Cell growth dynamics
Figure S1 shows the growth curves of three selected algal monocultures (D. tertiolecta, T. suecica, C. closterium) studied at four selected salinities (9, 19, 27, and 38). The initial number of inoculated cells in the growth media was similar for all species studied, approximately 4.0 × 104 cells mL−1. All selected microalgae persisted in the salinity range from 9 to 38. The calculated growth rates and doubling times of three algae in the exponential growth phase at the salinity studied are summarized in Table S1. All microalgae had the shortest doubling time and fastest growth at salinity of 9.
Confocal images of algal cells grown at the corresponding salinities are shown in Figures S2–S4. Microscopic observations of D. tertiolecta at salinity of 27 showed no changes in autofluorescence or actin composition compared to cells grown at salinity of 38 (Figure S2a and b). At salinity of 19, D. tertiolecta cells build up actin layer (Figure S2c), which is particularly pronounced at salinity of 9 (Figure S2d). In addition, as observed in the transmitted light channel, at salinity of 9, some cells lost their flagella and became rounder, and the actin layer appeared thicker than in the control. No changes in autofluorescence or actin composition were observed in T. suecica over the entire range of salinity examined (Figure S3a-d). However, as observed in the transmitted light images, T. suecica cells tend to lose their flagella as salinity decreases. This effect was most evident at salinities of 19 and 9, where almost all cells have lost their flagella and the detached flagella accumulate around the cells. The cells of C. closterium maintained both shape and autofluorescent properties throughout the salinity drop (Figure S4a-d). As observed in the transmitted light channel, droplets accumulated inside the cells with decreasing salinity, although no trend was noted.
Qualitative insights into the movement of D. tertiolecta cells at selected salinities are shown in Figure S5. At salinity of 9, approximately 66% of the cells (109 cells) were stationary or oscillating around a center (Tables S2.1), while the remainder (55 cells) exhibited considerable trajectories. At salinity of 19, a total of 68 cells were counted in the sample, of which about 57% (39 cells) were stationary. At salinity of 27, approximately 63% (41 cells) were stationary, while 37% (24 cells) exhibited considerable movement and were quantified. In contrast, at salinity of 38, most cells (79%) were in motion, while the minority (21%) were stationary.
Box plots of cell speeds of D. tertiolecta are shown in Fig. 1a.
The median of the speed at salinity of 9 was 29 µm s−1, while the medians at salinities of 19 and 27 were identical: 50 µm s−1. At salinity of 38, the median speed was significantly higher: 75 µm s−1. The Shapiro test for normality yielded p = 2.7 × 10−10, 1.5 × 10−9, 3.4 × 10−5, and 1.8 × 10−6, confirming that the density distributions of speeds were very far from normal. The Wilcoxon rank sum test showed that the density distributions of speeds were significantly different for cells grown at salinities of 9 and 19, 27 and 38, but not for cells grown at 19 and 27.
Because the group of cells that were stationary or vibrating around the center exhibited significantly different motion than the group of cells that were moving, it was important to note the speeds of the moving cells. The average speeds at salinities of 9, 19, 27, and 38 were 74 µm s−1, 103 µm s−1, 77 µm s−1, and 81 µm s−1, respectively (Table S2.2a).
Boxplots of the search radius of D. tertiolecta cells are shown in Fig. 1b. The median search radius at salinity of 9 was 2 µm, whereas the medians at salinities of 19 and 27 were 4 and 3 µm, respectively. At salinity of 38, the median search radius was significantly larger: 18 µm (Table S2.2.b). The Shapiro test for normality yielded p = 2.2 × 10−16, 7.3 × 10−16, 2.1 × 10−13, and 2.2 × 10−16, respectively, confirming that all density distributions of the search radius were very far from normal. The Wilcoxon rank sum test showed that the density distributions of the search radius were significantly different for cells grown at salinities of 9 and 19, 27 and 38, but again not for cells grown at 19 and 27 (p = 0.88). The group of cells that moved consistently had an average search radius of 12 µm, 27 µm, 79 µm, and 72 µm at salinities of 9, 19, 27, and 38, respectively. In the same order of salinity, the linearity of motion was 0.1, 0.09, 0.32, and 0.45, respectively. Thus, the linearity was the same at salinities of 9 and 19 and 3 to 4.5 times smaller than at 27 and 38.
Qualitative insights into the movement of T. suecica cells grown at selected salinities are shown in Figure S6. At salinity of 9, approximately 43% of cells (35 cells) were stationary or showed oscillatory movement in place (Tables S3.1), whereas the majority of cells (46 cells) were clearly moving. At salinity of 19, a total of 124 cells were counted, with approximately 59% being stationary (73 cells). At salinity of 27, approximately 27% (26 cells) were stationary, while 73% (69 cells) showed significant movement and were quantified. In contrast, at salinity of 38, only 6% of cells were stationary, while 94% of cells moved vigorously.
Box plots of cell speeds of T. suecica are shown in Fig. 2a.
The median of the speed was 78 µm s−1 at salinity of 9, while the medians were 50 µm s−1 at salinity of 19 and 112 µm s−1 at salinities of 27. At salinity of 38, the median speed was significantly higher: 201 µm s−1 (Table S3.2a). The Shapiro test for normality yielded p = 4 × 10−4, 1.8 × 10−10, 0.07, 8.8 × 10−5, respectively. Only the density distribution at salinity of 27 did not deviate significantly from normality. The Wilcoxon rank sum test showed that the density distributions of speed were significantly different from each other, except for the distributions at salinities of 9 and 27, for which p = 0.05 was determined. The group of uniformly moving cells grown at salinities of 9, 19, 27, and 38 had an average speed of 124 µm s−1, 155 µm s−1, 137 µm s−1, and 201 µm s−1, respectively (Table S3.2).
Boxplots of the search radius of T. suecica cells are shown in Fig. 2b. The median search radius at salinity of 9 was 6 µm, while the medians at salinities of 19 and 27 were 4 and 12 µm, respectively. At salinity of 38, the median search radius was an order of magnitude larger: 102 µm (Table S3.2b). The Shapiro test for normality yielded p = 4.6 × 10−16, 2.2 × 10−16, 6 × 10−16, and 1.4 × 10−15, respectively, confirming that all density distributions of the search radius are very far from normal. The Wilcoxon rank sum test showed that the density distributions of the search radii were significantly different from each other. The search radii of cells grown at salinities of 9 and 19 (p = 0.027) and cells grown at salinities of 9 and 27 (p = 0.22) were the narrowest. The group of significantly moving cells grown at salinities of 9, 19, 27, and 38 had an average search radius of 57 µm, 121 µm, 61 µm, and 174 µm, respectively. The linearity of moving cells with increasing salinity was 0.26, 0.58, 0.19, and 0.55, respectively, and reached the highest values at salinities of 19 and 38.
Electrochemical characterization of algal cells and released surface-active organic matter
The chronoamperometric curves for oxygen reduction recorded in the cell suspension of D. tertiolecta in seawater at potential of − 400 mV showed signals attributable to the adhesion of single cells to the charged interface (Fig. 3a).
The dependence of the signal frequency of D. tertiolecta grown at different salinities on the applied potentials is shown in Fig. 3b. The potential range of cell adhesion was defined with critical potentials of adhesion at the positively Ec+ and negatively charged interface, Ec−. The most negative and the most positive potentials, where at least one amperometric signal occurs per 10 consecutive I-t curves, correspond to the critical potentials (Žutić et al. 1993; Ivošević et al. 1994). The narrowest potential range of adhesion was recorded in the D. tertiolecta cell culture grown at salinity of 9, characterized by critical potentials of − 140 mV and − 990 mV in seawater, while the widest potential range of cell adhesion was recorded in the cell suspension grown at 38 salinity, from − 110 to − 1240 mV, corresponding to favorable growth conditions. The frequency of amperometric signals increased with decreasing salinity due to the lower ionic strength of the medium, which was reflected in the increase in oxygen reduction current, thus enhancing the amperometric signals. The maximum number of amperometric signals occurred at potential of − 400 mV for all four salinities studied, as the interfacial tension is close to the maximum value (electrocapillary maximum). At potential of − 400 mV, the mercury electrode was positively charged and there was an electrostatic attraction between the positively charged interface and the negatively charged D. tertiolecta cells. By changing the potential in either direction, the interfacial tensions decreased and the number of amperometric signals from the cells decreased accordingly. At potential of − 800 mV, the mercury was negatively charged and the signal frequency decreased due to electrostatic repulsion with the negatively charged D. tertiolecta cells. Conversely, the chronoamperometric curves recorded in the suspensions of T. suecica and C. closterium were perfectly regular because there was no adhesion to the charged liquid interface due to cell rigidity (Novosel and Ivošević DeNardis 2021).
The electrochemical characterization of the released surface-active organic matter in the growth medium was determined by recording polarograms (current–potential curve) of Hg(II), which is proportional to the surfactant activity in the sample. The surfactant activity of the sample corresponded to a quantitative measure of the physiological activity of the cells in the growth medium. The data showed that the surfactant activity of the cells gradually decreased as follows: T. suecica (19) > D. tertiolecta (9) > D. tertiolecta (19) ∼ T. suecica (27) (Fig. 3c).
Nanoscale imaging of algal cell and released extracellular biopolymers
Nanoscale imaging of single algal cells was performed at salinities of 9, 19, 27, and 38 (control). Regardless of salinity, all three species retained the same general cell shape. Dunaliella tertiolecta cells grown at all salinities tested had an ovoid shape with two flexible flagella. Tetraselmis suecica cells had an ellipsoidal shape at all salinities tested and the cell surface had granular structures corresponding to micropearls (Novosel et al. 2021). Most of the cells of T. suecica grown at salinity of 38 had flagella, and only half of the cells grown at salinity of 27 had flagella, whereas the cells grown at salinities of 9 and 19 had no flagella. The cells of C. closterium grown at all salinities tested had an elongated shape with flexible rostrae that could be clearly distinguished from the central part of the cell. Three morphologically distinct parts could be distinguished on the cell: the girdle band, the valve, and the raphe (Pletikapić et al. 2012; Novosel et al. 2021).
Based on AFM image analysis, the morphological parameters (length, width, height, and surface roughness) of cells grown at selected salinities are summarized in Table S4. The size of D. tertiolecta and T. suecica cells had the highest values at salinity of 38. The size of both, D. tertiolecta and T. suecica, grown at salinities of 9, 19, and 27 was similar and smaller than cells grown at 38. The roughness of D. tertiolecta cell surface was highest at salinity 38 and similar for salinities of 9, 19, and 27. The roughness of the cell surface of T. suecica was similar at all tested salinities. The length, width, and height range of C. closterium grown at salinities of 9, 19, and 27 were similar and greater compared with cells grown at salinity of 38.
The supramolecular organization of the released extracellular polymers (EPS) of D. tertiolecta, T. suecica, and C. closterium at selected salinities are shown in Fig. 4.
Around the cells of D. tertiolecta grown at salinities of 38 and 27, only globules and no fibrils or fibrillar networks were observed. Globules and some single fibrils were observed around the cells grown at salinity of 19. Around the cells grown at salinity of 9, a material consisting of globules, single fibrils and flat smooth structures was noted (Fig. 4a, b). The extracellular biopolymers around T. suecica grown at salinities of 9, 19, and 27 were in the form of a dense fibrillar networks and were located all around the cells, whereas no fibrillar material was observed at control salinity of 38 (Fig. 4c, d). The fibrils that formed the network ranged in height from 5 to 50 nm, with the highest network density found around the cells at salinity of 19 (Figure S7). The extracellular biopolymers of C. closterium were in the form of single fibrils, locally cross-linked fibrils, and globules. For C. closterium grown at salinities of 9 and 19, a denser EPS material around the cells was noted and the fibrils exhibited a higher degree of cross-linking (Fig. 4e, f), whereas C. closterium grown at salinity of 27 had a lower degree of cross-linked fibrils near the cells, while single fibrils were mostly observed further from the cells.
Nanomechanical characterization of algal cells by AFM
The local elastic properties (E) of algal cells were quantified using the apparent Young’s modulus calculated for the maximum indentation depth. At salinity of 38 (control), the cells of D. tertiolecta are characterized by the lowest E values. The cells of T. suecica are stiffer, while the local E values of C. closterium can be up to several MPa. The difference in mechanical response of these cells to compression could be due to differences in cell morphology. The cells of T. suecica are surrounded by close-fitting theca of fused organic scales. The cells of C. closterium contain stiff chloroplasts in the girdle band, and the cells of D. tertiolecta are covered only by the thick plasma membrane (Oliveira et al. 1980; Medlin and Kaczmarska 2004). Figure 5 shows the overlay of the box plots and Young’s modulus distributions obtained for the algal cells of D. tertiolecta, T. suecica, and C. closterium cultured at salinities of 9, 19, 27, and 38, respectively.
Because the distributions of Young’s moduli are broad and not symmetrical, we compared the median values of the cell populations studied. The median was accompanied by an interquartile range (IQR), which describes where the central 50% of the data lie (median (IQR). Statistical significance was determined using the Kruskal–Wallis ANOVA test (p < 0.05) to confirm differences between groups. Regardless of algal species, decreasing salinity increased the apparent Young’s modulus (Fig. 5). The statistical significance for all groups was less than 0.0001 at the 0.05 level (Kruskal–Wallis ANOVA test) compared to the control group. Median (IQR) values obtained for D. tertiolecta cells increased from 3.5 kPa (3.2 kPa) at 38 to 8.6 kPa (7.6 kPa) at 27, 6.6 kPa (4.8 kPa) at 19, and 8.1 kPa (4.4 kPa) at 9, respectively. Statistical significance (p < 0.05) was also found for the measurements at salinities of 9 and 19, and at 19 and 27, whereas no statistically significant difference was found at salinities of 9 and 27. A similar trend of elastic modulus changes was observed in T. suecica cells. The cells were stiffer at low salinities, and E (27) > E (9) > E (19). The corresponding medians were 138 kPa (190 kPa), 151 kPa (223 kPa), and 115 kPa (198 kPa), respectively. The value of E determined for T. suecica at salinity of 38 was 46 kPa (80 kPa). Weaker statistical significance was found between algal cells cultured at salinities of 9 and 27 (p = 0.0011), and 19 and 27 (p = 0.0019). There was no statistical significance between cells of T. suecica cultured at salinities of 9 and 27. For C. closterium, the salinity stress increased the Young’s modulus from 215 kPa (436 kPa) at normal conditions to 362 kPa (742 kPa) at salinity of 27. A further decrease in salinity to 19 and 9 was accompanied by an increase in the E to 553 kPa (532 kPa) and 537 kPa (759 kPa), respectively. The statistical significance for the 19 and 9 groups and 27 and 19 groups of C. closterium cells was less than 0.05, while the p value determined for the 9 and 27 groups was less than 0.001.
Adhesive and hydrophobic properties of algal cells
The adhesive properties, quantified by the maximum work of adhesion (Wadh), of algal cells grown at different salinities on chemically modified probes were studied. Cells were indented with hydrophilic (OTS −) and hydrophobic (OTS +) AFM probes. The change in hydrophobic properties (ΔWadh) of the algal cell surface was determined by subtracting the work of adhesion determined for bare and OTS-coated cantilevers (Table S5): ΔWadh = Wadh(no OTS) − Wadh(OTS). A positive value of ΔWadh indicated that hydrophilic interactions dominate, while negative values indicated that hydrophobicity smothered the hydrophilicity. The mean values of the maximum work of adhesion (± standard error of the mean) obtained from measurements with bare and OTS-coated cantilevers are shown in Table S5.
We hypothesized that algal cells change their adhesion properties under salinity stress. Indeed, the hydrophobic properties of the cells changed depending on the salinity studied (Fig. 6), and these changes were species-dependent.
The resulting chemical properties of microalgae are shown in Figure S8. The data show that in the case of D. tertiolecta, the hydrophobic and hydrophilic properties were rather balanced, being characterized by low values of ΔWadh (see Table S5). Only at salinities of 9 and 27 the cells became slightly more hydrophobic, as ΔWadh is − 0.018 ± 0.014 fJ and 0.047 ± 0.012 fJ, respectively. More pronounced salinity-dependent changes in surface properties were observed in T. suecica and C. closterium. At salinities of 38 and 27, ΔWadh of T. suecica was close to zero. A further decrease in salinity resulted in high negative ΔWadh values. In addition, we observed a significant decrease in the probability of adhesion to the bare AFM probe (PnoOTS), accompanied by an increase in POTS (see Figure S8 and Table S5). While the overall surface properties of T. suecica changed from balanced to hydrophobic with decreasing salinity, in the case of C. closterium we found that salinity can be a trigger between hydrophobic and hydrophilic surface properties. At salinity of 38, C. closterium cells had a hydrophilic surface (ΔWadh = 0.0583 ± 0.0072 fJ). When the salinity decreased to 27, the ΔWadh increased to 0.163 ± 0.085 fJ. Moreover, when the salinity decreased to 19, the relationship between the surface properties of the algal cells changed, and the cells of C. closterium became highly hydrophobic. At salinity of 9, the resulting work of adhesion was still negative. Moreover, the results of adhesion probability showed the affinity to hydrophilic materials depending on the structure and properties of the microalgal species (see Figure S8). Dunaliella tertiolecta showed a very low adhesion probability to the hydrophilic surfaces, in contrast to the cells of C. closterium, which preferentially adhered to the hydrophilic probe throughout the salinity studied. In the case of T. suecica, at salinities of 27 and 38, PnoOTS > POTS, while at salinities of 9 and 19, PnoOTS < < POTS.
Lipid characterization of algal cells
The changes in cellular content of membrane lipids, ST, GL, and PL, caused by the decrease in salinity in the microalgal cell cultures of D. tertiolecta, T. suecica, and C. closterium are shown in Fig. 7.
In general, the least changes in total membrane lipids and classes with decreasing salinity were observed in D. tertiolecta (Fig. 7a, d, g, and j). In T. suecica, decreasing salinity resulted in a decrease in cellular content of total membrane lipids with a decrease of GL, followed by an increase of PL. In the microalga C. closterium, a decrease in total membrane lipid concentration, GL and PL was observed with a decrease in salinity from 38 to 9, but without a particular trend of change. Parallelly, a statistically significant (p < 0.05) increase in the cellular content of sterols was observed with the decrease in salinity in all three microalgae. At salinity of 9, the sterol content in diatom C. closterium was increased 3.8-fold compared to salinity of 38. However, the highest cellular sterol content was observed in T. suecica (Fig. 7e).