving) before using wastewater microalgae biomass for butanol production. A biomass sterilization effect could be expected from the application of alkaline peroxide pretreatment, which has also shown high sugars release yields when used for 6 lignocellulosic materials (Monlau et al., (2012); Toquero and Bolado, (2014)). Compared with other chemical pretreatments, alkaline-peroxide pretreatment is carried out at mild temperatures, and it leads to a lesser formation of inhibitors than in other processes (Bolado-Rodríguez et al., 2016). Tijani et al., (2015) suggested this pretreatment as a suitable process for microalgae biomass rich in hemicellulose, thanks to its moderate operating conditions and its high efficiency releasing xylose. For macroalgae, its viability has just started to be tested. Li et al., (2016) optimized hydrogen peroxide as pretreatment for Ulva prolifera waste biomass, in order to improve ulterior enzymatic hydrolysis process. When applying optimum conditions (0.2% H2O2, 50ºC, 12h and pH 4.0) they obtained 420 mg/g biomass of reducing sugars. Nevertheless, to the authors’ knowledge, the potential of this pretreatment to enhance sugar release from microalgae biomass has never been explored. The aim of this work was the elucidation of the performance of enzymatic hydrolysis for saccharification of microalgae biomass cultivated in different types of wastewaters. An analysis of the influence of biomass composition and storage conditions, such as freeze-drying or cooling, on the released sugars yields and their transformation on other byproducts was conducted. Finally, the potential of alkalineperoxide pretreatment for hemicellulose solubilisation and biomass sterilization was herein assessed for the first time. 2. Materials and methods 7 2.1. Microalgae Freeze-dried microalgae biomass (A1) and the same biomass reconstituted with distilled water at a concentration of 150g/L (A2) were obtained from a thin-layer photobioreactor fed with domestic wastewater at a HRT (hydraulic residence time) of 3.3 days. Microalgae biomass was composed of Scenedesmus obliquus (95%), Scenedesmus quadricauda (4%) and Nitzschia sp. (1%). Freeze-dried (B1) and fresh (B2) microalgae biomass were also cultivated in a thin-layer photobioreactor at a HRT 3.3 days fed with pig manure wastewater diluted at 10%. The composition of B1 and B2 was Aphanothece sp. (61%) and Scenedesmus obliquus (39%). Biomass A1, A2, B1 and B2 were kindly supplied by Cajamar Foundation (Almeria, Spain). Finally, fresh microalgae biomass (C) was cultivated at the Department of Chemical Engineering and Environmental Technology of the University of Valladolid (Spain) in an anoxic–aerobic algal–bacterial photobioreactor with biomass recirculation (Alcántara et al., 2015). The photobioreactor was operated at a HRT 2 days and a sludge retention time (SRT) of 10 days using fresh domestic wastewater. Biomass C was composed of Scenedesmus obliquus (48%), Desmodesmus spinosus (45%) and Nitzschia palea (7%) and it was centrifuged for 10 min at 10000 rpm and maintained at 4ºC prior to use. 2.2. Enzymatic hydrolysis Enzymatic hydrolysis assays of untreated and pretreated microalgae were performed in 100 mL Erlenmeyer flasks containing 6% w/w dry solid and a mixture of 10 FPU g-1 (Celluclast 1.5L - Cellulase from Trichoderma reesei) and 20 CBU g-1 (Novozyme 188 – β-glucosidase from Aspergillus niger) of cellulose (dry basis) (Travaini et al., 2013). The pH was adjusted at 4.9 ± 0.1. The hydrolysis assays were carried out in a rotary 8 shaker at 50 ºC and 300 rpm for 48 h. Samples were drawn after hydrolysis and stored at 4ºC prior to the determination of the concentration of sugars (glucose, xylose, cellobiose and arabinose) and potential byproducts (oxalic, formic, acetic, butyric, succinic and levulinic acids, methanol and xylitol). 2.3. Alkaline-peroxide microalgae pretreatment Based on previously published experiments conducted with lignocellulosic materials (Toquero and Bolado, (2014); Karagöz et al., (2012)), H2O2 concentrations ranging from 1% to 7.5% were initially selected for the pretreatment of microalgae biomass A1 and A2. The high H2O2 concentrations used in A1 and A2 assays involved harsh reactions, which resulted in gas generation, biomass losses by splashing and even break of some bottles. Therefore, only H2O2 concentrations of 1% and 2.5% were later on applied to B1, B2 and C. Known mass of microalgae were placed in 1 L bottles and adequate volumes of H2O2 solutions (of the selected concentrations), were added to obtain 5% w/w suspensions. Then, the pH was adjusted to 11.5 with 2 M NaOH and the systems incubated in a rotatory shaker at 50ºC and 120 rpm for 60 min. The slurry was cooled down to room temperature, and the residual solid was separated by centrifugation (10 min, 10000 rpm). The experiments were conducted in duplicate. The liquid and solid fractions were stored at 4 ºC for further composition analysis of sugars (glucose, xylose, cellobiose and arabinose). In addition, the potential byproducts formed during biomass pretreatment (oxalic, formic, acetic, butyric, succinic, and levulinic acids, methanol and xylitol) were analysed in the liquid fraction. The solid fractions were used as a substrate in a subsequent enzymatic hydrolysis assay carried out as described above (Toquero and Bolado, 2014). 9 2.4. Analytical methods The identification, quantification and biometry measurements of microalgae were carried out by microscopic examination (OLYMPUS IX70) of microalgae samples (fixed with lugol acid at 5% and stored at 4 °C prior to analysis) according to Sournia, (1978). The absorbance ratio [(ABS at 680nm - ABS at750nm)/ ABS at 680nm)], measured in a GENESYS 20 visible spectrophotometer, was used as a qualitative estimation of the microalgae to bacteria ratio (Fairchild et al., 2005). The determination of the carbon and nitrogen content of the biomass was performed using a LECO CHNS-932 analyzer, while phosphorus and sulphur content analyses were carried out spectrophotometrically after acid digestion in a microwave according to the internal protocol of the Laboratory of Instrumental Analysis of Valladolid University. The starch content was measured following the 996.11 AOAC method. The protein and lipid content were determined using the Lowry method and Kochert method, respectively (Serejo et al., 2015). The content of moisture, extractives, ash and insoluble residue in raw biomass samples was analysed following NREL (National Renewable Energy Laboratory – USA) analytical procedures. The carbohydrate content in the raw and pretreated microalgae was determined by HPLC-RI using a modified NREL procedure. First, biomass was subjected to a concentrated acid hydrolysis for 1 h by adding 3 mL of H2SO4 (72% w/w) at 30ºC to a 300 mg dry biomass sample. Then, 84 mL of deionized water was added to dilute the acid concentration to 4% w/w prior to autoclaving at 121ºC for 1h. Then, solid and liquid fractions were separated by centrifugation (10 min, 10000 rpm). The liquid fraction was stored at 4ºC for the determination of sugars, 16 main byproducts found in the liquid fraction after pretreatment of A1 and A2 biomass were formic acid (~60%) and acetic acid (20%), with methanol and succinic acid detected at very low concentrations. On the other hand, acetic acid represented 50% of the total byproducts after pretreatment in the liquid fraction of B1, B2 and C, while formic, butyric, succinic and levulinic acids and xylitol were produced at trace levels. Methanol was only detected in the liquid fraction of sample C after pretreatment. Finally, and in agreement with the results reported by other authors when applying alkaline peroxide pretreatment for lignocellulosic materials (Karagöz et al., 2012), neither furfural nor HMF (inhibitory compounds) were detected in this work. Sugars solubilisation and transformation during the pretreatment of DWB represented a noteworthy loss of total sugar potential. The losses increased with H2O2 concentration, accounting for 35.4, 43.8, 45.3 and 61.0% in A1, 34.4, 40.7, 46.9 and 51.3% in A2, and 25.2 and 26.5% in C. These high sugar losses during pretreatment allowed foreseeing a final low sugar release yield during enzymatic hydrolysis in A1 and A2. At this point, it should be remarked that the final sugar content of the microalgae hydrolysate is critical for the economic sustainability of microalgae biorefineries devoted to ferment the released sugars. In our particular study, the low sugars concentration, along with the high concentration of byproducts and potentially inhibitory residues from alkaline-peroxide pretreatment would hinder the fermentation of the hydrolysates by a diauxic microorganism such as Pichia stipitis. On the other hand, these losses were barely noticeable in PMWB (3.8 and 3.7% in B1, and 2.1 and 2.7% in B2 at 1 and 2.5% H2O2, respectively). Again, the biomass from pig manure wastewater was more resistant in a chemical inhibitory medium. This finding highlighted the beneficial effect of alkaline-peroxide pretreatment on the further 17 utilization of biomass with high bacteria/algae ratios. On the other hand, the freezedrying and initial moisture content of the biomass exhibited a scanty effect on the sugar release and further bioconversion during H2O2 pretreatment. Thus, only slightly higher solubilisation yields and byproducts generation were obtained for freeze-dry biomass (A1 and B1) and reconstituted (A2) or fresh biomass (B2). Significant biomass losses during pretreatment of ~30% of the initial microalgae mass were estimated for samples A1 and A2 from the results in Table 3 (data not shown). These high values suggested a solubilisation of others components than sugars during pretreatment, whose determination was out of the scope of this study. In fact, alkaline-peroxide pretreatment is capable of supporting high lignin solubilisations in wheat straw at operating conditions compared to those used in this work (5% H2O2, pH 11.5, 1h, 50ºC) (Toquero and Bolado, 2014). In addition, a decrease in cellulose and hemicellulose content compared to the raw biomass was observed for all solid fractions of pretreated material. 3.3.2 Enzymatic hydrolysis of pretreated samples Table 2 shows the concentration of released sugars and byproducts resulting from the enzymatic hydrolysis of pretreated samples. No clear correlation between hydrogen peroxide concentration and the yields of glucose and xylose release was found, considering the different sugars concentrations in the pretreated materials before enzymatic hydrolysis (Table 3). These results were in agreement with Li et al., (2016), who reported an increase in the reducing sugar yield when increasing H2O2 concentration up to 0.5%, followed by a reduction of sugars yield when increasing H2O2 concentration to 2 %. 18 The concentration of released glucose from all pretreated samples was lower than that from untreated samples. The released glucose yield for A1 varied from 67.3 to 78.8% in pretreated samples, which was significantly lower than the 93.6% for untreated A1 biomass. Similar released glucose yields ranging from 63.7 to 70.7% were obtained for A2. However, comparable glucose yields (~65%) were found during the enzymatic hydrolysis of untreated and pretreated samples of biomass C. These glucose release yields recorded in pretreated biomass were very similar to the value of 64% reported by Harun and Danquah, (2011a) during the cellulose-based hydrolysis of Chlorococum sp. pretreated by ultrasounds. On the other hand, very low xylose release yields were obtained for all pretreated microalgae samples, despite most studies investigating the enzymatic hydrolysis of lignocellulosic materials pretreated with H2O2 under alkaline conditions reported an increase on released xylose yield. For example, this yield increased from 6.4% to 28.9% when sugarcane bagasse was pretreated (Yu et al., 2015) and from 9.3% to 48% when pretreating wheat straw (Toquero and Bolado, 2014). In general terms, the concentration of byproducts was similar in hydrolysates from pretreated samples and in those from raw materials, which suggests that H2O2 pretreatment did not exerted a significant disinfectant effect. In fact, the concentration of byproducts increased with H2O2 concentration likely due to a chemical mediated sugars oxidation. Enzymatic hydrolysis released almost the entire sugar content of pretreated B1 and B2 samples, which was transformed to byproducts at concentrations similar to those recorded in untreated biomass samples (e.g. ≈ 6 g/L acetic acid). Freeze-drying resulted in a higher concentration of byproducts in the hydrolysate compared to the hydrolysate of the pretreated fresh sample B2. In addition to acetic, 19 formic and butyric acid, succinic acids were obtained in the hydrolysate of microalgae C, although at lower concentrations than those recorded for A1 and A2. Biomass pretreatment promoted the generation of oxalic acid and increased methanol production in samples A1 and A2, along with the formation of acetic, formic and succinic acids. The concentration of sugars released from pretreated samples by successive acid hydrolysis is shown in Fig. 1. The pretreatment of biomass grown in domestic wastewater (A1, A2 and C) decreased the release of sugars in the first acid hydrolysis compared to untreated biomass. Extraction efficiencies of 58-69%, 24-33% and 6-10% were measured in the first, second and last cycle. Nevertheless, the sugar released in the first acid hydrolysis cycle increased with H2O2 concentration in the three DWB samples. These experimental observations could be attributed to the antagonistic effects of the pretreatment. Indeed, while H2O2 pretreatment disrupts biomass structure, it promotes the loss of easily releasable sugars by solubilization. On the other hand, the pretreatment of samples B1 and B2 increased the released sugar during the first acid hydrolysis compared to untreated PMWB samples, which resulted in yields of 60-70%. Sugar solubilisation during the pretreatment of PMWB was low and the disruption of the cell wall structure was dominant. Surprisingly, the first acid hydrolysis after pretreatment did not achieve the high values of sugar solubilisation obtained by enzymatic hydrolysis of untreated samples. It was hypothesis the disruption effect of bacteria is higher than that of the tested pretreatment, but the low pH values during acid hydrolysis inhibited the hydrolytic mechanisms of bacteria. In order to evaluate the overall performance of the process, solubilisation of glucose and xylose and their further oxidation during both pretreatment and enzymatic hydrolysis must be considered. The alkaline-peroxide pretreatment increased sugar 20 solubilisation from biomass by enzymatic hydrolysis, but at decreasing or similar sugar recovery yields due to the generation of byproducts. At the highest H2O2 concentration tested, only 10 % of the initial sugars present in A1 and A2 remained in the pretreated and hydrolysed biomass residues. Sugar extraction in samples B1 and B2 was however complete. No influence of H2O2 concentration on sugar solubilisation was found in sample C. Conclusions Enzymatic hydrolysis supported high efficiencies of glucose release from DWB but a low xylose release. Despite the efficient sugar solubilisation from PMWB mediated by the enzymatic method tested, the high bacterial content of this biomass promoted a rapid oxidation of the released sugars to organic acids and methanol. No significant influence of the biomass storage conditions was observed during enzymatic hydrolysis. Finally, alkaline-peroxide pretreatment increased the global sugar solubilisation, considering both, pretreated liquid fractions and hydrolysates from enzymatic hydrolysis. Overall, the evaluated alkaline-peroxide pretreatment increased sugar oxidation to organic acids and methanol regardless of the biomass type and storage conditions. 5. Acknowledgements This work was supported by the research unit UIC 071 of the regional government “Junta de Castilla y León – JCyL”, Spain. The authors thank “Instituto Nacional de Investigación y Tecnología Agraria y Alimentaria – INIA”, “Ministerio de Economía y 21 Competitividad – MINECO” (RTA2013-00056-C03-02) and “Junta de Castilla y León – JCyL” (VA094U14) for the financial support of this work. Judit Martin wish to thank “Junta de Castilla y León – JCyL” for providing her Doctorate Scholarship. 6. References 1. Acién F.G., Fernández, J.M., Molina-Grima E., 2014. 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