Microalgae settling agent and use thereof

Abstract

The present application belongs to a kind of microalgae sedimentation technology, more particularly to a kind of microalgae sedimentation agent and its application.The present application specifically discloses a kind of sedimentation agent and its composition, the sedimentation agent contains planarian body and / or planarian extract, the sedimentation agent is co-cultured with the microalgae cell of appropriate density, so that microalgae is naturally settled;The planarian extract is obtained by filtering the solution of planarian cultivation.The sedimentation agent and its composition of the present application are natural sedimentation agents for dunaliella salina, and do not introduce impurities and toxic substances;Therefore, the amount of pheromone used in the present application is very small, the efficiency of coagulation is high, the chemical interference is small, and as a natural sedimentation agent, there is no toxicity.

Description

Technical Field

[0001] This invention relates to a sedimentation technology for microalgae, and more particularly to a sedimentation agent for microalgae and its sedimentation method. Background Technology

[0002] Microalgae are single-celled algae widely distributed on land and in the ocean. They are autotrophic plants with high photosynthetic efficiency, short growth cycles, and strong adaptability to their environment. Microalgae not only possess rich nutritional value but are also a rich resource of high-value chemicals and pharmaceuticals. Their cells contain components difficult to obtain from other organisms, such as unsaturated fatty acids, astaxanthin, β-carotene, and various bioactive substances, giving them both nutritional and medicinal value. Currently, polyunsaturated fatty acids produced from microalgae, such as gamma-linolenic acid, EPA, and DHA (brain gold), can prevent and treat cardiovascular diseases and cancer, and regulate the function of the central nervous system and visual system. The polysaccharides, proteins, pigments, and other active substances produced by cell metabolism give them excellent development prospects in food, medicine, genetic engineering, liquid fuels, feed, environmental protection, and cosmetics.

[0003] Dunaliella salina is considered the best natural cell factory for beta-carotene and is a good nutritional health food. Under artificial intervention, beta-carotene accumulates significantly within Dunaliella salina cells, reaching 10-14% of its dry weight. In recent years, BASF in Australia has been cultivating Dunaliella salina using a runway pool production method, producing approximately 14 tons of dry beta-carotene annually. Globally, the production of beta-carotene from Dunaliella salina is approximately 1200 tons per year, which is expected to meet over 95% of beta-carotene demand. Dunaliella salina can also produce various other nutrients and bioactive substances with promising applications. The tocopherols, zeaxanthin, and polyunsaturated fatty acids provided by Dunaliella salina have antioxidant, antibacterial, and immunomodulatory functions, and can be used to enhance the flavor of low-salt foods and exert antibacterial and preservative effects. For example, adding the total dry matter of Dunaliella salina to pasta can increase its protein content, improve texture, and increase dietary fiber content; adding it to fruit and vegetable juices can increase total phenolic content, enhance antioxidant capacity, and improve sensory appeal. Therefore, Dunaliella salina, as a green additive in the food industry, has received increasing attention. With global population growth, food shortages, and deteriorating environmental quality, the commercial cultivation and production of Dunaliella salina demonstrates immense practical value and far-reaching significance. Dunaliella salina cells exhibit strong resilience, photosynthetic efficiency exceeding that of most terrestrial plants, and annual yields per acre more than 10 times that of the highest-yielding grain crops. Its land utilization rate is far higher than that of traditional livestock farming, and it can efficiently utilize wasteland in saline-alkali areas, offering high output value and effectively alleviating the food crisis. Currently, large-scale Dunaliella salina cultivation bases have been established in regions such as Inner Mongolia, which has profound significance for developing new high-quality food sources.

[0004] Currently, various methods exist for the pretreatment and enrichment separation stages in microalgae harvesting, including pre-oxidation, chemical flocculation, sedimentation, filtration, centrifugation, air flotation, and air flotation. However, each method has its advantages and disadvantages. Existing research results indicate that after pre-oxidation treatment, cells secrete extracellular products, which is beneficial for flocculation and sedimentation, but excessive use of chemicals can cause cell damage or even death. Chemical flocculation can greatly improve separation efficiency, but it easily causes water pollution. Due to the low concentration and small particle size of microalgae culture solutions, traditional methods such as sedimentation, filtration, and centrifugation for microalgae harvesting suffer from limitations such as low efficiency, high energy consumption, high cost, and cumbersome operation, which are not conducive to large-scale production. Air suspension is simple to operate, but the flocculation effect is not thorough. Currently, flocculation sedimentation is a traditional method for microalgae separation. This method is low in cost, easy to operate, and has a high recovery rate (Zhou Quan, Research on Salt Algae Culture and Air Flotation Harvesting Technology in Salt Production Mother Liquor. Seawater and Lake Salt and Chemical Industry 24, 11 (1995).). This method involves adding 500 mg / L FeCl3 and 50 mg / L polyacrylamide to the algal solution sequentially, then inserting a gas conduit into the bottom of the harvesting tank for aeration. The temperature is controlled at 25–35°C, and the pH at 7.5–8.2. Complete precipitation occurs within 1 hour, yielding an algal biomass of 0.64–0.96 g / L. American Microbial Resources Company harvests Dunaliella salina by introducing the algae into a dedicated harvesting and sedimentation tank, adding a certain proportion of aluminum sulfate under appropriate stirring to enrich the algae (Zheng Yi, Beijing University of Chemical Technology (2003)). The Qiquanhu Chemical Plant in Turpan, Xinjiang, my country, uses air flotation combined with flocculants to enrich Dunaliella salina (Duan Xuehui, Dunaliella salina and its biomass harvesting. Marine Salts and Chemicals 2722 (1998)). Chen Chunsheng et al. adjusted the pH of the algal solution to about 10.5 by adding about 720 mg / L of NaOH, causing the Dunaliella salina cells to self-flocculate. Then, the algal cells were separated from the culture medium by a high-efficiency air flotation device, which can achieve a recovery rate of 80% (Chen Chunsheng et al., Research on alkaline flocculation air flotation separation of Dunaliella salina [J]. Marine Development 20, 42 (2003).).

[0005] Patent CN103484373A discloses a method for concentrating and collecting microalgae in the field of algae separation technology. By adding sodium hydroxide and sodium carbonate to the algal solution, the microalgae cells and the precipitated insoluble salts co-precipitate, thus rapidly obtaining a concentrated microalgae solution. The added sodium carbonate can be obtained by absorbing carbon dioxide from waste gas using sodium hydroxide, thereby reducing greenhouse gas carbon dioxide emissions. The microalgae concentration rate can reach over 95%. Patent CN108865893B discloses a method for alkaline flocculation harvesting and circulating cultivation of microalgae. It utilizes a culture medium containing bicarbonate to cultivate microalgae, taking advantage of its high alkalinity, and adds a low concentration of flocculating ions for flocculation and sedimentation to harvest the microalgae; however, this method is costly. Patent CN103555586A discloses a method for collecting microalgae using the flocculation effect of protamine sulfate. The method of using protamine to flocculate microalgae in this invention is simple and effective. Protamine at a concentration of 5-40 mg / L can achieve a flocculation rate of about 75-98%. After precipitation, the algal cells still maintain good vitality and motility. The algal precipitate can be transferred to fresh liquid culture medium for further cultivation, with an algal cell precipitation rate of 76.8-90.4%. However, the protamine resources in this method are limited, which is not conducive to industrial-scale production.

[0006] Dunaliella salina thrives in high-salt environments, and large-scale outdoor cultivation using salt-producing mother liquor has been successfully achieved. Although Dunaliella salina can accumulate large amounts of β-carotene under stress conditions, harvesting its cells is challenging. The cells become brittle during dormancy, and the high viscosity of the culture medium makes them susceptible to breakage during harvesting, such as by centrifugation, which damages the active ingredients. Therefore, microalgae collection has always been a technological bottleneck in the industry's development. Harvesting Dunaliella salina is a crucial step in the entire production process, directly affecting the quality and production cost. Currently developed harvesting methods include centrifugation, filtration, flotation, sedimentation, and chemical flocculation. While centrifugation can quickly collect microalgae cells, it requires significant equipment and energy. Filtration is prone to membrane clogging, requiring constant cleaning. Natural sedimentation is slow. Flotation is energy-intensive. Chemical flocculation still requires further separation of the flocculant and the amorphous cell clusters formed by the adhesion of microalgae cells, such as filtration, centrifugation, or sedimentation. Furthermore, the extensive use of chemical flocculants such as ferric chloride, ferric sulfate, aluminum sulfate, and polyacrylamide can pollute the environment. The method of co-culturing filamentous fungi and microalgae to form mycelial balls can harvest 100% microalgae without adding any chemicals or requiring large-scale equipment. However, the co-culturing of filamentous fungi and microalgae can easily introduce biotoxins.

[0007] Patent CN103103133A discloses a method for harvesting Dunaliella salina: 1) In the later stage of Dunaliella salina cultivation, the algal cell density reaches (1~10)×10 6Harvesting is only possible when the algal cell count / mL is above 1; 2) Add flocculant mother liquor to the algal solution, stir well, let stand, remove the supernatant, collect the algal sludge, and calculate the flocculation efficiency; This method is simple and economical, utilizing inorganic flocculants, that is, through electrostatic force, intermolecular attraction and hydrogen bonding, the Dunaliella cells and flocculants adsorb each other, aggregate, flocculate and precipitate, thereby achieving the purpose of separating algal cells from the culture medium and enriching algal cells. The flocculant is added alone, which is low in cost, and the flocs form quickly and settle rapidly; however, the residue of inorganic flocculants such as PCA, alum or ferrous sulfate used is a major problem for the subsequent utilization of Dunaliella. Patent CN106399110B discloses a method for harvesting Dunaliella salina, comprising: a first part determining the pH value of the Dunaliella salina algal solution; a second part calculating the amount of sodium hydroxide used to adjust the pH value of a certain volume of Dunaliella salina algal solution to 12 to induce flocculation of the Dunaliella salina; a third part determining the time required for complete flocculation; and a fourth part harvesting the Dunaliella salina, with a harvesting efficiency greater than 80%. Patent CN107435027A discloses another method for harvesting Dunaliella salina, which involves... Under suitable environmental conditions, the algae are cultured to the specified values. The pH of the culture medium is adjusted to 11-13 using sodium hydroxide of appropriate concentration to the Dunaliella salina cells. The flocculation time is set at 2-4 hours. After flocculation, both the clear liquid and the algal sludge are adjusted to pH 7.8-9.3 using concentrated industrial hydrochloric acid. The clear liquid is reused to amplify the culture of Dunaliella salina cells. The algal sludge is washed with salt and dried to produce algal powder. Using the harvesting method of this invention, the purpose of concentrating Dunaliella salina cells can also be achieved, resulting in good harvesting results with a harvesting efficiency of 68%-87%.

[0008] Protozoa are a group of protozoa, many of which have close relationships with microalgae, primarily the following four: A. Predator-prey relationship: Protozoa feed on microalgae, approaching and consuming them through cilia, a crucial link in the aquatic food chain; microalgae, as primary producers, generate oxygen and organic matter during photosynthesis, providing food and energy for consumers, including protozoa; B. Symbiotic relationship: Some protozoa form symbiotic relationships with microalgae. For example, some protozoa contain photosynthetic microalgae within their bodies. The protozoa provide habitat and essential nutrients for the microalgae, while the microalgae provide organic matter for the protozoa. This symbiotic relationship is beneficial to both parties; a well-known example is... Examples include the green marine algae *Parasaurus*, which have a symbiotic relationship with *Chlorella*. *Chlorella* provides energy to the algae through photosynthesis, while the algae provide protection and a carbon source for *Chlorella*. C. Interdependence: In certain environments, microalgae and algae may form an interdependent relationship. For example, in the nitrogen cycle, algae decompose nitrogen produced by microalgae, a process that helps maintain nutrient cycling in the ecosystem. D. Competition: Under limited resources, algae and microalgae may sometimes compete, especially when resources such as light and nutrients are limited. Both require these resources for growth and reproduction, which can lead to competition.

[0009] Dunaliella salina cells have a diameter of approximately 5–10 μm, making traditional filtration systems prone to clogging and requiring frequent backwashing to maintain filtration efficiency. Therefore, the collection of Dunaliella salina has always been a major obstacle to its large-scale industrial cultivation. Currently, companies primarily employ mechanical, chemical, and electrochemical methods for collection, including gravity sedimentation, flocculation, flotation, filtration and screening, centrifugation, electroplating, ultrasonication, and solidification. Most of these techniques are energy-intensive, inefficient, time-consuming, and prone to introducing chemical impurities. Therefore, there is an urgent need to find environmentally friendly, efficient, and low-cost collection methods. Summary of the Invention

[0010] To address the aforementioned technical limitations, this invention provides a natural settling agent derived from the vegetative insect *Dunaliella salina*, which has a predatory effect on Dunaliella salina. The interaction between the vegetative insect and Dunaliella salina causes the algae to cease movement and thus settle. Based on this, the invention was completed.

[0011] In a first aspect, the present invention provides a settling agent and a composition thereof, wherein the settling agent contains worm bodies and / or worm extract, and the settling agent is co-cultured with microalgae cells at an appropriate density, so that the microalgae settle naturally.

[0012] Furthermore, the extract of the worm is obtained by filtering a solution from culturing worms.

[0013] Furthermore, when the settling agent contains swamp worms, the density of swamp worms in the algal solution is 1-8 worms / ml, preferably 2-7 worms / ml, and more preferably 2-5 worms / ml.

[0014] Furthermore, when the settling agent contains *Pleurotus ostreatus* extract, the concentration of *Pleurotus ostreatus* extract in the settling agent in the algal solution is 0.1-1 mL / mL, preferably 0.1-0.8 mL / mL, more preferably 0.2-0.7 mL / mL, and even more preferably 0.3-0.5 mL / mL.

[0015] Furthermore, the density of *Vaccinium* cells in the algal solution is 5 × 10⁻⁶. 6 —15×10 6 cell / mL, preferably 6×10 6 —14×10 6 cell / mL, more preferably 7×10 6 —12×10 6 cell / mL, more preferably 8×10 6 —12×10 6 cell / mL, further optimized to 9×10 6 —10×10 6 cell / mL.

[0016] In one embodiment, the preferred cell density of the *Vaccinium bracteatum* is 9.85 × 10⁻⁶. 6 cell / mL.

[0017] Furthermore, the worm pack is preferably Eulotes.

[0018] Furthermore, the microalgae is preferably Dunaliella salina.

[0019] Secondly, the present invention provides an application of a flocculant containing *Pterocarya stenoptera* or *Pterocarya stenoptera* extract in the natural sedimentation of microalgae; the application is to co-culture the flocculant containing *Pterocarya stenoptera* or *Pterocarya stenoptera* extract with microalgae, thereby causing the microalgae to settle naturally.

[0020] Furthermore, the microalgae include Spirulina, Chlorella, Haematococcus pluvialis, Dunaliella salina, Cryptodinium coccineum, Micrococcus pluvialis, Chlorella triangularis, Pleuronectiformis, Chaetoceros, and Leymus chinensis.

[0021] Furthermore, the microalgae is preferably Dunaliella salina.

[0022] Furthermore, the density of the parasitic insects in the algal solution in the settling agent is 1-8 insects / ml, preferably 2-7 insects / ml, and more preferably 2-5 insects / ml.

[0023] Furthermore, the concentration of the *Pleurotus ostreatus* extract in the settling agent is 0.1–1 mL / mL, preferably 0.1–0.8 mL / mL, more preferably 0.2–0.7 mL / mL, and even more preferably 0.3–0.5 mL / mL.

[0024] Furthermore, the density of *Vaccinium* cells in the algal solution is 5 × 10⁻⁶. 6 —15×10 6 cell / mL, preferably 6×10 6 —14×10 6 cell / mL, more preferably 7×10 6 —12×10 6 cell / mL, more preferably 8×10 6 —12×10 6 cell / mL, further optimized to 9×10 6 —10×10 6 cell / mL.

[0025] In one embodiment, the preferred cell density of the *Vaccinium bracteatum* is 9.85 × 10⁻⁶. 6 cell / mL.

[0026] Furthermore, the swamp worm is preferably Euplotes.

[0027] Thirdly, the present invention provides a method for the natural sedimentation of microalgae, the method comprising:

[0028] S1. Adjust the density of *Violetella* cells in the solution to achieve a suitable microalgae cell density;

[0029] S2. Add *Echinochloa crus-galli* and / or *Echinochloa crus-galli* extract to the solution with adjusted cell density;

[0030] S3. Co-cultivate for a period of time.

[0031] Furthermore, the density of the swamp worms in the algal solution in the settling agent is 1-8 worms / ml, preferably 2-7 worms / ml, and more preferably 2-5 worms / ml.

[0032] Furthermore, the concentration of the *Pleurotus ostreatus* extract in the settling agent is 0.1–1 mL / mL, preferably 0.1–0.8 mL / mL, more preferably 0.2–0.7 mL / mL, and even more preferably 0.3–0.5 mL / mL.

[0033] Furthermore, the density of *Vaccinium* cells in the algal solution is 5 × 10⁻⁶. 6 —15×10 6 cell / mL, preferably 6×10 6 —14×10 6cell / mL, more preferably 7×10 6 —12×10 6 cell / mL, more preferably 8×10 6 —12×10 6 cell / mL, further optimized to 9×10 6 —10×10 6 cell / mL.

[0034] In one embodiment, the preferred cell density of the *Vaccinium bracteatum* is 9.85 × 10⁻⁶. 6 cell / mL.

[0035] Furthermore, the swamp worm is preferably Euplotes.

[0036] Furthermore, the microalgae include Spirulina, Chlorella, Haematococcus pluvialis, Dunaliella salina, Cryptodinium coccineum, Micrococcus pluvialis, Chlorella triangularis, Pleuronectiformis, Chaetoceros, and Leymus chinensis.

[0037] Furthermore, the microalgae is preferably Dunaliella salina.

[0038] Beneficial effects

[0039] The method described in this application has two key advantages. First, the amount of pheromone used is extremely small. As a rapidly reproducing planktonic organism, pheromone is very low-cost for this amount of sedimentation. Second, as a common planktonic organism, pheromone is a natural sedimentation agent for Dunaliella salina and does not introduce impurities or toxic substances. Therefore, this sedimentation method uses very little pheromone, has high sedimentation efficiency, minimal chemical interference, and as a natural sedimentation agent, it is non-toxic. Attached Figure Description

[0040] Figure 1 A picture of a simple vacuum filtration device.

[0041] Figure 2 This represents the percentage of Dunaliella salina that has settled over time.

[0042] Figure 3 To evaluate the sedimentation activity and stability of the filtrate from the worm *Pleurotus ostreatus*.

[0043] Figure 4 This is an illustration of the final settling degree of *Velociraptor dulcis*. Detailed Implementation

[0044] The specific embodiments of the present invention will be further described below. It should be noted that these descriptions are for the purpose of aiding understanding the present invention, but do not constitute a limitation thereof. Furthermore, the technical features involved in the embodiments described below can be combined with each other as long as they do not conflict with each other.

[0045] Unless otherwise specified, the experimental methods used in the following embodiments are conventional methods, and the experimental materials used in the following embodiments are all available through conventional commercial channels.

[0046] The "harvest rate" involved in this invention is estimated by measuring the cell density and volume of the cells settling at the bottom, estimating the number of cells settling, and using this method to estimate the total number of cells. The ratio of the number of cells settling to the total number of cells is used as the harvest rate.

[0047] Example 1: Filtration of *Dunaliella salina* and Culture and Quantification of Dunaliella salina Cells

[0048] After filtering, sterilizing, and cooling the seawater, add f / 2 medium at a ratio of 1:1000 to the sterile seawater. Then, take Dunaliella salina cells, add them to the prepared medium, shake well, and culture in a light incubator for 5-7 days.

[0049] The density of the cultured Dunaliella salina was determined using a hemocytometer, and the density was increased to 5 × 10⁻⁶. 6 When the density reaches 1 / cell / mL, it indicates that the density is close to the maximum density of exponential growth, and sedimentation experiments are most effective at this point.

[0050] The fed Dunaliella salina were filtered through a 20-micron filter membrane, rinsed once with sterile seawater, and then rinsed back down. The filtered fluid was collected in a sterile culture flask and allowed to stand for one day. Before use, the fluid was shaken well, and 1 mL was taken from the middle of the flask and the number of Dunaliella salina was recorded under an inverted microscope. This process was repeated 3-5 times, and the average value was taken to determine the density of the Dunaliella salina. The number of Dunaliella salina added was then determined based on the amount of Dunaliella salina used.

[0051] Dunaliella salina has a concentration of 9.85 × 10⁻⁶. 6 At a concentration of cell / mL, the number of *Pteris vittata* was 5.

[0052] Example 2: Test of the Motility Effect of Dunaliella salina

[0053] Add *Dunaliella salina*, *Dunaliella salina* filtrate, and an equal volume of sterile seawater to *Dunaliella salina* that has reached near its maximum concentration. Cultivate for 6 days, and take 20 μL daily into a sperm motility detection system to detect its movement. Calculate the percentage of immobile cells in the exported data.

[0054] Processing of extracting *Pteris vittata*: Take the core of an empty centrifuge column, and cut the dropper tip according to the diameter of the end of the core. Then assemble a simple suction filtration device (such as...). Figure 1(As shown). Take the cultured *Pleurotus ostreatus* solution, use a dropper to add the solution to the core of an empty centrifuge column, and filter. After obtaining the *Pleurotus ostreatus* solution, use a sheared dropper to backwash with sterile seawater. If the cultured *Pleurotus ostreatus* solution contains many other algal cells, the filtration can be repeated several times. After filtration, observe the purity of the *Pleurotus ostreatus* under a microscope. Let the filtered *Pleurotus ostreatus* stand for one day, and observe its reproduction after one day. The entire operation is performed under sterile conditions.

[0055] Cells with a density of 9.85 × 10⁻⁶ were respectively... 6 Add the same volume of blank culture medium, *Dunaliella salina* filtrate (200 *Dunaliella salina* cultured in 50 mL of sterile seawater, allowed to stand for 24 h, shaken well, filtered to obtain the filtrate, and added at a ratio of 1 mL extract per 2 mL of algal solution, i.e., extract concentration of 0.33 mL / mL), and *Dunaliella salina* (density of 5 individuals / mL) to *Dunaliella salina*.

[0056] The results showed that in the experimental group with the addition of *Pterocarya spp.*, Dunaliella salina exhibited significant sedimentation on the third day; in the experimental group with the addition of *Pterocarya spp.* filtrate, significant sedimentation of Dunaliella salina was induced on the fourth day; and in the control group with the addition of blank culture medium, no significant sedimentation was observed over time.

[0057] By the third day, a significant difference in the immobile percentage had emerged between the experimental group with *Pteris vittata* and the control group. By the fourth day, both the *Pteris vittata* filtrate group and the *Pteris vittata* group showed significant differences from the control group, with an immobile percentage reaching 99% (e.g., ...). Figure 2 (As shown).

[0058] Example 3: Settlement Effect Test

[0059] Add Dunaliella salina (density 9.85 × 10⁻⁶) to a 5 mL centrifuge tube. 6 Cell density was measured at 4 mL / mL. Then, 5 *Paratomium spp.* (5 spp. / mL; 1 mL) and 1 mL of *Paratomium spp.* extract (1 mL of filtrate obtained after filtering 50 mL of sterile seawater containing nearly 200 *Paratomium spp.*) and 1 mL of sterile seawater were added as controls. The mixture was cultured for 6 days, and the cell density at the bottom was measured daily. The results are shown below. Figure 3 As shown.

[0060] On the sixth day after adding *Alternaria solani* and its extract, the density of cells settling at the bottom showed a significant difference compared to the control group, showing a significant increase (i.e., a significant increase in bottom cell density due to algal cell settling), and the increase was more significant in the group with *Alternaria solani* than in the group with *Alternaria solani* extract.

[0061] To further evaluate the stability of the sedimentation effect of the *Pheretima aspergillum* solution, 1 mL of *Pheretima aspergillum* solution (density 5 individuals / mL) was added to a total volume of 4 mL of *Dunaliella salina* solution (density 9.85 × 10⁻⁶). 6 The control group was given an equal volume of sterile seawater. The sedimentation of Dunaliella salina was observed daily. Five parallel culture experiments were conducted, and sedimentation was observed on the 6th day. The percentage of Dunaliella salina cells that remained stationary at the bottom reached 95% in all five experimental groups. There was no significant difference between the experimental groups, indicating that the autonomous movement of Dunaliella salina by the vegetative worms is stable. Therefore, it can be determined that the vegetative worm solution has relatively stable biological activity and is suitable for the sedimentation process of Dunaliella salina cells.

[0062] like Figure 4 The image shows the final sedimentation effect. Groups E and D represent the experimental groups with added *Dalbergia latifolia*. In these groups, 1 mL of *Dalbergia latifolia* solution (density 5 individuals / mL) was added to a total volume of 4 mL of *Dunaliella salina* solution (density 9.85 × 10⁻⁶). 6 (cells / mL); control was a control group that did not undergo any biological treatment and maintained the variable as unique (4 mL of Dunaliella salina solution, density 9.85 × 10⁻⁶ cells / mL); 6 (cells / mL, with 1 ml of sterile seawater added); after standing for three days, the algal cell density at the bottom of the control group was 8.9 × 10⁻⁶. 6 The algal cell density at the bottom of the experimental group was 1.98 × 10⁶ cells / mL. 8 The concentration of algal cells was measured at approximately 0.16 mL, yielding a concentrated liquid of about 0.16 mL. Based on the sedimentation cell density and volume, the recovery rate was calculated to be 66.9%. This sedimentation method uses very little pheromone, has high sedimentation efficiency, minimal chemical interference, and as a natural sedimentation agent, it is non-toxic.

[0063] The experiments conducted in this invention demonstrate that *Dunaliella salina* plays a remarkably significant role in the settling process of Dunaliella salina. Therefore, using *Dunaliella salina* has two major advantages: firstly, the amount of *Dunaliella salina* required is extremely small; as a rapidly reproducing planktonic organism, its cost is very low for this settling dosage. Secondly, as a common planktonic organism, *Dunaliella salina* acts as a natural settling agent for Dunaliella salina, without introducing impurities or toxic substances.

[0064] The above description is a general description of the invention. Variations in form and equivalent substitutions may be made as needed or in accordance with practical requirements. While specific terminology is used herein, it is intended for descriptive purposes and not for limitation. Various alterations or modifications can be made to the invention by those skilled in the art, and these equivalent forms also fall within the scope defined by the appended claims.

Claims

1. A co-culture system comprising Euplotes aedipus and / or Euplotes aedipus extract, wherein the Euplotes aedipus and / or Euplotes aedipus extract is co-cultured with microalgae cells at an appropriate density, so that the microalgae naturally settle; the microalgae is Dunaliella salina; and the Euplotes aedipus extract is obtained by filtering a solution of cultivated Euplotes aedipus.

2. The co-culture system of claim 1, wherein the cells are derived from the same species. When the co-culture system comprises Euplotes aedipus, the density of Euplotes aedipus in the co-culture system is 1-8 individuals / mL.

3. The co-culture system as described in claim 1, characterized in that, When the co-culture system comprises Euplotes aedipus extract, the density of Euplotes aedipus extract in the co-culture system is 0.1-0.8 mL / mL.

4. The co-culture system as described in claim 1, characterized in that, The density of the cells of the algae in the liquid was 5 x 10 6 -15 x 10 6 cells / mL.

5. Use of Euplotes aedipus or Euplotes aedipus extract in the preparation of a microalgae settling agent, wherein the use is by co-culturing Euplotes aedipus or Euplotes aedipus extract with microalgae, so that the microalgae naturally settle; the microalgae is Dunaliella salina; and the Euplotes aedipus extract is obtained by filtering a solution of cultivated Euplotes aedipus.

6. The use according to claim 5, wherein the compound is ###0002### When the microalgae settling agent comprises Euplotes aedipus, the density of Euplotes aedipus in the microalgae settling agent is 1-8 individuals / mL.

7. The use according to claim 5, characterized in that, When the microalgae settling agent comprises Euplotes aedipus extract, the density of Euplotes aedipus extract in the microalgae settling agent is 0.1-0.8 mL / mL.

8. The use according to claim 5, characterized in that, The density of the cells of the algae in the liquid was 5 x 10 6 -15 x 10 6 cells / mL.

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