Development of a Greenhouse Wastewater Stream Utilization System for On-Site Microalgae-Based Biostimulant Production

Abstract

The challenges to feed the world in 2050 are becoming more and more apparent. This calls for producing more with fewer inputs (most of them under scarcity), higher resource efficiency, minimum or zero effect on the environment, and higher sustainability. Therefore, increasing the circularity of production systems is highly significant for their sustainability. This study investigates the utilization of waste streams from greenhouse hydroponic drainage nutrient solutions for the cultivation of the microalgae Desmodesmus sp. The cultivation was done in an automatically controlled container-scale closed tubular Photo Bio-Reactor (PBR). The study included lab-scale open-pond system experiments and in situ container-scale experiments in the greenhouse wastewater system to assess biomass growth, optical density, nitrogen consumption, and the influence of enzymatic complexes on microalgae cell breakdown. A batch-harvesting process was followed, and the harvested microalgae biomass was pre-concentrated using FeCl₃ as a flocculant that has demonstrated efficient sedimentation and biomass recovery. Following microalgae sedimentation, the produced biomass was used for biostimulant production by means of a biocatalysis process. The enzymatic complexes, “EnzProt”, “EnzCell”, and “EnzMix” were tested for cell breakdown, with “EnzMix” at a dosage of 10% showing the most promising results. The results demonstrate successful biomass production and nitrogen uptake in the lab-scale open-pond system, with promising upscaling results within container-scale cultivation. The findings contribute to a better assessment of the needs of Desmodesmus sp. culture and highlight the importance in optimizing culture conditions and enzymatic processes for the production of biostimulants.

1. Introduction

Wastewater treatment and recycling is a crucial process to ensure human well-being in a sustainable environment. The ever-increasing volume of wastewater produced by the expansion of urbanization and industrialization poses a significant environmental and public health risk [1]. If left untreated, wastewater infused with chemicals, pathogens, and other various pollutants can lead to the contamination of natural water bodies, posing a serious threat to the ecosystem and human health [2].

The main goal of wastewater treatment systems is to purify contaminated water by domestic, industrial, agricultural or commercial activities. The processes the wastewater undergoes in order to be rendered clean are physical, chemical, and biological and aim to remove pathogens and pollutants [3]. Treated water that meets quality standards can then be released back into the environment.

The emerging contaminants and the need for sustainable and efficient practices make necessary the understanding of current treatment methodologies and the exploration of recent innovations in order to achieve environmentally friendly wastewater treatment solutions. A recent example of pertinent research is the work conducted by Silva [1], which focuses on the need for wastewater treatment and analyzes various wastewater treatment methods and their levels of sustainability, with the aim of proposing an appropriate wastewater treatment approach. Another example is a paper published by Silva [4] that aims to present and emphasize the importance of reuse as a water supply solution for cities. It also explains the evidence and the strategies that cities can use to boost water supply.

Microalgae and macroalgae are considered rich sources of plant biostimulants, presenting an attractive business opportunity in agriculture and agro-based industries. Despite their potential, the use of microalgae as biostimulants has encountered constraints, mainly attributed to insufficient research and high production costs. As a result, many microalgae species with potential biostimulant properties remain unexploited [5,6].

Microalgae effectively contribute to the achievement of all the Sustainable Development Goals (SDGs) and are considered a renewable and sustainable energy source that does not compete with food production, does not require fresh water, and offers multiple applications [7]. In particular, the integration of microalgae-based biotechnology signifies a reduction in the carbon footprint associated with agricultural practices, aligning with the imperative for environmentally conscious methodologies [8].

Microalgae cultivation in controlled environments such as closed Photo Bio-Reactors (PBRs) presents distinct advantages [9] compared to open-race ponds. Closed systems offer controlled conditions that mitigate external contamination, allowing precise regulation of critical parameters including light, temperature, pH, and nutrients, resulting in improved growth rates and higher biomass yields [10,11]. Furthermore, they enable efficient resource utilization, protection from adverse weather conditions, and optimal control over light exposure. Closed PBRs also facilitate higher algae concentrations due to their controlled culture volume, minimizing dilution effects while offering flexibility in location and scalability. These technological advancements [11] in algae cultivation technology have diverse applications in industries such as pharmaceuticals, nutraceuticals, biostimulants, biofuels, and aquaculture systems, showcasing their potential for varied sustainable uses.

PBRs employed in this system are meticulously designed for optimal microalgae growth, leveraging advanced lighting technologies such as LED or OLED [12] to enhance the cultivation process and maximize biostimulant production efficiency, taking into account optimal light penetration for photosynthesis and maximum biomass production [13,14]. Understanding the intricate dynamics of light distribution within PBRs is crucial for maximizing microalgae growth and productivity. For instance, Akach et al. [15] delved into the simulation of light distribution within PBRs, shedding light on the complexities of this phenomenon and its significance in photocatalytic wastewater treatment, which could potentially optimize microalgae growth. Adequate aeration and efficient mixing of the culture medium is also critical, supplying oxygen and nutrients to the microalgae and preventing sedimentation. Furthermore, the potential benefits of each system can be maximized while minimizing negative effects by implementing two-stage culture systems in both closed and open systems [16].

This study examines the potential of using wastewater streams from agricultural processes to cultivate microalgae species, specifically Desmodesmus sp., in various controlled environments and reactor systems for biostimulant production. This process is tested in different settings, including laboratory-scale Duran flasks and open-pond, containerized, and photo bioreactor systems under a range of conditions, such as light intensity and duration, temperature, aeration, and nutrient composition. Additionally, this research examines the utilization of a complex biocatalysis process to transform microalgae biomass into high-quality biostimulants. Furthermore, it explores the scalable application of these methods, examining the possibilities and constraints of producing microalgae biomass and subsequent biostimulant on a large scale. The study evaluates the potential of these methods as sustainable, cost-effective [9], and environmentally friendly solutions for agricultural waste management and biostimulant production.

As shown in Figure 1, the plant follows a circular economy approach by gathering rich-in-nutrients wastewater streams from hydroponic greenhouses in an innovative existing automation recycling system, which can transform the fabricated microalgae biomass into a biostimulant formulation based on a novel enzymatic hydrolysis procedure.

The aim of the present study is to present an innovative agro-wastewater management system, based on the circular economy of resources, with enormous environmental and economic potential. The main short-term impacts of the PBR plant include the creation of a circular economy model compatible with the agri-farm wastewater purification process and the production of an algae biomass purification process, allowing operators and managers to analyze the potential of such technologies, raising the awareness of the agricultural sector with regard to environmental impacts while transferring the concept of waste equals resource.

2. Materials and Methods

2.1. Development of an Automated Agro-Wastewater Treatment Plant

The agro-wastewater treatment plant is a containerized and automatically controlled system designed by STAM S.r.l. (Genoa, Italy) to treat different types of wastewater using closed photo-bioreactors. Compared to other microalgae cultivation methods, the system design is a versatile system that could also benefit from the use of an artificial lighting system. The design, components, and operational features of the treatment plant are illustrated in Figure 2.

The plant setup consists of two loops of closed PBRs, each with an approximate volume of 950 L, made out of 18 connected AlgaeFit^® poly-methyl methacrylate, highly resistant to photo-oxidation (Dext: 90 mm; Dint: 82 mm; Length: 10,000 mm). The PBRs are connected and then mounted on an AISI 304 stainless steel support structure (skid dimensions: length 10 m; width 1.5 m; height: 2 m). The pipes are connected in a closed system, and the water is pumped by two recirculation pumps and artificially illuminated by two sets of LED lamps (230V 24W NAT INDOOR WA 120°, C-LED S.R.L., Bologna, Italy) and an RB C-LED lamp (TL PLUS 230V 150W PURLPLE EVO, C-LED S.R.L., Bologna, Italy).

To facilitate independent control and synchronized communication during the different stages of algae growth, the system is equipped with dedicated shut-off valves and a recirculation pump. These shut-off valves allow for precise regulation of nutrient supply and gas exchange within each photo-bioreactor. The recirculation tank works as a heat exchanger connected to a special heat pump able to control the process temperature. In addition, the recirculation pump ensures efficient mixing of the algae culture, promoting uniform growth while maximizing productivity. Externally, the system incorporates four storage tanks, namely the Storage Wastewater, Medium Algae, Discharge/Sedimentation Tank, and Cleaning Water tanks, each serving a specific purpose in the treatment process. The indoor temperature of the plant is controlled by an electric coil heating system and a glycol cooling system managed by an air–water heat pump (12 kW Aermec, Rochford, UK) to ensure precise temperature regulation and stability. Additionally, LED artificial lighting is employed to support optimal microalgae growth. Furthermore, dedicated sensors monitor pH and temperature levels, while air supply valves facilitate the introduction of air into the system.

A control system is dedicated to managing the operation of the different components of the system (recirculation pumps, artificial lighting, system loading and unloading, automatic flushing and filling, and the control of solenoid valves). An important and innovative feature of this system is the incorporation of a tank dedicated to biomass conversion into the biostimulant. To ensure precise temperature control throughout the process, a customized profile temperature controller with data logging capabilities is employed. This temperature controller allows for real-time monitoring and adjustment of the temperature in the biostimulant tank, ensuring optimal conditions for biomass conversion. The wastewater treatment process is based on the growth of a microalgae inoculum inserted under optimal concentration conditions into the loop that contains the nutrient solution to be treated. By controlling the microalgae cultivation parameters, i.e., temperature, pH, and artificial lighting, the microalgae grow to the detriment of nitrogen- and phosphorus-based compounds present in the hydroponic nutrient solution to be treated. These compounds form, together with other elements, the nutritional basis for microalgae growth.

Working Phases and Algae Growth Cycles

As a result of the process, the water is partially depurated, and it can be used for irrigation purposes, with concentrations of nitrogen (N) and phosphorus (P) compliant with EU regulations. The integration of a biostimulant tank allows for the efficient utilization of microalgae biomass, maximizing the production of high-value bio-products such as plant enablers. Additionally, the continuous monitoring and adjustment of the system through sensor data ensures consistent and optimal performance, further enhancing the overall productivity and quality of the plant enablers produced. The biostimulant reactor or biostimulant tank has a volume equal to 40/50% of the sedimentation tank, according to the biostimulant production process diagram presented in Figure 3. The biostimulant tank has a stirring system, and it is resistant to physical and chemical impacts such as temperature and pH variations. In this study, the agro-wastewater plant was tested for the production of biostimulants, utilizing the wet biomass of the Desmodesmus sp. strain. This microalgae species was selected based in previous studies [17].

2.2. Optimizing Sedimentation Practices and Enzyme Complexes for Biostimulant Production

This study delves into the cultivation of Desmodesmus sp. microalgae for biostimulant production using advanced columnar vertical PBR systems. Desmodesmus sp., a freshwater microalgae species, was chosen for its high efficiency in producing crop biostimulants. The cultivation process began with laboratory-scale bottles and extended to vertical columnar PBRs, which are sophisticated closed vertical glass columns designed to optimize light exposure and growth conditions. To enhance biomass concentration, flocculation-coagulation techniques were employed, effectively aggregating the microalgae cells for easier harvesting. Additionally, the study focused on the development of enzymatic complexes crucial for the biocatalytic conversion of biomass into biostimulants. Three different enzymatic complexes were examined for their efficiency and effectiveness in this process.

2.2.1. Microalgae Cultivation and Pre-Concentration of Microalgae Biomass for Lab Biocatalysis

Desmodesmus sp. was selected as the microalgae species under investigation for its ability to produce crop biostimulants [18,19]. The strain was obtained from STAM S.r.l. (Genoa, Italy) and was stored in the facilities of Neoalgae Micro Seaweed Products (Gijón, Spain). Desmodesmus sp. was used to inoculate glass flasks (1–2 L) under controlled conditions, with a photoperiod set at 16:8 h (light–dark). The light was provided through lamps with a continuous photon flux of 80–100 μmol·s⁻¹·m⁻². Moreover, the temperature was also maintained constant at 25 °C while agitation was provided using aeration through air pumps with a flow rate of 100 L h⁻¹. This method ensured homogeneity and facilitated gas exchange within the system. The culture was then scaled up in 10 L flasks inside a greenhouse with uncontrolled conditions. Finally, the culture was transferred to the last station of the production scale, which was a columnar vertical PBR system. This industrial production PBR system consisted of closed vertical glass columns of 3 m height and 30 cm width, with a maximum volume of 100 L. Biomass separation was tested based on the flocculation-coagulation procedure to develop a standard pre-concentration protocol. For this reason, two different flocculants were added to Desmodesmus sp. cultures of 1 L in different concentrations. In particular, chitosan and Iron (III) chloride (FeCl₃) were added to the microalgae culture in two (60 and 70 mg L⁻¹) and three (50, 75, and 150 mg L⁻¹) different doses, respectively. To ensure the economic viability of the process, a limit was set on the amount of reagent used, focusing on the flocculant to produce an effective biostimulant. The efficiency of the flocculants was assessed by measuring sedimentation speed and biomass recovery.

2.2.2. Biocatalysis Process under Laboratory Conditions

Selection of the Optimum Enzyme Complex for the Biocatalysis Process

Three enzymatic complexes were developed by Neoalgae Micro Seaweed Products (Gijón, Spain), leveraging the expertise of the company and insights from previous studies [20,21,22]. The complex “EnzMix” is a mixture of cellulases and proteases, the complex “EnzCell” is composed mainly of cellulases, while the complex “EnzProt” mainly consists of proteases. The selection of these enzymes was based on their easy-to-operate nature, which was considered essential for their low requirements for optimum activity.

“EnzMix”, “EnzCell”, and “EnzProt” were added to 200 mL of culture at a ratio of 0.5% over the total volume of the assay and were exposed to agitation for 24 h to ensure an optimum breakdown of the cellular walls. Adjustments to the pH and temperature were made in accordance with the optimal operating conditions for each enzyme complex. Afterward, a visual measurement was performed through optical microscopy to evaluate the cellular degradation of the hydrolyzed sample. Moreover, samples were subjected to analysis of the chlorophylls present at 680 nm of optical density (OD680) under the assumption that broken cell walls cause photosynthetic systems to degrade while degrading the chlorophyll.

Selection of the Optimum Dose and Conditions of the Enzymatic Complex for the Biocatalysis Process

Based on the results, further steps of the experimental design were focused on the mixed activity (cellulase and protease) of the enzymatic complex “EnzMix”. The next stages of the experimental setup were focused on the optimization of the dosage and conditions of the enzymatic complex. “EnzMix” is an easy-operating enzymatic complex, with no temperature control or pH modifications needed during the biocatalysis process. The enzyme concentration was optimized in relation to the dry weight measurements of the Desmodesmus sp., since the enzymatic activity is improved along with the relation among enzyme molecules and substrate molecules. In addition, a sample from the cultures at the exponential-phase stage was sampled, with an initial dry weight content of 1.08 g L⁻¹. Moreover, to assess the optimum enzyme concentration, four increasing concentrations (1%, 4%, 8%, and 10%) of the selected enzymatic complex were tested and compared to a control treatment that did not include any enzyme. These doses were selected to ensure both the environmental and economic viability of the process, making the practical application of these enzymes feasible.

2.3. Desmodesmus sp. Cultivation for Field Biostimulant Production

2.3.1. Desmodesmus sp. Cultivation at Laboratory Scale for On-Site Biostimulant Production

A microalgae culture of the Desmodesmus sp. strain was established in the Laboratory of Agricultural Constructions and Environmental Control of the University of Thessaly, Greece, using a stock inoculum provided by Neoalgae Micro Seaweed Products (Gijón, Spain). The initial inoculum volume (1 L) was divided into five Duran flasks with a capacity of 2 L each. The cultivation conditions were as follows: illumination intensity: 80,000 lux, RGB lights, and 20,000 lux day lights; temperature: 27.0 ± 1.0 °C; illumination duration: 16:8 h (light–dark). The culture was maintained for 14 days in the flasks to allow proper biomass growth and subsequently enabling the inoculation of 20 additional flasks with a total culture volume of 40 L. Moreover, agro-wastewaters, i.e., hydroponic drainages collected from a soilless tomato greenhouse crop, were used as a growth medium for the microalgae culture. The nutrient composition of the drainages was 14.7 mmol NO₃⁻ L⁻¹, 0.02 mmol NH₄⁺ L⁻¹, 0.64 mmol PO₄³⁻ L⁻¹, 4.19 mmol K L⁻¹, and 4.91 mmol Ca²⁺ L⁻¹.

In order to meet the nutritional requirements essential for the growth of microalgae, the deficient nutrients present in the agro-wastewaters were supplemented using the BG-11 medium formulation (

Table 1. Macronutrient and trace metal compounds (mM) in the BG-11 medium recipe.

Macronutrient Compound	Concentration (mM)
NaNO3	17.60
K2HPO4	0.23
MgSO4·7H2O	0.30
CaCl2·2H2O	0.24
Citric Acid·H2O	0.03
Ferric Ammonium Citrate	0.02
Na2EDTA·2H2O	0.003
Na2CO3	0.19
Sodium Thiosulfate Pentahydrate	1.00
BG-11 Trace metals	Concentration (mM)
H3BO3	46.00
MnCl2·4H2O	9.00
ZnSO4·7H2O	0.77
Na2MoO4·2H2O	1.60
CuSO4·5H2O	0.30
Co(NO3)2·6H2O	0.17

), which was further adjusted according to the drainage analysis to ensure adequacy. Both the BG-11 medium and the nutrient solutions contained within the Duran flasks were subjected to sterilization using an autoclave sterilizer at 121 °C for 20 min prior to utilization. This sterilization process was employed to eliminate any potential microbial contaminants, thereby ensuring aseptic conditions for the cultivation of microalgae.

To increase volume and, therefore, microalgae biomass production, an open-pond bioreactor of 1.18 m² (diameter of 123 cm and height of 25.5 cm) surface area was established, equipped with mechanical stirring and a pump aeration system. The culture obtained from the Duran flasks was used to inoculate the open-pond bioreactor with an initial volume of 200 L. The culture lasted 51 days in total. However, due to the absence of significant growth observed during the first five experimental days, a total volume of 80 L was removed from the culture to reduce the initial 200 L volume and enhance the photosynthetic rate. This adjustment was necessary due to the large inoculum volume in relation to the available surface area of the open pond. In addition to volume reduction, NaHCO₃ was added to the culture at a concentration of 0.5 g L⁻¹ to provide a carbon boost.

During the experimental period, culture contamination by parasitic protozoa and rotifers was observed in the open-pond bioreactor system. To mitigate this issue, daily pH treatments were implemented. The pH value was adjusted to 4.0 using HCl (18.5% a.r.) for 10 min and then to 7.0 using NaOH (40%). This adjustment ensured the survival and growth of the culture without complications. Upon achieving a microalgae dry biomass of 1.425 g L⁻¹ on Day 51, the biomass was used to feed the biostimulant tank for the production of the 1st biostimulant batch (batch A), with a total volume of 100 L.

2.3.2. Desmodesmus sp. Cultivation at Container Scale for On-Site Biostimulant Production

To produce sufficient biomass to inoculate the container (STAM S.r.l, Genoa, Italy) with a loop capacity of 950 L, a new culture (100 L) was cultivated in the open-pond bioreactor installed at the premises of the University of Thessaly, following the same methodology used in the Desmodesmus sp. production under lab-scale conditions. The culture in the open-pond bioreactor lasted 19 days. After reaching a certain value of optical density and dry weight, the inoculum was transferred and scaled up from the open-pond to the closed (container) bioreactor scale.

Upon achieving a microalgae of a certain dry biomass on Day 34 in the PBR, the wet biomass was collected after being left for 24 h in the sedimentation tank and used to feed the biostimulant tank for the production of the 2nd biostimulant batch (batch B). In this experimental setup, the efficiency of the previously investigated flocculant FeCl₃ was also assessed at the container scale. Therefore, on Day 35, an additional volume of 500 L of the inoculum was transferred to the sedimentation tank following the protocol used in all other cases, but in this case, FeCl₃ was added to the tank to speed up the sedimentation process. In particular, 0.075 g L⁻¹ of FeCl₃ was added to the inoculum. Then, the wet microalgae biomass precipitated in the base of the conical sedimentation tank was collected and transferred to the biostimulant tank for the 3rd biocatalysis process (batch C). However, it is imperative to note that scaling up the inoculum from the open pond to the PBR did not yield the expected results. Both the optical density and the dry weight of microalgae did not exhibit a significant increase over time.

2.3.3. Desmodesmus sp. Production at Laboratory Scale for On-Site Biostimulant Production

Following the Desmodesmus sp. cultivation at the container scale for the field biostimulant production trial, approximately 100 L of the culture developed inside the container was transferred from the container loop to the open-pond bioreactor in the laboratory for further growth. This transfer aimed to investigate the factors contributing to the lack of growth in the container culture. It was hypothesized that lighting conditions, specifically the duration and intensity of daylight, as well as the lack of carbon dioxide (CO₂) played a crucial role in biomass growth. In the laboratory, a fixed photoperiod of 16:8 h (light–dark) was implemented. The culture followed the same protocol according to the previously established process and was transferred to the biocatalysis tank on Day 20 for a further biocatalysis process (batch D).

2.3.4. Desmodesmus sp. Production at Container Scale through CO₂ Infusion and Increased Lighting for On-Site Biostimulant Production

Firstly, 140 L of inoculum was initially produced in the Laboratory of Agricultural Constructions and Environmental Control at the University of Thessaly, employing the open-pond bioreactor and adhering to the same methodology employed for Desmodesmus sp. production under laboratory conditions. Upon attaining a substantial biomass concentration, ten Duran flasks, each with a capacity of 2 L and collectively amounting to 20 L, were employed for inoculation. Then, the inoculum of a total volume of 140 L was transferred to the PBR system (Figure 4).

The culture medium within the container was meticulously adjusted to align with the desired nutrient concentrations, specifically nitrate (NO₃⁻), potassium (K), phosphate (PO₄³⁻), and calcium (Ca²⁺), in accordance with the drainage characteristics. To achieve the targeted nutrient concentrations essential for the culture, supplementation was carried out for the remaining nutrients following the BG-11 recipe (Table 1). To optimize light intensity, all lights from both loops were incorporated into a singular loop, with only one of the two loops designated for the experiment. Furthermore, LED strip lights were also adjusted into the system to remedy the previously inadequately illuminated back side, thereby improving the overall distribution of light throughout the environment. The cumulative light intensity from both white and RGB lights reached 1320 uW/cm². The photoperiod for microalgae was established at a ratio of 16 h of light to 8 h of darkness (16:8). On Day 12 of the cultivation period, CO₂ was introduced into the system. This strategic addition serves a dual purpose: firstly, to supply an essential carbon boost that promotes robust growth, and secondly, to regulate the pH level, ensuring optimal conditions for the thriving microalgae culture. Daily and for a fixed duration, the culture received a controlled supply of CO₂. At the same time, continuous monitoring of the pH of the culture was carried out, given the inherent ability of CO₂ to cause a decrease in the pH. Notably, during the growth phase, there was a distinct tendency for pH to exhibit an upward trajectory. In response to this phenomenon, CO₂ supply assumed a central role in moderating the observed upward trend, thus serving as an essential countermeasure during the growth phase. During this trial, two more batches were produced, namely batch E and batch F, on Day 26 and Day 40, respectively.

2.3.5. Measurements

The content of nitrogen in the culture medium was measured using the HACH analytics (HQ40D Portable MultiMeter, Manchester, UK). Optical density (OD) values were measured using a spectrophotometer (DR3900 Spectrophotometer, HACH, Manchester, UK). To measure the dry weight of the culture, a centrifuge (Universal 320R Hettich, Tuttlingen, Germany) was used to separate the biomass from the rest of the aqueous nutrient solution, followed by a drying process where the collected biomass was oven-dried at 70 °C (Raypa^®, Barcelona, Spain), and then weighted using a high-accuracy weight scale (Adam Equipment, Oxford, CT, USA).

2.3.6. Biocatalysis Process

The following protocol was used to perform the biocatalysis process: At first, the Desmodesmus sp. culture was introduced directly into the biostimulant tank after being pre-concentrated in the sedimentation tank for at least 24 h. From this point onwards, continuous agitation was employed until the completion of the hydrolysis process. The temperature was raised to 50 °C, followed by the addition of citric acid to establish an optimum culture pH within the range of 6.0–6.5. Subsequently, the enzyme complex “EnzMix” was introduced into the biostimulant tank and left to incubate overnight for 24 h. In cases where microalgae were cultivated without the supplementation of CO₂, the enzymes were applied at a concentration of 10% relative to the biomass within the culture. Conversely, in the scenarios involving CO₂ supplementation, denoted by batches “E” and “F”, the concentration of “EnzMix” was increased to 15%. The final product was stored in a shaded area with an average air temperature of approximately 22 °C. Moreover, an additional step was included in the process during the production of batches “C”, “E”, and “F”. In particular, the culture was first transferred to the sedimentation tank, where 150 g L⁻¹ FeCl₃ was added to expedite the sedimentation process. The resulting sediment was then transferred to the biostimulant tank following the biocatalysis protocol as referenced above. In addition, after the completion of the biocatalysis, the pH was lowered to 3.80, and the final biostimulant was stored for future use.

3. Results

3.1. Determining the Best Sedimentation Method and Enzyme Complex for Biostimulant Production

3.1.1. Pre-Concentration of Microalgae Biomass

The selected strain Desmodesmus sp. exhibited different behavioral patterns depending on both the flocculant type and the final concentration added to the culture. As shown in Figure 5, cultures were added to different glass graduated cylinders and mixed with the two flocculants under investigation.

Based on the results, both chitosan and FeCl₃ flocculants presented promising results regarding sedimentation. However, the doses were definitive in identifying FeCl₃ as the most suitable substance at the maximum concentration (150 mg L⁻¹) in terms of sedimentation speed (9.9 cm min⁻¹) and biomass recovery (12.9 g L⁻¹) of wet and pre-concentrated biomass, as compared to the rest of the tested doses (75 and 50 mg L⁻¹). The biomass recovery results obtained from the addition of chitosan were relatively good, featuring suitable biomass recovery, with significant differences found after the comparison with the results obtained from the FeCl₃ samples. However, in the case of the chitosan flocculant, lower sedimentation speed was recorded (5.1 cm min⁻¹ at 70 mg L⁻¹) (

Table 2. Sedimentation speed (cm min⁻¹) and biomass recovery (g L⁻¹) results after adding two flocculants (chitosan and FeCl₃) to Desmodesmus sp. culture at different doses (mg L⁻¹).

Flocculant Name	Added Dose of Flocculant (mg L−1)	Sedimentation Speed of Flocculant (cm min−1)	Biomass Recovery from Culture (g L−1)
FeCl3	50.0	8.7	10.1
	75.0	9.5	11.8
	150.0	9.9	12.9
Chitosan	60.0	4.0	11.5
	70.0	5.1	10.1

).

3.1.2. Biocatalysis Process under Laboratory Conditions

Selection of the Optimum Enzyme Complex for the Biocatalysis Process

Samples of Desmodesmus sp. culture were subjected to each enzymatic complex and agitated for 24 h. A control sample was also agitated for the same time without the addition of any enzymes. After 24 h, absorbance measurements performed on the control treatment showed an OD680 of approximately 1.1, consistent with the values recorded prior to exposure to the agitation process. Moreover, the microscopic analysis also demonstrated the integrity of the Desmodesmus sp. cells after 24 h of agitation. On this basis, it was assumed that all of the variations in chlorophyll content detected through the spectrophotometric analysis and the visual anomalies found under microscopic analysis were due to the hydrolytic activity of the enzymatic complexes. Furthermore, microscopy studies revealed the integrity of the cell walls and the bright green color of the observed cells, showcasing the presence of intact cells.

As shown in Figure 6 and in accordance with the results obtained by the tested enzyme “EnzProt”, a slight but not significant reduction was recorded in chlorophyll concentration. “EnzProt” did not produce a variation in the value measured, while the microscopic analysis showed how “EnzProt” cells presented a complete cell wall, with no visible ruptures. On the same basis, a colorimetric analysis showed that these cells possessed a bright green color, similar to what was observed in the control treatment. This result can be attributed to the protease activity of this enzymatic complex, which cannot degrade any intern protein due to the inability to penetrate through the cellular walls, especially in the absence of wall-breaking activity.

On the other hand, the “EnzCell” assay showed clearly a decrease in the chlorophyll concentration, as observed in the OD680 analysis. This reduction was related to the cellulase nature of the enzyme complex, resulting in a reduction of chlorophyll concentration by 30.8% compared to the control treatment results. The cell walls of microalgae are mainly composed of cellulose, and therefore, the high rate of broken walls by the cellulase-type enzymatic complex is an expected result. Moreover, the microscopic analysis showed that Desmodesmus sp. cells partially lost the bright green color compared to that observed on cells of the control treatment. Loss of the color was not accompanied by visible degradation of the cell walls. However, the colorimetric variation observed is correlated with the degradation measured during absorbance studies. Based on the obtained results, ”EnzCell” is demonstrated to be an efficiency enzyme for breaking down the cell walls of Desmodesmus sp. and thus should considered valuable for specific studies.

On the same basis, the “EnzMix” assay demonstrated a clear decrease in the concentration of the present chlorophylls, resulting in a maximum reduction of chlorophylls by 40.7% compared to the control treatment measurements. Moreover, this descent was accompanied by a clear rupture of the cell walls of the microscopically analyzed samples. Figure 6 clearly depicts the colorimetric variation among cells of the “EnzMix” and the control treatment, highlighting the loss of the bright green characteristic color, as well as a clear thinning of the cell wall of the former complex.

Selection of the Optimum Dose and Conditions of the Enzymatic Complex for the Biocatalysis Process

As shown in Figure 7, the initial chlorophyll measurement in terms of absorbance conducted on the control treatment sample was 0.918 at a maximum OD680. Moreover, the addition of 1% of the enzymatic complex “EnzMix” resulted in a slight, albeit not significantly important, increase in the absorbance level. This outcome could be attributed to the natural deviation of the selected measuring method rather than the influence of the hydrolytic activity on the culture sample. On the other hand, the addition of 4% of enzymatic complex “EnzMix” resulted in a non-statistically significant decrease in the OD680 values compared to the control treatment. However, the addition of 8% of the ”EnzMix” complex did not follow a similar trend to the other percentages tested, since its application resulted in a significant reduction in chlorophyll values, possibly due to the optimal interaction among substrate and enzymatic molecules. Finally, the addition of 10% of “EnzMix” to the Desmodesmus sp. culture also resulted in a clear, albeit slightly greater, reduction compared to that observed in the case of the 8% addition. Hence, the optimal concentration of the enzymatic complex “EnzMix” for the most effective biocatalysis process was found to be 10%, correlating with the biomass present. The reduction in chlorophyll absorbance readings was significantly higher when “EnzMix” was applied to Desmodesmus sp. culture at 10% concentration.

3.2. Desmodesmus sp. Production Using Agro-WasteWaters for Field Biostimulant Production

A series of experiments was conducted at both laboratory and container scales using agro-wastewaters for field biostimulant production. These experiments were based on previous studies regarding the selection of the most suitable enzyme complex and its optimal dose, which was determined to be “EnzMix” at a concentration of 10%.

3.2.1. Desmodesmus sp. Production at Lab Scale for On-Site Biostimulant Production

The optical density measurement, specifically at a wavelength of 750 nm, was used to assess the efficiency of microalgae biomass growth, as it correlates with the absorption of visible radiation (with the chlorophyll absorption peak at 680 nm). As shown in Figure 8a, the optical density of Desmodesmus sp. grown in laboratory conditions at 680 and 750 nm exhibited an increasing trend until Day 50 of the culture, resulting in 3.21, and 2.46 OD values, respectively. This increase in OD was followed by an increase in the dry biomass weight up to 1.42 g L⁻¹, showcasing a significant biomass productivity of Desmodesmus sp. during the 50-day culture period (Figure 8b).

Nitrogen content measurements (NO₃⁻-N) in the culture medium were associated with the rate of biomass growth, with higher NO₃⁻-N consumption corresponding to increased culture growth. As shown in Figure 8c, NO₃⁻-N levels decreased as microalgae biomass increased. In particular, NO₃⁻-N content in the last stage of the culture (Day 50) was reduced by 40% as compared to the initial NO₃⁻-N concentration found in the culture at Day 0.

3.2.2. Desmodesmus sp. Production at Container Scale for On-Site Biostimulant Production

During the second experimental trial, the Desmodesmus sp. culture was first grown in the open-pond system for a total of 19 days, to increase both the optical density and the dry biomass before inoculating the container. The initial optical density of the culture on Day 0 was 1.545 at 750 nm, while the dry weight was 0.842 g L⁻¹. As presented in Figure 9a, a substantial increase in optical density was initially recorded, which continued until experimental Day 19, reaching a value of 3.25 and 2.55 at 680 and 750 nm, respectively. Similarly to the optical density increase, a corresponding increase in dry microalgae biomass productivity was also observed during the 19-day open-pond experiment, reaching a value of 1.413 g L⁻¹ (Figure 9b).

After reaching the aforementioned values, 100 L of the culture was transferred inside the container loop to produce higher microalgae biomass using drainages of a hydroponic crop as a growth medium to be further used as a biostimulant. The culture was then monitored inside the loop for 35 days. However, the culture within the container did not present any significant increase in either the optical density or the dry microalgae biomass. Both optical density (Figure 10a) and dry biomass (Figure 10b) decreased, despite some intermittent days where an increase in the dry weight was recorded. The total dry biomass decreased from 0.842 g L⁻¹ to 0.184 g L⁻¹, followed by a decrease in OD750 from 0.516 to 0.326.

This phenomenon is likely to occur due to suboptimal conditions prevailing in the container, such as the low lighting, temperature, and lack of CO₂ supply, which degraded the culture’s continued growth, although the cells remained viable. As a result, in this case study, the biostimulant production was performed using a microalgae biomass of significantly lower productivity compared to that of the first case study, where the dry microalgae biomass used for the biocatalysis process was 87% higher.

3.2.3. Desmodesmus sp. Production at Lab Scale for On-Site Biostimulant Production

Since the microalgae remained viable despite their inability to grow under the container scale experiment, the culture was collected and then transferred to the same open-pond system that was originally grown under lab conditions for further testing. As depicted in Figure 11, an increase in optical density coincided with the culture’s growth. The optical density of the microalgae culture in the open-pond system increased from 0.463 to 2.856 at 750 nm, whilst the dry weight value increased from 0.263 g L⁻¹ to 1.392 g L⁻¹.

3.2.4. Desmodesmus sp. Production at Container Scale through CO₂ Infusion and Increased Lighting for On-Site Biostimulant Production

The inoculum culture of Desmodesmus sp. was initiated in the open-pond system, spanning from 21 September 23 (Day 0) to 11 October 2023 (Day 20). The open-pond bioreactor culture initiated on Day 0 exhibited an initial optical density of 0.491 (at 750 nm) and a dry weight of 0.312 g L⁻¹ when reaching 120 L. The optical density was systematically measured daily at two distinct wavelengths, namely 750 nm and 680 nm. Figure 12a depicts the optical density trends, while Figure 12b illustrates the corresponding dry weight measurements of the open-pond culture. The optical density within the open pond exhibited a consistent and gradual increase, culminating in a peak value of 2.05, with the dry weight reaching 1.143 g L⁻¹ by Day 20. This observation is indicative of robust cellular proliferation within the open-pond system. Upon attaining a substantial biomass concentration, ten Duran flasks, each with a capacity of 2 L and collectively amounting to 20 L, were employed for inoculation. The inoculum, characterized by an OD of 3.25 (at 750 nm) and a corresponding dry weight of 1.715 g L⁻¹, was utilized to feed a single loop of the container bioreactor.

The inoculation of the container started with an initial optical density of 0.298 (at 750 nm) and a dry weight of 0.177 g L⁻¹, since the 140 L produced in the lab was diluted with drainages of soilless crops serving as a growth medium for microalgae, leading to a significant decrease in OD and DW. As shown in

Table 3. Nutrient savings (%) for the Desmodesmus sp. culture when using hydroponic drainages.

Macronutrients	BG-11 (mmol L−1)	Drainage (mmol L−1)	Nutrient Saving (%)
NO3−	12.9	9.5	73.6
K	0.05	3.81	100.0
PO43−	0.13	0.76	100.0
Ca2+	0.09	4.73	100.0

, significant savings in nutrient concentrations were realized, exemplified by an impressive 74% reduction in NO₃⁻. In addition, complete elimination of the need for supplementation of K, PO₄³⁻, and Ca²⁺ was observed, reflecting a 100% savings in these essential elements in the culture medium.

Initially, at the onset of the container culture, despite the utilization of all available lights from both loops and additional LED stripes, no discernible growth was observed. On Day 0, the initial optical density of the container culture was recorded at 0.298 when measured at 750 nm, while the dry weight was documented at 0.177 g L⁻¹. As depicted in Figure 13a, a notable absence of a significant increase in OD persisted until Day 12 of the experiment. The optical density values at 680 and 750 nm reached 0.47 and 0.36, respectively, indicating relatively modest progress within this time frame.

Following the introduction of CO₂, a discernible augmentation in dry microalgae biomass productivity was evident up to Day 26 (fifth harvesting period), and this positive trend extended into the subsequent sixth and final harvest. The observed increase highlights the favorable influence of CO₂ supplementation on microalgae cultivation, underscoring its potential to enhance overall productivity over successive harvesting cycles. The results, depicted in Figure 13, outline a steady increase in dry weight over the initial 26 days of cultivation, culminating at 0.971 g L⁻¹ and accompanied by an optical density at 750 nm of 1.604 on Day 26. The harvested biomass was then used as the main feedstock for the biocatalysis tank intended for the production of the fifth biostimulant (Batch E). After reaching the targeted biomass, a total volume of 300 L was collected from the microalgae culture. The harvested volume was then directed to a sedimentation tank, where it underwent a sedimentation process for 24 h. During this settling phase, FeCl₃ was used as a flocculating agent to facilitate the settling of suspended particles, helping to separate the biomass from the liquid phase. This strategy helps to clarify and concentrate the harvested microalgae biomass, enhancing the efficiency of downstream processing for biostimulant production. The biocatalysis process was carried out in a tank with a total volume of 130 L. Subsequently, the loop was replenished with 300 L of drainage from the greenhouse hydroponic production, and crop cultivation continued. Based on the production of 130 L of biostimulant over a span of 26 days, it can be inferred that approximately 5 L of biostimulant is generated per day with the aid of CO₂ supplementation.

The developmental phase spanned a duration of 14 days (Day 40). Upon attaining a dry microalgae biomass concentration of 0.876 g L⁻¹ and an OD750 reading of 1.41, a total volume of 500 L was extracted from the culture. This harvest was subsequently conveyed to the conical sedimentation tank, where it underwent a sedimentation process over a 24-h period. Following the sedimentation phase, 150 L of the harvested material was meticulously transferred to the biocatalysis tank. This marked the initiation of the sixth batch (Batch F) in the production sequence of the biostimulant, resulting in a 10.7 L day⁻¹ biostimulant production.

Throughout the growing season, pH levels were monitored daily. Figure 14 shows the pH dynamics, indicating an initial pH of 7.09, which quickly increased to 11.21. The extensive rise in pH was controlled by applying daily CO₂ injections and lowering the pH to 6.0–6.3 as monitored with a pH meter. In addition, increased CO₂ concentrations showed the potential to lower the pH of the culture medium, leading to a subsequent impact on the activities of critical enzymes that play a role in the photosynthesis process. Strains showed sensitivity to elevated CO₂ levels, and this sensitivity was observed in association with initial low pH levels (around pH 5–6), although it was not the sole determinant. Throughout the active growth phase, pH levels showed an increase of up to 11, gradually decreasing towards the end of the growing season.

The concentrations of various nutrients, namely nitrogen, potassium, and calcium, were measured in the present study. Both calcium and potassium showed a gradual decrease until the first harvest on Day 26 (Figure 15a,b). Subsequently, after the first harvest, there was a noticeable increase in both calcium and potassium levels, especially after CO₂ addition to the loop. This increase is also attributed to the time required to achieve homogeneity after the initial inoculation, which involves incorporating a draining solution into the culture. Over the following days and up to the final harvest, Ca²⁺ and K levels showed a gradual decline until the second inoculation, which included the introduction of 500 L of drainage. The observed dynamics in nutrient concentrations provide valuable insights into the temporal variations and the interaction of nitrogen, calcium, and potassium throughout the experiment.

At the same time, the nitrate consumption profile is graphically depicted in Figure 15c. Notably, a steady decrease in nitrate levels was observed, reaching a minimum value of 13.85 mmol L⁻¹. The observed nitrogen dynamics underscore the vital role of nitrates in supporting the microalgae’s nutritional requirements during their developmental stages. Hence, as shown in Table 3, the initial NO₃⁻ concentration of the nitrates needed as a growth medium of Desmodesmus sp. was 12.9 mmol L⁻¹. The original nitrate content of the inoculum, however, was not taken into consideration for this value; therefore, the initial content in the container was higher (18.3 mmol L⁻¹) than that of the desired content. The nitrate content decreased by 24.2%, resulting in a biostimulant rich in nitrogen.

3.2.5. On-Site Biostimulant Production

A total of six biocatalyses were performed, producing six different biostimulants, which were analyzed for their amino acid concentration (

Table 4. Amino acid (%) profile of the biostimulants produced after six batches in both open-pond and closed bioreactors.

Amino Acids	Batch A	Batch B	Batch C	Batch D	Batch E	Batch F
Alanine	0.049	0.027	0.027	0.027	<0.25	<0.25
Arginine	0.010	0.016	0.015	0.016	<0.25	0.42
Asparagine	0.008	0.055	0.056	0.053	<0.25	<0.25
Aspartic Acid	0.462	0.011	0.011	0.014	<0.25	<0.25
Cysteine	0.012	0.016	0.016	0.017	0.54	0.67
Glutamic acid	0.048	0.042	0.047	0.042	<0.25	0.25
Glutamine	0.019	0.024	0.023	0.024	<0.25	<0.25
Glycine	0.015	0.013	0.013	0.011	0.31	0.53
Histidine	0.022	0.024	0.025	0.029	<0.25	<0.25
Isoleucine	0.010	0.026	0.028	0.024	<0.25	<0.25
Leucine	0.019	0.007	0.008	0.010	<0.25	<0.25
Lysine	0.009	0.017	0.018	0.020	<0.25	<0.25
Methionine	0.012	0.007	0.007	0.009	<0.25	<0.25
Phenylalanine	0.005	0.015	0.015	0.014	<0.25	<0.25
Proline	0.190	0.064	0.066	0.067	0.34	0.29
Serine	0.062	0.076	0.075	0.074	<0.25	<0.25
Threonine	0.146	0.154	0.155	0.153	<0.25	<0.25
TOTAL	1.098	0.594	0.605	0.604	1.190	1.910

). The first was performed using biomass obtained directly from a Desmodesmus sp. culture grown in the laboratory (batch A), the second from biomass obtained at the container scale (batch B), the third from container-scale biomass with FeCl₃ (batch C), and the fourth from biomass which, after a failed container-scale adaptation, was transferred back to the laboratory scale (batch D). Finally, two more batches (batches E and F) were produced using CO₂ as a carbon booster at the container scale.

As shown in Table 4, the total amino acid concentration of the biostimulant batches that entered the container in the final trial, i.e., Batch E and Batch F, presented significantly higher concentrations when compared to all other batches.

Table 5. NPK (%) analysis of the biostimulants produced after six batches, in both open-pond and closed bioreactors.

NPK (%)	Batch A	Batch B	Batch C	Batch D	Batch E	Batch F
N	0.284	0.205	0.234	0.245	0.100	0.100
P	0.462	0.367	0.386	0.399	0.209	0.199
K	0.302	0.245	0.222	0.239	0.140	0.220

presents the concentrations of total nitrogen, phosphorus, and potassium, revealing the potential bioactivity of the developed products as biostimulants. It is, however, crucial to consider both the application method of the biostimulant and the sustainability of the process. Focus was given to creating a process for in situ recycling of wastewater from hydroponic crops, aiming for sustainability and cost-effectiveness while yielding a biostimulant from microalgae biomass.

4. Discussion

The cultivation of microalgae necessitates a considerable water input, comprising 10–20% of the overall production expenditure [23]. Consequently, the integration of algae biomass production with agro-wastewater treatment presents a financially advantageous strategy for the microalgae-based biostimulant sector. Specifically, this integration can reduce the overall costs associated with microalgae cultivation by utilizing agro-wastewater as a nutrient source, thereby minimizing the need for fresh water and additional synthetic nutrients [24]. This approach can lower the production costs of microalgae-based biostimulants, making the process more sustainable and economically viable.

Microalgae present a promising alternative to seaweed, boasting distinct advantages such as rapid growth and adaptability for large-scale cultivation. This can be achieved through sophisticated setups like photobioreactors or open-pond systems, allowing for meticulous control over key variables such as pH, temperature, nutrient composition, and illumination [25,26,27]. However, the endeavor of producing substantial quantities of microalgal biomass for commercial use remains a formidable challenge [25]. Strategies aimed at mitigating the associated costs encompass several approaches: transitioning from batch cultures to semi-continuous ones, exploiting wastewater resources, and implementing media and nutrient recycling [25]. In the present study, the cultivation of microalgae within a closed photobioreactor system was evaluated, utilizing recycled nutrient-rich drainage from soilless crops as a growth medium. The aim of the present study was to present an innovative agro-wastewater management system based on a circular economy of resources by studying the production of an on-site biostimulant while mitigating nutrient leaching and conserving resources. The observed differences in microalgae biomass productivity between the open-pond system and the closed photobioreactor highlight the importance of environmental control and management in cultivation systems. The average daily increment of the dry biomass in the open pond was 0.05 g L⁻¹ and varied from 0.01 to 0.15 g L⁻¹ day⁻¹. This increment was in line with the work of Li et al. [28], who reported a biomass productivity of Desmodesmus sp. microalgae ranging from 0.15 to 0.35 g L⁻¹ day⁻¹ in an open-pond system using a modified BG-11 medium. Similarly, Xia et al. [29], while conducting an outdoor experiment in 5 L bioreactors with Desmodesmus sp., revealed a 66.3 mg L⁻¹ increase of the daily dry biomass when using the BG-11 recipe. In addition, based on the results of a recent study [30], higher growth of Desmodesmus sp. was recorded, reaching 1.95 g L⁻¹ as compared to S. obliquus when cultivated using leachate as the substrate. This consistency suggests that the open pond can achieve reasonable productivity levels under optimal conditions while changing the growth medium from standard solutions (BG-11) to agro-wastewaters. However, in the present study, the lower biomass production is associated with the rotifer and protozoa infestation detected in the open-pond culture on Day 5, which reduced the rapid growth rate of microalgae biomass production but did not inhibit growth completely. The production of microalgae in commercial raceway ponds and other systems is adversely affected by pathogens, including bacteria, fungi, and viruses. Although culture crashes induced by these pathogens and invaders can be mitigated, they cannot be entirely avoided. Effective management of biological contamination relies on stringent hygiene practices, robust quality control measures, and tailored prevention protocols. Chemical control agents are particularly effective, provided they do not compromise the final product, are environmentally acceptable, and remain cost-effective [31]. Despite this challenge, the microalgae continued to grow, albeit at a reduced rate, showcasing their resilience to environmental stressors. Conversely, the closed PBR system, with the introduction of CO₂, exhibited significantly higher biomass increment rates compared to the open-pond trials. This improvement underscores the importance of controlled environments in maximizing microalgae growth. By regulating parameters such as CO₂ concentration, temperature, and light intensity, the closed system provides an ideal setting for microalgae cultivation, leading to enhanced productivity. Adding CO₂ to microalgae cultivation systems can indeed enhance the growth and productivity of microalgae, improving both the quality and quantity of the biomass produced. However, this process also comes with increased costs. The additional expenses stem from the need for equipment to supply and monitor CO₂, as well as the operational costs associated with maintaining the desired CO₂ levels in the cultivation system. However, the utilization of otherwise harmful gases with high CO2 concentrations, such as flue gases, for microalgae cultivation has been proposed as an alternative method to reduce operational costs while achieving adequate microalgae growth [32,33,34]. The mean daily dry biomass increment of 0.07 g L⁻¹ in the closed PBR system not only surpasses that of the open pond but also indicates a faster and more efficient production process. In particular, in a 50-day open-pond trial, the estimated dry biomass produced over 14 days amounts to 339 mg L⁻¹, whereas in a comparable 14-day experiment utilizing CO₂ in the PBR, the yield escalates to 683 mg L⁻¹. This accelerated growth rate translates to quicker turnaround times and increased output, making the closed PBR system a favorable option for microalgae biomass production. Overall, these findings emphasize the significance of environmental factors and cultivation techniques in optimizing microalgae productivity. While open-pond systems offer a cost-effective approach, they are susceptible to external influences, as evidenced by the infestation issue. In contrast, closed systems provide greater control over growth conditions, leading to superior performance and higher-quality biomass output.

Cultivating microalgae using agro-wastewaters derived from extensive soilless cultivation systems with the aim of producing wastewater low in nitrogen and phosphorus presents a dual benefit, as it not only reduces the costs associated with supplementing nutrients in microalgae culture but also contributes to the maintenance of water resources. Nitrate is a crucial nutrient for microalgae growth, serving as a primary nitrogen source. In the open-pond system, where environmental conditions are less controlled, nitrate depletion may occur more rapidly due to factors such as microbial activity, nutrient leaching, and fluctuations in temperature and light intensity. As a result, the rate of nitrate consumption tends to be higher, leading to a more pronounced decrease in nitrate content over time. Conversely, the closed PBR system provides a controlled environment where nutrient levels can be maintained within optimal ranges for microalgae growth. By regulating factors such as light, temperature, and CO₂ availability, as well as implementing efficient nutrient cycling mechanisms, such as recirculating water and maintaining balanced nutrient input, the PBR system minimizes nutrient losses and enhances nutrient uptake efficiency by the microalgae. In the present study, an average consumption of 5.7 mg L⁻¹ day⁻¹ NO₃⁻-N was recorded in one day of open-pond culture, with this value varying between 1.2 and 10.0 mg L⁻¹ day⁻¹, that is, 3.5 to 30.0 mg L⁻¹ NO₃⁻-N in three days. In the case of the culture grown in the PBR system, a lower decrease in nitrate content (2.5 mg L⁻¹ day⁻¹ NO₃⁻-N) was observed throughout the experimental period. The lower decrease in nitrate content (2.5 mg L⁻¹ day⁻¹ NO₃⁻-N) observed in the closed PBR system throughout the experimental period indicates that nitrate is being utilized more efficiently by the microalgae, resulting in a slower rate of depletion. This efficient nutrient utilization contributes to sustained growth and productivity, ultimately leading to higher biomass yields and a more stable cultivation process. Similarly to the results of the present study, Samori et al. [35] showed that a Desmodesmus communis culture was reported to consume 22.49 mg L⁻¹ NO₃⁻-N in 3 days when using effluent generated by a local wastewater reclamation facility as a growth medium. The strain exhibited an average consumption rate of 30.35 mg L⁻¹ day⁻¹ of NO₃⁻, with peak consumption recorded on Day 22 at 80.6 mg L⁻¹ day⁻¹. Similarly to the work of Wang and Lan [36], who studied the removal of nitrates in an N. oleoabundans culture when exposed to wastewater streams at an initial concentration of 218 mg N-NO₃⁻ L⁻¹, this study reveals that the total nitrate consumption throughout the experimental season did not reach a near-zero level.

Moreover, in a recent study [37], the growth performance of Desmodesmus sp. was investigated under varying CO₂ concentrations (0.03%, 5%, and 10% v/v) within a small-scale photobioreactor loop. The results of our study revealed a prolonged lag phase of approximately 20 days prior to CO₂ introduction into the loop, contrasting with the brief lag phase of 1 to 3 days reported in the study by Anand et al. [37]. Despite this disparity, both studies showcased a sustained increase in microalgae growth during the exponential phase over 14 days following the introduction of CO₂. Remarkably, these findings corroborate previous studies [38,39], further underlining the consistent influence of CO₂ concentration on microalgae biomass productivity. This growth trend, observed in both dry biomass and optical density, was attributed to optimal nutrient levels conducive to cell proliferation within the reactor. However, the differences between the two studies can be attributed to the significantly lower volume of the PBR loop used in our study, approximately 1000 L compared to the 34 L volume used by Anand et al. [37]. Consequently, the extended lag phase observed in our study prior to CO₂ introduction is primarily attributed to the larger volume of the PBR, requiring additional time for proper solution mixing during inoculation, resulting in lower initial dry biomass levels.

FeCl₃ and chitosan were also evaluated for their ability to increase the sedimentation of the strain Scenedesmus sp. in a recent study [40]. More specifically, they found that the flocculation efficiency of FeCl₃ increased by 45% when increasing the application dosage from 0.1 to 0.15 g L⁻¹. In addition, several studies [40,41] have shown a linear correlation between the required dose and microalgae biomass concentration, highlighting an increase in dose as microalgae biomass increases. This is likely to occur due to the flocculation mechanism, as an increase in the amount of suspended algal cells is recorded with increasing biomass, making the application of higher concentration flocculants necessary. The findings of this study are supported by Guillaume et al. [42], who also investigated enzyme treatments on plant cells. Both studies investigate the impact of enzyme treatments on chlorophyll concentration and cell wall integrity in plant cells. In the present study, utilizing the “EnzMix” assay resulted in a significant decrease in chlorophyll concentration, with a maximum reduction of 40.7% compared to control treatments. This reduction correlates with a noticeable rupture in the cell walls of examined samples. Similarly, Guillaume et al.’s [42] study reveals a synergistic effect between protease and cellulase enzymes, resulting in enhanced biocatalytic activity compared to when these enzymes are used individually. Both studies underscore the potential of enzyme treatments in altering chlorophyll concentration and cell wall structure, with implications for various applications in biotechnology and agriculture. The current study’s findings align with those of Álvarez et al. [17], who found that a 10% concentration of the “EnzMix” complex is optimal for efficient degradation of cell walls. This suggests consistency in the efficacy of the enzyme complex across different research investigations, reinforcing its potential as a reliable tool for cell wall degradation processes.

A key aspect of the approach of the present work is the elimination of the centrifugation step, which reduces both the final cost and energy consumption. Consequently, the biostimulant process, utilizing enzymatic hydrolysis, commences with a lower biomass concentration compared to conventional methods. This results in a slightly lower amino acid concentration (0.6–2%) compared to other commercial biostimulant products (>3%). However, this disparity is insignificant considering that commercial biostimulants typically require significant dilution (6 to 8 times) during application. In contrast, the produced biostimulant either requires minimal dilution or can be diluted at a much lower rate. Closed bioreactors with hydroponic drainages represent a sustainable and efficient approach to producing microalgae-based biostimulants. By harnessing the benefits of controlled cultivation and nutrient recycling, these systems offer potent biostimulants with consistent efficacy while minimizing environmental impact. Embracing such innovative technologies is crucial for advancing sustainable agriculture and meeting the challenges of global food security.

5. Conclusions

This work elucidates the intricate dynamics influencing algae growth and productivity, emphasizing the pivotal roles played by species selection, nutrient management, and innovative photobioreactor techniques. Utilizing circular systemic solutions in agriculture, such as repurposing agro-wastewater for algae cultivation, holds promise for addressing environmental challenges while maximizing resource utilization. Reusing drainage from soilless crops to grow microalgae presents a promising opportunity to address multiple environmental challenges while harnessing valuable resources. By recycling nutrient-rich drainage water, water conservation is promoted, and the reliance on synthetic fertilizers is reduced, mitigating nutrient runoff and eutrophication. Additionally, microalgae cultivation can serve as a sustainable wastewater treatment method, effectively removing pollutants and improving water quality.

This study emphasizes the introduction of an automated self-controlled system for on-site biostimulant production through microalgae valorization. The scalability of this approach, demonstrated from laboratory conditions to industrial-scale fabrication, underscores its efficacy, facilitated by enzymatic complexes. Among the parameters evaluated, FeCl₃ emerged as the most effective flocculant in terms of sedimentation and biomass recovery, demonstrating its potential as a viable option in algal culture systems. In addition, EnzMix 10% demonstrated remarkable efficacy in reducing chlorophyll concentration and cell wall rupture. In the context of microalgae growth, the incorporation of a closed tubular PBR, particularly with CO₂ supplementation, showed superior performance. This configuration not only facilitated enhanced microalgae growth but also resulted in significant resource savings, with a remarkable 74% reduction in nitrate levels and full retention of K, P, and Ca. These findings highlight the importance of tailored strategies to optimize algal cultivation processes. The identified optimal conditions hold significant promise for enhancing both the efficiency and sustainability of microalgae-based systems, paving the way for future developments in this field.