The collection method is crucial for a successful establishment of an axenic culture. Damage of algal cells during collection leads to failure. When sampling an unknown environment or target organism, multiple collection methods should be addressed. Sampling techniques may include syringe sampling, scraping, brushing, and the inverted Petri dish method. However, the most reported sampling method is simply water collection from a water body. If the target species is usually found in-depth, a Niskin bottle or rosette sampler can be used. When sampling, it is fundamental to record factors such as light, water temperature, dissolved O₂ and CO₂, nutrient concentration, pH, and salinity to mimic these conditions at the laboratory and establish the location coordinates to replicate sampling if needed [2, 3, 5].

The sampling conditions will not play the main role in their posterior axenicity as long as the aseptic technique is applied. However, if the genera or species to be isolated is targeted or previously known, the conditions of the collection should be adjusted to favour it, so the axenic culturing can be achieved more easily.

Once a sample is taken to the lab, adding nutrients to the natural sample will potentiate microalgal growth. Organic substances should be added in small amounts because they can cause undesirable bacterial proliferation. The enriched culture is incubated and examined every few days for the growth of the target species, and then individual cells are isolated. Sometimes the target species grow successfully after isolation but die after several transfers, indicating that the culture medium lacks an element or compound or that the organism accumulates wastes that poison their environment [2]. Enrichment of samples should be targeted for different kinds of microalgae; for example, delicate algae and flagellates could be favored by enriching samples with ES enriched seawater; chrysophytes, chlorophytes, diatoms, and cryptophytes grow well under the addition of F/2 media to water samples. On the other hand, cyanobacteria growth is facilitated by adding BG11SW media [6].

This nutrient addition process will facilitate the growth of the targeted genera or species, easing the posterior purification techniques by favoring their growth. This will allow the number of microalgal cells to overthrow the number of fungi or bacterial cells, or even the cells of other microalgal species that are not of interest to the researcher.

Microalgae usually secrete a heterogeneous matrix called extracellular polymeric substances (EPS), which mainly consists of polysaccharides, proteins, nucleic acids, and lipids. This way, algal cells interact with each other, mediate their adhesion to surfaces, and generate cell protection and energy storage reservoirs. Bacteria frequently attach and feed on this EPS layer. Vortexing, sonication, and surfactant treatment prior to isolation methods assist in detaching bacteria and increasing the separation efficiency [7, 8].

Detaching the bacteria from the EPS will increase the effectiveness of techniques such as filtration, centrifugation, and antimicrobial treatment, by allowing the proper separation of cells by their difference in size and density and making the bacteria more susceptible to the chemical agents added.

Essentially, samples are separated into two portions mainly based on particle size differences [7]. Recently, Haoujar (2020) applied this method by filtering samples from Moroccan Mediterranean seawater through a series of membranes decreasing mesh (33, 20, and 0.45 μm) then placing membranes on agar plates, thus achieving the isolation of 32 microalgae (Mostly coccoid) [9]. This method allows the separation of microorganisms by their cell size only, but by itself, it is usually not enough to obtain axenic or unialgal cultures since different organisms can have similar cell sizes. However, this technique can be applied in such a way that the microalgae cells, which are usually bigger than bacteria, can be retained in a membrane or filter to be posteriorly cultivated in a media in which most parts of the bacterial cells were removed.

Centrifugation works by separating organisms by cell size using gravity. Low-density cells, such as bacteria, tend to stay in the supernatant fraction and are discarded, while microalgae and higher-density cells normally remain in the pellet fraction. After several centrifuging cycles, a higher concentration of a desired microalgal strain will be achieved. Using a silica or percol density gradient is useful for separating the different cells into different bands. The separation efficiency of centrifugation depends on speed and time; therefore, these parameters should be optimized for a given species. Also, excessive centrifugal force can sometimes damage fragile cells [5, 7, 10]. Lee (2021) applied density gradient centrifugation to dinoflagellate species Karenia mikimotoi and Alexandrium tamarense using a 90%, 90–50%, 90–50–30%, and 90–50–30–10% Percoll solution and harvesting the algal cells, reducing the bacterial population from 5.79 ± 0.22 log10 CFU/mL to 1.13 ± 0.07 log10 CFU/mL [11]. As seen in several studies, centrifugation is helpful in concentrating algal cells and reducing the bacterial population. However, this technique does not generate an axenic algal culture by itself, but it will facilitate posterior techniques that will ensure the purity of cultures.

This method can be performed with a micropipette, a pasteur pipette, or a glass capillary. The objective is to collect a single cell from the sample, deposit it in a sterile droplet of culture medium, pick up the cell again, and transfer it to a second sterile droplet. This process is repeated until a single algal cell can be placed into the final isolation vessel [2]. Novel techniques for single-cell isolation include micromanipulation by employing stereomicroscope, microcapillary tubes, and optical tweezers, which greatly reduces labor and cell damage. However, they require expensive equipment and skilled lab personnel. These techniques perform cell separation with a high level of accuracy, making it an ideal tool for screening and isolation. Microcapillaries are usually made of glass with an outer diameter of 1 mm. On the other hand, optical tweezers (OT) is a non-invasive method for physical manipulation that uses a highly focused laser beam to trap and manipulate microscopic objects using only the properties of light, applying usually infrared (IR) region wavelengths between 700 and 1300 nm that generates two kinds of forces, the scattering force produced by the photons striking the cell along their propagation direction and the gradient force produced by a gradient of field intensity. These forces act as optical tweezers and depend on the wavelength of the laser beam and the particle size. This allows for the micromanipulation of a single object that ranges in size from nanometres to micrometres, including living cells. Such optical forces can stop and drag biological swimmers, such as microalgae. Application of OT for single-cell manipulation may be based on fluorescence activation or cells’ shape, size, and refractive index difference, providing an opportunity to sort cells to a compartment without any pre-treatment. However, very few studies have reported on this specific technique [5, 12-16]. Pilat et al. concluded that non-invasive optical manipulation of living cells of photosynthetic microorganisms could be performed using laser wavelengths longer than 935 nm. They also observed that a 1064 nm wavelength did not cause any damage to the microalgae, and it is sufficient for reliable 3D manipulation of cells. Lim (2012) applied the serial dilution and posterior micromanipulator technique to obtain pure cultures of Nannochloropsis sp., Dunaniella salina, Chlorella sp., and Tetraselmis sp [17]. Ota (2019) designed a glass microfluidic device to perform isolation, cultivation, and time-course observation of Euglena gracilis by using semi-closed microchannels. Then the cells were identified by Raman microscopy, a vibrational spectroscopic technique that relies on the inelastic scattering of monochromatic light produced by a laser that interacts with the sample, which causes a shift in the energy of the photon to a higher or lower frequency that can be detected by the equipment. Biomolecules in microalgae have their own specific functional groups and show characteristic peak frequencies that appear in Raman spectra, which allows researchers to identify specific biomolecules for specific microalgae species. In this case, they manage to identify the formation of paramylon, a unique metabolite of E. gracilis [16, 18].

In order to achieve successful isolation using this technique, the algae must be able to grow on solid media. Coccoid cells and most diatoms grow well on agar, some flagellates do not grow on agar, and dinoflagellates rarely grow on agar. After agar culturing, a micropipette tip is driven into the agar to pick up the selected cells and then discharged into a liquid culture medium. This technique can also be used to maintain axenic cultures by perpetual transfer with repeated use of the agar pour technique [2]. It is common to use more than one method for microalgae isolation. Jahn (2014) isolated 24 unialgal cultures by filtering samples through a 100 μm filter, enriching the samples with f/2-medium and streaking the enriched microalgal communities on f/2-medium solidified plates with agar, then picking colonies from the plates and transferring them to a liquid f/2-medium in 12 well plates [3]. It is also common for researchers to use several medium cultures to obtain different genera of microalgae. Thao (2017) isolated oleaginous microalgae. After sampling, they used BG11 for fresh, f/2B for brackish, and f/2 for marine water samples. Then, for all three media, a solid or semi-solid medium was prepared by adding 10 g/L or 5 g/L agarose, respectively. Two methods of isolation were applied: 1) Streaking on solid agarose plates; 2) Adding 0.1 mL of enriched cultures to 5 mL of semi-solid medium, vortexed and poured over the solid medium to form a semi-solid overlay, incubated, and repeated until pure isolates were obtained [19]. Certain features of microalgae, such as phototaxis, can be combined with agar cultivation to maximize results. Banerjee (2012) isolated Chlamydomonas sp. from Chlorella sp. mixtures by passing light (6,000 lux for 7 h) through agar, given that Chlamydomonas has flagella and the rhodopsin pigment is attracted towards the light [20]. Isolation in agar by itself may lead to an axenic microalgae culture, as long as the targeted microalgae to be isolated does not grow on consortia with other microorganisms and does not have bacterial cells attached to the EPS. Posterior tests and microscopic observation may be required to ensure the axenicity of the culture. If, after isolation in agar, the targeted microalgae remain contaminated, further techniques should be applied. However, this technique allows the separation of targeted microalgae from most of the microorganisms from a sample and drastically reduces the contaminants.

This is done by transferring only one cell into a vessel by diluting a sample or culture. This technique is usually performed by preparing and filling test tubes with 9 mL of sterile saline solution or adequate culture media. Then, 1 mL of microalgae culture or an enriched sample is transferred into the first test tube and mixed, obtaining a 1:10 dilution. Then, 1 mL from the first dilution is transferred into the second tube and repeated five to six times. It is important to consider inoculating many tubes from the last dilution because some single cells will die and some inocula will contain two or more species or no cells at all. Enrichment substrates can be added to isolation tubes to specifically grow a target species or incubate under certain specific conditions. Axenic isolates are not often obtained with the dilution technique because bacteria are commonly more abundant than algae. The dilution technique is often used when attempting to culture random algal species from field samples with the goal of discovering and studying new species [2, 21]. Dilution techniques will allow obtaining cultures which are mostly composed of some microalgal species but do not ensure the axenicity of the culture. Contamination tests and microscopy observation is needed to detect other microorganisms present in the culture. Further techniques will likely have to be applied in order to generate the axenic microalgal culture.

The main objective of antibiotic treatment is to reduce bacterial contamination of an algal culture to a number that will allow a small inoculum to transfer enough algal cells without viable bacteria. The efficacy and toxicity of the antibiotic treatment depend on the concentration and period of exposure and vary depending on the microalgae and bacteria species. Viable bacteria tend to decline drastically after 48 h of exposure to the microalgal culture to the antibiotic treatment. Then a small portion can be transferred to an adequate medium without antibiotics. Antibiotics will work in one of two ways: bactericidal (e.g., penicillin, vancomycin) will kill bacteria by interfering with the formation of the bacterial cell wall, or bacteriostatic (e.g., chloramphenicol, tetracycline), which interferes with the bacterial metabolism but does not necessarily kill them. Theoretically, antibiotics will destroy all the contaminants without harming algal cells, but this is rarely achieved in practice. A well-studied and basic antibiotic treatment is 100/25/25 mg/L of penicillin, streptomycin, and gentamycin which is usually tolerated by most algae. Purification may also be achieved by the sequential transfer of the algal culture through a series of different antibiotics. This method has lower toxicity to the algae. Also, one antibiotic mix may be fatal to some bacteria but only suppress the growth of others, while using sequential antibiotics may kill the surviving bacteria [2, 7, 22].

Many combinations and techniques for purification with antibiotics can be applied, from simple antibiotic addition like Bonett (2020), who managed to obtain axenic cultures from Chlorella sp., C. sorokiniana and Desmodesmus sp using only a BG11 medium added with ampicillin in concentrations of 50, 70, 80, 90 – 100 mg/ L [23], to using combinations such as the one reported by Wu (2013) in which he obtained axenic cultures of Desmodesmus sp. by continual sub-culturing in BG11 agar plates supplemented with ampicillin and kanamycin [24]. More complex antibiotic combinations can be used when bacterial contamination persists. Han (2016) obtained purified strains of Nannochloropsis sp., Cylindrotheca sp., Tetraselmis sp., and Amphikrikos sp after treating them with a mix of ampicillin, gentamycin, kanamycin, neomycin, and streptomycin (600 mg/L each) for 3 days and then transferring them to an antibiotic-free medium for 5 days to posterior spread on solid f/2 media to isolate colonies. They also reported that microalgae treated with antibiotics at high concentrations resume their growth fast when transferred to an antibiotics-free medium [25]. Lee (2021) generated axenic cultures for both armored and unarmored dinoflagellate species Karenia mikimotoi and Alexandrium tamarense. After gradient centrifugation they used an antibiotic treatment by spreading 0.1 mL xenic cultures on agar containing: penicillin (100 U), streptomycin (100 μg/mL), gentamicin (100 μg/mL) and tetracycline (1 μg/mL). After 14 days of incubation, no signs of bacterial colonies were obtained [11].

Regarding antibiotic toxicity to microalgae, Youn and Hur found that chloramphenicol and penicillin G showed lethal effects on microalgae, being higher for chloramphenicol and that the optimum quantity for antibiotic treatments without microalgal toxicity was 2,000 ppm of dihydrostreptomycin for Chlorella ellipsoidea, neomycin 500 ppm for Isochrysis galbana and Heterosigma ahashiwo, chloramphenicol 500 ppm for Cyclotella didymus, and dihydrostreptomycin sulphate and neomycin 6,000 ppm for Thalassiosira allenii [20]. Bashir and Cho (2016) also studied the effect of antibiotics on microalgae, comparing the growth and protein synthesis of chlorophytes Dictyosphaerium pulchellum and Micractinium pusillum in the presence of kanamycin and tetracycline and reported a significant effect of both antibiotics in the measured parameters [26].

Antibiotics are usually applied once the sample has been enriched and previous isolation techniques have been applied to the culture, so a unialgal culture has been obtained. This treatment will allow the elimination of bacteria and/or fungi present in the culture or reduce it in a way that will ease the obtention of the axenic culture. Antibiotic treatment is one of the most used techniques by researchers for obtaining axenic microalgal cultures. It is usually economical, and most laboratories have the facilities and equipment to carry out the methodology.

Obtaining axenic cyanobacteria culture using an antibiotic treatment usually has certain limitations and sometimes requires an alternative. Microalgae tend to be more resistant to lysozyme digestion than bacteria. Another option for generating an axenic culture is using a lysozyme base. Axenic cyanobacterial cultures have previously been obtained by treating cultures with the minimum lethal concentration for up to 90 min [7]. Also, combinations of enzymatic treatment and antibiotics can lead to promising results. Tale (2014) used a combination of lysozyme (20 μg/mL) and antibiotic mixture (cefotaxime-500 μg/mL and tetracycline 50 μg/mL) to obtain axenic cultures of the genera Chlorella and Monoraphidium [27].

Sodium hypochlorite, sulfonamides, potassium tellurite, detergents, pesticides, salts, aldehydes, and peroxides have also been used to obtain axenic cultures. However, these compounds tend to have higher toxicity on microalgae. Recently the triiodide ion (-3 I) has been applied with promising results. This is adsorbed onto a resin surface and then filled into a column. The contaminated unialgal cell suspension is usually run through this column several times for axenization. This technique was applied to a Nannochloropsis gaditana culture contaminated with bacteria. Observations lead to the conclusion that the treatment remarkably reduced the number of viable bacteria and had few effects on the microalgae. There are a few reports on the use of natural agents in algae cultures for contamination control, such as cinnamaldehyde, geraniol, carvacrol, and cinnamic acid, that were evaluated to eliminate fungi. Natural chemical agents are generally less polluting, more easily degradable, and may comply with disposal regulations. Also, the resistance of biological contaminants to currently used chemical agents enhances the importance of finding new alternatives. Still, these natural substances need research and regulation in order to be used [7, 26-28].

For the optimal chemical agent control of contamination and purification of microalgae cultures, the screening of antibiotics, chemicals, or enzymes which inhibit biological pollutants without damaging the target microalgae is the preferred route [29, 30].

Microalgae are traditionally maintained by periodical sub-culture due to the feasibility of the procedure. Conventional sub-culturing is made by moving an inoculum from a late log/stationary phase into a pre-sterilized, fresh culture medium using the aseptic technique. The transfer frequency is determined by the growth characteristics of the strain. Continuous maintenance over long periods of time is labor-intensive. However, it is often the only procedure available in some laboratories or the only way for the preservation of microalgae recalcitrant to cryopreservation or freeze-drying. A principal limitation of perpetual transferis the media and incubation regimens, that in contrast to the natural conditions may cause morphological and physiological changes in some species. Other concerns include the possibility of contamination or handling mistakes [35].

Freeze-drying requires the elimination of water from a frozen algal suspension by sublimation under reduced pressure for which specialized equipment is needed. Freeze-dried microalgae maintain their original shape and texture, and it has been found to be successful in preserving mostly cyanobacteria. This procedure minimizes the hazard of genetic drift, contamination, and accidental mislabeling. Also, the cultures require smaller storage space and reduced labor. It is also the best way of shipping cultures. However, freeze-drying has not been found to be a successful maintenance method for many microalgae, which exhibit very low levels of viability after the freeze-drying process [35, 36].

Cryopreservation may be defined as the storage of an organism at an ultralow temperature, so it remains capable of surviving the defrosting process. Many microalgae can be cryopreserved but typically with lower post-thaw viability levels. Cryopreservation is still largely an empirical science process. Despite this, hundreds of species of cyanobacteria and eukaryotic microalgae have been successfully cryopreserved by commonly applying methanol (MeOH), dimethylsulfoxide (DMSO), or glycerol as cryoprotective additives (CPAs). This type of methodology reduces the routine costs for the culture collection, ensuring the stability and purity of samples and reducing the risks of contamination during the handling of samples. The specific procedure and CPA used for a particular species or even a strain may vary. Some trial and error may be required for new isolates. Cryopreservation often includes placing 1.0 ml of the culture (preferably actively growing under suitable conditions) at the desired temperature of choice, either a -20°C, -80 °C, or -170 °C (vapor phase of liquid nitrogen) with the chosen CPA. The usage of a two-step protocol may be necessary for some strains. This requires the addition of the CPA to the culture and cooling it to a specified subzero temperature to facilitate dehydration, then cooling rapidly to the final storage temperature. For the revival of algal cultures, the cryogenic vials are usually placed in a 35 °C water bath for 5 minutes and then transferred into 50 ml of an appropriate growing medium and incubate at appropriate conditions. Generally, strains that display specific cell characteristics such as unicellular, small, spherical, with no spines, and without vacuoles are usually highly viable after thawing. This cryopreservation methodology is currently applied by The Culture Collection of Freshwater Microalgae based at Universidade Federal de São Carlos, which currently has the largest microalgae collection in Brazil, holding around 700 strains of freshwater microalgae. Some disadvantages are that cryopreservation requires specialized equipment, special consumables, and training. Errors resulting from improper thawing under uncontrolled conditions, even briefly, may lead to the death of culture [35-39].

After cryopreservation, viability should be determined by detecting the metabolic activity and/or reproduction capacity of the cells. Vital staining employing fluorescene diacetate or re-growth of cells are the most practicable approaches. Usually, a minimum acceptable viability level is around 30%. In addition, there should be no significant differences in genotype or phenotype between control and post-thaw cultures. These conditions are currently problematic to achieve and may not be technically feasible for some taxa and for non-axenic material [40].