Microbial lipid production by oleaginous yeasts grown on Scenedesmus obtusiusculus microalgae biomass hydrolysate.

Due to increasing oil prices and climate change concerns, biofuels have become increasingly important as potential alternative energy sources. However, the use of arable lands and valuable resources for the production of biofuel feedstock compromises food security and negatively affect the environment. Single cell oils (SCOs), accumulated by oleaginous yeasts, show great promise for efficient production of biofuels. However, the high production costs attributed to feedstocks or raw materials present a major limiting factor. The fermentative conversion of abundant, low-value biomass into microbial oil would alleviate this limitation. Here, we explore the feasibility of utilizing microalgae-based cell residues as feedstock for yeast oil production. We developed an efficient, single-step enzymatic hydrolysis to generate Scenedesmus obtusiusculus hydrolysate (SH) without thermo-chemical pretreatment. With this eco-friendly process, glucose conversion efficiencies reached 90-100%. Cutaneotrichosporon oleaginosus, Cryptococcus curvatus and Rhodosporidium toruloides were cultivated on SH as sole nutrients source. Only C. oleaginosus was able to accumulate intracellular lipids, with a 35% (g lipid/g DCW) content and a yield of 3.6 g/L. Our results demonstrate the potential valorization of algal biomass into desired end-products such as biofuels.

Introduction

The ever-increasing energy demand in today’s industrial world led to the widespread use of non-renewable fossil fuels such as petroleum. The transition from a society with waste generating, linear production routes to one cyclic valorization path in conjunction with renewable resource management is one of the most demanding technological goals for establishing sustainable bioeconomy [1, 2]. This scenario particularly applied renewable energy supply routes that demand a switch from finite fossilto sustainable platform solutions. Moreover, dwindling of fossil resources, escalating enviroclass="Chemical">nmeclass="Chemical">ntal pollutioclass="Chemical">n, surgiclass="Chemical">ng class="Chemical">n class="Chemical">CO2 and greenhouse gas emissions, in addition to climate change have collectively driven the search for alternative energy sources [3]. Accordingly, technological innovations that enable a more sustainable lifestyle are coveted [4].

Biofuels have garnered great interest in recent years as alternatives for fossil fuel. In fact, plant-derived biofuel offers a partial solution to the ever-increasing energy demand, due to their renewability. However, this first-generation of biofuels, generated from edible crops, impacts agricultural activity and jeopardizes food security [1] To meet the current annual global-demand of biodiesel, more than double of the currently arable land would be required to grow crops that are explicitly grown for fuel production [5]. Consequently, alternative sources for biofuel production that do not affect food security are in high demand [1]. One of those alternatives is the use of oleaginous microorganisms such as n class="Species">algae aclass="Chemical">nd class="Chemical">n class="Species">yeast [6].

Oleaginous microorganisms accumulate class="Chemical">lipid at a miclass="Chemical">nimum of 20% (g class="Chemical">n class="Chemical">lipids/g dry cell weight (DCW)) [7]. However, lipid accumulation in oleaginous microorganisms, such as yeast, fungi and microalgae, is not a constitutive feature, but rather an adaptive response to particular environmental factors [8]. In environmental conditions abundant in carbon source and deficient in specific nutrients such as nitrogen, phosphorus or sulfur, oleaginous microorganisms convert excess carbon into fatty acids and incorporate them into triglycerides (TAGs) as a form of energy storage [9]. TAGs are stored in specialized organelles called lipid bodies (also known as lipid droplets) [10]. Single Cell Oils (SCOs) can be efficiently converted into biodiesel and biofuel [11, 12]. Various oleaginous yeasts have been subject to extensive investigations such as Yarrowia lipolytica, Rhodosporidium toruloids and Lipomyces starkeyi, with reports of lipid accumulation in excess of 70% (g lipid/g DCW) [13, 14]. The potential biotechnological applications of these oleaginous yeasts, utilizing various carbon sources have been previously reported [9, 15, 16].

Moon et al. first class="Disease">isolated Cutaneotrichosporon oleaginosus (class="Chemical">n class="Species">ATCC 20509) in 1978 from factory drain samples of the Iowa State University Dairy Farm. C. oleaginosus readily utilizes glucose, galactose, cellobiose, xylose, sucrose, and lactose as carbon source [1, 17–19]. Furthermore, this yeast is able to metabolize glycerol, N-acetylglucosamine, volatile fatty acids and ethanol and 4-hydroxymethylfurfural [20-22]. To improve the sustainability of SCOs from socio-economic aspects, Y. lipolytica has undergone extensive genetic engineering aimed at simultaneous sugar uptake (hexoses and pentoses) from complex and wastewater hydrolysates, which is an inherent ability in C. oleaginosus [8]. Depending on carbon, nitrogen sources and stress conditions (nitrogen, phosphate or sulfate limitation), cellular lipid accumulation in C. oleaginosus can reach up to 85% (g lipid/g DCW) [23, 24]. In addition to fast growth rate, this oleaginous yeast exhibits a fatty acid profile that mimics that of vegetable oils, specifically palm oil, with palmitic, stearic and oleic acid as dominant fatty acids [25]. McCurdy et al. reported that biodiesel B20 derived from C. oleaginosus TAGs meet the ASTM (D6751) certification [1, 9]. Through recycling and finding appropriate industrial sink for bio-compounds, a recent study touching on the socio-economic sustainability of C. oleaginosus SCOs has been recently prepared in our group [4].

Other oleaginous species that can be exploited for the biofuel sector include microclass="Species">algae. Iclass="Chemical">n coclass="Chemical">ntrast to iclass="Chemical">ndustrial crops, such as palm or caclass="Chemical">nola placlass="Chemical">nts, biomass geclass="Chemical">neratioclass="Chemical">n from microclass="Chemical">n class="Species">algae has high space and time yields. Globally, around 280 tons/ha of algae dry biomass and 3.9 tons/ha of forest biomass are produced every year [26]. Additionally, microalgae display high CO2 fixation ability (513 tons of sequestered CO2 per hectare per year). Specifically, 1.6-2 grams of CO2 is captured for every gram of algal biomass produced, at an efficiency of 80-99% [27, 28]. While microalgae can provide renewable oils by photosynthetically converting atmospheric CO2 to lipids, yields are conventionally lower compared to oleaginous yeast species [29-31]. In most algae oil production processes, the extracted cell residue is not contributing to the overall process economy [29, 31].

Yet recently, several value-adding outlets for this residue have been achieved, either by feeding it back into renewable production (class="Chemical">oil, food, feed, etc.) or by recycliclass="Chemical">ng of resources [4, 32]. Similar waste-free bioreficlass="Chemical">nery approaches have beeclass="Chemical">n coclass="Chemical">nsidered iclass="Chemical">n the desigclass="Chemical">n aclass="Chemical">nd optimizatioclass="Chemical">n of biogas productioclass="Chemical">n processes [33]. Iclass="Chemical">n that respect, the residual biomass, which is rich iclass="Chemical">n fermeclass="Chemical">ntable class="Chemical">n class="Chemical">sugars, can be used as feedstock for oleaginous yeasts cultivation [34]. Specifically, Scenedesmus spp. belong to the most common freshwater green algae. Scenedesmus obtusiusculus A189, a newly isolated member of the Chlorophyta genus, is characterized by high growth rates in fresh and saline media, in addition to high inherent carbohydrate content [35]. Fermentative conversion of various biomass sources into SCOs and subsequent biofuel via oleaginous yeasts has recently received an increasing interest in the scientific and industrial community [2, 4]. Specifically, microalgae biomass does not contain recalcitrant lignin, making it suitable for eco-friendly and cost-effective hydrolysis methods [35]. To that end, efficient hydrolysis of algae biomass could provide a sustainable stream of monomeric hexose and pentose sugars for fermentative growth. However, chemical biomass hydrolysis, which is most commonly applied in industry, tarnishes the eco-friendly aspect of biofuel production [2]. Alternatively, enzymatic hydrolysis efficiently generates sugar-rich fermentation media, yet necessitates thermo-chemical pretreatment steps to break down the lignocellulosic biomass components into monomeric hexose and pentose sugars [36]. These pretreatment steps release inhibitory compounds that hinder growth of subsequent oleaginous yeast inoculum in the prospective hydrolysate. Removal of these inhibitory compounds imposes additional costs, time and effort for the production process [37].

In this study, we examine the enzymatic hydrolysis of microclass="Species">algae-based cell residues aclass="Chemical">nd the subsequeclass="Chemical">nt use of its hydrolysate (SH) for the cultivatioclass="Chemical">n of oleagiclass="Chemical">nous class="Chemical">n class="Species">yeasts (C. oleaginosus). A single-step enzymatic hydrolysis process was devised and optimized, allowing efficient hydrolysis and saccharification of microalgae biomass, without thermo-chemical pretreatment. The resulting S. obtusiusculus hydrolysate (SH), barring expensive additives, was utilized as the sole fermentative media for C. oleaginosus, Cryptococcus curvatus and Rhodosporidium toruloides. The accumulated lipids, deposited as intracellular lipid bodies (LBs), were relatively quantified by Nile red analysis. Moreover, C. oleaginosus growth, dry biomass and lipid weight were evaluated.

Materials and methods

Algae strains and biomass determination

class="Species">Scenedesmus obtusiusculus (A189) residues were obtaiclass="Chemical">ned from Pharmaceutical Biology Group, Erclass="Chemical">nst Moritz Arclass="Chemical">ndt Uclass="Chemical">niversity (EMAU), Griefswald, Germaclass="Chemical">ny. class="Chemical">n class="Chemical">Water content was determined following the milling and drying of the algal samples at 60 °C overnight. Total carbohydrate concentration was determined by the thymol-sulfuric acid method [38]. The standard Kjeldahl procedure was utilized to determine the amount of protein in the algae biomass [39]. Total lipids were extracted according to Folch et al., and determined gravimetrically after solvent evaporation [40]. Biomass ash content was determined by following the AOAC procedure [41]. Biological replicates ensured reproducible measurements.

Enzymatic hydrolysis and preparation of SH

class="Species">Scenedesmus obtusiusculus Hydrolysate (SH) was prepared by hydrolyziclass="Chemical">ng autoclaved class="Chemical">n class="Species">algae biomass. Briefly, dry biomass samples weighing 50 g were transferred to 2 L glass bottles containing 1 L of 50.0 mM sodium acetate buffer, pH 5.0. Different hydrolytic enzyme mixtures were examined, including Cellic-Ctec2 (Novozymes, Denmark), Celli-Htec2 (Novozymes, Denmark), Pectinex (Novozymes, Denmark) and Fungamyl (Novozymes, Denmark). Hydrolytic Reactions were initiated by adding the enzyme solution and incubating the mixture at 50 °C for 72 h. Buffer and enzymes were sterile-filtered prior to hydrolysis. Samples were then spun down for 30 min. Cross-filtration using a 10 kDa membrane made from regenerated cellulose was completed under the following parameters: Inlet-Pressure (P1) of 2 bar, Repentant-Pressure (P2) of 0.3–0.5 bar and permeate was open to atmospheric pressure. Flow-Rates of retentate and permeate were adjusted to 2 L/min and 0.1 L/min respectively. A 0.2 µm filter capsules were installed at the outlet to sterilize the resulted hydrolysate. Biological triplicates of the SH were prepared.

Sugar analysis

class="Chemical">Sugar compositioclass="Chemical">n of the hydrolysate was determiclass="Chemical">ned by aclass="Chemical">n Agileclass="Chemical">nt 1100 series HPLC with a Refractive Iclass="Chemical">ndex (RI) detector (Shodex, RI101) aclass="Chemical">nd Ultraviolet Iclass="Chemical">ndex (Sedere-Fraclass="Chemical">nce, Sedex75). Followiclass="Chemical">ng cross-filtratioclass="Chemical">n, 5 µL sample was iclass="Chemical">njected oclass="Chemical">n aclass="Chemical">n class="Chemical">n class="Chemical">Aminex HPX-87P column (8% cross-linked resin, lead ionic, Bio-Rad) and separated at 70 °C with double-distilled water as mobile phase. Run parameters were set to a duration of 30 min, a flow rate of 0.4 mL/min and detection at 50 °C. Samples’ RI signal was aligned with that of internal standard curves.

Yeast strains and culture conditions

class="Species">Yeast straiclass="Chemical">ns Cutaclass="Chemical">neoclass="Chemical">n class="Species">trichosporon oleaginosus (ATCC 20509), Cryptococcus curvatus (CBS 5324) and Rhodosporidium toruloides (NP11) were maintained on yeast peptone dextrose (YPD) agar (20 g/L peptone, 10 g/L yeast extract, 20 g/L glucose, 20 g/L agar) at 4 °C for short-term storage. Minimal nitrogen media (MNM) (30 g/L glucose, 1.5 g/L yeast extract, 0.5 g/L NH4Cl, 7.0 g/L KH2PO4, 5.0 g/L Na2HPO4·12H2O, 1.5 g/L MgSO4·7H2O, 0.08 g/L FeCl3·6H2O, 0.01 g/L ZnSO4·7H2O, 0.1 g/L CaCl2·2H2O, 0.1 mg/L MnSO4·5H2O, 0.1 mg/L CuSO4·5H2O, 0.1 mg/L Co(NO3)2·6H2O; pH 5.5) was adopted from Suutari et al. for induction of lipogenesis [42].

Following pre-culturing in YPD broth for 24 h, class="Species">yeast cells were ceclass="Chemical">ntrifuged, washed with class="Chemical">n class="Chemical">PBS buffer (8 g/L NaCl, 0.2 g/L KCl, 1.44 g/L Na2HPO4, 0.24 g/L KH2PO4; pH 7.4) and inoculated in 250 mL baffled flasks, containing 50 mL of MNM, at an initial seeding OD600 of 0.5. Incubation lasted for 4 days in a rotary shaker at 120 rpm and 28 °C. To evaluate the capability of utilizing hydrolysates for yeast growth and lipid accumulation, the selected oleaginous strains were cultured solely on SH lacking any additives or carbon supplementation. Biological cultures of the oleaginous yeasts were carried out in triplicates.

Nile red staining

Samples were analyzed in technical triplicates using a modified protocol from Sitepu et al. [43]. Briefly, 225 µL of each class="Species">yeast culture was traclass="Chemical">nsferred to a 96-well black microtiter plate. Serial dilutioclass="Chemical">ns were performed iclass="Chemical">n triplicates to eclass="Chemical">nsure aclass="Chemical">n optical declass="Chemical">nsity < 1 before 50 µl class="Chemical">n class="Chemical">DMSO was added to each well. Initial absorbance readings were taken at 600 nm and for growth monitoring and correction of fluorescence readings for growth variation. A volume of 25 µL Nile red was then added to each well (final concentration of 50 µg/mL). Fluorescence measurements (recorded before and after Nile red addition) at excitation at 530/25 nm; emission at 590/35 nm; and kinetic reading for 5 min with 30 s interval were taken. Maximal emission values were determined and fluorescence measurements were corrected for variation in cell density by dividing the fluorescence unit by background optical density OD600 values.

Gravimetric analysis

Technical triplicates of the total class="Chemical">lipid coclass="Chemical">nteclass="Chemical">nt of the oleagiclass="Chemical">nous class="Chemical">n class="Species">yeasts were determined by a modified method by Folch et al. [40]. Briefly, 15 mL of yeast cultures were washed and homogenized using an Avestin Emulsiflex at a sample port pressure of 1200 bar and a chamber pressure of 8 bar. Lipids from the homogenate were extracted with 6 mL of Folch solution (2:1 chloroform: methanol). Lipid extraction continued overnight at room temperature and shaking speed of 120 rpm. Subsequently, 1.2 mL of 0.9% NaCl were added to aid phase separation. The lower phase was aspirated using a syringe and added to pre-weighed glass vessels. The chloroform was fully evaporated under a nitrogen stream and glass vials were weighed again. The extracted lipid samples were used to calculate lipid content as total lipid weight and as percent of dry yeast weight.

Dry biomass determination

A volume of 2 mL of each n class="Species">yeast culture was traclass="Chemical">nsferred iclass="Chemical">n triplicates to pre-weighed 2 mL Eppeclass="Chemical">ndorf tubes. The tubes were weighed agaiclass="Chemical">n the followiclass="Chemical">ng ceclass="Chemical">ntrifugatioclass="Chemical">n at 14,000 g for 5 miclass="Chemical">n, washiclass="Chemical">ng aclass="Chemical">nd dryiclass="Chemical">ng at 60 °C overclass="Chemical">night. Measuremeclass="Chemical">nts were recorded iclass="Chemical">n triplicates by subtracticlass="Chemical">ng the weight of the sample tubes from their respective pre-weights.

Results

Algae biomass analysis

In this study, the biomass composition of S. obtusiusculus was determined (Table 1). This green class="Species">algae displays high amouclass="Chemical">nts of class="Chemical">n class="Chemical">carbohydrates and crude proteins comprising 34% and 49% (g/g DCW) respectively. Other components encompassed water, lipids and ash measured at 3.7, 8.3 and 1.9% (g/g DCW), respectively. The acquired biomass data suggest that S. obtusiusculus can be quantitatively hydrolyzed by chemical and enzymatic systems to release monomeric pentose and hexose sugars, which could serve as a carbon source for microbial cultivation.

Table 1

Biochemical composition of S. obtusiusculus, calculated as percent of total dry weight

Biomass component	Content %
	% (g/g dry biomass weight)
Water	3.7
Carbohydrates	33.8
Proteins	48.7
Lipids	8.3
Pigments, secondary metabolites	3.6
Ash	1.9

Relative standard deviation for all given numbers is ≤ ± 2%

Biochemical composition of S. obtusiusculus, calculated as percent of total n class="Disease">dry weight

Hydrolysis of S. obtusiusculus dry biomass

Various commercial hydrolase enzyme mixtures were tested for their biomass liquefaction efficiencies including a cellulase mix (Cellic-Ctec2, class="Chemical">Novozymes), a class="Chemical">n class="Chemical">hemicellulose-mix (Cellic-Htec2, Novozymes), a pectinase mix (Pectinex, Novozymes), and an amylase mix (Fungamyl, Novozymes). Biomass to glucose conversion ratios from the various enzyme mixtures is presented in Fig. 1a. The cellulase mix (Cellic CTec 2) exhibited the best activity. Glucose monomerization reached a saturation level between 12 and 14 g/L starting from a 50 g algae biomass, which translates to a glucose yield of 0.24-0.28 g/g of DCW. Furthermore, a 1% (v/v) of cellulase mixture Cellic-Ctec2 was combined with varying concentrations of the remaining enzyme mixtures, none of which yielded a significantly better conversion ratio (Fig. 1b).

Fig. 1

Glucose concentration of SH displayed as a factor of various enzymes mixes and concentrations (a). Glucose concentration of SH displayed as a factor of the combination of enzyme mixtures with Cellic CTec 2 (b)

n class="Chemical">Glucose coclass="Chemical">nceclass="Chemical">ntratioclass="Chemical">n of SH displayed as a factor of various eclass="Chemical">nzymes mixes aclass="Chemical">nd coclass="Chemical">nceclass="Chemical">ntratioclass="Chemical">ns (a). class="Chemical">n class="Chemical">Glucose concentration of SH displayed as a factor of the combination of enzyme mixtures with Cellic CTec 2 (b)

To assess the efficiency of class="Chemical">glucose liberatioclass="Chemical">n from the class="Chemical">n class="Species">algae biomass, measurements of enzymatic hydrolysis were compared with those of acidic hydrolysis. Table 2 confirms that glucose conversion efficiencies reached (90–100%), with glucose constituting up to two-thirds (g/g of DCW) of the total carbohydrate content of SH.

Table 2

A comparison of monosaccharide content % (g/g dry biomass weight) resulting from acidic and enzymatic hydrolysis

Sugar	Acidic hydrolysis	Enzymatic hydrolysis	Conversion
	% (g/g dry biomass weight)	% (g/g dry biomass weight)	Efficiency (%)
Glucose	22	20–22	90–100
Mannose		–	–
Galactose	10	2–2.5	20–25
Rhamnose	~ 1	0	0
Fucose	~ 1	0	0
Ribose	~ 1	0	0

Relative standard deviation for all given numbers is ≤ ± 2%

A comparison of class="Chemical">monosaccharide coclass="Chemical">nteclass="Chemical">nt % (g/g class="Chemical">n class="Disease">dry biomass weight) resulting from acidic and enzymatic hydrolysis

The combined enzymatic conversion rate of class="Chemical">mannose aclass="Chemical">nd class="Chemical">n class="Chemical">galactose, which were not distinguished by HPLC, was limited to only 20–25% (g/g DCW).

However, total class="Chemical">sugar coclass="Chemical">nceclass="Chemical">ntratioclass="Chemical">n iclass="Chemical">n the hydrolysate was still relatively low (12–14 g/L class="Chemical">n class="Chemical">glucose) given a carbohydrate content of 34% (g/g DCW). Accordingly, hydrolysis was repeated with higher amounts of biomass retaining a 1% (v/v) concentration of the cellulase mixture. As a result, glucose concentration in SH reached 48 g/L starting with a 200 g Scenedesmus dry biomass. This accounts fora glucose yield of 0.24 g/g of DCW, maintaining the high conversion efficiencies of (90–100%). Subsequent experiments were conducted with SH comprising this high glucose concentration (48 g/L).

Yeast growth and lipid production

High throughput class="Chemical">Nile red screeclass="Chemical">niclass="Chemical">ng was employed to determiclass="Chemical">ne the class="Chemical">n class="Chemical">lipid yield of three oleaginous yeast strains C. oleaginosus, C. curvatus and R. toruloides. Following a 4-day fermentative growth on SH as the sole carbon source, Nile red analysis revealed low lipid content in C. curvatus and R. toruloides. Contrarily, C. oleaginosus was able to accumulate a considerable amount of lipid bodies when cultivated on the same media (Fig. 2).

Fig. 2

Rapid estimation of lipid contents in C. oleaginosus, C. curvatus and R. toruloides following 4 days cultivation on SH, determined by Nile red assay

Rapid estimation of class="Chemical">lipid coclass="Chemical">nteclass="Chemical">nts iclass="Chemical">n C. oleagiclass="Chemical">nosus, class="Chemical">n class="Species">C. curvatus and R. toruloides following 4 days cultivation on SH, determined by Nile red assay

Thus, C. oleaginosus underwent subsequent experiments to determine growth rate and class="Chemical">lipid absolute quaclass="Chemical">ntitatioclass="Chemical">n wheclass="Chemical">n cultivated oclass="Chemical">n Mclass="Chemical">n class="Chemical">NM and SH. To evaluate C. oleaginosus growth-rate in MNM and SH media, optical density at 600 nm was measured over 4 days (Fig. 3). SH media resulted in highest final growth for C. oleaginosus measured at OD600 of about 30, compared to OD600 of about 22 in MNM.

Fig. 3

C. oleaginous growth trend when grown on MNM and SH over 4 days in a shake flask fermentation

C. oleaginous growth trend when grown on Mn class="Chemical">NM aclass="Chemical">nd SH over 4 days iclass="Chemical">n a shake flask fermeclass="Chemical">ntatioclass="Chemical">n

Gravimetric analysis was performed to determine total class="Chemical">lipid coclass="Chemical">nteclass="Chemical">nt iclass="Chemical">n C. oleagiclass="Chemical">nosus followiclass="Chemical">ng fermeclass="Chemical">ntatioclass="Chemical">n oclass="Chemical">n Mclass="Chemical">n class="Chemical">NM and SH in shake flasks (Fig. 4). The yeast accumulated nearly 61% and 35% (g lipid/g DCW) lipids when grown on MNM and SH, respectively. After 4 days fermentation, lipid yield in C. oleaginosus reached about 5.3 g/L of culture when cultivated on MNM, and 3.6 g/L of culture when cultivated on SH media.

Fig. 4

Lipid content (%, g lipid/g DCW) and lipid yield (g/L) of C. oleaginous cultivated in MNM and SH media for 4 days in shake flask fermentation

class="Chemical">Lipid coclass="Chemical">nteclass="Chemical">nt (%, g class="Chemical">n class="Chemical">lipid/g DCW) and lipid yield (g/L) of C. oleaginous cultivated in MNM and SH media for 4 days in shake flask fermentation

Discussion

For years, microclass="Species">algae have beeclass="Chemical">n exploited as a source for value-added products, with class="Chemical">numerous commercial applicatioclass="Chemical">ns that iclass="Chemical">nclude eclass="Chemical">nhaclass="Chemical">nciclass="Chemical">ng the class="Chemical">nutritioclass="Chemical">nal value of food aclass="Chemical">nd aclass="Chemical">nimal feed, as well as beiclass="Chemical">ng iclass="Chemical">ncorporated iclass="Chemical">nto cosmetics [44]. The sigclass="Chemical">nificaclass="Chemical">nt properties of microclass="Chemical">n class="Species">algae biomass as raw material for microbial cultivation include high carbohydrates contents and lack of recalcitrant lignin [45]. The use of microbes as a platform for lipid and subsequent biofuel and biodiesel production offers: (1) renewability and potential sustainability, (2) requires less labor and fewer raw materials, (3) is easier to scale up, (4) does not compete with edible-plants for land, (5) generates less waste and (6) is not affected by season or climate [46]. Recently, the valorization of seagrass and brown macroalgae biomass as feedstock for C. oleaginosus lipid production, in addition to the techno-economic feasibility of the bioprocesses have been conducted in our group [2, 4]. In this study, S. obtusiusculus biomass was chosen as feedstock for oily yeast growth, due to its high carbohydrates content 34% (g/g dry biomass weight). In comparison, Scenedesmus obliquus, Chlorella vulgaris, Chlamydomonas rheinhardii, and Dunaliella salina algae species exhibit sugar content per dry biomass weight of 10%, 12%, 17% and 32% respectively [44].

Complete chemical hydrolysis (class="Chemical">H2SO4) have beeclass="Chemical">n regularly implemeclass="Chemical">nted for the productioclass="Chemical">n of hydrolysate from ligclass="Chemical">nocellulosic biomass [47, 48]. Lately, two-stage hydrolysis processes starticlass="Chemical">ng with mild chemical treatmeclass="Chemical">nt (class="Chemical">n class="Chemical">dilute sulfuric acid) and followed by enzymatic hydrolysis have gained popularity amongst industrial applications [49]. Corn-stover biomass hydrolysis required a pretreatment of the biomass with 0.5 M NaOH at 80 °C for 75 min [50]. However, these methods often generate inhibitory substances that might hinder or completely abolish the growth of microorganisms cultivated in the resulting hydrolysates. Furfural was found to elongate the lag-phase; while benzoic acid reduced growth rate and biomass yield [51]. Thus, complex detoxification step would be necessary prior to the fermentation, ensuing additional costs and tarnishing the eco-friendly aspect of the biofuel production process [36]. Accordingly, we opted to use a single-step enzymatic approach in this study that would allow efficient hydrolysis of algae biomass without the need for any pretreatment steps. Sterilization of S. obtusiusculus biomass was performed in a laboratory-scale autoclave at 120 °C for 15 min to eliminate the microbial contaminants present within the microalgae residue. This not considered as a pretreatment step since hydrolysis of hemicellulose and cellulose only starts at temperatures greater than 150 °C [37]. Efficient saccharification of S. obtusiusculus biomass by single-step enzymatic hydrolysis using a cellulase mix was possible. In fact, the Cellic CTec 2 combines a number of different enzymatic activities (exo-, endo-glucanase activity and proteinase activity). The optimal activity was obtained at an industrially relevant concentration of 1% (v/v) at 50.0 °C and pH 5.0 in 50 mM, sodium acetate buffer for 72 h. Quantitative biomass to glucose conversion ratio remained high, even when raising substrate amounts up to 200 g/L. Beyond this point, viscosity was too high for effective hydrolysis. Notably, the diverse heteropolymeric structure of algal cell wall might account for the low conversion efficiency of mannose and galactose [52]. Commonly available enzyme mixtures including Celli-Htec2 (Novozymes), Pectinex (Novozymes) and Fungamyl (Novozymes) failed to liberate total monosaccharides from these structures.

class="Chemical">Lipid accumulatioclass="Chemical">n iclass="Chemical">n oleagiclass="Chemical">nous class="Chemical">n class="Species">yeasts is usually triggered upon excess carbon and nutrient deficiency (e.g., nitrogen phosphate or sulfur). Lipid yields and fatty acid profile vary depending on the type and concentration of the carbon and nitrogen source [1, 7, 53]. Single-step enzymatic hydrolysis generated glucose-rich hydrolysate, the preferred monomeric sugar for microbial fermentation. A 10 KDa cross-filtration was subsequently implemented, and the permeate product (SH), exhibiting now nitrogen limitation and high C/N ratio, was used as fermentation media for high lipid accumulation in yeast.

The oleaginous class="Species">yeast Cutaclass="Chemical">neotrichosporoclass="Chemical">n oleagiclass="Chemical">nous is able to metabolize a broad class="Chemical">n class="Chemical">monosaccharide spectrum including hexoses and pentoses into intracellular TAGs [23]. This yeast was also able to grow well in a model medium with a carbohydrate mixture that resembled a typical microalgae derived-hydrolysate [54]. In this work, C. oleaginosus, as well as C. curvatus and R. toruloides were cultivated in S. obtusiusculus hydrolysate. Most interestingly, in contrast to the other two oleaginous yeasts, high-throughput Nile red analysis indicated that only C. oleaginosus was able to accumulate significant amounts of intracellular lipids when grown in SH.

Without any nutritional addition to the hydrolysate (class="Chemical">biotin, class="Chemical">n class="Species">yeast extract, pure glucose…), SH was utilized as the sole carbon source for lipid production in C. oleaginosus. The assessment was conducted on the basis of lipid accumulation. The results were evaluated along with the data from cultivation in the synthetic MNM—a medium known to induce lipid biosynthesis in oleaginous yeasts [55]. C. oleaginosus grew faster in SH media, in comparison with MNM. The yeast yielded 61% (g lipid/g DCW) of intracellular lipid when grown on MNM, and about 35% (g lipid/g DCW) when grown on SH. Following fermentation, C. oleaginosus achieved total lipid yield of 3.6 g/L when cultivated on SH media. In scaled-up experiments, SH would prove a cost-effective alternative for the relatively expensive synthetic MNM media.

To establish nutrient limitation, the class="Chemical">microalgae hydrolysate uclass="Chemical">nderweclass="Chemical">nt ultrafiltratioclass="Chemical">n thus elimiclass="Chemical">naticlass="Chemical">ng proteiclass="Chemical">ns aclass="Chemical">nd peptides aclass="Chemical">nd establishiclass="Chemical">ng a high C/class="Chemical">n class="Chemical">N ratio. However, other factors besides nitrogen-limitation could induce lipogenesis in C. oleaginosus. Effect of phosphate and sulfur limitation on lipid accumulation in oleaginous yeasts have been previously reported [56]. High C/P ratio prompted high lipid yield in R. toruloides even in the presence of excess nitrogen [22]. For future work, soluble phosphates could be precipitated and removed by interaction with metal ions, such as Ca2+, Mg2+, or Fe3+ [57], and the resulting hydrolysate—now exhibiting high C/N and C/P ratios—could allow for even higher for lipid accumulation by C. oleaginosus.

Cutaneoclass="Species">trichosporon oleaginosus cultivated iclass="Chemical">n S. obtusiusculus hydrolysate achieved a high growth rate aclass="Chemical">nd accumulated substaclass="Chemical">ntial amouclass="Chemical">nt of iclass="Chemical">ntracellular class="Chemical">n class="Chemical">lipids. Previous research showed that this yeast accumulates lipid in the form of triacylglycerides, with a fatty acid (FA) profile consisting mainly of C16 and C18 FA [58]. Palmitic acid, stearic acid and oleic acid constitute the major raw material for downstream processing and subsequent conversion into green biofuels [59]. Furthermore, chemo-catalytic conversion of lipids produced by C. oleaginosus into biodiesel was achieved with a 98.9% w/w recovery [9]. The physical properties of resulting B20 - comparable to Soybean B20 - meet the ASTM requirements [9]. This FA profile makes SCOs from C. oleaginosus a suitable alternative for plant and vegetable oils.

Conclusion

This study demonstrated that class="Species">Scenedesmus obtusiusculus biomass could be valorized as a substrate for microbial class="Chemical">n class="Chemical">lipid production. A single-step enzymatic hydrolysis was implemented that efficiently released monomeric sugars from the biomass without the need for any pretreatment. This approach alleviated the need for detoxification steps, reduced upstream processing costs and maintained the eco-friendly aspect of biofuel production. The oleaginous yeast C. oleaginosus was able to grow fast and accumulate 3.6 g/L of lipids when cultivated on the microalgae hydrolysate, and the resulting microbial oil could be converted to high-grade biodiesel. Microalgae biomass offer value-added biofuel yield potential as compared to terrestrial plantation; biomass-to-fuel conversion processes are improved by necessitating no agricultural land, alleviating direct competition with food security and requiring low water and resource demand. Furthermore, the integration of yeast and algae species in a single SCO platform towards “zero concepts” with respect to emission and excess resources has recently been reported. In one study, the oleaginous microalgae Phaeodactylum tricornutum was supplemented with CO2 supplied from the oleaginous yeast C. curvatus in a co-fermentation approach [32]. In another study focused on the holistic valorization of unexploited marine biomass, a waste-free, microbial oil-centered cyclic bio-refinery approach integrated the production of yeast lipids and animal feed with precious metal biosorbents [4]. In that respect, the algal effluent resulting from the ultra-filtration of the algae hydrolysate should be further characterized and profiled for possible added-value in the process described in this study.