Microplate-based high throughput screening procedure for the isolation of lipid-rich marine microalgae
© Pereira et al; licensee BioMed Central Ltd. 2011
Received: 5 August 2011
Accepted: 22 December 2011
Published: 22 December 2011
We describe a new selection method based on BODIPY (4,4-difluoro-1,3,5,7-tetramethyl-4-bora-3a,4a-diaza-s-indacene) staining, fluorescence activated cell sorting (FACS) and microplate-based isolation of lipid-rich microalgae from an environmental sample. Our results show that direct sorting onto solid medium upon FACS can save about 3 weeks during the scale-up process as compared with the growth of the same cultures in liquid medium. This approach enabled us to isolate a biodiverse collection of several axenic and unialgal cultures of different phyla.
Microalgae are aquatic photosynthetic microorganisms able to transform carbon dioxide into biochemicals that can later be processed into biofuels, food, feed and high-value bioactive compounds . With regard to biofuels, algal biomass is considered likely to be one of the most important sources of renewable energies in the near future . Although biodiesel production from microalgae is a proven technology, it still faces several technical and economical constraints that need to be addressed [3, 4] in order to scale up production and thus lower the final production costs .
Extraction of bioactive compounds with potential applications in pharmacology and biomedicine is a relatively new trend in microalgal biotechnology. Microalgal biomass presents natural active compounds responsible for distinct biological activities, such as cytotoxic, antibiotic, antioxidant, antifungal, anti-inflammatory and antihelminthic compounds [6–9].
The rise of interest in these microscopic organisms for biotechnological applications is due to the unique biochemical features and their vast biodiversity, which to date is almost entirely unexploited . Although many culture collections of microalgae have been established, the variety of unknown species and strains present in the environment with potential application in the production of biofuels and/or as a source of bioactive compounds is very high [8, 10, 11]. Thus, easy and feasible high throughput screening procedures are essential in order to isolate novel species and strains for specific purposes.
Although several techniques for microalgae isolation have been described previously, such as single-cell isolation in liquid and solid media, serial dilutions, medium enrichment, gravimetric separation, micromanipulation and atomized cell spray [12, 13], flow cytometry has recently shown significant potential in improving microalgal strains for lipid production in an expedited fashion [14, 15]. Fluorescence activated cell sorting (FACS) enables the selection of particular strains of microalgae and subsequent isolation . The characterization of different populations within any mixture of cells is performed through direct measurement of optical cell properties (for example, light scatter and multicolor fluorescence emission), which in turn enables FACS of defined cell populations that can be cultured separately at a later stage [17–19].
Several authors have reported successful sorting procedures for microalgae. Reckermann  described the sorting and culturing of a variety of unicellular species isolated from an environmental water sample. More recently, Doan et al.  reported the isolation from Singapore waters of microalgal strains for the purpose of biofuel production. However, FACS has been considered to be a technique displaying low efficiency for the isolation of unialgal cultures, especially those of fragile species such as dinoflagellates . Therefore, there is a need to develop simpler and faster methods allowing the isolation of fast-growing strains.
In the present work, a combination of two methods was tested: FACS combined with growing cells in 96-well plates containing solid agar growth medium to accelerate both the isolation procedure and culture scale-up. This combination resulted in a high throughput screening procedure to isolate and screen for lipid-rich strains by means of BODIPY 505/515 (4,4-difluoro-1,3,5,7-tetramethyl-4-bora-3a,4a-diaza-s-indacene) staining that can also be used to isolate fast-growing microalgae. These cells can then be further tested for bioactivities. Via this approach several strains of microalgae were isolated and easily scaled up to higher volumes at a later stage.
Results and discussion
Selection of fast growing strains
Algal strains intended for biotechnological applications need to be produced as fast as possible in large-scale systems in order to ensure a sustainable process. Therefore microalgae displaying high growth rates are essential. In this context, water samples were supplemented with Algal growth medium and incubated for 7 days using growth conditions as described in Methods. This enrichment step facilitates the isolation procedure, allowing fast-growing strains to dominate, by competition, other microalgae of less interest under a set of desired growth conditions. If the enrichment step is omitted, fast-growing cells might be overlooked during the isolation procedure due to their low concentration. Thus, this stage is a key step in the selection of microalgae isolated by the present method.
Isolation by FACS
Although BODIPY, phycoerythrin and phycocyanin fluorescence were used in the strain isolation for this specific environmental sample, different combinations of channels can be chosen to produce a similar two-dimensional plot as shown in Figure 2. In this particular case, FL2 and FL4 produced the best clustering for the chosen gating method. Though the other channels gave relevant information, they were not used in the procedure for separation of the different naturally occurring cell populations since they formed overlapped clusters. The use of fluorescence emission due to BODIPY staining in the gating procedure should be done with care since the concentration of lipids in the cells varies with the culture growth stage. Cells normally present higher lipid concentrations during stationary phase as compared with cells growing exponentially. As in the present work environmental samples underwent a pre-enrichment step to isolate fast-growing cells able to withstand competition from other microalgae, it is possible that lipid-rich strains growing actively might have been overlooked.
To assess cell morphology, the gated clusters were then sorted directly to microscope slides and observed in a Zeiss Axio Imager Z2 fluorescence microscope. Cells from the P1 cluster did not survive the cell sorting procedure, since upon microscopic observation only disrupted cells were found. The same problem was described by Reckermann  with Fibrocapsa japonica that disrupted soon after sorting. Microscopic observations suggested that the established gates were able to isolate monoalgal cultures since all observed fields from the same cluster showed the same strain. As expected, P5 was an exception and microscopic observation showed different cyanobacterial strains and debris.
Efficiency of cell sorting (solid medium vs liquid medium)
The efficiency of the sorting procedure obtained in this work (70%) was considerably higher than that reported by Sensen et al.  for the removal of bacteria from unialgal cultures (20% to 30%). Global recovery, however, was lower than reported by Doan et al.  in novel isolates from an environmental water sample (82% to 100%). Still, sorting efficiency is highly dependent on the original sample, including the starting group of species and their abundance, thus demonstrating the importance of the initial enrichment step. For instance, Sinigalliano et al.  compared the isolation efficiency of dinoflagellates between FACS and manual picking using a micropipette and the former approach appeared to be less efficient than the latter (0.5% vs 2%, respectively). This could be due to the frailness of algae belonging to this taxonomic group. The enrichment step can also lead to a selective enrichment of some species. In our work, Algal growth medium, which contains low levels of silica, was used . Therefore, domination of mixed cultures by diatoms and other silicate-requiring algae was not favored during the enrichment process.
To ensure algal growth and estimate the best starting number of cells for an expedited recovery, different numbers of events (cell-containing droplets) were tested. For each well 1, 2, 10 or 20 events were sorted. Interestingly, the starting number of cells per well was not a constraint in the scale-up procedure on solid medium. Regardless of the number or events sorted per well, visual growth was obtained after a 2-week incubation period. Single cell sorting in solid medium (one event) yielded a single axenic colony in most wells (Figure 6A), which was easily transferred onto a Petri dish with solid medium. Wells with > 1 event (2, 10, 20 events) gave rise to a large number of colonies; however, all of them presented visible bacterial growth (Figure 6B-D).
In liquid medium, after a 2-week incubation period algal growth was only visible in the wells with 10 or 20 events. Most of the obtained cultures were not unialgal and showed abundant bacterial contamination as verified by microscopic observation (data not shown). Algal growth in wells with one or two events was only visible after 4 weeks. A few of these cultures were, however, axenic, though at a very low frequency (5%). Therefore, in liquid medium, increased culture growth can be achieved by sorting more events, which results in a faster scale-up, but the use of several replicates is recommended when sorting to liquid medium to isolate axenic cultures as this improves the scale-up procedure efficiency. Overall, the scale-up procedure was considerably faster in solid medium (4 weeks) than the required period for achieving the same growth and culture volume using liquid medium (7 weeks).
The fact that BODIPY cell staining did not affect cell recovery in both solid and liquid medium is essential for present and future work in the detection and isolation of lipid hyper-producing algae. Although it has already been reported that culturing cells after Nile red staining is possible [14, 15], BODIPY presents several advantages when compared with Nile red: (1) BODIPY is able to stain a wide range of algae groups without the need of using high concentrations of dimethylsulfoxide (DMSO) or acetone to carry the dye in, which can be key to wider and faster cell recovery and scale-up; and (2) BODIPY preferentially traces lipid bodies instead of other cytoplasmic compartments .
Identification of microalgae
Strains isolated during the sorting procedure
Species and strains isolated throughout this work, corresponding phylum and methods applied in the identification of each strain
18S rDNA sequencing
18S rDNA sequencing
18S rDNA sequencing
18S rDNA sequencing
18S rDNA sequencing
18S rDNA sequencing
18S rDNA sequencing
18S rDNA sequencing
18S rDNA sequencing
Growth and lipid assessment
Flow cytometry coupled with fluorescence activated cell sorting is an efficient tool for isolating strains of microalgae, which can be used either as biodiesel raw material, or as a source for bioactive compounds. Our FACS approach is a user-friendly, fast procedure than most common methods for the isolation of microalgae, with the advantage of being able to obtain axenic, unialgal cultures in a matter of weeks. This resulted in a 4-week culture scale-up (instead of a 7-week scale-up) if cells were sorted directly onto solid rather than liquid medium.
A wide range of microalgae groups has already been isolated by this method. This suggests that the method described here is a promising high throughput procedure to isolate lipid-rich strains as well as microalgae for other biotechnological purposes. The gating procedure and culture medium added in the initial enrichment step are key to favor the isolation of fast-growing microalgal taxa with high lipid contents.
Sampling and microalgal growth
Water sampling was performed in aquaculture ponds at the facilities of Atlantik Fish SA on the south-eastern coast of Portugal. In every sampling spot the water was collected and stored in 1 l bottles and kept at room temperature. At a later stage, the water samples were transferred to 80 ml test tubes containing liquid Algal growth medium, which was prepared with sterile seawater supplemented with an Algal stock solution concentrated 1,000 × . Cultures were kept in an incubator for 7 days at 21°C, with a 12:12 h dark/light photoperiod, at a photon flux density of 80 μmol/m2/s. Growth on solid medium was carried out in either Petri dishes or 96-well plates containing Algal growth medium solidified with 1.5% agar.
Lipid bodies were stained with a solvatochromic fluorochrome, BODIPY 505/515 (Life Technologies Europe BV, Porto, Portugal), as described by Cooper et al. . Cells were stained with a 1 mM aqueous solution of BODIPY dissolved in DMSO (0.2%) to attain a final concentration of 1 μM. Upon addition of the fluorochrome, tubes were vortexed for 1 minute and incubated at room temperature in darkness for 10 minutes.
The flow cytometer used in our studies was a Becton Dickinson FACS Aria II (BD Biosciences, Erembodegem, Belgium). Fluorescence readings were performed by excitation with a blue and red laser (488 and 633 nm, respectively). The emission signal was measured in three channels upon excitation with the blue laser: FL1 channel centered at 530/30 nm; FL2 centered at 585/42 nm; and FL3 centered at 695/40 nm. A fourth channel, FL4, registered the emission at 660/20 nm after excitation with the red laser.
Samples were acquired with the software FACSDiva version 6.1.3 (BD Biosciences, Erembodegem, Belgium). After the acquisition of samples, images were treated with the analysis software, Infinicyt 1.5.0 (Cytognos S.L., Santa Marta de Tormes, Spain).
The settings and compensations of all channels and lasers were the same for all sorting procedures. The flow cytometry sheath fluid used in all experiments was sterile filtered seawater. Filters (PALL) used had a pore size of 0.2 μm. Sorting was performed at 2,000 events/s flow rate using 'single cell' sort precision mode, with a 100 μm nozzle.
Cells were sorted directly into wells of 96-well plates containing 250 μl of either liquid or solid (agar) Algal growth medium. In order to assess the best number of cells needed to achieve visible culture growth in a feasible time, sorting conditions were set in order to direct 1, 2, 10 or 20 droplets into each well. After the cell sorting procedure, cells were incubated for 2 weeks in the same incubator and growth conditions as described above.
Microscopic images were acquired in a Zeiss AXIOMAGER Z2 microscope, with a coollSNApHQ2 camera and AxioVision software version 4.8 (Carl Zeiss MicroImaging GmbH, Gõttingen, Germany), using the 100 × lens. For the fluorescence images, we used Zeiss 38 He filter set (Carl Zeiss MicroImaging GmbH, Gõttingen, Germany) for fluorescein isothiocyanate (FITC) and the transmitted light images were acquired using differential interference contrast. Z stacks were acquired and the resulting image was a maximum intensity projection of the stacks. For the plate cultures the images where acquired in a Zeiss SteREO Lumar.V12 stereoscope, equipped with an Axiocam MRC, using AxioVision software release 4.8 (Carl Zeiss MicroImaging GmbH, Gõttingen, Germany). Images were treated using Image J software (Research Service Branch, NIH, Bethesda, MD).
The scale-up procedure was carried out by streaking a single colony from a well that did not show any signs of bacterial growth onto a Petri dish containing solid Algal growth medium. After 1 week axenic plates were scrapped to 100 ml Erlenmeyer flasks with liquid Algal growth medium. A week later the 100 ml culture was transferred to 1 l reactors with aeration. In liquid medium, the scale-up was performed by transferring the 250 μl of the 96-well plates into 5 ml test tubes containing 1 ml of liquid Algal growth medium under minor aeration. Cells were allowed to grow for 1 week and were then transferred successively to 20 ml test tubes, 100 ml test tubes and finally 1 l reactors with aeration.
Gravimetric determination of total lipids
Lipid extraction was performed according to a modified protocol by Bligh and Dyer . Briefly, the obtained algal biomass was homogenized at room temperature with an IKA Ultra-Turrax disperser (IKA-Werke GmbH, Staufen, Germany), in a mixture of chloroform, methanol and water (2:2:1). The mixture was centrifuged to allow phase separation, and a known volume of the organic phase was pipetted into a new preweighed tube. The extract was then evaporated until dryness in a warm bath (50°C) and weighed carefully to estimate lipid contents.
Primers used in the identification of the isolated strains (18S rDNA) and determination of axenicity status of established microalgal cultures (16S rDNA)
Sequence (5' to 3')
The MarBiotech research group belongs to CCMAR http://www.ccmar.ualg.pt, an institution dedicated to carry out research in marine sciences belonging to ASSEMBLE (the Association of European Marine Biological Laboratories; http://www.assemblemarine.org; CCMAR is named as 'Faro'). MarBiotech is led by JV, who has published several works in the elucidation of the biosynthesis pathway of lipophyllic pigments (carotenoids) in the microalga Dunaliella salina. Recently JV has secured the collaboration of one postdoctoral fellow (LC) and a faculty member (LB), who provided funding and expertise concerning analytical methods for lipid detection. HP and CVD are recipients of research grants for graduate students. AM and CF are members of the University of Algarve (where CCMAR is located) and are technical staff specialized in flow cytometry and microscopy, respectively.
This work was supported by the SEABIOMED project (PTDC/MAR/103957/2008), funded by the Foundation for Science and Technology (FCT) and the Portuguese National Budget. LC is an FCT postdoctoral research fellow (SFRH/BPD/65116/2009). CP is an exchange student funded by the European Erasmus mobility program. We also would like to acknowledge AquaExam for the identification of cyanobacteria and Rodrigo Costa (CCMAR) and his research group for the help provided with the successful PCR amplification of 16S rDNA.
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