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Assessment of Nutrients Recovery Capacity and Biomass Growth of Four Microalgae Species in Anaerobic Digestion Effluent

Laboratory of Technologies for Environmental Protection and Utilization of Food By-Products, Department of Food Science and Technology, International Hellenic University (IHU), Sindos, 57400 Thessaloniki, Greece
Laboratory of Biochemical and Biotechnological Processes (LB2P), Department of Chemical Engineering, Aritotle University of Thessaloniki (AUTH), 54124 Thessaloniki, Greece
Cyanolab, Department of Botany, School of Biology, Aristotle University of Thessaloniki (AUTH), 54124 Thessaloniki, Greece
Author to whom correspondence should be addressed.
Water 2022, 14(2), 221;
Received: 29 November 2021 / Revised: 26 December 2021 / Accepted: 10 January 2022 / Published: 12 January 2022


Four microalgae species were evaluated for their bioremediation capacity of anaerobic digestion effluent (ADE) rich in ammonium nitrogen, derived from a biogas plant. Chlorella vulgaris, Chlorella sorokiniana, Desmodesmus communis and Stichococcus sp. were examined for their nutrient assimilation efficiency, biomass production and composition through their cultivation in 3.7% v/v ADE; their performance was compared with standard cultivation media which consisted in different nitrogen sources, i.e., BG-11NO3 and BG-11ΝH4 where N-NO3 was replaced by N-NH4. The results justified ammonium as the most preferable source of nitrogen for microalgae growth. Although Stichococcus sp. outperformed the other 3 species in N-NH4 removal efficiency both in BG-11NH4 and in 3.7% ADE (reaching up to 90.79% and 69.69% respectively), it exhibited a moderate biomass production when it was cultivated in diluted ADE corresponding to 0.59 g/L, compared to 0.89 g/L recorded by C. vulgaris and 0.7 g/L by C. sorokiniana and D. communis. Phosphorus contained in the effluent and in the control media was successfully consumed by all of the species, although its removal rate was found to be affected by the type of nitrogen source used and the particular microalgae species. The use of ADE as cultivation medium resulted in a significant increase in carbohydrates content in all investigated species.

1. Introduction

Microalgae have been recognized as a promising alternative source for biomass production that can be allocated in a wide range of products for different consumer markets, such as animal/human feed, aquaculture, biofertilizers/biostimulants, biofuels and nutraceuticals. Their potential to photosynthetically assimilate large amounts of carbon and nutrients and accumulate intracellularly commercially exploitable products such as lipids, proteins, carbohydrates, and pigments makes them emerging candidates for a variety of applications [1]. Despite the attractiveness of microalgae exploitation for clean energy production, large-scale plants cannot yet be considered as a commercially available option, as there are certain constraints hindering their economic competitiveness [2]. Among them the necessity for large amounts of inorganic nutrients and CO2 supply, crucial for their efficient production, seems to be major bottleneck for the economic viability of the industrial development of microalgae-based technologies [3]. Many authors have highlighted the necessity of integrating microalgae culture systems in industrial infrastructures, exploiting their wastewater streams rich in inorganic nutrients and flue gases rich in CO2 and heat, in an efficient manner as cheap input to microalgae system. This approach will contribute to the reduction in the environmental footprint of the industrial process, generating a distinct economic benefit for the entire plant [4,5,6]. The utilization of wastewater streams as a nutrient source for microalgae culture not only promotes the sustainability of their production, but also addresses the urgent need for wastewater reuse and nutrients recovery, as the discharge of inadequately treated effluents, loaded with nutrients, mainly nitrogen and phosphorous, represents a critical environmental concern [7].
Use of microalgae for wastewater bioremediation has been tested in many different types of effluents, such as municipal, industrial, food and livestock. Although satisfactory results have been recorded, certain constraints have been identified related to installation and operation costs, increased costs for post utilization of produced biomass, or competition of microalgae for areas used for food crops cultivation [8,9]. Among a broad spectrum of available effluent sources, a stream of particular interest is the liquid fraction obtained by the anaerobic digestion process, derived from biogas plants. The global industry of biogas production has significantly developed over the past few years, with the European Biogas Association reporting that the current installations in Europe are more than 18,000 [10]. Anaerobic digestion (AD) is a feasible and straightforward way to harvest bioenergy from the co-digestion of residues from agricultural and agri-food activities, such as animal manures (cattle slurry, chicken manure), food waste (cheese whey, olive mill waste) and horticultural wastes (maize silage) [11]. However, the increasing number of biogas plants unavoidably results in a severe increase in the amount of the anaerobic digestion effluent (ADE), which constitutes the main by-product of the procedure [12]. The liquid fraction of this stream, representing 80–90% of its total mass, has a high moisture content (81–98%) and consequently a high volume compared to its fertilizing value, making its management, in terms of transportation and storage, difficult [13,14]. Furthermore, the liquid ADE contains high levels of nitrogen (mostly in the form of NH4-N) and phosphorus, and thus its discharge to the receiving bodies is an inappropriate management practice, as it leads to severe eutrophication of aquatic ecosystems [15]. On the other hand, the use of the digestate in agricultural lands, which represents the current management practice, is restricted due to nitrogen surplus and low annual loading thresholds: EU Regulation 2092/91 strictly limits the input of farmyard manure to a maximum level of 170 kg N/ha/year [16]. Advanced biological and physicochemical processes have been tested for nutrients removal, such as chemical precipitation, ammonia stripping, aerobic biological processes, and denitrification. However, these processes, besides presenting high operating costs and energy demand, result in by-products, such as sludge, that further complicate their application [17]. Nevertheless, these methods are focused on nutrients removal rather than on their recovery from the effluents. Thus, the integration of ADE treatment and microalgae biomass production could be a key factor in aiding the development of the AD sector, making microalgae-based technology economically feasible and environmentally beneficial. An additional benefit can be derived by further variolization of produced biomass, through the utilization of microalgae lipids to obtain bio-oil [18], or their co-digestion in the AD reactor aiming to the enhancement of methane content in biogas up to about 66% [19].
Different types of ADEs have been already explored as potential feedstock for microalgae cultivation [14,20,21]. Although the majority of the published works in the literature contains encouraging results, the data are not comparable, since different microalgae species, substrates, operational conditions and technologies are used in these studies. The utilization of algae for the treatment of ADEs has been mainly studied in effluents collected from anaerobic digesters treating the excess sludge of municipal wastewater treatment plants. However, the treatment of ADE from biogas plants receiving wastes from animal husbandry units represents a challenging concept due to the high concentration of ammonium nitrogen, exceeding in some cases 3 g/L, and the variability of their chemical composition [22,23]. Nevertheless, the properties of influents to anaerobic digesters greatly affect the composition of digestate effluents and especially their nitrogen content. AD plant operators are looking for low-cost biomass sources, in their efforts to reduce anaerobic process operation costs. Especially, chicken manure represents a low-cost substrate for the AD process, which, however, has a high content of nitrogen mainly in the form of ammonium. Furthermore, great variation in the composition of ADEs is observed due to the different batches being employed in the AD process, according to the availability and the seasonality of the feedstocks, or the different animal diet elaborated in each live-stock plant [22,23]. On the other hand, derived effluents contain compounds, such as urea, organic acids, and phenols, originating from the feedstock substates at concentrations that potentially inhibit microalgae growth [22], or endogenous organisms (protozoa, fungi, bacteria, and rotifers) that might affect the biomass growth rate [24]. As such, the successful implementation of microalgae cultures for bioremediation of liquid effluents derived from biogas plants primarily requires the identification of robust and dominant microalgae species, capable of proliferation in this harsh ADE environment containing a variety of biological and chemical contaminants and efficiently assimilating ammonium nitrogen and phosphorous.
The primary objective of this study is to evaluate the response of four microalgae species cultivated in unsterilized effluent of a live-stock anaerobic digestion unit, in order to identify the most suitable candidate in terms of biomass productivity and nutrients recovery efficiency. Specific tasks include the assessment of the effect of nitrogen source on biomass growth rate. Furthermore, the quantification of microalgae-derived biomolecules, such as carbohydrates, proteins, lipids and pigments was conducted as a means of correlating carbon storage mechanism with the physiology state of microalgae cells under different culture conditions, aiming to evaluate the capacity of each species to be further utilized in the context of multiple commodity production, aligned with the biorefinery concept. This holistic screening of microalgae response to a well-characterized ADE and the comparison with the reference media is claimed to be key contribution of the present study to: (i) the increase in the technology maturity of microalgae processes in wastewater treatment, (ii) the assessment of the microalgae capacity in waste attenuation in real industrial streams with biologic load compared to model streams, (iii) the assessment of the potential use of the produced microalgae biomass in biorefineries, and (iv) the generation of the required knowledge for the scale-up studies of the microalgae-based waste water treatment processes.

2. Materials and Methods

2.1. Microalgae Species

The microalgae species investigated in the current study are Chlorella vulgaris, Chlorella sorokiniana, Desmodesmus communis and Stichococcus sp. All of the strains were grown as unialgal, non-axenic cultures.
C. sorokiniana was isolated from the anaerobic digestion effluent employed in the present study according to the following procedure: diluted ADE samples in a concentration of 1%, 2% and 5% v/v were incubated in 500 mL glass volumetric cylinders at 25 °C. The samples were continuously supplied with atmospheric air at a flowrate of 0.5 L/min, achieving intensive agitation of the content. Cultures were symmetrically exposed to illuminance of 1000 lux, provided by cool white LED strips, with a photoperiod of 16/8 h light/dark. The samples were incubated for 18 days until the presence of microalgae was visually apparent (green color) in the cylinders. The microscopic observation confirmed the presence of microalgae population with monoculture characteristics. A volume of 3 mL of the solution was then transferred into 250 mL flasks containing sterilized BG-11 medium (composition in Section 2.3) and maintained under the same light and temperature conditions, using air supply of 0.25 L/min. Τhe species were characterized based on a polyphasic approach, using both morphological and molecular methods, following the protocol described in [25]. Briefly, morphological examination was performed using a Zeiss Axio imager z2 microscope (Carl Zeiss, Oberkochen, Germany), whilst the main diacritical features of Chlorella were identified. The genetic identification of the strain was based on the PCR amplification and sequencing of the molecular markers, 18S rRNA gene and rbcL gene, which are used widely in phylogenetic analyses of Chlorophyta. Direct sequences comparison, through Genbank database, indicated Chlorella sorokiniana species as the closest relative, with an identity up to 99%.
C. vulgaris and D. communis strains were obtained from the MicroAlgae and Cyanobacteria Culture Collection (TAU-MAC) [26] while Stichococcus sp., already cultivated in the lab, was isolated from the Crete coastal area in Southern Greece.

2.2. ADE Collection

The ADE employed in this study was collected from the effluent of an 1 MWel biogas plant through the anaerobic treatment of the waste streams from local animal husbandries. About 20 L of ADE were collected in a plastic tank and transferred to the laboratory. The sample was immediately centrifuged at 5000× g for 10 min, filtered (Whatman Inc., 150 mm, Grade 1, pore size 11 μm) and stored at −20 °C to avoid variation of wastewater composition.

2.3. Pre-Cultures Maintenance

Microalgae species were stocked at 25 °C in 500 mL Erlenmeyer flasks with a working volume of 300 mL in an incubating room. The entire incubating area was uniformly illuminated by LED strips producing cool white light and all of the flasks were exposed at illuminance of 1000 lux. Atmospheric air was sparged through filter (Whatman PTFE syringe filters, pore size 0.2 μm) at a flow rate of 0.2 L/min. A photoperiod of 16 h lighting followed by 8 h darkness was applied. Stock cultures were regenerated every 15 days in new flasks by inoculating fresh medium with the 15-day-old pre-culture at a volume ratio of approximately 1/10. The maintenance and regeneration of the stock cultures at standard frequency ensured the inoculation of all main cultures with a cell population of roughly the same physiology state and metabolic activity. All species were maintained in BG-11 medium, containing nutrients (in g/L): NaNO3 1.5, K2HPO4·3H2O 0.04, MgSO4·7H2O 0.075, CaCl2·2H2O 0.036, Citric Acid 0.006, Ammonium ferric citrate 0.006, Na2EDTA 0.001, Na2CO3 0.02; and trace elements (in mg/L): H3BO3 2.86, MnCl2·4H2O 1.81, ZnCl2 0.222, Na2MoO4·2H2O 0.391, CoCl·6H2O 0.05, CuSO4·5H2O 0.079. All of the nutrients and conical flasks were autoclaved at 121 °C for 20 min in separate solutions to prevent contamination in the early stage of growth. Ammonium ferric citrate and trace elements solutions were filter-sterilized (Whatman PTFE syringe filters, pore size 0.2 μm).

2.4. Species Cultivation

2.4.1. Media Composition

Microalgae species were cultivated in 3 different culture media, properly modified to account for the same elemental nitrogen concentration of 124 mg/L: (a) BG-11NO3 was prepared with standard chemical composition reported above, though with NaNO3 at a concentration of 0.75 g/L instead of 1.5 g/L; (b) BG-11NH4 was prepared by the replacement of sodium nitrate by ammonium chloride (NH4Cl) at a concentration of 0.472 g/L; (c) non-sterilized ADE was diluted with sterilized distilled water up to a final concentration of 3.7% v/v. All of the chemicals and glasses were autoclaved at 121 °C for 20 min.

2.4.2. Culture Conditions

The experiments were conducted in 250 mL Erlenmeyer flasks. Approximately 150–155 mL of each culture medium were inoculated with 15–20 mL of preculture, resulting in cultures of total initial volume of 170 mL and initial Optical Density (OD600nm) of approximately 0.35. Note that this value refers only to the cultures in BG-11NO3 and BG-11NH4, while the respective value for cultures in ADE medium was significantly higher due to the inherent coloring of the effluent. The flasks were placed in a in a shaking incubator (GFL 3031, GFL Gesellschaft für Labortechnik mbH, Burgwedel, Germany), operating at 110 rpm shaking rate and at a constant temperature of 25 °C. 250 mL/min of atmospheric air passed through 0.2 μm pore size Whatman PTFE syringe filters were sparged into each flask (EK-2LR, Kytola Instruments, Lahti, Finland). In addition, 2.5 mL/min of CO2 (purity ≥ 99.9%) were supplied in each flask using precision flowmeters (FL-3845G-HVR, Omega Engineering, Norwalk, CT, USA). Cool white LED strips (6000 K, 7.2 W/m) were employed to uniformly illuminate the cultures with 1200 lux light intensity measured by a portable light meter (SP 200K, Sauter, Kern & Sohn, Balingen, Germany). A daily photoperiod of 16 h lighting followed by 8 h darkness was applied. Species were cultivated in 17-day batches.

2.4.3. pH Control

The employed air-CO2 gas-mixture, other than the culture aeration and provision of inorganic carbon, serves for the efficient pH regulation. However, this regulation was not feasible only with the CO2 stream in cultures with BG-11NH4 as nutrient medium and they were buffered using 12 g/L Piperazine-1,4-bis (2-ethanesulfonicacid, PIPES, pKa 6.8), in order to prevent the sharp decrease in pH, due to acidification of the medium [27].

2.5. Analytical Measurements

2.5.1. Growth Determination

The progress of the microalgae culture growth in each experiment and the respective assimilation of nutrients were monitored by sample collection from cultures every 48 h. Optical density (OD) at 600 nm was measured using a UV-vis spectrophotometer (DR 3900, HACH, Loveland, CO, USA) while pH was daily measured to ensure sufficient pH control of the cultures (SevenMulti pH meter, Mettler Toledo, Greifensee, Switzerland). Determination of biomass concentration, measured as dry cell weight (DCW), was carried out by filtering 5 mL of samples cultures through a pre-weighted glass microfiber filter (Whatman 934-AH, pore size 1.2 μm), dried at 50 °C overnight, and weighted at a high precision micro-balance (XP 105, Mettler Toledo, Greifensee, Switzerland). Microscopic observations of the culture were carried out in an optical microscope (DM 2000, Leika, Wetzlar, Germany). Cell numbers were quantified using a Neubauer hemocytometer. Prior to counting, culture samples were properly diluted so as the final number of cells counted in the 5 squares of the chamber (1 central square and 4 corner squares) not to exceed 300. Cell counts for each sample were performed in triplicates.

2.5.2. Nutrient Analysis

Concentrations of ammonium nitrogen (N-NH4), nitrate nitrogen (N-NO3), total nitrogen (TN), soluble phosphorus (P-PO4) and chemical oxygen demand (COD) were determined in the liquid phase of filtrate samples using standard HACH cuvette tests in a UV–Vis spectrophotometer (DR 3900, HACH, Loveland, CO, USA). The characterization of digestate in macronutrients and heavy metals was performed via Atomic Absorption Spectroscopy (AAS) (AA-7000, Shimadzu, Kyoto, Japan).
The net amount of the produced microalgae biomass (DCWp, g/L), the average productivity (AP, mg/L/day), the nutrients removal efficiency (REi, %), the average removal rate (RRi, mg/L/day) and the specific removal efficiency (SREi, mg/g DCWp) were calculated according to the following equations:
D C W p = D C W f D C W 0
A P = ( D C W p / Δ t ) 1000
R E i = ( C 0 C f ) / C 0
R R i = ( C 0 C f ) / ( Δ t )  
where C0 and Cf correspond to nutrient concentration (mg/L) in the culture medium at the start (Day 0) and at the end of the experiment (or the day that the measured concentration was zeroed), respectively; Δt (day) is the elapsed time from the beginning to the end of the experiment (or the day that the measured concentration was zero), and DCWf and DCW0 represent the biomass concentrations at the start (Day 0) and at the end (Day 17) of the experiment (g/L), respectively.

2.5.3. Pigments Determination

The determination of pigments was carried out in samples collected at the end of each experiment according to the following protocol [28,29]. A 2 mL culture sample was centrifuged (7000× g for 10 min) and the precipitate was washed with distilled water 3 times. Then, 2 mL of pure methanol (99.8%) was then added to the cell pellet and the mixture was vortexed and left in dark at room temperature for 20 min. The concentration of pigments was calculated (in μg/L) through the following equations with prior measurement of the supernatant absorbance (A) at 470, 652 and 665 nm (DR 3900, HACH, Loveland, CO, USA):
C h l o r o p h y l l a ( C h l a ) = 16.72 × A 665 9.16 × A 652
C h l o r o p h y l l b ( C h l b ) = 34.09 × A 652 15.28 × A 665
C a r o t e n o i d s = ( ( 34.09 × A 470 ) ( 1.63 × C h l a ) ( 104.96 × C h l b ) ) / 221
The intracellularly accumulated macromolecular metabolites were determined at the end of each experiment. Biomass was harvested via centrifuge at 7000× g for 10 min and washed twice with distilled water. Cell pellets stored at −20 °C and then lyophilized.

2.5.4. Lipids Determination

Lipids were extracted from lyophilized cells and lipid content was gravimetrically determined according to the Bligh and Dyer extraction protocol [30]. Briefly, 5 mg of the freeze-dried algal biomass were treated with 2 mL methanol, 1 mL chloroform and 0.8 mL distilled water (2:1:0.8 v/v) and subjected to sonication within an ice-bath for 20 min, at 50% of sonicator’s maximum amplitude (Vibra Cell VC-505, Sonics & Materials, Newtown, CT, USA). The solution was centrifuged, and the solvent was collected. The procedure was repeated three subsequent times to ensure total lipids extraction. Volumes of 3 mL chloroform and 3 mL distilled water were added in the liquid solvent phase (2:2:1.8 v/v), and the mixture was centrifuged once more, so as the formation of two phases is ensured. After a complete separation of the two phases, the upper phase was removed, and the derived single-phase mixture (lower phase) containing the extracted lipids was collected. The lipids were dried overnight at 45 °C and weighed at a precision microbalance (XP 105, Mettler Toledo, Greifensee, Switzerland).

2.5.5. Proteins Determination

Proteins were extracted from 2 mg freeze-dried biomass upon treatment with 9.6 mL of 0.5 M NaOH aqueous solution, containing 5% v/v methanol and 0.4 mL 0.05 M phosphate buffer followed by sonication in an ice-bath for 10 min, at 50% of sonicator’s maximum amplitude to ensure cell breakage and protein release [31]. After homogenization, an additional 5 mL of the aqueous solution (NaOH 0.5 M, 5% v/v MeOH) was added, and samples were heated at 100 °C for 30 min under continuous stirring. The protein content was measured with a Micro-BCA kit (Thermo Scientific, Waltham, MA, USA) at a microplate spectrophotometer (ELx808 microplate Reader, BioTek, Winooski, VT, USA). The calibration curve was obtained with Bovine Serum Albumin (BSA) solutions of known concentrations.

2.5.6. Carbohydrates Determination

For the measurement of the carbohydrates content, 2 mg of lyophilized algal biomass were re-dispersed in 1 mL 2.5 M HCl solution and subsequently incubated at 100 °C for 3 h under continuous stirring, to achieve cell membrane breakage and reduction in polysaccharides, oligosaccharides, and disaccharides to monosaccharides. After neutralization with 2.5 M NaOH and centrifugation, the phenol-sulfuric method was used to quantify the content of the neutral monomers as glucose equivalent, by treating the unknown samples with 1 mL 1% (w/v) phenol solution and 5 mL 96% (w/w) H2SO4 and measurement of the absorption at 483 nm (Lamda 35, Perkin Elmer, Akron, OH, USA) [32]. Solutions of D-glucose of known concentrations were used as reference standards for calculating the glucose concentration-absorption calibration curve.

2.6. Statistical Analysis

All of the experiments were performed in duplicates and reported results are expressed as the respective mean values ± SD. One-way analysis of variance (ANOVA) was applied to determine whether there were any statistically significant differences between variables in Minitab (Minitab LLC, State College, PA, USA). The significance of differences between the analyzed variables was determined with a Tukey’s test. In all of the tests, the adopted level of significance was p < 0.05.

3. Results and Discussion

The study aims at the identification of the appropriate microalgae species for the reduction in the contained ammonium in the ADE of anaerobic digestion effluent. Towards this direction, the effect of different nitrogen sources on microalgae growth was examined, through the comparison of the growth of the employed microalgae species in diluted ADE rich in N-NH4 and chemically formulated media, i.e., standard BG-11 containing nitrates (BG-11NO3) and modified BG-11 utilizing ammonia nitrogen instead of nitrates (BG-11NH4). This task aims at developing a more comprehensive understanding of the nutrients utilization by microalgae and justify their adaptation to the effluent.
Four microalgae species were selected based on their origin, their growth rates in wastewater streams and their implementation in similar investigations. C. sorokiniana was isolated from the corresponding ADE, representing a native microorganism already adapted to the particular environment. Chlorella and Desmodesmus are two morphologically different genera that are commonly used in a variety of wastewater treatments applications [8,33,34,35]. On the other hand, Stichococcus sp. has been poorly investigated in terms of its capacity to grow in wastewater streams, even if high biomass production rates in culture media (3N-BBM-V) have been reported in the literature [36].

3.1. ADE Composition

Physicochemical analysis of the anaerobic digestion effluent was carried out to explore the adequacy of the effluent for the cultivation of microalgae in terms of the contained nutrients (Table 1). As shown, the effluent contained the majority of elements usually present in standardized culture media. However, a high concentration of ammonium nitrogen is measured (exceeding 3 g/L), mainly attributed to the introduction of poultry manure in the feedstock of the anaerobic digestion unit. Such a high ammonia content is inhibitory to microalgae cultures and dilution of the effluent up to a certain level is required prior to be used as culture medium.
A dilution of the raw ADE to a final concentration of 3.7% v/v was selected in an effort to provide a nutrient medium with desired nitrogen content, less than 150 mg/L, to prevent a potential inhibition effect on microalgae growth [37]. In addition, this dilution renders the ADE based cultivation medium comparable to the chemically defined artificial media (BG-11NO3 and BG-11NH4), primarily in terms of elemental nitrogen content (Table 1). Nitrogen in BG-11NO3 medium is exclusively in the form of nitrates, whereas in the other two media different types of nitrogen are expected: diluted ADE contains various nitrogenous sources, mostly accounting to ammonia (124 mg/L), with low nitrates content (3.3 mg/L). BG-11NH4 contains ammonia nitrogen (124.92 mg/L N-NH4) and it is also supplied with 12 g/L PIPES (C8H18N2O6S2), in order to achieve pH control. However, nitrogen contained in PIPES is not considered as an available N source for microalgae cells [38]. Regarding phosphorous availability in the media, BG-11 contains 5.5 mg/L P-PO4, while a reduced concentration was estimated in the diluted ADE, corresponding to 3 mg/L P-PO4. Among the inorganic components, potassium is significantly higher in 3.7% ADE than BG-11 media. However, elevated potassium values can benefit microalgae growth and stimulate ammonium uptake [39]. On the other hand, low concentrations of heavy metals that are measured in the digestate are not expected to restrict the growth of the tested species. Overall, the major abnormality of the digestate phase is the imbalance in terms of the two major microalgae growth components i.e., high nitrogen concentration along with low phosphorus values.
Thermal sterilization of ADE is a major issue when it is intended to be used as microalgae cultivation medium: a compromise should be made between its role as nutrient source on one side and the potential of culture contamination on the other side [39]. Steam sterilization of digestate has been proved to affect sample composition and characteristics, resulting in ammonia reduction due to stripping at elevated temperature, pH and total COD concentration increase, and phosphorus precipitation [40,41]. Moreover, sterilization of the digestate stream is expected to greatly increase the operating costs of full-scale microalgae plant used for effluent treatment. In the present study, preliminary experiments justified the reduction in ammonia concentration by more than 43% compared to the raw sample, when ADE subjected to autoclave treatment (data not shown); therefore, un-sterilized digestate samples were used in this study. Filtration trough 0.2 μm filters might address potential contamination issues, however, un-sterilized ADE was used in this study in order to evaluate the proposed process performance under industrial scale conditions where filter sterilization of large volumes of ADE might be costly and cumbersome.

3.2. Screening of Microalgae Species

3.2.1. Optical Density and Biomass Production

The growth of four selected microalgae species, C. vulgaris, C. sorokiniana, D. communis and Stichococcus sp. is examined in three different cultivation media, aiming to assess the potential to grow in liquid ADE solution derived from biogas plants. In general, optical density and biomass (expressed as Dry Cell Weight, DCW) measurements revealed that all tested microalgae species are adapted to the particular effluent and increase their population. Their growth in 3.7% ADE was further compared with reference culture media (BG-11), containing nitrate and ammonium as nitrogen source, in order to justify the ability of the species to assimilate different nitrogen sources.
Initial optical density in BG-11NO3 and BG-11NH4 in all experiments was the same for all species and approximately equal to 0.35, while for the 3.7% ADE it was 0.9, since the diluted ADE prior to inoculum addition is colored. However, the initial DCW and cells number were not the same for all microalgae species, due to their different structure, cell size/weight and pigments concentration in each species. As shown in Table 2, D. communis culture had almost double the initial DCW compared to the other three species and a noticeable lower cell number. This is attributed to the different cell morphology of the particular species. Both Chlorella species’ cells employed in the present study have spherical formulation with an average diameter between 3 and 9 μm [25] while Stichococcus sp. cells are rod-like with cell size ranging from 3 to 18 µm (microscopic observations). On the other hand, Desmodesmus communis is characterized by large ellipsoidal cells, 11–21 μm long and 3.4–9 μm wide, usually arranged in four-celled coenobia (Figure 1) [25]. Furthermore, as observed in the present study, D. communis has a significantly lower pigment concentration (chlorophyll a, chlorophyll b and carotenoids) compared to the other three species, as will be shownlater).
Figure 2 depicts the time profile of the optical density of the 4 species in the 3 different media, through the 17-day cultivation period together with the net biomass production measured at the last day of the cultivation period for each treatment. It is evident that all species demonstrated the highest growth rates, both in terms of OD600nm and DCWp when cultivated in BG-11NH4, where ammonium nitrogen was provided as nitrogen source. Significant differences between results were statistically proved by the ANOVA test (p < 0.05). These findings are aligned to the corresponding values of nitrogen consumption in different culture media as discussed in Section 3.2.2.
Specifically, Chlorella sorokiniana exhibited the maximum biomass production (DCWp), when cultivated in BG-11NH4 (2.33 g/L), followed by Chlorella vulgaris (2.26 g/L), Stichococcus sp. (1.96 g/L) and Desmodesmus communis (1.94 g/L) (Figure 2e and Table S1). However, a similar behavior is not depicted in the respective optical density trajectories (Figure 2b), as C. vulgaris has the maximum OD600nm at the end of the cultivation period followed by Stichococcus sp., Chlorella sorokiniana and Desmodesmus communis. The inconsistency between these values is explained by taking into consideration the concentration of pigments in each species (Figure 3 and Table S1). More specifically, C. vulgaris has the maximum chlorophyll a concentration (42.93 mg/L), compared to other 3 species, when BG-11NH4 was used as cultivation medium, while extremely poor concentration of chlorophyll a measured in D. communis cultures (1.81 mg/L) justifying the low OD values of this species, although its biomass concentration is comparable with Stichococcus sp. When nitrate was provided as nitrogen source (BG-11NO3), lower DCW and OD values were recorded for all studied species. However, the observed comparative performance of each species changed when ammonium nitrogen was used. The cultures of C. vulgaris and Stichococcus sp. in BG-11NO3 achieved the highest biomass production (1.03 and 1 g/L respectively), followed by C. sorokiniana (0.82 g/L) and D. communis (0.77 g/L) (Figure 2d and Table S1). In general, ammonium is considered the most accessible nitrogen source for algae metabolism, as the energy cost to reduce N-NH4 to organic matter is lower than the cost for other nitrogen forms reduction [42]. By using ammonium, the microalgae cells avoid energy needed for the nitrate/nitrite reduction, as NH4+ is directly incorporated into amino acids by condensation with glutamate to form glutamine, catalyzed by glutamine synthetase. It is possible that the lower energy requirement for the ammonium utilization, in contrast to nitrate uptake, allows a higher rate of nitrogen assimilation by microalgae and enables energy allocation to other metabolic processes, such as photosynthesis and growth [43]. This assumption is confirmed in the present study, as both the consumed nitrogen and the produced biomass, recorded in cultures of BG-11NH4 medium were much higher than the corresponding ones in cultures with BG-11N03 medium (Figure 4 and Table S1).
Successful growth of the 4 microalgae species was observed in 3.7% ADE. The growth curve and the final biomass of the four microalgae species is presented in Figure 2c,f and Table S1. These data are further supported by cells number significant incensement at the final day of cultivation period, justifying that DCW and OD augmentation reflects the microalgae growth. Furthermore, microscopic observations did not support any bacteria presence or other type of contamination. Among the 3 species, Chlorella vulgaris exhibited the highest values of OD and DCWp (3.11 and 0.89 g/L respectively), followed by C. sorokiniana and D. communis with the same DCWp values (0.7 g/L) and finally Stichococcus sp. (0.59 g/L). Chlorella sorokiniana cultivated in 3.7% ADE resulted in higher chlorophyll (a and b) concentration than the other 3 species, with concomitant higher OD values compared to D. communis, although these 2 species achieved the same biomass values (Figure 2 and Table S1). Many Chlorella species have been previously examined for their bioremediation capacity in centrates compared with control media and different results have been concluded in terms of biomass production or average productivity [15,23,40,44]. In the present study, D. communis achieved the lowest biomass production (DCWp) compared to the other 3 species when cultivated in BG-11NO3 and BG-11NH4, but its performance in terms of biomass production in 3.7% ADE was much higher than the result of similar studies [44,45].
From a literature review, it has been revealed that few articles have been published referring to Stichococcus sp. cultures in effluents [46,47]. However, in the present study the particular species exhibited an interesting behavior: during the first six days of the cultivation period in the 3.7% ADE medium the cells were organized in aggregates rather in unicellular structures and flocculated. This naturally occurred flocculation response of the Stichococcus sp. to the stress caused by the harsh environment of the effluent is considered as a survival strategy of the species which upon the adaptation period exhibited a high nitrogen need. However, after that extended adaptation period, the culture exhibited a growing period during the following days, resulting in a final produced biomass of 0.59 g/L (Figure 2f). From the screening of the performance of the studied microalgae species it can be concluded that maximum biomass production efficiency is attained in BG-11NH4 while the corresponding value in 3.7% ADE was about 60–70% lower. When the adequacy of the cultivation in 3.7% ADE is compared against the BG-11NO3 medium, the respective values of DCWp attained accounted to 74% for the Stichococcus sp., 86% for the C. vulgaris, 85% for the C. sorokiniana and 91% for the D. communis.

3.2.2. Nutrient Assimilation

The nutrient uptake of ammonium (N-NH4), nitrate (N-NO3), total nitrogen (TN), -phosphate (P-PO4) and chemical oxygen demand (COD) were measured during the cultivation of the 4 species in 3.7% ADE, BG-11N03 and BG-11NH4. The assessment of the growth of microalgae in the three different culture media and the corresponding nutritional utilization (with emphasis in nitrogen consumption) is crucial, since these two operational traits determine the decision making over the most appropriate microalgae species for bioremediation purposes.
  • Nitrogen uptake
Nitrogen participates in a variety of vital biological compounds, such as peptides, proteins, enzymes, chlorophylls, energy transfer molecules (ADP, ATP), and genetic materials (RNA, DNA) [48,49]. Microalgae cultivated in BG-11 supplemented with ammonium nitrogen instead of nitrate achieved higher nitrogen assimilation rates. More specifically, Stichococcus sp., was found to be an excellent N-NH4 consumer, achieving the highest nitrogen uptake in BG-11NH4, i.e., 90.79% of the initial nitrogen corresponding to 105.32 mg N-NH4 in 17 days of cultivation (Figure 4b,e). However, the performance of the same species when nitrate was used as N-source was lower, consuming only 43.06% of the initially provided nitrogen (52.1 mg N-NO3), representing the lowest nitrogen consumption when compared to the other 3 species (Figure 4a,d). In BG-11N03, representing a universally suggested medium for the cultivation of many non-marine microalgae species, C. vulgaris and C. sorokiniana achieved an increased nitrogen uptake efficiency (65.23% and 51.95% of the initially loaded nitrogen respectively), while D. communis exhibited lower capacity (46.4%) (Figure 4d). As mentioned, Stichococcus sp. consumed the higher amount of ammonium nitrogen n BG-11NH4, followed by C. vulgaris (89.46%), C. sorokiniana (79.18%) and D. communis (73.26%) (Figure 4e).
Despite the unambiguously improved response of microalgae cultures to media containing ammonium as nitrogen source, the vast majority of the cultivation media used for algae cultivation contain nitrate instead of ammonium nitrogen. This is attributed to the fact that when ammonia is used as N-source, the addition of biological buffers or pH controllers is required in order to prevent the sharp decrease in the pH. Several studies have demonstrated that growth of microalgae in unbuffered media supplemented with N-NH4 resulted in pH values lower than 4 after few days of cultivation, with concomitant inhibition of the cells proliferation [27,41,50,51]. The use of N-NH4 in microalgae cultures medium causes acidification of the buffer presumably because of photons translocation out of microalgae cells, in order to maintain cell neutrality during uptake of the cations [27]. However, when ammonia is the fundamental N-source in biological buffers with high alkalinity, as it happens in ADE, the continuous air supply for mixing of the cell cultures can result to pH increase due to media alkalinity consumption and carbon dioxide equilibria, followed by the formation of gaseous ammonia and volatilization. Under these conditions, pH adjustment is required to prevent the air stripping of ammonia [52,53]. On the other hand, when nitrate is used as nitrogen source the pH of the buffer is increased when air is supplied, because of hydroxyl production [27,50,51,54]. In the present study, pH adjustment of cultures in BG-11N03 and ADE was achieved by the addition of 1 % v/v CO2, while pH control in BG-11NH4 cultures (which were also supplemented by CO2 for conformity reasons) is carried out by the addition of PIPES in concentration 12 g/L.
Ammonium nitrogen bioremediation of the ADE by microalgae is depicted in Figure 4c,f. All of the investigated species successfully assimilate the available nitrogen of the effluent producing biomass. More specific, Stichococcus sp. cultivated in 3.7% ADE performed particularly well with respect to nitrogen removal, in terms of both consumption rate and efficiency, yielding an average N-NH4 uptake of 4.94 mg/L/day and consuming 69.7% of the initial amount of N (Figure 4f and Table S1). This behavior is in complete alignment with its performance in BG-11NH4, where maximum N-NH4 was observed. Note that, ammonium uptake efficiency by Stichococcus sp. in 3.7% ADE was even higher than nitrate nitrogen consumption in BG-11NO3 for the same culture conditions, confirming that this species is capable of assimilating N-NH4 extremely efficiently compared to nitrate.
The N-NH4 removal efficiency of C. vulgaris, C. sorokiniana and D. communis in the ADE is comparable, ranging from 49.1% to 53.1% (Figure 4f). Previous studies on microalgae bioremediation capacity in anaerobic digestion wastewaters suggest similar findings. Kim et al. [15] and Kobayashi et al. [55], investigated the response of different microalgae species in ADE derived from wastewater treatment plants and anaerobic digester fed with manure slurry respectively, and observed comparable N-NH4 removal rates for C. vulgaris and C. sorokiniana; however, the initial nitrogen concentration in these studies was lower than in the present one. When Desmodesmus sp. was tested in 5% ADE from a biogas plant in batch mode cultivation, it yielded the same nitrogen removal rate compared with the present study (i.e., 3.72 mg/L/day), even though a lower N-NH4 concentration was used [45].
It should be underlined that efficient NH4 removal is observed in this study, although the AD effluent contained substantially higher initial levels of nitrogen, as well as phosphorus, compared to other reported works. In general, in most of the studies using centrate as cultivation media for microalgae cultures, N-NH4 concentration does not exceed 100 mg/L [14,56,57]. Many authors have reported that ammonium nitrogen concentration higher than this threshold is probably toxic to microalgae cultivation. A recent study by Zheng et al. (2019) tried to elucidate the effect of different N-NH4 concentrations on microalgae cell viability maintaining the same effluent concentration, and it was concluded that ammonium nitrogen in a concentration of 110 mg/L was favorable in terms of photosynthetic efficiency and nutrient consumption, while cell viability was significantly decreased when N-NH4 was increased from 110 to 220 mg/L [58]. On the other hand, an assessment of different concentrations of digestates as cultivation media for Chlorella sp. showed an inhibitory effect of ammonia nitrogen concentration over 160 mg/L, when maize silage/swine slurry and cattle manure media was used, while the same behavior was not observed in sewage sludge and dairy wastewater [56]. Meanwhile, C. sorokiniana tested in the same ammonia concentration (i.e., 160 mg/L N-NH4) exhibited the maximum growth rate and removal efficiency of nitrogen and phosphorus compared to lower initials concentrations [50]. Ji et al. (2014) reported that Desmodesmus sp. was not able to survive in anaerobic digestion wastewater in a 12% dilution corresponding to 100 mg/L N-NH4, although in the current study, Desmodesmus communis successfully proliferated in 3.7% ADE despite the fact that ammonium concentration was higher than 120 mg/L [57]. Therefore, microalgae tolerance is not exclusively linked to ammonium concentration alone, but potentially to a number of parameters with synergetic effects such as sample turbidity and macro- and micro-nutrients concentration.
When N-NH4 removal efficiency by microalgae is assessed, it is very important to ensure that the pH is efficiently adjusted. Based on hydrolysis of ammonia, nitrogen removal is possibly taking place abiotically (ammonia volatilization/stripping) than via a microalgae bioremediation process. There is a well-known chemical equilibrium between NH3 and NH4+ in water, according to the following formula: NH4+ + OH ⇔ NH3 (g) + H2O (pKa = 9.26 at 25 °C). Stability of the equilibrium between ammonium (NH4+; ionized form) and ammonia (NH3; non-ionized gaseous form) is highly dependent on temperature and pH; nitrogen exists exclusively in the ammonium form (NH4+) under acidic conditions (below pH 7), about 0.5% is in gaseous form (NH3) at pH 7, while the fraction of NH3 increases in alkaline conditions. Temperature is also a determining factor in this process, since ammonia is dominant at lower pH at high temperature as pKa value decreases with temperature [42,52]. Many authors have reported abiotic loss of ammonia in microalgae cultivation in digestates [40,59,60]. As stated by González et al., [61], ammonia stripping constitutes the main removal mechanism in open photo-bioreactors, and NH4 uptake up to 98% is easily attained. In the present study, all data support the assumption that ammonium nitrogen removal was not caused by abiotic transformation but by microalgae uptake, given that the pH of the cultures in ADE was maintained very close to 7 by CO2 gas injection and at temperature of 25 °C (data not shown).
The NH3-N removal mechanism may include not only the assimilation by microalgae, but also the conversion of ammonia into nitrates by endogenous nitrifying bacteria present in non-sterilized wastewater. Several authors have reported that nitrates increase during microalgae cultivation in wastewater streams [24,62]. In the present study, N-NO3 concentration in 3.7% ADE is 3.3 mg/L, while a certain number of nitrates are added to the medium with the inoculum, given that the precultures were exclusively prepared in BG-11NO3. As a result, the initial concentration of N-NO3 in ADE experiments ranged from 18 to 36 mg/L, depending on the residual NaNO3 concentration in the pre-cultures. However, as shown in Table 3, during the cultivation of the 4 microalgae species in 3.7% ADE, only a slight variation of the initial nitrate concentration was observed. Therefore, the presence of nitrifiers is excluded, and the N-utilization by the microalgae was highly selective towards the ammonium form, which the preferred N substrate. It is assumed that nitrate consumption does not occur as long as ammonium nitrogen was not completely consumed [63]. Similar data were obtained in the BG-11NH4 experiment (data not showed).
In addition to the above nitrogen sources, microalgae can utilize nitrogen derived from organic compounds, such as urea, glycine, amino acids, purines, and pyrimidines etc. [52,53]. Depending upon the availability of nitrogen in the environment, algal cells control nitrogen using nitrogen transcription regulators [54]. Most of the nitrogen in the digestate was in the form of ammonium nitrogen (approximately 70–78%), and therefore was readily available to microalgae [20,56]. Other forms of nitrogen include nitrates (approximately 3%, as referred before), nitrites and organic N. Concentration of N-NO2 is measured to zero, so residual nitrogen forms in the ADE is mainly organic nitrogen [40,59,64]. However, as presented in Table 3 nitrogen consumption by all of the tested microalgae species in the ADE included exclusively N-NH4 uptake, while residual nitrogen (corresponding to N-organic) was not reduced, advocating the suggestion that microalgae do not use other nitrogen sources until all the ammonium is utilized [52]. Several authors reported the removal tendency of total and ammonia nitrogen by different microalgae species cultivated in ADEs, suggesting nitrate and organic nitrogen playing a very weak role in algae growth on centrates [40,65].
  • Phosphorus uptake
Phosphorus is a key factor in the energy metabolism of algae and is present in nucleic acids, lipids, proteins, and the intermediates of carbohydrate metabolism. Inorganic phosphates play a significant role in algae cell growth and metabolism. During microalgae metabolism, phosphorus, preferably in the forms of H2PO4 and HPO4, is incorporated into organic compounds through phosphorylation, much of which involves the generation of ATP from adenosine diphosphate (ADP), accompanied by a form of energy input [48,57]. Similarly to nitrogen, phosphorus removal might occur by abiotic processes, as elevated pH and Ca2+ or Mg2+ concentration in the buffer can lead to P precipitation and co-precipitation, such as calcium phosphate, magnesium phosphate and/or struvite (MgNH4PO4·6H2O) [15,54,66]. However, in the present study, the maintenance of the culture pH below 7.5 and temperature at 25 °C, confines the mechanisms of phosphorus removal far and away to algae uptake.
The results of phosphorus removal by the different species cultivated in the three media are shown in Figure 5 as a function of time. The cellular need in phosphorus is not similar to that of nitrogen, thus an entirely different assimilation policy was implemented. Precisely, phosphorous uptake was found to occur much faster in Desmodesmus communis cultures compared to the other 3 species, regardless the employed cultivation medium (i.e., BG-11NO3, BG-11NH4 and 3.7% ADE), the N source (NO3 or NH4) and the initial P-PO4 concentration (3 or 5.5 mg/L) (Figure 5 and Table 1). A depletion of orthophosphates was observed by the fifth day of cultivation, resulted in D. communis presenting the highest P-PO4 removal rate in each experiment, compared to the other species. Other studies, assessing the bioremediation capacity of different Desmodesmus species in centrates reported lower P removal rates [45,57]. However, the respective performance of the other three species seems to be related with the culture media used. C. vulgaris exhibited the lowest P-PO4 average removal rate (RR), among the studied species when cultivated in BG-11NO3, but the highest N-NO3 RR.
On the other hand, C. sorokiniana and Stichococcus sp. presented significantly lower phosphorus uptake rate cultivated in 3.7% ADE, compared to their performance in BG-11NO3 and BG-11NH4 (Table S1). The experimental results indicate that higher P-PO4 RRs are not necessarily combined with higher biomass accumulation by these species.
Given that all of the tested media were relatively poor in phosphorus, complete phosphorus consumption considerably earlier than the 17 days of cultivation was attained by all of the studied species. In order to ensure the simultaneous utilization of both nitrogen and phosphorus, the N/P ratio should be within a proper range [48]. The optimal ratio differs among different microorganisms, due to species-varying metabolic pathways, but it appears that in most cases it varies between 5 and 30 [67]. In the present study, initial N/P ratio in BG-11 media was 22.3, while in ADE the corresponding ratio was 41, with respect to ammonium nitrogen. Compared to previously reported anaerobic digestion effluents the phosphorous contained in the centrate was not in an adequate proportion, taking into consideration the high concentrations of nitrogen [45,56,57,59,68]. This nutrient lack of balance might explain the moderate performance of microalgae in 3.7% ADE, compared to BG-11 media; it is expected that an adjusted N/P ratio would increase both the culture growth rate and the removal rates of nitrogen and phosphorus [8,50].
  • COD removal
The reduction in organic carbon in wastewater is as important as nutrient uptake in order to meet the effluent discharge standard. Effluents purification with emphasis on COD bioremediation by microalgae has widely been investigated [5,69,70]. Microalgae are capable of growing mixotrophically and simultaneously assimilate CO2 and organic carbon with both respiratory and photosynthetic metabolism operating concurrently [54].
In the present study, soluble COD is effectively consumed during microalgal growth, enhancing the remediation efficiency of the process. As presented in Figure 6, COD removal varied from 557 to 317 mg/L, with C. sorokiniana achieving the highest COD RE (63%) and D. communis the lowest one (35%). A great fraction of COD was consumed during the first ten days, while additional COD reduction was rather negligible at longer times. This can be attributed to the fact that carbon compounds that can be consumed by microalgae are utilized at short times, while the remaining organics are not easily assimilated and remain in the solution [71]. Thus, part of the organic carbon compounds present in ADE are utilized by microalgae as carbon source simultaneously with the supplied CO2, potentially shifting their metabolism to a mixotrophic mode. It is worth mentioning that, in the present study, despite the relatively high value of the initial COD compared to other similar works in the literature [3,70] no inhibitory effect was identified on the culture performance.

3.2.3. Effect of Nutrient Media on Biomass Composition

The primary target of this study is the investigation of the potential of microalgae for the treatment of an effluent with high nitrogen content; however, the examination of microalgae composition represents a challenging approach for the following biomass valorization in various applications. It is known that microalgae biomass composition is significantly affected by the cultivation conditions, nitrogen source and growth media composition [72]. The biochemical composition of the four species, cultivated in the three different media, is presented in Figure 7. The results of the present work justified that both cultivation media and nitrogen source had a considerable impact on the accumulation of the three major intracellular metabolites (proteins, carbohydrates, and lipids) of the selected species. Protein production was slightly decreased in all species when 3.7% ADE was used as cultivation medium (not statistically different), while during cultivation in BG-11 media, negligible effect was recorded by the type of the nitrogen source on the proteins content. Previous reports suggested that cultivation of microalgae in anaerobic digestion effluents resulted in lower proteins content when compared to cultivation under control media [11,55]. However, in these studies, complete consumption of all available nitrogen took place, and the reduction in the protein content in microalgae cells is occurred under N- starvation conditions [36,49]. Ιn the present study, concentration of the residual nitrogen (N-NH4) was ranged from 10.6 to 31.9 mg/L, indicating nitrogen availability until the end of the experiments.
On the other hand, it was clearly observed that 3.7% ADE favored the synthesis of carbohydrates by all studied species compared to BG-11 media. Specifically, D. communis presented the most remarkable increase in its polysaccharide content corresponding to 83.1% (compared to the corresponding value on BG-11 media), followed by C. sorokiniana (53.3%), Stichococcus sp. (40.7%) and C. vulgaris (13.6%). It has been proved that stressful conditions (e.g., nutrients starvation, variable light policies, exposure to toxic substances) might cause an alteration on microalgae metabolism turning the flow of the photosynthetically fixed carbon from protein synthesis to lipid or carbohydrate synthesis pathways [49,73]. Cultivation of species in ADE can be accounted as a stress factor, associated with the most moderate biomass production. Moreover, taking into account that carbohydrates serve as carbon and energy sinks in microalgae cells [46,74] accumulation of sugars could be justified as a stress response for microalgae cells. These observations align with other studies where microalgae metabolism is shifted towards the accumulation of sugars under cultivation in anaerobic digestion effluents [11,55,65].
As shown in Figure 7 proteins and carbohydrates synthesis pathway was not affected by the type of nitrogen source used in the standardized BG-11 growth medium (nitrate or ammonium). However, a different behavior was recorded in lipids accumulation, that presented a direct correlation to ammonium nitrogen (BG-11NH4): as depicted in Figure 7c, lipids content was maximized in all microalgae species under cultivation in BG-11 supplemented with ammonium nitrogen (p < 0.05). The same observations have been previously reported in other studies related to the effect of different nitrogen sources in microalgae performance [38,53,74]. Although limited data regarding the effect of ammonia nitrogen in artificial growth media on microalgae biochemical composition have been reported, it seems that cellular accumulation of lipids in microalgae is usually enhanced by applying factors that limit or prohibit cells’ proliferation. Nevertheless, elevated lipid content in microalgae cells using ammonium media is often associated with moderate cell growth of the cultures, and consequently, can be considered as a result of stress response. However, in the present work, the microalgae cultures in BG-11NH4 resulted in maximum values of microalgae biomass and lipid content. A similar behavior was not observed during the species cultivation in the digestate containing ammonia. It can be assumed that the use of ammonia as nitrogen source, can trigger the lipid overexpression; a tendency that possibly suppressed in the presence of the effluent because of the cells’ metabolism alteration towards carbohydrates accumulation.

4. Conclusions

Four microalgae species were cultivated in ammonium rich effluent derived from biogas plant and two synthetic cultivation media supplemented with both nitrate (BG-11NO3) and ammonium nitrogen (BG-11NH4). The results of the study proved that ADE could serve as growth medium for microalgae cultivation, and both biomass production and wastewater purification are feasible in a simultaneous and synergetic manner, since all examined species showed substantial proliferation in 3.7% ADE.
Examination of microalgae response in different nitrogen sources revealed that ammonium is the preferable nitrogen form for all investigated species, as (1) cultivation in BG-11NH4 resulted in the most elevated values of cell mass and nitrogen removal efficiency and (2) no consumption of other nitrogenous sources was recorded when N-NH4 was available (i.e., N-NO3 and organic nitrogen). However, the performance of each species in terms of biomass production and nutrients assimilation differed in each medium. Chlorella vulgaris achieved the highest biomass production when cultivated both in BG-11 containing nitrates and in 3.7% ADE, compared to the other 3 species, while Chlorella sorokiniana exhibited the highest value of cell mass in BG-11NH4. Stichococcus sp. was found to assimilate ammonium nitrogen more rapidly in 3.7% ADE and in BGNH4 than the other 3 candidates, even if this trend is not accompanied by high values of cell mass weight.
Even though microalgae P-PO4 assimilation capacity was found to be associated with the nitrogen source and the cultivation medium, Desmodesmus communis stood out among the other four investigated species, recording the highest phosphorus removal rates in any of the three cultivation media that were tested. However, low biomass production (DCWp), pigment concentration and COD removal efficiency values of the particular species recorded both in ADE and reference media, mean that D. communis is not a sustainable option for ADE bioremediation.

Supplementary Materials

The following supporting information can be downloaded at:, Table S1: Microalgae performance and productivities, nutrients bioremediation capacity and biochemical composition at the end of the cultivation period of the four species in the three culture media.

Author Contributions

P.P. and S.-N.S. performed and validated the experiments and collected the data; U.L. and S.G. contributed to the strain identification. P.P wrote and edited the manuscript. P.S C.C. and S.G. reviewed the manuscript; P.S. and C.C. supervised the experimental work, and conceptualized the project. P.S is the project administrator and responsible for funding acquisition. All authors have read and agreed to the published version of the manuscript.


This research has been co-financed by the European Union and Greek national funds through the Operational Program Competitiveness, Entrepreneurship and Innovation, under the call RESEARCH—CREATE—INNOVATE (project code: T1EDK-00406 -title «Innovative technologies for the elimination of NH3 toxicity in anaerobic fermentation to increase methane production»).

Data Availability Statement

Data are contained within the article.


The authors would like to thank the Laboratories of research and analysis Q-Lab for the characterization of digestate in macronutrients and heavy metals.

Conflicts of Interest

The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.


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Figure 1. Microalgae species used in the present study: (a) C. vulgaris, (b) C. sorokiniana, (c) D. communis and (d) Stichococcus sp.
Figure 1. Microalgae species used in the present study: (a) C. vulgaris, (b) C. sorokiniana, (c) D. communis and (d) Stichococcus sp.
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Figure 2. Optical density (OD600nm) (ac) and biomass concentration values (DCWp) at the end of the culture period (df) of the four microalgae species in the three different media: blue bar: C. vulgaris, orange bar: C. sorokiniana, green bar: D. communis and red bar: Stichococcus sp.
Figure 2. Optical density (OD600nm) (ac) and biomass concentration values (DCWp) at the end of the culture period (df) of the four microalgae species in the three different media: blue bar: C. vulgaris, orange bar: C. sorokiniana, green bar: D. communis and red bar: Stichococcus sp.
Water 14 00221 g002aWater 14 00221 g002b
Figure 3. Pigment concentration at the end of the culture period (17th day) of the four microalgae species in the three different media (ac): green bar: Chlorophyll a, blue bar: Chlorophyll b orange bar: Carotenoids.
Figure 3. Pigment concentration at the end of the culture period (17th day) of the four microalgae species in the three different media (ac): green bar: Chlorophyll a, blue bar: Chlorophyll b orange bar: Carotenoids.
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Figure 4. Nitrogen concentration in the three different media by microalgae (ac) and nitrogen RE at the end of the culture period (df) of the four microalgae species in the three different media: C. vulgaris, orange bar: C. sorokiniana, green bar: D. communis and red bar: Stichococcus sp.
Figure 4. Nitrogen concentration in the three different media by microalgae (ac) and nitrogen RE at the end of the culture period (df) of the four microalgae species in the three different media: C. vulgaris, orange bar: C. sorokiniana, green bar: D. communis and red bar: Stichococcus sp.
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Figure 5. Phosphorus consumption in the three different media by microalgae: (a) BG-11N03; (b) BG-11NH4 and (c) 3.7% ADE.
Figure 5. Phosphorus consumption in the three different media by microalgae: (a) BG-11N03; (b) BG-11NH4 and (c) 3.7% ADE.
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Figure 6. COD uptake in 3.7% ADE by microalgae.
Figure 6. COD uptake in 3.7% ADE by microalgae.
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Figure 7. Biochemical composition of the four microalgae species under cultivation in the 3 different media (ac): blue bar: BG-11NO3, orange bar: BG-11NH4 and green bar: 3.7% ADE.
Figure 7. Biochemical composition of the four microalgae species under cultivation in the 3 different media (ac): blue bar: BG-11NO3, orange bar: BG-11NH4 and green bar: 3.7% ADE.
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Table 1. Physicochemical composition of raw ADE.
Table 1. Physicochemical composition of raw ADE.
ParametersBG-11NO3BG-11NH43.7% ADERaw ADE
N-NH4 (mg/L)ND123.921243351.35
Ν-ΝO3 (mg/L)123.92ND3.389.19
TN (mg/L)123.92123.921704594.6
P (mg/L)5.505.50381.1
COD (mg/L)NDND895.6324,200
BOD (mg/L)NDND2386430
K (mg/L)13.7013.70116.963161
Ca (mg/L)9.819.8113.66369.1
Na (mg/L)212.289.4169.731884.57
Fe (mg/L)
Cl (mg/L)18.02331.5860.441633.6
Mg (mg/L)6.986.988.3225.14
Mn (mg/L)0.500.500.236.32
Zn (mg/L)
Cu (mg/L)
S (mg/L)9.829.82NMNM
Co (mg/L)0.010.01NMNM
Mo (mg/L)0.160.16NMNM
B (mg/L)0.500.50NMNM
Citric acid (mg/L)6.006.00NDND
ND = Not Detected, NM = Not Measured.
Table 2. Initial conditions applied in the three growth experiments.
Table 2. Initial conditions applied in the three growth experiments.
BG-11NO3BG-11NH43.7% ADE
Initial OD600nm
C. vulgaris0.34 ± 0.000.34 ± 0.000.91 ± 0.01
C. sorokiniana0.35 ± 0.010.35 ± 0.000.91 ± 0.01
D. communis0.35 ± 0.010.35 ± 0.010.88 ± 0.00
Stichococcus sp.0.37 ± 0.010.37 ± 0.010.97 ± 0.01
Initial DCW (g/L)
C. vulgaris0.12 ± 0.000.12 ± 0.000.12 ± 0.00
C. sorokiniana0.11 ± 0.000.11 ± 0.000.11 ± 0.00
D. communis0.27 ± 0.010.27 ± 0.010.27 ± 0.01
Stichococcus sp.0.14 ± 0.010.14 ± 0.010.14 ± 0.01
Initial cell number (cells/mL)
C. vulgaris4.17 × 106 ± 3.16 × 1054.21 × 106 ± 3.05 × 1054.09 × 106 ± 2.22 × 105
C. sorokiniana4.36 × 106 ± 1.21 × 1054.13 × 106 ± 2.16 × 1054.32 × 106 ± 1.19 × 105
D. communis3.367 × 105 ± 5.92 × 1043.561 × 105 ± 1.98 × 1043.489 × 105 ± 3.01 × 104
Stichococcus sp.3.47 × 106 ± 2.17 × 1053.29 × 106 ± 1.77 × 1053.52 × 106 ± 1.09 × 105
Table 3. Consumption of different nitrogen forms of the four microalgae species cultures in 3.7% ADE.
Table 3. Consumption of different nitrogen forms of the four microalgae species cultures in 3.7% ADE.
C. vulgarisC. sorokinianaD. communisStichococcus sp.
Nitrogen (mg/L)Day 0Day 17Day 0Day 17Day 0Day 17Day 0Day 17
ΤΝ169 ± 1.42104.25 ± 14.01176 ± 0.97114.5 ± 15.81177 ± 0.56115 ± 13.45186 ± 1.19102 ± 10.09
N-NH4 (mg/L)122.94 ± 0.2557.72 ± 12.03125.8 ± 0.6964 ± 13.77122.6 ± 0.3460.4 ± 12.53120.5 ± 0.1436.52 ± 8.72
N-NO3 (mg/L)18.06 ± 0.1118 ± 0.1015.5 ± 0.0415.8 ± 0.1424.1 ± 0.0224.1 ± 0.0634.9 ± 0.0336 ± 0.72
N-organic28 ± 1.5028.53 ± 2.6534.7 ± 0.3434.7 ± 3.0830.3 ± 0.3430.5 ± 1.2230.6 ± 1.4429.48 ± 0.92
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Psachoulia, P.; Schortsianiti, S.-N.; Lortou, U.; Gkelis, S.; Chatzidoukas, C.; Samaras, P. Assessment of Nutrients Recovery Capacity and Biomass Growth of Four Microalgae Species in Anaerobic Digestion Effluent. Water 2022, 14, 221.

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Psachoulia P, Schortsianiti S-N, Lortou U, Gkelis S, Chatzidoukas C, Samaras P. Assessment of Nutrients Recovery Capacity and Biomass Growth of Four Microalgae Species in Anaerobic Digestion Effluent. Water. 2022; 14(2):221.

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Psachoulia, Paraskevi, Sofia-Natalia Schortsianiti, Urania Lortou, Spyros Gkelis, Christos Chatzidoukas, and Petros Samaras. 2022. "Assessment of Nutrients Recovery Capacity and Biomass Growth of Four Microalgae Species in Anaerobic Digestion Effluent" Water 14, no. 2: 221.

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