Mixotrophic growth of the polar marine microalga Chlamydomonas sp. RCC2488 (malina) using potato peel hydrolysates as carbon source

doi:10.21203/rs.3.rs-5321861/v1

Download PDF

Research Article

Mixotrophic growth of the polar marine microalga Chlamydomonas sp. RCC2488 (malina) using potato peel hydrolysates as carbon source

https://doi.org/10.21203/rs.3.rs-5321861/v1

This work is licensed under a CC BY 4.0 License

Journal Publication

published 22 Feb, 2025

Read the published version in BioEnergy Research →

You are reading this latest preprint version

Chlamydomonas malina is a polar microalga with high PUFA production under phototrophic conditions. In this study, C. malina, strain RCC2488, was cultivated on glucose and potato peel hydrolysates (PPH), at 120 µmol photons m^-2 s^-1 of light intensity, to investigate the feasibility of growing under mixotrophic conditions. Potato peels were subjected to an acid-hydrothermal pretreatment. The resulting material was separated into three fractions, each of which was prepared for the subsequent enzymatic reaction with a-amylase and amyloglucosidase: PPH1, supernatant + paste collectively; PPH2, only paste, H₂SO₄ – removed with water; PPH3, only supernatant. Transmembrane glucose transport, growth kinetics, macromolecular composition, as well as lipid types, and fatty acid profile of C. malina were determined. The microalga was able to grow and transport glucose under mixotrophic but not heterotrophic conditions. The highest concentration of reducing sugar (glucose) was found in PPH1 and PPH3. However, the lowest biomass content was found in those hydrolysates, likely due to the presence of furfural and hidroxymethylfurfural (HMF). In contrast, C. malina had the highest biomass productivity in glucose and PPH2. The PPHs promoted lipid accumulation in C. malina but with different lipid composition. PPH1 and PPH3 promoted the synthesis of TAG, while PPH2 allowed the accumulation of polar lipids with high PUFA content. Therefore, pretreatment and hydrolysis optimization are necessary to use potato peel as an efficient carbon source without toxic by-products. Mixotrophic cultivation of C. malina was possible but not ideal since higher growth is attained under phototrophic conditions.

Polar microalgae

mixotrophic

potato waste

furfural

PUFA

Microalgae are natural single-cell ‘factories’ for a tailored production of a wide variety of economically valuable chemical compounds with high complexity, such as pigments and polyunsaturated fatty acids (PUFA). PUFAs, such as omega-3 fatty acids, are used by the pharmaceutical industry due to their high therapeutic significance in clinical and epidemiological studies [1]. These compounds are typically not synthesized by humans and animals [2], but they are naturally synthesized by polar microalgae. This is because these microorganisms need to modify their cellular membrane to adapt to cold environments by increasing the abundance of PUFAs [3–7], which help maintain membrane fluidity and flexibility during low or freezing temperatures. In addition, PUFA synthesis protects the membrane against damage caused by cold due to its high content of double bonds, which scavenge free radicals [8]. Some good examples of high PUFAs polar microalgae producers are Chloromonas platystigma, Chlamydomonas sp. RCC2488 malina, and Macrochloris rubreoleum [3, 4, 7]. An interesting feature of polar microalgae is their metabolic flexibility, which switches from phototrophic to mixotrophic/heterotrophic mode, depending on the environmental polar conditions, which are usually oscillating and extreme. The mixotrophic mode – which uses an organic carbon source under light conditions – may be associated with a two to three-fold increase in omega-3 fatty acid productivity compared with phototrophic cultivation systems [1, 9]. Indeed, cell growth is higher under mixotrophic conditions because of the higher energy density of the carbon source in comparison with CO₂ [1, 9]. However, not all the microalgae species can switch from phototrophic to mixotrophic metabolism and metabolize an organic carbon source in the presence of light. Therefore, the microalgae species that can grow under mixotrophic mode are limited [9].

Glucose is one of the preferred carbon sources of most living cells on Earth [2, 10]. Hence it is often used in many mixotrophic microalgae cultures [1, 2, 10]. However, the high cost of glucose is the major commercial concern of mixotrophic cultures [11]. Therefore, multiple studies have focused on improving the economic feasibility of biomass and metabolite production from microalgae. One of these methods is to decrease production costs by using alternative carbon sources, like organic wastes. Low-cost substrates such as empty palm fruit, sorghum juice and coconut water, among others, have been tested for this purpose [12]. Although several organic wastes have been successfully tested for microalgal cultivation, evaluating the use of locally available wastes is important to improve the process’s economic and environmental feasibility. In this context, in the North of Norway, potato peel (a high glucose-containing processed potato waste) is an interesting potential candidate as a carbon source for microalgae growth. Potato peels had previously been used as the carbon source for cultivating Saccharomyces cerevisiae and Bacillus [13–17]. The hydrolysis of this bio-waste resulted in high yields of reducing sugars when an acid or hydrothermal pretreatment followed by an enzyme hydrolysis was used [18–23]. Usually, a pretreatment step to modify the structure of the lignocellulosic matrix followed by an enzymatic hydrolysis is employed. A crucial consideration is that this treatment should not generate toxic byproducts that can inhibit microalgal growth. In this study, we evaluated the capability of the polar microalga C. malina to grow under mixotrophic conditions. We tested whether C. malina can transport glucose through its membrane under mixotrophic conditions using a ¹⁴C tracer. Then, we assessed whether C. malina could grow using pure glucose as the carbon source in mixotrophic mode. Different potato peel hydrolysate pretreatments as the carbon source for the growth, biomass productivity, macromolecular composition, lipid classes and profile, under mixotrophic conditions were also evaluated.

Strain and culture conditions

Chlamydomonas sp. RCC2488 (strain Chlamydomonas sp. MALINA FT89.6 PG5, hereafter referred to as C. malina) is a marine microalga isolated from the Beaufort Sea, within the Arctic Ocean [24] and kept at the Roscoff Culture Collection. The algal stock was maintained in agar plates containing modified f/2 medium. For all experiments, the inoculum was prepared in 250 mL shake-flasks containing 50 mL of modified f/2 medium at 4 °C and 100 rpm, maintained at 120 mmol m^-2 s^-1 of light intensity for 5 days. Modified f/2 medium was composed by the following salts (in mM): NaNO₃ 31.8, NaH₂PO₄H₂O 1.32, FeCl₃·6H₂O 0.105, Na₂EDTA·2H₂O 0.105, the following trace elements (in mM): CuSO₄·5H₂O 0.36, Na₂MoO₄·2H₂O 0.234, ZnSO₄·7H₂O 0.69, CoCl₂·6H₂O 0.378, MnCl₂·2H₂O 8.19, and the following vitamins (in mM): thiamine HCl 26.6, biotin 0.18 and cyanocobalamin 0.036 [4]. For the cultivation experiments, the preparation of this medium included the use of seawater from the North Atlantic shoreline of Tromsø (Norway) with a salinity of approximately 35. For the glucose transport experiments, the salinity was adjusted with 35 g/L NaCl in distiller water.

All experiments were conducted in 500 mL shaking flasks at 4 °C and 100 rpm. The positive control experiment contained additionally 5 g/L pure glucose. PPH1 and PPH3 (see below) cultivation media were diluted 1:10 for the cultivation experiments (i.e. 10 mL of PPH diluted in 90 mL of f/2 for PPH1 and PPH3 cultivations). The flasks were inoculated at a cell concentration of ~ 2 x 10⁵ cell mL^-1. Cultures were kept at 120 mmol m^-2 s^-1 of light intensity for 10 days. At this time, cells were harvested by centrifugation at 2000 x g for 5 min, washed with 0.5 M ammonium formate, centrifuged again as before, and the pellets were stored at -70 °C for further analyses. All experiments were carried out in tetraplicates.

Potato peel hydrolysates (PPH)

Fresh potato peel waste from different Norwegian potato varieties was collected from the kitchen of the lab members and purchased at the local market. This waste was immediately stored at -20 °C until use. The frozen potato peel waste was washed with distilled water and dried in an air oven (ThermoFisher Scientific, HeraTherm, Waltham, MA, USA) at 85 °C for 3 days. Then it was ground in a mortar to get a powder with a particle size between 0.1 and 1 mm. An acid-thermal pretreatment was applied to 10 g of potato peel powder by adding 100 mL of 1% H₂SO₄, mixed for 1 hour and then autoclaved at 121 °C for 20 minutes. The resulting material was separated into three fractions, each of which was prepared for the subsequent enzymatic reaction in a sodium acetate – calcium chloride buffer with the following differences:

PPH1) Potato peel paste and supernatant collectively.

For preparation, 0.58 mL glacial acetic acid and 0.074 g CaCl₂ was added to 100 mL of the suspension and the pH was adjusted to 5 with 3M NaOH.

PPH2) Potato peel paste, H₂SO₄ – free.

This paste was separated from the supernatant by filtration with Whatman filter paper no. 1 and rinsed several times with distilled water. To this paste (free of sulfuric acid), 100 mL of a solution containing 0.58 mL glacial acetic acid and 0.074 g CaCl₂ was added, and the pH was adjusted to 5 with 3M NaOH.

PPH3) Potato peel supernatant.

This material was separated from the paste by filtration with Whatman filter paper no. 1. To this supernatant (~100 mL), 0.58 mL glacial acetic acid and 0.074 g CaCl₂ were added, and the pH was adjusted to 5 with 3M NaOH.

Finally, for all treatments, the enzymatic hydrolysis started by adding 1 mL of 𝛼-amylase (2800 U, Megazymes Ltd.), mixed by vortexing and incubated at 100 °C for 15 min with periods of mixing at 5, 10, and 15 minutes. Afterward, samples were cooled down in a water bath at 50 °C. Then 1 mL of amyloglucosidase (3300 U, Megazymes Ltd.) was added and incubated at 50 °C for 30 minutes with mixing periods of 5 minutes. The reaction was stopped by cooling it down in a cold-water bath. At the end, the pH was adjusted to 7 with 3M NaOH and stored at 4 °C for further experiments.

The presence of furfural, hydroxymethylfurfural (HMF), and acetic acid in the three PPH treatments was determined (Table 1). As reference, we measured the concentrations of glucose, furfural, HMF, and acetic acid in potato peel powder in water solution (10% w/v) incubated for 1 hour at room temperature (control 1), followed by hydrothermal treatment (without H₂SO₄) at 121 °C for 20 minutes in an autoclave (control 2) and followed by enzymatic hydrolysis as previously described (control 3).

Growth measurement

Cell growth was assessed using a hematocytometer (cell count) and dry cell weight (DCW) measurement. A correlation was made between the optical cell density and the dry weight using the equation:

W = (0.884 × A₇₅₀) + 0.0117

Where, W is the dry cell weight (g L^-1), and A₇₅₀ is the absorbance measured at 750 nm. In the interval of 0.1 ≤OD₇₅₀nm≤0.8, the OD at 750 nm of C. malina and dry cell weight (g L^-1) showed a linear relationship with an R² = 0.9987. Culture samples (0.5 – 1 mL) were collected daily to measure the absorbance at 750 nm in a 1 cm micro-cuvette using a spectrophotometer (Hach-Lange DR3900, Hach, International). The algal dry cell weight (DCW) was evaluated gravimetrically by filtrating 5-10 mL of culture broth through a pre-dried and pre-weighed 0.45 mm pore size nitrocellulose membrane filter (Merck Millipore, MA, USA).

Initial glucose transport rate

The kinetics of the transmembrane glucose transport was conducted as previously described [10, 25]. Briefly, C. malina cells were cultivated in f/2 medium containing glucose in shaking flasks under hetero (darkness) and mixotrophic conditions (120 mmol m^-2 s^-1) until mid-log phase (5 days), then the cells were centrifuged at 14,000 x g for 10 min at 4 °C. The supernatant was discarded, and the cellular pellet was kept on ice, washed in f/2 medium twice, and dissolved in the same medium without a carbon source at a cell density of 1 x 10⁸ cells mL^-1. For the [¹⁴C]-glucose uptake assay, 540 𝜇L of cell suspension kept on ice was incubated for 15 min at 4 °C, then the kinetics started by adding 60 𝜇L of [¹⁴C]-glucose (0.5 mM, 5 mCi/mmol). The suspension was maintained at 4 °C with shaking. At 0, 5, 10, 20, 40, and 60 min intervals, 50 𝜇L of the sample was taken, filtered it immediately through a membrane filter (pore size 0.45 𝜇m) and washed the filter three times with f/2 medium. The filters were dried and placed in vials with 5 mL of Ecolite scintillation cocktail (ICN Biomedicals, Costa Mesa, CA, USA). Radioactivity was measured using a scintillation counter (Beckman LS6000IC, Fullerton, CA, USA). The [¹⁴C]-glucose uptake rates were calculated from the initial, linear data trend, based on the relation of intracellular [¹⁴C]-glucose versus time.

Carbohydrate, reducing sugar, and protein analyses

For biomass analysis, the total carbohydrate content was determined using the phenol-sulfuric acid method [26] after hydrolysis of the biomass with 1N HCl to yield simple sugars. The glucose, furfural, HMF, and acetic acid content of each PPH treatment and the controls were quantified using an HPLC. Briefly, 1 mL aliquots from each PPH treatment sample and from the controls were centrifuged at 14,000 x g for 10 min at 4 °C. The supernatant was filtered through a 0.2 µm nylon filter into an HPLC vial and stored at 4 °C until analysis. A Waters HPLC system (Millipore, Co., Milford, MA, USA) comprised of a 600E quaternary bomb, 717 automatic injector was used to process the samples. Refractive index detector was used for the identification of glucose and acetic acid (Model 2410 Waters, Millipore Co., Milford, MA, USA), while photodiode array detector for furfural and HMF detection (Model 996, Waters, Millipore Co., Milford, MA, USA); both using an Aminex HPX-87H column (Bio-Rad, Hercules, CA). As mobile phase, 5 mM sulfuric acid at a flow rate of 0.6 mL min^-1 and 60 °C was used [27].

For protein determination in the biomass, the Lowry method was used [28] after alkaline hydrolysis of the sample with 1N NaOH.

Lipid extraction, quantification, and analyses

Lipids from C. malina were extracted using organic solvents, such as chloroform and methanol, according to previous very well-established protocols [4, 7, 29]. These extracts were separated into neutral lipids (triacylglycerols, TAGs) and polar lipids by using solid-phase extraction as described previously [4, 7, 29]. After separation, the lipids of each fraction were derivatized to fatty acid methyl esters (FAMEs) with 5% H₂SO₄ in methanol and heated at 70 °C for 3 h. Phases were separated using a mix of hexane:H₂O (1:1). FAMEs from both neutral (TAGs) and polar lipids were identified and quantified by gas chromatography (GC) fitted with a Flame Ionization Detector (Scion 436, Bruker, USA) and an Agilent CP-Wax 52 CB column (Agilent Technologies, USA) using a splitless injector [4, 7, 29]. External Supelco® 37-component standards (Sigma-Aldrich, USA) were used to identify the FAMEs.

Calculations and statistical analysis

Cell growth kinetics and productivities were calculated accordingly with a 4-parameter logistic function previously characterized for C. malina [4, 7].

The Shapiro-Wilk test was used to validate the normal distribution of the data, and the Brown-Forsythe test was applied to confirm the homogeneity of the variance between treatments. One-way analysis of variance (ANOVA) and post-hoc Tukey’s multiple comparison test were applied to each set of experiments to determine statistical differences among treatments. P values smaller than 0.05 were considered statistically significant. Details of each test are described in the supplementary information Statistic Procedures S1.

Chlamydomonas malina is able to transport glucose into the cells under mixotrophic conditions

We aimed to investigate the ability of C. malina to transport glucose under mixotrophic conditions. Previous studies in our laboratory demonstrated that C. malina cannot grow using glucose as a carbon source under heterotrophic conditions, but it can grow mixotrophically with glucose (Figure S1), similar to the tropical model organism and genus sister Chlamydomonas reinhardtii [30]. Therefore, we wondered how the algal cells were able to uptake glucose under mixotrophic conditions. For this, we performed experiments to determine the transmembrane glucose transport in algal cells growing under heterotrophic and mixotrophic conditions (120 µmol m^− 2 s^− 1) with 3 g/L glucose and [¹⁴C]-glucose tracer. Figure 1 presents the kinetics of glucose transport into the cells.

The microalga was able to transport glucose mixotrophically inside its cells at a rate of 0.015 𝜇mol g⁻¹ min⁻¹ determined during the linear phase of transport in the graph. After 20 minutes, the stable level of [¹⁴C]-glucose tracer inside the cells suggests either saturation of the transporters or that the consumption rate was similar or lower than the transport rate. Clearly, C. malina do not transport glucose under heterotrophic condition. Apparently, under mixotrophic conditions the light promoted glucose uptake into the cells since under dark conditions (heterotrophic) there was no transport (Fig. 1, gray squares). However, the transmembrane glucose transport rate of C. malina is over 300-times slower than observed in efficient heterotrophic microalga, like as Neochloris oleoabundans, grown under strict dark conditions [10]. Besides, C. malina presented higher growth rates and biomass content under phototrophic conditions. This could be due to an imbalance of light energy distribution between the photosystems (PS) under mixotrophic conditions. Liu et al. [32] demonstrated higher ratio of energy distribution in PSI as compared to PSII in Phaeodactylum tricornutum under mixotrophic conditions. In this study, P. tricornutum cells were more susceptible to photodamage under the same light intensity compared to those under phototrophic conditions [32]. Hence, they concluded that the increased electron flux to PSI have a substantial effect on the consumption of protons to possibly protect microalgal cells from photodamage. Light protection, adjusting the gradient of protons on both sides of the thylakoid membrane, could be mediated by the cyclic phosphorylation of the PSII [33]. Besides, it is possible that the presence of glucose reduces the requirement of light energy capture and transmission from PSII [34].

It is tempting to assume that C. malina catabolizes glucose under mixotrophic conditions, as suggested for C. reinhartii [30], where the energy for glucose metabolism is obtained from the light via photosynthesis through the photosystems I and II (PSI and PSII), generating reducing power (NADPH) and ATP via photophosphorylation [35]. However, we only confirm the transmembrane transport, not the catabolism of glucose. Therefore, if confirmed, additional studies are necessary to investigate the pathways needed for the catabolism of this carbon source in C. malina, and the type of glucose transporters. In its genus sister, C. reinhardtii, the sole insertion of a hexose uptake protein (HUP1) in the plasma membrane induced glucose metabolism under heterotrophic conditions [36]. This suggests that cytosolic glucose could be imported and broken down in the chloroplasts. Glucose could enter either using a direct glucose transporter (hexose transporter HXT) or being converted to glucose 6-phosphate and imported through a hexose-phosphate (GPT) translocator catalyzed by a hexokinase in the cytosol. Hexokinase has been found in the chloroplasts of C. reinhardtii but has not confirmed in the cytosol yet, though a dual localization could be possible [37].

PPH1 and PPH3 yielded the maximum reducing sugar concentration with the concomitant presence of furfural, HMF and acetic acid

Treatments PPH1 and PH3 resulted in similar (p > 0.05) and the highest reducing sugar (glucose) concentration, yielding 0.46 and 0.39 g g^− 1, respectively. Both treatments yielded the highest content of furfural, HMF, and acetic acid (Table 1). PPH2 resulted in around 4 g/L of reducing sugar, corresponding to a yield of 0.04 g g^− 1 and almost no furfural and HMF, but similar concentrations of acetic acid as PPH1 and PPH3. These results suggest that an acid-hydrothermal pretreatment can considerably influence the efficiency of the hydrolysis step demonstrating a higher release of reducing sugars compared to no acid addition (control 2). However, toxic by-products are also present. Potato peel waste has a high potential as a carbon source for cultivating microorganisms, but an optimization of the hydrolysis treatment is necessary. For example, Ben Taher et al. [20] obtained 77 g L^− 1 of reducing sugars from potato peel residues applying a hydrothermal pretreatment followed by hydrolysis using a crude enzyme system composed of cellulase, amylase, and hemicellulase. Using a response surface methodology to optimize enzyme hydrolysis revealed that substrate concentration, pH, and temperature significantly affect on the enzymatic conversion of polysaccharides contained in the pretreated potato peel residues [20]. We also tried using a hydrothermal pretreatment followed by enzymatic hydrolysis (control 3), but the sugar conversion was substantially lower than the results obtained by Ben Taher et al. [20], resulting in only 0.875 g L^− 1 of glucose with no furfural, HMF or acetic acid content. However, in contrast to Ben Taher et al. [20], we did not optimize the substrate concentration, pH, or temperature and moreover used a different enzymatic mixture. Then, we adapted the methodology published by Kumar et al. [23] using an acid (3% H₂SO₄) + hydrothermal (steam + pressure) pretreatment followed by an enzyme hydrolysis with a mixture composed of 𝛼-amylase and amyloglucosidase. However, we obtained a two-fold decreased reducing sugar yield than Kumar et al. [23], probably due to the lower acid concentration (1% H₂SO₄) in the pretreatment and the different enzyme mixture used in our enzymatic hydrolysis.

Table 1

Reducing sugar (glucose), furfural, and HMF concentration contained in the PPHs and controls. Product composition and concentration after pretreatment (control 1, 2, and 3) and hydrolysis. All the PPH treatments were subjected to a hydrothermal + sulfuric acid (1%) pretreatment before enzymatic hydrolysis with 𝛼-amylase and amyloglucosidase. Control 1 consisted of potato peel powder in water solution (10% w/v) incubated for 1 hour. Control 2 was treated identically as control 1, but then hydrothermally treated at 121 °C for 20 minutes in an autoclave (without H₂SO₄). Control 3 was treated as control 2 followed by an enzymatic hydrolysis with 𝛼-amylase and amyloglucosidase as previously described in the material and methods section. Data are presented as means ± SD of three independent measurements. Different superscript lowercase letters indicate a significant difference among means of groups (one-way ANOVA with post hoc Tukey HSD test, p < 0.05).
Treatment	Glucose (g/L)	Furfural (g/L)	HMF (g/L)	Acetic acid (g/L)
Control 1	0^a	0 ^a	0 ^a	0 ^a
Control 2	0.213 (± 0.003) ^a	0 ^a	0 ^a	0 ^a
Control 3	0.875 (± 0.005) ^a	0 ^a	0 ^a	3.64 (± 0.42) ^b
PPH1	46.34 (± 5.9) ^b	4.71 (± 0.08) ^b	8.53 (± 0.056) ^b	3.53 (± 0.37) ^b
PPH2	4.19 (± 0.6) ^a	0 ^a	0.001 (± 0.0001) ^a	3.22 (± 0.26) ^b
PPH3	39.79 (± 3.07) ^b	4.02 (± 0.06) ^c	7.01 (± 0.01) ^b	3.18 (± 0.55) ^b

PPH1: Potato peel paste and supernatant collectively.

PPH2: Potato peel paste, H₂SO₄ – free.

PPH3: Only supernatant.

The presence of furfural and HMF in the pretreatment of the potato peel inhibited growth and decreased the biomass productivity in C. malina

The highest biomass concentration (0.4 and 0.38 g L^− 1), cell numbers (1.4 x 10⁷ and 1.5 x 10⁷ cells mL^− 1), and maximum productivities (37 and 39 mg L^− 1 day^− 1) were achieved using glucose and PPH2, respectively as carbon sources under mixotrophic conditions (Figs. 2A, B and C). Under these conditions, the cell number increased by almost two orders of magnitude (from 2.5 x 10⁵ to 1.5 x 10⁷ cells mL^− 1). There was no apparent growth (p > 0.05) when C. malina was cultivated in PPH1 and PPH3 (Figs. 2A and B), indicated by a slight increase in the cell weight (~ 0.07 g L^− 1 in both cases) and cell numbers (from ~ 2.5 x 10⁵ to 6.2 x 10⁵ cells mL^− 1 in both cases), therefore, compared to growth with PPH2, the lowest productivity was obtained in these treatments (~ 5-fold lower; Fig. 2C). In contrast to PPH1/PPH3, the PPH2 paste was washed with water, where most of the furfural and HMF were removed (Table 1). Using the PPH directly after the acid pretreatment and enzymatic hydrolysis for algae cultivation was investigated to explore possible avoidance of extra steps. However, sulfuric acid pretreatment followed by high temperature generates toxic by-products such as furfural, HMF (Table 1), and phenolic compounds that can affect microalgae growth [38, 39]. Therefore, it is crucial to optimize the acid concentration for optimal pretreatment of organic wastes while avoiding the formation of unwanted by-products that would reduce the usefulness of syrups obtained from organic wastes [38]. PPH1 and PPH3 were diluted 1:10, and the pH was adjusted to 7 at the beginning of the cultivation. Nonetheless, the inhibitory compounds (furfural and HMF) remained in the cultivation medium, inhibiting growth. Furan derivatives, 2-furaldehyde (furfural), and HMF are the main degradation compounds generated from pentoses and hexoses, respectively [40]. The majority of the fermenting microorganisms can reduce furans to their corresponding less toxic alcohol. HMF is reduced to 2,5-bis-hydroxymethylfuran and furfural to furfuryl alcohol, and both could also be oxidized to formic acid under anaerobic conditions [41]. If furans are present at high concentration – which is usual when acids are used as hydrolytic agents at high temperatures – they produce an inhibitory effect hampering with glycolytic enzymes and synthesis of macromolecules, causing a prolonged lag phase and reducing growth [42, 43] as we observed in cells growing in PPH1 and PPH3. Though, these effects depend on furan concentration but are highly related to yeast strains [39]. In addition, interaction effects between inhibitory compounds have been observed [44]. Evidences of growth inhibition caused by furfural and derivatives in algae and other microorganisms have been documented. For instance, when Chlorella pyranoidosa FACHB-10 was grown mixotrophically using furfural wastewater diluted two and five times (the furfural concentrations were not shown but it was produced from a furfural production plant in China), the growth of algal cells was inhibited because sulfuric acid, furfural and acetic acid where present in the medium [45]. In another study, Kriechbaum et al. [46] observed a subsequent decrease in Chlorella vulgaris (UTEX 2714) biomass productivities due to an increase in the concentration of furfural from 0.011 to 0.016 g L^− 1, HMF from 0.004 to 0.006 g L^− 1 and acetic acid from 0.46 to 0.64 g L^− 1 in a cultivation medium produced from a wheat straw hydrolysate. The lipid-rich alga Aurantiochytrium limacinum SR21 was found to be highly sensitive to concentrations of furfural up to 0.72 g L^− 1, and concentrations of HMF up to 3.7 g L^− 1 in sugarcane bagasse hydrolysate. Thus, furfural was identified as the primary harmful substance in the sugarcane bagasse hydrolysate, and removing it by activated charcoal treatment enabled efficient lipid production [47]. Furfural and HMF exerted an inhibitory effect on Spirulina maxima growth with a complete inhibition by furfural at 0.67 g L^− 1 and for HMF at 1.13 g L^− 1. Inhibition of photosynthesis was detected and shown by the decrease in oxygen production. Therefore, it was concluded by the authors, that furans could interfere with metabolic processes and cause lysis of Spirulina cells [48]. It has also been reported that furfural at 0.6 g L^− 1 can cause 30% biomass reduction during mixotrophic acetate-based cultivation of C. reinhardtii [49]. This furfural concentration is close to the concentrations contained in the PPH1 and PPH3 cultivation medium (0.47 ± 0.08 and 0.40 ± 0.06 g L^− 1, respectively) for its genus sister alga C. malina in the present study. Concluding, we suggest that the furfural and HMF content in the PPH1 and PPH3 inhibited the growth of C. malina.

For C. malina, the PPH2 alternative was similar to using pure glucose in terms of growth. We attribute this to the removal of furfural derivatives, which favored the growth of this algal strain; despite this, C. malina grows at a rate 1.8-times higher under phototrophic conditions [4]. Interestingly, in PPH2 C. malina was able to grow in the presence of acetic acid (~ 3 g L^− 1), such as other microalgae like Aurantiochytrium limacinum SR21 (up to 12 g L^− 1 of acetic acid) [47] and C. reinhardtii that grows regularly in a medium containing 1 g L^− 1 of acetic acid (TAP medium) [50] and assimilates up to 10 g L^− 1 of acetate, which yielded the highest biomass concentration in a study performed by Moon et al. [30]. However, our study cannot confirm the catabolism of this carbon source in C. malina. For instance, acetate catabolism has been demonstrated in C. reinhardtii [51], where exogenous acetate was assimilated via the TCA cycle. Reactions of the pentose phosphate and glycolysis pathways, which are inactive under phototrophic conditions, presented substantial flux under mixotrophic conditions. Moreover, increased flux through the cyclic electron flow, which enhanced the growth on acetate, was observed [51].

Syrup from the saccharified potato peel promoted lipid accumulation in C. malina

The protein content in cells of C. malina is shown in Fig. 2D. Cells cultivated in glucose attained the highest protein content (36%) followed by PPH2 (28.4%). Cells grown on PPH1 and PPH3 obtained the lowest protein content (12.3 and 10.6%, respectively), but the cells in these media obtained the highest carbohydrate content (Fig. 2D; 33.8 and 34.6%, respectively), followed by cells grown in glucose (28.7%). The lowest carbohydrate content was detected in cells cultivated in PPH2 (13.6%). In the case of the lipid content (Fig. 2D), the PPH treatments promoted lipid accumulation in C. malina, resulting in around 45% of lipid content, and was similar for the three PPH treatments (p < 0.05). Cells of C. malina in pure glucose had a similar macromolecular composition to the cells cultivated under phototrophic conditions [7], which produced the maximal biomass content, even higher than in mixotrophic conditions (~ 12.5-times higher) [7]. Protein was the major component; but carbohydrates and lipid synthesis were not promoted [7]. Under mixotrophic conditions with glucose (this study) and phototrophic conditions [7], the cells produced proteins as building blocks for cellular synthesis without energy reserve metabolites. However, cells cultivated in PPH1 and PPH3 were lower in protein content than in lipid and carbohydrate content, suggesting a stress condition caused by the PPH environment. Normally, when microalgal cells grow under some environmental or nutritional stress, the carbon flux redirects towards the synthesis of energy reserve metabolites, such as carbohydrates, in the short term, and lipids in the long term [7, 10, 52–56]. An intermedium behavior can be observed in PPH2, where the cells still grew similarly to those in glucose and synthesized higher amounts of proteins than PPH1 and PPH3 but lower amounts than in glucose, and more carbon flux was channeled to the synthesis of lipids, probably at the expense of the carbohydrate synthesis. While it is plausible that this difference is due to the acetic acid environment in PPH2 that was not present in the glucose medium, additional studies are necessary to clarify this phenomenon.

PUFA synthesis in the polar fraction was favored in cells cultivated in glucose and PPH2 while PUFA synthesis in the TAG fraction was promoted in the presence of furfural and HMF

Under phototrophic conditions, nutrient-replete, and optimal temperature conditions, the C. malina cells synthesize PUFA in their polar lipid fraction as a natural mechanism to adapt to cold environments [3–7], which helps to maintain the membrane fluidity and flexibility during low or freezing temperatures. However, at temperatures above 4 °C and nitrogen-deplete conditions, microalgal cells accumulate neutral lipids (TAG) as lipid droplets, which can be an indicator of stress [7].

Figure 3 shows the fatty acid types in the polar (A) and neutral (B; TAG) fractions of the total lipid content. Maximum total polar lipid content (252.2 mg g^− 1) was found in cells cultivated in glucose. This polar fraction of fatty acids was composed of similar amounts of MUFA and PUFA but with a lower content of SFA (Fig. 3A). Similar results were found in cells growing in PPH2 (p < 0.05), where most of lipids were MUFA and PUFA in the polar fraction (Fig. 3A), which we can assume were located in the cell membrane. On the contrary, most lipids found in cells cultivated in PPH1 and PPH3 were present in the TAG fraction (total ~ 225 mg g^− 1). Cells in PPH1 had the highest fatty acid content as MUFA (114.8 mg g^− 1). Interestingly, cells in the PPH3 had the highest fatty acid content as PUFA (122.5 mg g^− 1) found in the TAG fraction (Fig. 3B). It is tempting to suggest that, apart from using these PUFA as energy reservoirs, PUFA in the TAG pool of cells growing in PPH3 were used as scavengers of reactive oxygen species (ROS) in this stressful environment. The increase of PUFA, especially omega-3 fatty acids such as C18:3n-3 caused by an alleviation of ROS and lipid peroxidation, was observed in other algal species under stress conditions [57]. However, the reason for PUFA in the TAG pool being high only in PPH3 remains unclear.

The fatty acid profile (Fig. 4) was similar for cells cultivated in glucose and PPH2 where the major content of fatty acids was oleic acid (C18:1n-9), followed by 𝛼-linolenic acid (C18:3n-3) and hexadecatetraenoic acid (C16:4n-3). Cells cultivated in PPH1 had the highest fatty acid content of the type C18:1n-9, but then it was followed by palmitoleic acid (C16:1n-7) and C18:3n-3. In cells growing in PPH3 the major fatty acid type contained in C. malina cells was C18:3n-3, followed by types C18:1n-9 and C16:4n-3.

These results suggest that cells in glucose and PPH2 were probably not stressed, as they primarily synthesized PUFA in the polar fraction (cellular membrane). PUFA are essential to keep the membrane's fluidity at low temperatures in polar microalgae under favorable growth conditions [58, 59], besides their functions as scavengers for ROS. However, cells cultivated in glucose and PPH2 under mixotrophic conditions had a lower growth rate (1.3 and 1.8-times lower, respectively) and biomass content (12.5-times) than cells cultivated under phototrophic conditions [7], suggesting that the mixotrophic mode was not favorable for this microalga but not to an extent to produce TAG as indicator of stress. For example, cells cultivated in PPH1 and PPH3 had similar growth, macromolecular composition, and lipid types and profile than cells grown at a stressful temperature condition (15°C), as previously reported [7]. Wherein that’s study, C. malina cells did not grow significantly and mainly accumulated TAGs composed by the monounsaturated fatty acids C16:1n-7 and C18:1n-9, as a consequence of stress. Apart from nutrient stress, the accumulation of TAG is also observed in other stressing conditions, such as temperature, pH, salinity, light intensity, and temperature [53, 60–66].

Interestingly, the fatty acid C16:4n-3, always present in phototrophically cultivated cells, was found also in all PPH treatments studied in the present work (at a higher proportion in PPH3), highlighting the importance of this fatty acid also under mixotrophic conditions in a polar microalga such as C. malina. This polyunsaturated fatty acid short-chain length C16:4n-3 is nearly exclusively present in the monogalactosyldiacylglycerol (MGDG) molecule, which can be found in a high proportion in the lipid fraction of the Chlamydomonas chloroplast membrane, contributing to the transition from liquid-crystalline to gel phase [54, 67, 68]. The specific roles of this molecular species of MDGD in the photosynthetic membrane are still unclear, therefore it is still subject to investigations [68].

Feasibility of using potato peel hydrolysates as a carbon source for microalgae cultivation

The polar microalga C. malina has previously been demonstrated to be highly productive under phototrophic conditions at low temperatures, obtaining high quantities of PUFA and biomass productivities comparable with other mesophilic high-productive algae [7]. However, C. malina was not able to grow under strict heterotrophic conditions (Figure S1) and may not be suitable for mixotrophic cultivation. The biomass content under mixotrophic conditions, either using glucose or PPH2, was 12.5-times lower than the ones obtained under phototrophic conditions. This was unexpected because C. malina is a polar microalga that was isolated from the Arctic, where the environmental conditions are extreme, having for example 24 hours of light during summer but also 24 hours of darkness during winter. Therefore, it is likely possible that C. malina cells are able to switch their metabolism to mixotrophic mode to use organic carbon sources present in the ocean (sugars and starches) [69]. This should be particularly important as a mechanism to thrive when the light is limited, e.g. during early/late winter in the polar areas [69]. Indeed, through this study, we can confirm that C. malina can at least thrive under mixotrophic conditions using a carbon source, but with lower biomass productivities as compared with other microalgae that are known for their high performance under mixotrophic conditions – i.e., Chlorella, Galdieria, Neochloris – with productivities up to 100-times higher than the productivities obtained with C. malina (Table 2).

On the other hand, potato peel has a high potential as a carbon source for microbial cultivation. However, conversion to reducing sugars depends on the pretreatment (if applied) and the enzyme mixture, for example, Arapoglou et al. [21] used a combination of three commercial enzymes for the hydrolysis of potato peel waste, obtained 18.5 g L^− 1 (92% conversion) of reducing sugars. Khawla et al. [22] obtained 69 g L^− 1 (conversion not reported) of reducing sugars from the hydrolysis of potato peel residues when they used a combined mixture of commercial amyloglucosidase and an onsite-produced amylase. Ben Taher et al. [20] obtained 77 g L^− 1 (84% conversion) when they applied a response surface methodology to optimize the enzyme hydrolysis, substrate concentration, pH, and temperature in a hydrothermal pretreatment followed by enzymatic hydrolysis using crude enzyme system composed of cellulase, amylase, and hemicellulase. The highest concentration of reducing sugar released (141 g L^− 1; 98% conversion) was reported by Soni et al. [23] in experiments combining an acid (3% H₂SO₄), hydrothermal (steam + pressure), and enzyme cocktail (cellulases, hemicellulases, pectinases, and amylases). In the present study, we obtained 46 g L^− 1 (46% conversion) of reducing sugars when an acid-hydrothermal pretreatment followed by enzymatic hydrolysis using commercial amylase and amyloglucosidase was applied. Therefore, yield improvements can be achieved if optimization of the hydrolysis process is first conducted. In addition, we suggest using this resource (previously optimized) with other mixo/heterotrophic microalgal cells such as the ones in Table 2, and preferably avoid the presence of inhibitory compounds such as furfural and HMF in the cultivation medium.

Although C. malina’s mixotrophic growth was limited, we want to highlight the versatility and flexibility of its metabolism and point out the potential of this and other polar microalgae as models of study and for biotechnology applications.

Table 2

Comparison of biomass productivities of *C. malina* cultivated under mixotrophic conditions with other microalgae strains at similar conditions.
Microalgal strain	Productivity (mg L^− 1 Day^− 1)	Carbon source	Reference
Chlamydomonas malina RCC2488	37.03	Glucose	This study
Chlamydomonas malina RCC2488	39.3	PPH2	This study
Chlorella pyrenoidosa C-212	3600	Glucose	[70]
Phaeodactylum tricornutum UTEX640	1476	Glycerol	[71]
Galdieria sulphuraria 074G	721	Fructose	[71]
Chlorella minutissima	296	Glucose	[72]
Chlorella vulgaris UTEX2714	260	Crude glycerol	[73]
Chlorella protothecoides UTEX256	500	Dairy waste	[74]
Neochloris oleoabundans UTEX1185	230	Cassava wastewater	[75]
Chlorella protothecoides UTEX25	667	Rice straw hydrolysate	[76]
Nannochloropsis sp. BR21	63.2	Sugarcane bagasse hydrolysate	[77]
Scenedesmus dimorphus NT8c	119.5	Sugarcane bagasse hydrolysate	[78]
Chlorella vulgaris	137.4	Sugarcane molasses	[79]

Polar C. malina is a microalga that is able to transport glucose through the cell membrane under mixotrophic conditions. Accordingly, C. malina can grow under mixotrophic conditions using glucose as carbon source. However, its growth was not higher than under phototrophic conditions. Cells of C. malina using glucose as a carbon source accumulated protein, and its lipid types were mainly polar with a high PUFA content as in optimal phototrophic conditions.

An acid-hydrothermal pretreatment of potato peel waste can considerably influence the efficiency of the hydrolysis, demonstrating a higher release of reducing sugars as compared with no acid addition. However, toxic by-products in a high concentration such as furfural, HMF and acetic acid are also present.

These toxic by-products present in the PPH1 and PPH3 treatments inhibited the growth, decreased the biomass productivity of C. malina, and promoted lipid accumulation, especially TAG, which was a stress indicator. A washing step with water was sufficient to remove the sulfuric acid in the potato peel paste (PPH2) but resulted in 10-times lower reducing sugar content. Nonetheless, this extra step resulted in similar growth, biomass productivities, and lipid content, types and profile than the cultivation with glucose.

Potato peel waste has a high potential as carbon source for the cultivation of microorganisms, other than C. malina, but an optimization of the hydrolysis treatment is necessary to yield a higher percentage of reducing sugar conversion without toxic by-products.

The authors declare that they have no known competing financial interests or personal relationships.

ACKNOWLEDGMENTS

We acknowledge technical support by Mario A. Caro Bermudez, Luz María Martínez Mejia, Georgina Hernández, Shirley Ainsworth, Roberto P. Rodríguez Bahena, David S. Castañeda Carreon, and Juan Manuel Hurtado Ramirez.

CONTRIBUTIONS

DM-S designed the research that resulted in this manuscript. SRA and DM-S collected the data of the cultivations and performed the biochemical composition analysis. SRA and DM-S conducted the hydrolysis analysis. DM-S designated and conducted the lipid analysis. LB and DMS performed glucose transport experiments. SRA, LB, LS-C, AM and DM-S contributed to the manuscript drafting, discussion, and critical revision of the article for important intellectual content. All authors contributed to the article and approved the submitted version.

FUNDING

This work was funded by The National Autonomous University of Mexico through its program DGAPA-PAPIIT (Grant number [IA202124]) and the IBT UNAM with the project 11053 P-11053 for DMS and by the Arctic University of Norway through the project BFE370010153 designated to DMS.

DATA AVAILABILITY

The datasets generated during and/or analyzed during the current study are available from the corresponding author on reasonable request.

Morales-Sánchez D, Martinez-Rodriguez OA, Martinez A (2017) Heterotrophic cultivation of microalgae: production of metabolites of commercial interest. Journal of Chemical Technology and Biotechnology 92:925–936. https://doi.org/10.1002/jctb.5115
Chen GQ, Chen F (2006) Growing phototrophic cells without light. Biotechnol Lett 28:607–616. https://doi.org/10.1007/s10529-006-0025-4
Hulatt CJ, Berecz O, Egeland ES, et al (2017) Polar snow algae as a valuable source of lipids? Bioresour Technol 235:338–347. https://doi.org/10.1016/j.biortech.2017.03.130
Morales-Sánchez D, Schulze PSC, Kiron V, Wijffels RH (2020) Production of carbohydrates, lipids and polyunsaturated fatty acids (PUFA) by the polar marine microalga Chlamydomonas malina RCC2488. Algal Res 50:102016. https://doi.org/10.1016/j.algal.2020.102016
Schulze PSC, Hulatt CJ, Morales-Sánchez D, et al (2019) Fatty acids and proteins from marine cold adapted microalgae for biotechnology. Algal Res 42:101604. https://doi.org/10.1016/j.algal.2019.101604
Komárek J, Nedbalová L (2007) Green Cryosestic Algae. Springer, Dordrecht, pp 321–342
Morales-Sánchez D, Schulze PSC, Kiron V, Wijffels RH (2020) Temperature-Dependent Lipid Accumulation in the Polar Marine Microalga Chlamydomonas malina RCC2488. Front Plant Sci 11:2080. https://doi.org/10.3389/fpls.2020.619064
Adarme-Vega T, Lim DKY, Timmins M, et al (2012) Microalgal biofactories: a promising approach towards sustainable omega-3 fatty acid production. Microb Cell Fact 11:96. https://doi.org/10.1186/1475-2859-11-96
Meng TK, Kassim MA, Cheirsilp B (2020) Mixotrophic Cultivation: Biomass and Biochemical Biosynthesis for Biofuel Production. In: Microalgae Cultivation for Biofuels Production
Morales-Sánchez D, Tinoco-Valencia R, Kyndt J, Martinez A (2013) Heterotrophic growth of Neochloris oleoabundans using glucose as a carbon source. Biotechnol Biofuels 6:100. https://doi.org/10.1186/1754-6834-6-100
Gouveia L, Oliveira AC (2009) Microalgae as a raw material for biofuels production. J Ind Microbiol Biotechnol 36:269–274. https://doi.org/10.1007/s10295-008-0495-6
Hakobyan L, Gabrielyan L, Trchounian A (2012) Yeast extract as an effective nitrogen source stimulating cell growth and enhancing hydrogen photoproduction by Rhodobacter sphaeroides strains from mineral springs. Int J Hydrogen Energy 37:6519–6526. https://doi.org/10.1016/J.IJHYDENE.2012.01.077
Mahmood AU, Greenman J, Scragg AH (1998) Orange and potato peel extracts: Analysis and use as Bacillus substrates for the production of extracellular enzymes in continuous culture. Enzyme Microb Technol. https://doi.org/10.1016/S0141-0229(97)00150-6
Ben Taher I, Fickers P, Chniti S, Hassouna M (2017) Optimization of enzymatic hydrolysis and fermentation conditions for improved bioethanol production from potato peel residues. Biotechnol Prog 33:397–406. https://doi.org/10.1002/BTPR.2427
Arapoglou D, Varzakas T, Vlyssides A, Israilides C (2010) Ethanol production from potato peel waste (PPW). Waste Management 30:1898–1902. https://doi.org/10.1016/J.WASMAN.2010.04.017
Khawla BJ, Sameh M, Imen G, et al (2014) Potato peel as feedstock for bioethanol production: A comparison of acidic and enzymatic hydrolysis. Ind Crops Prod 52:144–149. https://doi.org/10.1016/J.INDCROP.2013.10.025
Soni SK, Sharma B, Sharma A, et al (2023) Exploring the Potential of Potato Peels for Bioethanol Production through Various Pretreatment Strategies and an In-House-Produced Multi-Enzyme System. Sustainability (Switzerland) 15:. https://doi.org/10.3390/su15119137
Mahmood AU, Greenman J, Scragg AH (1998) Orange and potato peel extracts: Analysis and use as Bacillus substrates for the production of extracellular enzymes in continuous culture. Enzyme Microb Technol. https://doi.org/10.1016/S0141-0229(97)00150-6
Malakar B, Das D, Mohanty K (2020) Optimization of glucose yield from potato and sweet lime peel waste through different pre-treatment techniques along with enzyme assisted hydrolysis towards liquid biofuel. Renew Energy 145:2723–2732. https://doi.org/10.1016/j.renene.2019.08.037
Ben Taher I, Fickers P, Chniti S, Hassouna M (2017) Optimization of enzymatic hydrolysis and fermentation conditions for improved bioethanol production from potato peel residues. Biotechnol Prog 33:397–406. https://doi.org/10.1002/BTPR.2427
Arapoglou D, Varzakas T, Vlyssides A, Israilides C (2010) Ethanol production from potato peel waste (PPW). Waste Management 30:1898–1902. https://doi.org/10.1016/J.WASMAN.2010.04.017
Khawla BJ, Sameh M, Imen G, et al (2014) Potato peel as feedstock for bioethanol production: A comparison of acidic and enzymatic hydrolysis. Ind Crops Prod 52:144–149. https://doi.org/10.1016/J.INDCROP.2013.10.025
Soni SK, Sharma B, Sharma A, et al (2023) Exploring the Potential of Potato Peels for Bioethanol Production through Various Pretreatment Strategies and an In-House-Produced Multi-Enzyme System. Sustainability (Switzerland) 15:. https://doi.org/10.3390/su15119137
Balzano S, Gourvil P, Siano R, et al (2012) Diversity of cultured photosynthetic flagellates in the northeast Pacific and Arctic Oceans in summer. Biogeosciences 9:4553–4571. https://doi.org/10.5194/bg-9-4553-2012
Utrilla J, Gosset G, Martinez A (2009) ATP limitation in a pyruvate formate lyase mutant of escherichia coli MG1655 increases glycolytic flux to d-lactate. J Ind Microbiol Biotechnol 36:1057–1062. https://doi.org/10.1007/s10295-009-0589-9
Thompson AR (1950) A colorimetric method for the determination of esters. Aust J Chem 3:128–135. https://doi.org/10.1071/CH9500128
Utrilla J, Licona-Cassani C, Marcellin E, et al (2012) Engineering and adaptive evolution of Escherichia coli for d-lactate fermentation reveals GatC as a xylose transporter. Metab Eng 14:. https://doi.org/10.1016/j.ymben.2012.07.007
Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193:265–75
Breuer G, Evers WAC, de Vree JH, et al (2013) Analysis of fatty acid content and composition in microalgae. J Vis Exp. https://doi.org/10.3791/50628
Moon M, Kim CW, Park WK, et al (2013) Mixotrophic growth with acetate or volatile fatty acids maximizes growth and lipid production in Chlamydomonas reinhardtii. Algal Res 2:352–357. https://doi.org/10.1016/J.ALGAL.2013.09.003
Liu X, Duan S, Li A, et al (2009) Effects of organic carbon sources on growth, photosynthesis, and respiration of Phaeodactylum tricornutum. J Appl Phycol 21:239–246. https://doi.org/10.1007/s10811-008-9355-z
Ruban A, Lavaud J, Rousseau B, et al (2004) The super-excess energy dissipation in diatom algae: Comparative analysis with higher plants. Photosynth Res 82:. https://doi.org/10.1007/s11120-004-1456-1
Yang C, Hua Q, Shimizu K (2000) Energetics and carbon metabolism during growth of microalgal cells under photoautotrophic, mixotrophic and cyclic light-autotrophic/dark-heterotrophic conditions. Biochem Eng J 6:87–102. https://doi.org/10.1016/S1369-703X(00)00080-2
Castillo T, Ramos D, García-Beltrán T, et al (2021) Mixotrophic cultivation of microalgae: An alternative to produce high-value metabolites. Biochem Eng J 176
Doebbe A, Rupprecht J, Beckmann J, et al (2007) Functional integration of the HUP1 hexose symporter gene into the genome of C. reinhardtii: Impacts on biological H2production. J Biotechnol 131:27–33. https://doi.org/10.1016/j.jbiotec.2007.05.017
Johnson X, Alric J (2013) Central carbon metabolism and electron transport in chlamydomonas reinhardtii: Metabolic constraints for carbon partitioning between oil and starch. Eukaryot Cell 12:. https://doi.org/10.1128/EC.00318-12
Pradhan N, Kumar S, Selvasembian R, et al (2023) Emerging trends in the pretreatment of microalgal biomass and recovery of value-added products: A review. Bioresour Technol 369:128395. https://doi.org/10.1016/J.BIORTECH.2022.128395
Tomás-Pejó E, Alvira P, Ballesteros M, Negro MJ (2011) Pretreatment Technologies for Lignocellulose-to-Bioethanol Conversion. Biofuels 149–176. https://doi.org/10.1016/B978-0-12-385099-7.00007-3
Wyman CE (2007) What is (and is not) vital to advancing cellulosic ethanol. Trends Biotechnol 25:153–157. https://doi.org/10.1016/J.TIBTECH.2007.02.009
Taherzadeh MJ, Gustafsson L, Niklasson C, Lidén G (1999) Conversion of furfural in aerobic and anaerobic batch fermentation of glucose by Saccharomyces cerevisiae. J Biosci Bioeng 87:169–174. https://doi.org/10.1016/S1389-1723(99)89007-0
Almeida JRM, Modig T, Petersson A, et al (2007) Increased tolerance and conversion of inhibitors in lignocellulosic hydrolysates by Saccharomyces cerevisiae. Journal of Chemical Technology and Biotechnology 82
Klinke HB, Thomsen AB, Ahring BK (2004) Inhibition of ethanol-producing yeast and bacteria by degradation products produced during pre-treatment of biomass. Appl Microbiol Biotechnol 66:10–26. https://doi.org/10.1007/S00253-004-1642-2
Larsson S, Reimann A, Nilvebrant NO, Jönsson LJ (1999) Comparison of different methods for the detoxification of lignocellulose hydrolyzates of spruce. Applied Biochemistry and Biotechnology - Part A Enzyme Engineering and Biotechnology 77–79:91–103. https://doi.org/10.1385/ABAB:77:1-3:91/METRICS
Cheng P, Huang J, Song X, et al (2022) Heterotrophic and mixotrophic cultivation of microalgae to simultaneously achieve furfural wastewater treatment and lipid production. Bioresour Technol 349:126888. https://doi.org/10.1016/J.BIORTECH.2022.126888
Kriechbaum R, Loaiza SS, Friedl A, et al (2023) Utilizing straw-derived hemicellulosic hydrolysates by Chlorella vulgaris: Contributing to a biorefinery approach. J Appl Phycol 1:1–16. https://doi.org/10.1007/S10811-023-03082-0/FIGURES/5
Watanabe K, Nishijima M, Mayuzumi S, Aki T (2022) Utilization of Sugarcane Bagasse as a Substrate for Lipid Production by Aurantiochytrium sp. J Oleo Sci 71:1493–1500. https://doi.org/10.5650/jos.ess22206
Journal BP, Yu S, Forsberg Å, et al (2007) Furfural and Hydroxymethylfurfural inhibition of growth and photosynthesis in Spirulina. Phycological Journal 25:141–148. https://doi.org/10.1080/00071619000650131
Liang Y, Zhao X, Chi Z, et al (2013) Utilization of acetic acid-rich pyrolytic bio-oil by microalga Chlamydomonas reinhardtii: Reducing bio-oil toxicity and enhancing algal toxicity tolerance. Bioresour Technol 133:500–506. https://doi.org/10.1016/J.BIORTECH.2013.01.134
Harris EH (1989) Culture and Storage Methods. The Chlamydomonas Sourcebook 25–63. https://doi.org/10.1016/B978-0-12-326880-8.50007-9
Chapman SP, Paget CM, Johnson GN, Schwartz JM (2015) Flux balance analysis reveals acetate metabolism modulates cyclic electron flow and alternative glycolytic pathways in Chlamydomonas reinhardtii. Front Plant Sci 6:. https://doi.org/10.3389/fpls.2015.00474
Pruvost J, Van Vooren G, Cogne G, Legrand J (2009) Investigation of biomass and lipids production with Neochloris oleoabundans in photobioreactor. Bioresour Technol 100:5988–5995. https://doi.org/10.1016/j.biortech.2009.06.004
Msanne J, Xu D, Konda AR, et al (2012) Metabolic and gene expression changes triggered by nitrogen deprivation in the photoautotrophically grown microalgae Chlamydomonas reinhardtii and Coccomyxa sp. C-169. Phytochemistry 75:50–59. https://doi.org/10.1016/j.phytochem.2011.12.007
Li-Beisson Y, Shorrosh B, Beisson F, et al (2013) Acyl-Lipid Metabolism. In: The Arabidopsis Book. BioOne, p e0161
Morales-Sánchez D, Tinoco-Valencia R, Caro-Bermúdez MA, Martinez A (2014) Culturing Neochloris oleoabundans microalga in a nitrogen-limited, heterotrophic fed-batch system to enhance lipid and carbohydrate accumulation. Algal Res 5:61–69. https://doi.org/10.1016/j.algal.2014.05.006
Siaut M, Cuiné S, Cagnon C, et al (2011) Oil accumulation in the model green alga Chlamydomonas reinhardtii: Characterization, variability between common laboratory strains and relationship with starch reserves. BMC Biotechnol 11:7. https://doi.org/10.1186/1472-6750-11-7
Udayan A, Pandey AK, Sirohi R, et al (2023) Production of microalgae with high lipid content and their potential as sources of nutraceuticals. Phytochemistry Reviews 22
D’Amico S, Collins T, Marx J-C, et al (2006) Psychrophilic microorganisms: challenges for life. EMBO Rep 7:385–389. https://doi.org/10.1038/sj.embor.7400662
Morgan-Kiss RM, Ivanov AG, Modla S, et al (2008) Identity and physiology of a new psychrophilic eukaryotic green alga, Chlorella sp., strain BI, isolated from a transitory pond near Bratina Island, Antarctica. Extremophiles 12:701–711. https://doi.org/10.1007/s00792-008-0176-4
Converti A, Casazza AA, Ortiz EY, et al (2009) Effect of temperature and nitrogen concentration on the growth and lipid content of Nannochloropsis oculata and Chlorella vulgaris for biodiesel production. Chemical Engineering and Processing: Process Intensification 48:1146–1151. https://doi.org/10.1016/j.cep.2009.03.006
Draaisma RB, Wijffels RH, Breuer G, et al (2013) Effect of light intensity, pH, and temperature on triacylglycerol (TAG) accumulation induced by nitrogen starvation in Scenedesmus obliquus. Bioresour Technol 143:1–9. https://doi.org/10.1016/j.biortech.2013.05.105
Renaud SM, Thinh L Van, Lambrinidis G, Parry DL (2002) Effect of temperature on growth, chemical composition and fatty acid composition of tropical Australian microalgae grown in batch cultures. Aquaculture 211:195–214. https://doi.org/10.1016/S0044-8486(01)00875-4
Xin L, Hong-ying H, Ke G, Ying-xue S (2010) Effects of different nitrogen and phosphorus concentrations on the growth, nutrient uptake, and lipid accumulation of a freshwater microalga Scenedesmus sp. Bioresour Technol 101:5494–5500. https://doi.org/10.1016/j.biortech.2010.02.016
Jiang Y, Laverty KS, Brown J, et al (2014) Effects of fluctuating temperature and silicate supply on the growth, biochemical composition and lipid accumulation of Nitzschia sp. Bioresour Technol 154:336–344. https://doi.org/10.1016/j.biortech.2013.12.068
Pal D, Khozin-Goldberg I, Cohen Z, Boussiba S (2011) The effect of light, salinity, and nitrogen availability on lipid production by Nannochloropsis sp. Appl Microbiol Biotechnol 90:1429–1441. https://doi.org/10.1007/s00253-011-3170-1
Liu J, Yuan C, Hu G, Li F (2012) Effects of light intensity on the growth and lipid accumulation of microalga Scenedesmus sp. 11-1 under nitrogen limitation. Appl Biochem Biotechnol 166:2127–2137. https://doi.org/10.1007/s12010-012-9639-2
Dolhi JM, Maxwell DP, Morgan-Kiss RM (2013) Review: The Antarctic Chlamydomonas raudensis: An emerging model for cold adaptation of photosynthesis. Extremophiles 17:711–722
Liu B, Benning C (2013) Lipid metabolism in microalgae distinguishes itself. Curr Opin Biotechnol 24:300–309. https://doi.org/10.1016/j.copbio.2012.08.008
Lyon B, Mock T (2014) Polar Microalgae: New Approaches towards Understanding Adaptations to an Extreme and Changing Environment. Biology (Basel) 3:56–80. https://doi.org/10.3390/biology3010056
Yang C, Hua Q, Shimizu K (2000) Energetics and carbon metabolism during growth of microalgal cells under photoautotrophic, mixotrophic and cyclic light-autotrophic/dark-heterotrophic conditions. Biochem Eng J 6:87–102. https://doi.org/10.1016/S1369-703X(00)00080-2
Ceron Garcia MC, Fernandez Sevilla JM, Acien Fernandez FG, et al (2000) Mixotrophic growth of Phaeodactylum tricornutum on glycerol : growth rate and fatty acid profile. Journal of Applied Phycology 12: 12:239–248. https://doi.org/10.1002/bit.22257
Sloth JK, Jensen HC, Pleissner D, Eriksen NT (2017) Growth and phycocyanin synthesis in the heterotrophic microalga Galdieria sulphuraria on substrates made of food waste from restaurants and bakeries. Bioresour Technol. https://doi.org/10.1016/j.biortech.2017.04.043
Dubey KK, Kumar S, Dixit D, et al (2015) Implication of Industrial Waste for Biomass and Lipid Production in Chlorella minutissima Under Autotrophic, Heterotrophic, and Mixotrophic Grown Conditions. Appl Biochem Biotechnol 176:1581–1595. https://doi.org/10.1007/S12010-015-1663-6/FIGURES/6
Ma X, Zheng H, Addy M, et al (2016) Cultivation of Chlorella vulgaris in wastewater with waste glycerol: Strategies for improving nutrients removal and enhancing lipid production. Bioresour Technol 207:252–261. https://doi.org/10.1016/J.BIORTECH.2016.02.013
Patel AK, Joun J, Sim SJ (2020) A sustainable mixotrophic microalgae cultivation from dairy wastes for carbon credit, bioremediation and lucrative biofuels. Bioresour Technol 313:123681. https://doi.org/10.1016/J.BIORTECH.2020.123681
Ghiggi Sorgatto V, Ricardo Soccol C, Tatiana Molina-Aulestia D, et al (2021) Mixotrophic Cultivation of Microalgae in Cassava Processing Wastewater for Simultaneous Treatment and Production of Lipid-Rich Biomass. Fuels 2021, Vol 2, Pages 521-532 2:521–532. https://doi.org/10.3390/FUELS2040030
Joe MH, Kim JY, Lim S, et al (2015) Microalgal lipid production using the hydrolysates of rice straw pretreated with gamma irradiation and alkali solution. Biotechnol Biofuels 8:1–9. https://doi.org/10.1186/S13068-015-0308-X/TABLES/3
Manzoor M, Jabeen F, Ahmad Q ul A, et al (2021) Sugarcane Bagasse Hydrolysate as Organic Carbon Substrate for Mixotrophic Cultivation of Nannochloropsis sp. BR2. Waste Biomass Valorization 12:2321–2331. https://doi.org/10.1007/S12649-020-01185-0/FIGURES/6
Manzoor M, Ahmad Q ul A, Aslam A, et al (2020) Mixotrophic cultivation of Scenedesmus dimorphus in sugarcane bagasse hydrolysate. Environ Prog Sustain Energy 39:e13334. https://doi.org/10.1002/EP.13334
Laraib N, Manzoor M, Javid A, et al (2021) Mixotrophic cultivation of Chlorella vulgaris in sugarcane molasses preceding nitrogen starvation: Biomass productivity, lipid content, and fatty acid analyses. Environ Prog Sustain Energy 40:e13625. https://doi.org/10.1002/EP.13625

Download PDF

Journal Publication

published 22 Feb, 2025

Read the published version in BioEnergy Research →

Editorial decision: Reconsider pending major revisions
05 Dec, 2024
Reviewers agreed at journal
27 Oct, 2024
Reviewers invited by journal
27 Oct, 2024
Editor assigned by journal
23 Oct, 2024
First submitted to journal
23 Oct, 2024

You are reading this latest preprint version

Mixotrophic growth of the polar marine microalga Chlamydomonas sp. RCC2488 (malina) using potato peel hydrolysates as carbon source

Status:

Journal Publication

Version 1

Abstract

Figures

INTRODUCTION

MATERIALS AND METHODS

RESULTS AND DISCUSSION

Feasibility of using potato peel hydrolysates as a carbon source for microalgae cultivation

CONCLUSIONS

Declarations

References

Status:

Journal Publication

Version 1