Open Access

Surface physicochemical properties of selected single and mixed cultures of microalgae and cyanobacteria and their relationship with sedimentation kinetics

  • Ana L Gonçalves1,
  • Carla Ferreira1,
  • Joana A Loureiro1,
  • José CM Pires1 and
  • Manuel Simões1Email author
Bioresources and Bioprocessing20152:21

DOI: 10.1186/s40643-015-0051-y

Received: 23 February 2015

Accepted: 22 April 2015

Published: 2 May 2015

Abstract

Background

Microalgae are photosynthetic microorganisms presenting a diversity of biotechnological applications. However, microalgal cultivation systems are not energetically and economically feasible. Possible strategies that can be applied to improve the feasibility of microalgal production include biofouling control in photobioreactors, the use of attached growth systems and bioflocculation. These processes are ruled by surface physicochemical properties. Accordingly, the surface physicochemical properties of Chlorella vulgaris, Pseudokirchneriella subcapitata, Synechocystis salina and Microcystis aeruginosa were determined through contact angle and zeta potential measurements. Additionally, mixed cultures of the selected microorganisms were performed. Sedimentation kinetics of the studied cultures was also evaluated to understand how surface physicochemical properties influence microalgal recovery.

Results

All studied microorganisms, except S. salina, presented a hydrophilic surface. The co-culture of S. salina with the other studied microorganisms resulted in a more hydrophobic algal suspension. Regarding zeta potential determinations, all studied suspensions presented a negatively charged surface (approximately -40.8 ± 4.4 mV). Sedimentation experiments have shown that all microalgal suspensions presented low microalgal recovery efficiencies. However, a negative linear relationship between microalgal removal percentage and free energy of hydrophobic interaction was obtained.

Conclusions

The evidence of a relationship between microalgal removal percentage and free energy of hydrophobic interaction demonstrates the importance of surface physicochemical properties on microalgal settling. However, the low recovery efficiencies achieved, as well as the high net zeta potential values determined, indicate that another factor to consider in microalgal settling is the ionic strength of the culture medium, which play an important role in suspensions’ stability.

Keywords

Cyanobacteria Microalgae Mixed cultures Sedimentation kinetics Surface physicochemical properties

Background

Microalgal culturing has been the focus of several research studies worldwide, due to the wide variety of biotechnological applications described for these photosynthetic microorganisms [1,2]. When growing autotrophically, microalgae convert CO2 (from atmosphere or flue gas emissions) into organic carbon compounds, thus reducing the CO2 accumulation in the atmosphere [3-6]. Additionally, microalgae assimilate other compounds, such as nitrogen and phosphorus, frequently found in wastewaters, meaning that these microorganisms may play an important role in wastewater treatment processes [7-10]. Finally, microalgal biomass has several applications [1,11-13]: (i) human food and animal feed; (ii) production of cosmetics, drugs and functional food; and (iii) biofuels. However, microalgal cultivation still presents high process costs, which are mainly due to the low biomass productivities and the associated harvesting costs, accounting for 20% to 30% of biomass production costs [14]. Moreover, it requires large amounts of water and nutrients, which is the reason to be considered a process with high environmental impact [15].

To improve biomass productivities in microalgal photobioreactors (PBRs), new strategies should be adopted to avoid biofouling (the attachment of cells to the walls of PBRs). This phenomenon is responsible for lower biomass productivities, since the amount of light (the energetic source required for photosynthesis) that effectively penetrates into microalgal suspension is significantly reduced [16-18]. On the other hand, several authors have reported the growth of microalgae in biofilms. These immobilized growth systems facilitate further harvesting processes [7,16,19,20]. For both promotion and control of microalgal attachment, knowledge about surface physicochemical properties is essential [16,21,22]. Knowing the surface characteristics may also be very helpful in metabolite extraction. For example, the use of solvents with polarities similar to those of target metabolites may facilitate the contact between both, increasing extraction efficiency [23].

Regarding the harvesting process, currently applied methods include chemical coagulation/flocculation, gravity sedimentation, flotation, filtration, centrifugation and electrical-based processes [14,24,25]. Selection of an appropriate harvesting method depends on the end product, namely its value and properties [14]: it is important to consider the acceptable level of moisture, salt concentrations, cell damage and strain features, such as their density and size. Additionally, selection of an effective harvesting procedure must take into account that microalgal biomass must be further processed. Therefore, these procedures must not be toxic or contaminate microalgal biomass. It is also desirable that the selected harvesting method allows the recycling of the culture medium [25]. In this sense, bioflocculation appears as a viable alternative for the commonly used procedures. This method consists in the addition of other microorganisms or microbial metabolites to microalgal cultures, trying to stimulate microbial aggregation and flocs formation. The use of bioflocculation as harvesting method is more energetically efficient and reduces the impact of the addition of chemical flocculants to microalgal biomass, which tend to be an expensive and toxic alternative [26]. The ability of microorganisms to form flocs strongly depends on their surface physicochemical properties.

Information about the surface physicochemical properties of microalgae and about their interaction mechanisms may be very helpful in the development of energetically and economically feasible biomass production systems and harvesting procedures, as well as in the development of new strategies to prevent biofouling. This study provides information about the surface physicochemical properties of the microalgae Chlorella vulgaris and Pseudokirchneriella subcapitata and the cyanobacteria Synechocystis salina and Microcystis aeruginosa determined experimentally, according to the method proposed by van Oss et al. [27-29]. Selection of these microorganisms was based on the following factors [30-33]: (i) these microalgae and cyanobacteria can be easily grown in laboratory cultures and (ii) several authors have reported the use of these microorganisms in a wide variety of biotechnological applications, such as CO2 capture, wastewater treatment, biofuel production and synthesis of bioactive compounds. Additionally, algal suspensions resulting from the co-culture of different associations between the selected microorganisms were evaluated, to determine if these associations have influence on surface physicochemical properties and on their ability to form aggregates. Sedimentation kinetics of the studied cultures was also determined, to assess the influence of surface physicochemical properties on the recovery of microalgal biomass.

Methods

Microorganisms and culture medium

The microalgae C. vulgaris CCAP 211/11B and P. subcapitata CCAP 278/4 were obtained from the Culture Collection of Algae and Protozoa (UK), while the cyanobacteria S. salina LEGE 06079 and M. aeruginosa LEGE 91344 were obtained from the Laboratory of Ecotoxicology, Genomic and Evolution - CIIMAR (Centre of Marine and Environmental Research of the University of Porto, Portugal). Stock solutions of these microorganisms were prepared in OECD (Organisation for Economic Co-operation and Development) test medium [34], with the following composition (per litre): 250 mg NaNO3, 12 mg MgCl2·6H2O, 18 mg CaCl2·2H2O, 15 mg MgSO4·7H2O, 45 mg KH2PO4, 0.08 mg FeCl3·6H2O, 0.1 mg Na2EDTA·2H2O, 0.185 mg H3BO3, 0.415 mg MnCl2·4H2O, 3 μg ZnCl2, 1.5 μg CoCl2·6H2O, 0.01 μg CuCl2·2H2O, 7 μg Na2MoO4·2H2O and 50 mg NaHCO3. The culture medium was sterilized by autoclaving at 121°C for 15 min. The cells were incubated in 500-mL flasks at room temperature (24.0 ± 1.0°C), under continuous fluorescent light with an irradiance of 120 μE m−2 s−1 at the surface of the flasks. Agitation was obtained by bubbling atmospheric air (filtered through 0.22-μm cellulose acetate membranes, Orange Scientific, Belgium) in the bottom of the flasks.

Microalgal culturing in single and mixed cultures

Previously described stock solutions were used to prepare microalgal suspensions for further characterization of surface physicochemical properties and sedimentation kinetics determination. In these experiments, single cultures, as well as different associations between the selected microorganisms, were used. Batch experiments were performed in 500-mL flasks (VWR, Carnaxide, Portugal) with a working volume of 450 mL. Cells were cultivated for 7 days with an initial cell concentration of approximately 1.0 × 106 cells mL−1 (in mixed cultures, testing all possible associations of two microorganisms, the initial cell concentration was approximately 2.0 × 106 cells mL−1). Temperature, light and agitation conditions were the same as for the stock solutions preparation. All the experiments were performed in duplicates under aseptic conditions.

Surface physicochemical properties

Zeta potential measurements

In the last day of each culture, microalgal suspensions were harvested, washed twice and resuspended in MilliQ water (Millipore, Billerica, MA, USA) to obtain a final concentration of about 5.0 × 106 cells mL−1. Zeta potential was measured using a ZetaSizer Nano ZS (Malvern Instruments, Worcestershire, UK). All the determinations were performed in a clear disposable zeta cell at approximately 25°C. Mean values of each studied suspension were obtained by at least triplicate measurements of two independent experiments.

Surface contact angle measurements

In the last day of each culture, microalgal suspensions were harvested, washed twice and resuspended in saline solution (0.85% w/v NaCl) to obtain a final concentration of about 5.0 × 106 cells mL−1. Algal lawns were prepared by filtering the previously washed suspensions using 0.45-μm nitrocellulose membrane filters (Advantec MFS, Inc., Tokyo, Japan) until complete clogging of the membranes. Contact angle measurements were performed using the sessile drop method, as described by Busscher et al. [35]. The measurements were carried out at room temperature using water, formamide and α-bromonaphthalene (Sigma-Aldrich, Sintra, Portugal) as the reference liquids. Determination of contact angles was performed automatically using a model OCA 15 Plus (DataPhysics, Filderstadt, Germany) video-based optical contact angle measuring instrument, allowing image acquisition and data analysis. Contact angle measurements (at least 12 determinations for each liquid and for each algal suspension) were performed for two independent experiments.

Surface parameters and hydrophobicity determinations

After contact angle measurements, the values of surface hydrophobicity of the studied algal suspensions were determined using the approach of van Oss [36], which allows the assessment of the absolute degree of hydrophobicity of any surface in comparison with their interaction with water. In this approach, the degree of hydrophobicity of a given surface (s) is expressed as the free energy of hydrophobic interaction between two entities of that surface when immersed in water (w): \( \varDelta {G}_{sws}^{\mathrm{TOT}} \), in mJ m−2. When \( \varDelta {G}_{sws}^{\mathrm{TOT}}<0 \), the interaction between the two entities is stronger than the interaction of each entity with water and the material is considered hydrophobic. Alternatively, if \( \varDelta {G}_{sws}^{\mathrm{TOT}}>0 \), the material is hydrophilic. \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) can be calculated through the surface tension components of the interacting entities, according to Equation 1 [27-29]:
$$ \varDelta {G}_{sws}^{\mathrm{TOT}}=-2{\left(\sqrt{\gamma_s^{\mathrm{LW}}}-\sqrt{\gamma_w^{\mathrm{LW}}}\right)}^2+4\left(\sqrt{\gamma_s^{+}{\gamma}_w^{-}}+\sqrt{\gamma_s^{-}{\gamma}_w^{+}}-\sqrt{\gamma_s^{+}{\gamma}_s^{-}}-\sqrt{\gamma_w^{+}{\gamma}_w^{-}}\right) $$
(1)

where γ LW accounts for the Lifshitz-van der Waals (LW) component of the surface free energy and γ + and γ are the electron acceptor and electron donor parameters, respectively, of the Lewis acid-base component (γ AB), being \( {\gamma}^{\mathrm{AB}}=2\sqrt{\gamma^{+}{\gamma}^{-}} \). LW forces, usually attractive, result from instantaneous asymmetrical distribution of electrons in molecules (the higher the value of LW component, the more apolar is the surface and, therefore, the lower would be its affinity for polar liquids) [37]. Electron donor and acceptor parameters give information about the molecules present in the studied surface: higher γ + indicates the presence of positively charged molecules and higher γ indicates the presence of negatively charged molecules [36]. Acid-base (AB) forces result from electron transfer interactions between polar components of the involved surfaces. These interactions can be attractive (hydrophobic attraction) or repulsive (hydrophilic repulsion), depending on the free energy of hydrophobic interaction [37,38].

The surface tension components of a surface (s) were obtained by measuring the contact angles of three pure liquids (l), water and formamide (both polar) and α-bromonaphthalene (apolar), followed by the simultaneous resolution of three equations of the form Equation 2:
$$ \left(1+ \cos \theta \right){\gamma}_l^{\mathrm{TOT}}=2\left(\sqrt{\gamma_s^{\mathrm{LW}}{\gamma}_l^{\mathrm{LW}}}+\sqrt{\gamma_s^{+}{\gamma}_l^{-}}+\sqrt{\gamma_s^{-}{\gamma}_l^{+}}\right) $$
(2)

where θ is the contact angle and γ TOT = γ LW + γ AB. The surface tension of liquid components was obtained from the literature [39]. The contact angle of the apolar liquid, α-bromonaphthalene, was used to quantify the apolar energy component \( {\gamma}_s^{\mathrm{LW}} \), since \( {\gamma}_l^{-} \) and \( {\gamma}_l^{+} \) for this probe liquid are equal to zero. On the other hand, contact angles measured with the other probe liquids were used to determine the other surface parameters, \( {\gamma}_s^{+} \) and \( {\gamma}_s^{-} \).

Sedimentation kinetics

Microalgal recovery

In the last day of each culture, microalgal suspensions were also used to evaluate sedimentation kinetics. 1-mL samples were allowed to settle in polystyrene cuvettes at room temperature. Microalgal recovery within the settling period (8 h) was determined by measuring the turbidity (optical density (OD)) of the samples at 750 nm using a V-1200 spectrophotometer (VWR, Carnaxide, Portugal), according to the method proposed by Salim et al. [40]. Turbidity of the samples was measured at the same height in the cuvette. Microalgal removal percentage or microalgal recovery was calculated according to Equation 3 [40]:
$$ \%R=\frac{{\mathrm{OD}}_{750}\left({t}_0\right)-{\mathrm{OD}}_{750}(t)}{{\mathrm{OD}}_{750}\left({t}_0\right)}\cdot 100 $$
(3)

where OD750(t 0) is the turbidity of the sample determined in the beginning of microalgal settling and OD750(t) is the turbidity of the sample determined at time t.

Modelling microalgal sedimentation

Kinetic constants associated to the recovery of microalgal biomass were determined by fitting the experimental data, corresponding to the time-course evolution of microalgal removal percentages, to the Gompertz model, expressed by Equation 4 [41]:
$$ y=a\cdot \exp \left(- \exp \left(b-ct\right)\right) $$
(4)

where y is the output value or microalgal removal percentage (%), a is the upper asymptote (%), b (b > 0) sets the displacement along the x-axis and corresponds to the lag time observed in the beginning of settling experiments (h) and c (c > 0) sets the tangent at the inflection point and corresponds to the sedimentation rate (h−1). The kinetic parameters, a, b and c, were determined by minimizing the sum of squared residuals using the Solver supplement of Microsoft Excel 2013. The quality of the model fit was evaluated through analysis of the coefficient of determination (R 2).

Statistical analysis

Contact angles, surface tension parameters, zeta potential values and microalgal removal percentages were analysed using paired-samples t-test from the statistical software SPSS 22.0 (SPSS Inc., Chicago, IL, USA). Statistical tests were carried out at a significance level of 0.05.

Results and discussion

Surface physicochemical properties

The surface physicochemical properties determined using the approach of van Oss et al. [27-29] for the studied suspensions are shown in Table 1. Data is divided into single and mixed cultures, and the results are presented in the decreasing order of free energy of hydrophobic interaction (from the most hydrophilic to the most hydrophobic suspensions). Except for the microalga C. vulgaris and for the suspensions P. subcapitata + M. aeruginosa and P. subcapitata + S. salina, the contact angles measured with the polar probe liquids (water and formamide) were in agreement with the values of free energy of hydrophobic interaction, \( \varDelta {G}_{sws}^{\mathrm{TOT}} \). Higher contact angles were measured for microorganisms presenting more hydrophobic surfaces (lower \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) values). On the other hand, contact angles measured with the probe liquid α-bromonaphthalene for the different algal suspensions were not statistically different (p > 0.05).
Table 1

Contact angles and surface physicochemical properties determined for each of the studied suspensions

Microalgal suspensions

Contact angles (°)

Surface tension parameters and free energy of hydrophobic interaction (mJ m −2 )

Zeta potential (mV)

θ W

θ B

θ F

\( {\boldsymbol{\gamma}}_{\boldsymbol{s}}^{\mathbf{LW}} \)

\( {\boldsymbol{\gamma}}_{\boldsymbol{s}}^{\mathbf{AB}} \)

\( {\boldsymbol{\gamma}}_{\boldsymbol{s}}^{-} \)

\( {\boldsymbol{\gamma}}_{\boldsymbol{s}}^{+} \)

\( \varDelta {\boldsymbol{G}}_{\boldsymbol{sws}}^{\mathbf{TOT}} \)

Single cultures

         

C. vulgaris

46.5 ± 2.8 a

37.1 ± 1.5 a

61.0 ± 0.3 a

35.0 ± 1.5 a

0.0 ± 0.0 a

48.7 ± 6.9 a

0.1 ± 0.1 a

52.5 ± 14.3 a

−35.4 ± 0.4 a

M. aeruginosa

44.8 ± 5.5 a

37.1 ± 1.6 a

36.4 ± 8.4 b

35.9 ± 0.7 a

10.7 ± 2.8 b

57.4 ± 7.6 a

0.5 ± 0.4 a

43.4 ± 15.5 a

−40.0 ± 2.4 a

P. subcapitata

48.7 ± 1.1 a

37.3 ± 2.2 a

45.8 ± 3.6 b

35.8 ± 1.0 a

5.4 ± 1.5 b

35.9 ± 1.7 b

0.2 ± 0.2 a

13.8 ± 3.8 b

−48.1 ± 0.9 b

S. salina

51.9 ± 8.0 a

37.3 ± 1.0 a

73.6 ± 1.1 a

35.8 ± 0.4 a

0.0 ± 0.0 a

22.6 ± 3.2 b

0.0 ± 0.0 a

−10.2 ± 6.6 b

−43.1 ± 2.4 b

Mixed cultures

         

C. vulgaris + M. aeruginosa

41.9 ± 2.8 a

34.2 ± 4.4 a

51.6 ± 3.6 c

37.0 ± 1.8 a

0.5 ± 0.3 a

53.0 ± 8.9 a

0.0 ± 0.0 a

40.6 ± 11.8 a

−35.4 ± 1.0 a

C. vulgaris + S. salina

76.8 ± 4.2 b

35.3 ± 2.1 a

57.0 ± 3.1 c

36.6 ± 0.8 a

0.0 ± 0.0 a

52.2 ± 1.5 a

0.0 ± 0.0 a

40.1 ± 2.5 a

−45.1 ± 1.4 b

P. subcapitata + M. aeruginosa

41.9 ± 2.8 a

37.3 ± 0.9 a

35.5 ± 1.5 d

35.8 ± 0.4 a

11.7 ± 0.8 b

37.0 ± 4.8 b

0.9 ± 0.3 a

13.4 ± 7.2 b

−42.7 ± 1.4 b

C. vulgaris + P. subcapitata

57.0 ± 0.1 a

39.6 ± 0.1 a

56.0 ± 0.2 c

34.0 ± 0.1 a

0.0 ± 0.0 a

32.7 ± 0.3 b

0.0 ± 0.0 a

10.5 ± 0.5 b

−42.7 ± 1.5 b

P. subcapitata + S. salina

54.2 ± 1.9 a

37.6 ± 0.3 a

50.7 ± 1.1 c

35.6 ± 0.1 a

2.8 ± 0.5 c

32.0 ± 1.9 b

0.1 ± 0.0 a

8.3 ± 3.3 b

−40.5 ± 0.5 b

S. salina + M. aeruginosa

93.5 ± 3.3 c

35.0 ± 3.2 a

84.0 ± 1.3 a

36.7 ± 1.3 a

0.0 ± 0.0 a

7.6 ± 2.2 c

0.0 ± 0.0 a

−50.4 ± 8.6 c

−34.9 ± 1.0 a

Values are presented as the mean ± standard deviation of two independent experiments. Different letters within the same column represent statistically different values (p < 0.05). θ W, contact angle with water; θ B, contact angle with α-bromonaphthalene; θ F, contact angle with formamide; \( {\gamma}_s^{\mathrm{LW}} \), Lifshitz-van der Waals component of the surface free energy; \( {\gamma}_s^{\mathrm{AB}} \), Lewis acid-base component of the surface free energy; \( {\gamma}_s^{-} \), electron donor component; \( {\gamma}_s^{+} \), electron acceptor component; \( \varDelta {G}_{sws}^{\mathrm{TOT}} \), free energy of hydrophobic interaction.

Looking at electron donor and acceptor parameters, the values determined for the studied microorganisms ranged from 22.6 ± 3.2 to 57.4 ± 7.6 mJ m−2 and from 0.0 ± 0.0 to 0.5 ± 0.4 mJ m−2, respectively. For the studied consortia, \( {\gamma}_s^{-} \) and \( {\gamma}_s^{+} \) values ranged from 7.6 ± 2.2 to 53.0 ± 8.9 mJ m−2 and from 0.0 ± 0.0 to 0.9 ± 0.3 mJ m−2, respectively. These components were not statistically different from those obtained for single cultures (p = 0.61 and p = 0.99, respectively). Higher values determined for \( {\gamma}_s^{-} \) indicate that the studied suspensions are electron donors [42] and may be attributed to the presence of excessive molecules of oxygen in the surface of the microorganisms and to the neutralization of \( {\gamma}_s^{+} \) sites by the dominant \( {\gamma}_s^{-} \) sites through intermolecular and intramolecular interactions [36]. On the other hand, \( {\gamma}_s^{+} \) values close to zero are in accordance with the van Oss et al. [27-29] model, which predicts almost the non-existence of electron acceptor parameters [43]. According to Volpe and Siboni [43], the assumption of a monopolar surface is unrealistic. However, the authors concluded that the van Oss et al. [27-29] approach can be applied when the aim is to compare the ability of different microorganisms to interact with other microorganisms or surfaces. When the aim is to perform molecular interpretations, these parameters (determined through the van Oss et al. [27-29] method) should not be used. Additionally, plotting \( {\gamma}_s^{-} \) as a function of \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) resulted in a coefficient of determination (R 2) of 0.94, indicating that the van Oss et al. [27-29] approach can be correctly applied to determine the surface physicochemical properties of the selected algal suspensions. As for the contact angle measurements with the polar probe liquids, \( {\gamma}_s^{-} \) values correlated well with the free energy of hydrophobic interaction, since lower values of the electron donor parameter were observed for the microorganisms presenting lower \( \varDelta {G}_{sws}^{\mathrm{TOT}} \).

Regarding the Lifshitz-van der Waals component of the surface free energy, \( {\gamma}_s^{\mathrm{LW}} \), the values determined for the studied species were not statistically different (p > 0.05), ranging from 35.0 ± 1.5 to 35.9 ± 0.7 mJ m−2. Similar results determined for this component indicate that cell wall composition of the studied microorganisms may not have significant differences. In the study performed by Ozkan and Berberoglu [44], differences in the LW component determined for green algae and diatoms were attributed to differences in the chemical composition of their cell walls. Several authors have reported that green algae, such as C. vulgaris and P. subcapitata, present cellulose-based cell walls, whereas diatoms present silica-based ones [45-48]. \( {\gamma}_s^{\mathrm{LW}} \) values determined in this study for microalgae were very similar to those determined by Ozkan and Berberoglu [44] for green algae (ranging from 20.7 to 37.8 mJ m−2). Additionally, \( {\gamma}_s^{\mathrm{LW}} \) values determined for the studied cyanobacteria were also similar to those obtained for the cyanobacteria Synechocystis sp. and Anabaena variabilis (28.3 and 37.0 mJ m−2, respectively). \( {\gamma}_s^{\mathrm{LW}} \) values determined for the different combinations of microalgae and cyanobacteria ranged from 34.0 ± 0.1 to 37.0 ± 1.8 mJ m−2. These values were not statistically different (p = 0.24) from those determined for each individual strain, and in addition, no statistical differences (p > 0.05) were observed between the different studied consortia. As previously stated, similar results determined for this component indicate that cell wall composition of the studied microorganisms, even when grown in mixed cultures, may not have significant differences.

Values of \( {\gamma}_s^{\mathrm{AB}} \) determined for the studied microalgae ranged from 0.0 ± 0.0 to 5.4 ± 1.5 mJ m−2, whereas the same value determined for the cyanobacteria ranged from 0.0 ± 0.0 to 10.7 ± 2.8 mJ m−2. The values obtained for microalgae were similar to those obtained by Ozkan and Berberoglu [44] for green algae (ranging from 0.0 to 5.1 mJ m−2). Regarding the values obtained by the authors for cyanobacteria, \( {\gamma}_s^{\mathrm{AB}} \) determined in this study for the cyanobacterium S. salina was equal to the one reported for Synechocystis sp. A \( {\gamma}_s^{\mathrm{AB}} \) of 5.4 ± 1.5 mJ m−2 determined for the cyanobacterium M. aeruginosa, which is statistically higher than the one determined for S. salina (p = 0.01) may be related to the hydrophilic character of this cyanobacterium (\( \varDelta {G}_{sws}^{\mathrm{TOT}}>0 \)). The \( {\gamma}_s^{\mathrm{AB}} \) values determined for the microalgae and for the cyanobacterium M. aeruginosa are a measure of the hydrophilic repulsion, since these microorganisms presented a hydrophilic surface. On the other hand, \( {\gamma}_s^{\mathrm{AB}} \) determined for S. salina is a measure of hydrophobic attraction, due to the negative value of \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) determined for this cyanobacterium. The magnitude of AB interactions increases with increasing surface hydrophobicity (decreasing \( {\gamma}_s^{+} \) and \( {\gamma}_s^{-} \)), and species having hydrophilic surface properties experience repulsive acid-base interactions [49]. \( {\gamma}_s^{\mathrm{AB}} \) values determined for the studied consortia were not statistically different (p = 0.64) from those determined for microorganisms grown in single cultures, ranging from 0.0 ± 0.0 to 11.7 ± 0.8 mJ m−2. Since positive \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) values were determined for all studied consortia, except for the one composed by both cyanobacteria, \( {\gamma}_s^{\mathrm{AB}} \) values are a measure of hydrophilic repulsion. In the mixed culture of both cyanobacteria, \( {\gamma}_s^{\mathrm{AB}} \) indicates the degree of hydrophobic attraction.

The values determined for the free energy of hydrophobic interaction have demonstrated that the microalgae and the cyanobacterium M. aeruginosa presented hydrophilic surfaces, while the surface of S. salina was considered hydrophobic. According to the determined values of \( \varDelta {G}_{sws}^{\mathrm{TOT}} \), the microorganisms can be listed by increased degree of hydrophobicity, as follows: C. vulgaris, M. aeruginosa, P. subcapitata, and S. salina. However, no statistical differences (p = 0.60) were observed in \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) values determined for C. vulgaris and M. aeruginosa (52.5 ± 14.3 and 43.4 ± 15.5 mJ m−2, respectively). Negative values of \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) determined for S. salina suggest the potential of this microorganism to form flocs and, therefore, improve harvesting processes and also its ability to grow as sessile cells, which may have an application in biofilm formation for wastewater treatment and metabolite production. In mixed cultures, these values ranged from −50.4 ± 8.6 to 40.6 ± 11.8 mJ m−2. Three distinct groups can be defined among the studied consortia: (i) the consortia composed of C. vulgaris + M. aeruginosa and of C. vulgaris + S. salina constitute the most hydrophilic group, with \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) values of about 40 mJ m−2; (ii) the consortia of P. subcapitata + M. aeruginosa, C. vulgaris + P. subcapitata and P. subcapitata + S. salina constitute the group presenting a slightly hydrophilic surface (\( \varDelta {G}_{sws}^{\mathrm{TOT}} \) values close to 10 mJ m−2); and (iii) the more hydrophobic group, represented by the consortium of S. salina + M. aeruginosa (\( \varDelta {G}_{sws}^{\mathrm{TOT}} \) of −50 mJ m−2). These results have shown that the co-culture of a more hydrophobic microorganism with another one presenting a more hydrophilic surface results in a decrease in free energy of hydrophobic interaction values and, hence, in an increase in the degree of hydrophobicity. For example, the co-culture of the hydrophobic cyanobacterium S. salina with C. vulgaris, M aeruginosa and P. subcapitata resulted in a reduction of \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) values from 52.5 ± 14.3 to 40.1 ± 2.5, 43.4 ± 15.5 to −50.4 ± 8.6 and 13.8 ± 3.8 to 8.3 ± 3.3 mJ m−2, respectively. The huge decrease of \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) observed for the co-culture of both cyanobacteria may be related to the high contact angles measured with polar probe liquids (θ W and θ F of 93.5° ± 3.3° and 84.0° ± 1.3°, respectively), which are considered unusual [44]. The co-culture of the most hydrophilic microorganisms (C. vulgaris + M. aeruginosa) resulted in a hydrophilic surface, with \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) of 40.6 ± 11.8 mJ m−2.

Regarding zeta potential values determined for the studied microorganisms, these values ranged from −35.4 ± 0.4 to −48.1 ± 0.9 mV. Zeta potential measurements give information about the charge of cell surfaces: negative or positive values of zeta potential depend on the functional groups present on cell surfaces and also on the pH of the culture medium [44]. Functional groups commonly found on cell surfaces include hydroxyl (-OH), carboxyl (-COOH) and amine (-NH2) [44,50,51]. Since these groups are ionisable, when cells are exposed to low pH values, functional groups are protonated and net surface charge becomes positive, and on the other hand, when cells are exposed to high pH values, functional groups are deprotonated and the resulting net surface charge is negative. At the point of zero charge (PZC), corresponding to an intermediate pH, some groups are protonated while others are deprotonated and the surface charge is neutralized [52,53]. Negative values observed for the studied microalgae and cyanobacteria were expected, since the pH of the culture medium measured when the samples were collected was high (9.64 ± 0.65) and PZC reported for algae was approximately pH 3 [52], indicating that functional groups on the microorganisms’ surface were deprotonated. Additionally, net zeta potential values give information about suspensions’ stability. When absolute value of zeta potential is high, repulsive forces prevail over van der Waals forces, and hence, particles/cells are stable in the dispersed form. On the other hand, for low net zeta potential values, van der Waals forces (usually attractive) are higher than repulsive ones and the stability of the suspension is affected, resulting in the formation of aggregates and further settling [54,55]. Net zeta potential values determined for the studied microalgae and cyanobacteria are substantial, and therefore, it is expected that the microorganisms keep stable in suspension. These results are in accordance with the non-evidence of flocs formation in the studied cultures, especially in that of the hydrophobic cyanobacterium S. salina. Therefore, zeta potential measurements are very important for a better understanding of the interactions among microalgae and cyanobacteria. Although microorganisms may present hydrophobic surfaces, their ability to form aggregates or to attach to surfaces may be strongly affected by the ionic strength of the culture medium. As for single cultures, zeta potential values determined for the studied consortia were negative, indicating an overall surface charge negative. In fact, the pH of the culture medium in these cultures was about 9.50 ± 0.33, which is higher than the reported PZC for microalgae (pH 3). Therefore, functional groups on microorganisms’ surface may be deprotonated, conferring a negative surface charge. Zeta potential values determined ranged from −34.9 ± 1.0 to −45.1 ± 1.4 mV. These values were not statically different (p = 0.66) from those determined for single cultures, which may be related to similar ionic strengths of the culture medium, since both single and mixed cultures were cultured in OECD test medium [34] with the same composition. Additionally, no statistical difference (p > 0.05) was observed in zeta potential values for the studied microalgal associations. Net zeta potential values determined were also high, determining the stability of the suspensions and, therefore, the non-evidence of aggregates formation. This information may be very useful in the design of new strategies to control biofilm formation and development and to improve microalgal harvesting.

Determining the surface physicochemical properties of different consortia has shown that the co-culture of microorganisms with different degrees of hydrophobicity or hydrophilicity can alter the physicochemical properties of the single-cultured microorganisms. These findings can be very important in several applications, such as promoting aggregates formation and, therefore, bioflocculation and biofilm formation and, on the other hand, prevent and control biofouling. However, it is very important to take into account the ionic strength of the culture medium, since this parameter influences surface charge and, hence, the higher or lesser extent of electrostatic repulsions.

Sedimentation kinetics

Microalgal removal percentages determined for single and mixed cultures within the 8-h settling period are presented in Figure 1A and B, respectively. Analysis of these data indicates that sedimentation rates of the studied algal suspensions are very low. After 8 h of settling, maximum removal percentages achieved in single and mixed cultures were 86.6% ± 6.0% and 89.6% ± 2.2%, respectively. Additionally, as it is possible to see from Figure 1A and B, all microalgal suspensions presented a lag time, starting to sediment after, at least, 1 h of the beginning of the experiment. These results are in accordance with the zeta potential values determined for these suspensions. As it was referred in the ‘Surface physicochemical properties’ section, when absolute value of zeta potential is high, repulsive forces prevail over van der Waals forces, and hence, particles/cells are stable in the dispersed form. Similar removal percentages have been reported by Salim et al. [40] and Manheim and Nelson [56] when harvesting microalgae through bioflocculation. Higher removal percentages can be achieved using chemical coagulants/flocculants. In the study performed by Papazi et al. [57], chemical coagulants, such as aluminium and ferric sulphate, were used to harvest the microalga Chlorella minutissima, resulting in a removal percentage of 80% after 2 to 4 h of settling. In the study performed by Garzon-Sanabria et al. [58], after 1 h of settling, approximately 90% of the microalga Nannochloris oculata was recovered from the culture medium. These authors used aluminium chloride as coagulant.
https://static-content.springer.com/image/art%3A10.1186%2Fs40643-015-0051-y/MediaObjects/40643_2015_51_Fig1_HTML.gif
Figure 1

Microalgal removal percentages determined within the settling period for each of the studied suspensions. (A) Single cultures and (B) Mixed cultures. Error bars correspond to the standard deviation of two independent experiments. Dashed lines represent the model fit of the Gompertz model to the experimental data.

Relationship between surface physicochemical properties and microalgal settling

Table 2 presents microalgal removal percentages determined after 8 h of settling and the kinetic parameters obtained through the model fit of the Gompertz model to the experimental data. Looking at R 2 values determined through the model fit (values close to 1) and to the model curves superimposed to the experimental data (in Figure 1A and B), it is possible to conclude that the Gompertz model can be correctly applied to describe microalgal recovery in the studied conditions. The sedimentation kinetic parameters determined through this model confirm the low settleability of the studied microalgal suspensions, which may be related to the high net values determined for zeta potential. According to these data, lag times, b, observed in the beginning of the sedimentation process were high, ranging from 1.0 to 3.0 h for single cultures and from 1.7 to 4.0 h for mixed cultures. Additionally, low sedimentation rates, c, were determined for both single and mixed cultures: 0.33 to 0.63 h−1 and 0.24 to 0.94 h−1, respectively. Although microalgal recovery using these suspensions was not effective, the values determined through contact angle measurements can be correlated with sedimentation results. From Tables 1 and 2, it is possible to observe that, in general, an increase in the degree of hydrophobicity (lower \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) values) corresponds to an increase in microalgal removal percentages, % R, and in sedimentation rates, c. Higher removal percentages for the most hydrophobic surfaces were expected since, as it was stated in the ‘Surface parameters and hydrophobicity determinations’ section, when \( \varDelta {G}_{sws}^{\mathrm{TOT}} \) decreases, the interaction between the two entities is stronger than the interaction of each entity with water. Therefore, particles in suspension tend to form aggregates, improving settling rates. In fact, plotting removal percentages determined after 8 h of settling as a function of the free energy of hydrophobic interaction suggests a negative relationship between these parameters. By excluding the values corresponding to the three most hydrophobic surfaces and to the most hydrophilic one, a linear relationship between microalgal removal percentage and free energy of hydrophobic interaction was obtained (R 2 = 0.91). These results show the importance of studying surface physicochemical properties to understand the flocculation process and improve harvesting procedures.
Table 2

Sedimentation kinetics determined for each of the studied suspensions

Microalgal suspensions

%R a

Gompertz model parameters

a (%)

b (h)

c (h −1 )

R 2

Single cultures

     

C. vulgaris

73.6 ± 5.2 a

100.0

2.0

0.35

0.990

M. aeruginosa

66.1 ± 4.8 a

100.0

2.8

0.33

0.965

P. subcapitata

86.6 ± 6.0 b

89.7

1.0

0.63

0.999

S. salina

85.0 ± 7.5 b

100.0

3.0

0.56

0.997

Mixed cultures

     

C. vulgaris + M. aeruginosa

58.6 ± 0.8 c

100.0

1.7

0.24

0.993

C. vulgaris + S. salina

69.6 ± 13.7 a

95.3

2.7

0.41

0.999

P. subcapitata + M. aeruginosa

85.2 ± 2.2 b

100.0

2.1

0.45

0.999

C. vulgaris + P. subcapitata

89.6 ± 2.2 b

91.8

2.2

0.94

1.000

P. subcapitata + S. salina

72.4 ± 8.7 a

82.1

1.7

0.38

0.998

S. salina + M. aeruginosa

86.1 ± 2.7 b

100.0

4.0

0.66

0.994

Microalgal removal percentage was determined after 8 h of settling, and recovery kinetic parameters were determined through the Gompertz model. aValues are presented as the mean ± standard deviation of two independent experiments. Different letters within the same column represent statistically different values (p < 0.05). % R, microalgal removal percentage determined after 8 h of settling; a, upper asymptote; b, displacement along the x-axis or lag time (b > 0); c, tangent at the inflection point or sedimentation rate (c > 0); R 2, coefficient of determination obtained through the model fit of Gompertz model to the experimental data.

Conclusions

This study presented a broad characterization of surface physicochemical properties of microalgae and cyanobacteria, as well as how these properties vary when these microorganisms are co-cultured. Regarding the surface properties of single cultures, S. salina presented a hydrophobic surface, suggesting the ability of this microorganism to form aggregates. However, macroscopic observations have shown no evidence of aggregates formation, which was confirmed through the study of sedimentation kinetics. This might be due to the high net zeta potential values determined, which are responsible for the stability of the suspensions in their dispersed form. The co-culture of different microalgae and cyanobacteria has shown that when a more hydrophobic microorganism is co-cultured with another one presenting a more hydrophilic surface, free energy of hydrophobic interaction tends to decrease, resulting in a more hydrophobic surface. This study has also shown a negative linear correlation between microalgal removal percentages and free energy of hydrophobic interaction, reinforcing the importance of surface physicochemical properties on the sedimentation process. Due to the low recovery efficiencies achieved, another factor to consider in the prevention or promotion of cell attachment is the ionic strength of the culture medium, since this parameter strongly influences surface charge.

Declarations

Acknowledgements

This work was funded by FEDER funds through the Operational Programme for Competitiveness Factors - COMPETE, ON.2 - O Novo Norte - North Portugal Regional Operational Programme and National Funds through FCT - Foundation for Science and Technology under the projects: PEst-C/EQB/UI0511, NORTE-07-0124-FEDER-000025 - RL2_Environment&Health, Novomar - ref. 0687-Novomar-1-P, SFRH/BD/88799/2012, FEUP-ON2-25MC-EngªQuimicaBiologica and SFRH/BPD/66721/2009. The authors would like to thank CIIMAR (Centre of Marine and Environmental Research of the University of Porto) for providing the cyanobacteria Synechocystis salina LEGE 06079 and Microcystis aeruginosa LEGE 91344.

Authors’ Affiliations

(1)
LEPABE, Departamento de Engenharia Química, Faculdade de Engenharia, Universidade do Porto

References

  1. Brennan L, Owende P (2010) Biofuels from microalgae - a review of technologies for production, processing, and extractions of biofuels and co-products. Renew Sust Energ Rev 14:557–77View ArticleGoogle Scholar
  2. Demirbas A (2011) Biodiesel from oilgae, biofixation of carbon dioxide by microalgae: a solution to pollution problems. Appl Energy 88:3541–7View ArticleGoogle Scholar
  3. Bilanovic D, Andargatchew A, Kroeger T, Shelef G (2009) Freshwater and marine microalgae sequestering of CO2 at different C and N concentrations - response surface methodology analysis. Energy Conv Manag 50:262–7View ArticleGoogle Scholar
  4. Gonçalves AL, Simões M, Pires JCM (2014) The effect of light supply on microalgal growth, CO2 uptake and nutrient removal from wastewater. Energy Conv Manag 85:530–6View ArticleGoogle Scholar
  5. Ho S-H, Chen C-Y, Lee D-J, Chang J-S (2011) Perspectives on microalgal CO2-emission mitigation systems - a review. Biotechnol Adv 29:189–98View ArticleGoogle Scholar
  6. Sayre R (2010) Microalgae: the potential for carbon capture. Bioscience 60:722–7View ArticleGoogle Scholar
  7. Boelee N, Temmink H, Janssen M, Buisman C, Wijffels R (2011) Nitrogen and phosphorus removal from municipal wastewater effluent using microalgal biofilms. Water Res 45:5925–33View ArticleGoogle Scholar
  8. Park JBK, Craggs RJ, Shilton AN (2011) Wastewater treatment high rate algal ponds for biofuel production. Bioresour Technol 102:35–42View ArticleGoogle Scholar
  9. Rawat I, Ranjith Kumar R, Mutanda T, Bux F (2011) Dual role of microalgae: phycoremediation of domestic wastewater and biomass production for sustainable biofuels production. Appl Energy 88:3411–24View ArticleGoogle Scholar
  10. Silva-Benavides A, Torzillo G (2012) Nitrogen and phosphorus removal through laboratory batch cultures of microalga Chlorella vulgaris and cyanobacterium Planktothrix isothrix grown as monoalgal and as co-cultures. J Appl Phycol 24:267–76View ArticleGoogle Scholar
  11. Hu Q (2004) Industrial production of microalgal cell-mass and secondary products - major industrial species. In: Richmond A (ed) Handbook of microalgal culture: biotechnology and applied phycology. Blackwell Science Ltd., Oxford, UK, pp 268–71Google Scholar
  12. Singh S, Kate B, Banerjee U (2005) Bioactive compounds from cyanobacteria and microalgae: an overview. Crit Rev Biotechnol 25:73–95View ArticleGoogle Scholar
  13. Spolaore P, Joannis-Cassan C, Duran E, Isambert A (2006) Commercial applications of microalgae. J Biosci Bioeng 101:87–96View ArticleGoogle Scholar
  14. Molina Grima E, Belarbi EH, Acién Fernández FG, Robles Medina A, Chisti Y (2003) Recovery of microalgal biomass and metabolites: process options and economics. Biotechnol Adv 20:491–515View ArticleGoogle Scholar
  15. Lardon L, Hélias A, Sialve B, Steyer J-P, Bernard O (2009) Life-cycle assessment of biodiesel production from microalgae. Environ Sci Technol 43:6475–81View ArticleGoogle Scholar
  16. Ozkan A, Berberoglu H (2013) Adhesion of algal cells to surfaces. Biofouling 29:469–82View ArticleGoogle Scholar
  17. Posten C (2009) Design principles of photo-bioreactors for cultivation of microalgae. Eng Life Sci 9:165–77View ArticleGoogle Scholar
  18. Pulz O (2001) Photobioreactors: production systems for phototrophic microorganisms. Appl Microbiol Biotechnol 57:287–93View ArticleGoogle Scholar
  19. Irving TE, Allen DG (2011) Species and material considerations in the formation and development of microalgal biofilms. Appl Microbiol Biotechnol 92:283–94View ArticleGoogle Scholar
  20. Roeselers G, Van Loosdrecht M, Muyzer G (2008) Phototrophic biofilms and their potential applications. J Appl Phycol 20:227–35View ArticleGoogle Scholar
  21. Finlay JA, Bennett SM, Brewer LH, Sokolova A, Clay G, Gunari N et al (2010) Barnacle settlement and the adhesion of protein and diatom microfouling to xerogel films with varying surface energy and water wettability. Biofouling 26:657–66View ArticleGoogle Scholar
  22. Ista LK, Callow ME, Finlay JA, Coleman SE, Nolasco AC, Simons RH et al (2004) Effect of substratum surface chemistry and surface energy on attachment of marine bacteria and algal spores. Appl Environ Microbiol 70:4151–7View ArticleGoogle Scholar
  23. Mercer P, Armenta RE (2011) Developments in oil extraction from microalgae. Eur J Lipid Sci Technol 113:539–47View ArticleGoogle Scholar
  24. Christenson L, Sims R (2011) Production and harvesting of microalgae for wastewater treatment, biofuels, and bioproducts. Biotechnol Adv 29:686–702View ArticleGoogle Scholar
  25. Uduman N, Qi Y, Danquah MK, Forde GM, Hoadley A (2010) Dewatering of microalgal cultures: a major bottleneck to algae-based fuels. J Renew Sustain Energy 2:012701–15View ArticleGoogle Scholar
  26. Vandamme D, Foubert I, Muylaert K (2013) Flocculation as a low-cost method for harvesting microalgae for bulk biomass production. Trends Biotechnol 31:233–9View ArticleGoogle Scholar
  27. van Oss C, Chaudhury M, Good R (1987) Monopolar surfaces. Adv Colloid Interface Sci 28:35–64View ArticleGoogle Scholar
  28. van Oss C, Ju L, Chaudhury M, Good R (1989) Estimation of the polar parameters of the surface tension of liquids by contact angle measurements on gels. J Colloid Interface Sci 128:313–9View ArticleGoogle Scholar
  29. van Oss CJ, Good RJ, Chaudhury MK (1988) Additive and nonadditive surface tension components and the interpretation of contact angles. Langmuir 4:884–91View ArticleGoogle Scholar
  30. Chinnasamy S, Ramakrishnan B, Bhatnagar A, Das KC (2009) Biomass production potential of a wastewater alga Chlorella vulgaris ARC 1 under elevated levels of CO2 and temperature. Int J Mol Sci 10:518–32View ArticleGoogle Scholar
  31. McLarnon-Riches CJ, Rolph CE, Greenway DL, Robinson PK (1998) Effects of environmental factors and metals on Selenastrum capricornutum lipids. Phytochemistry 49:1241–7View ArticleGoogle Scholar
  32. Philippis R, Vincenzini M (1998) Exocellular polysaccharides from cyanobacteria and their possible applications. FEMS Microbiol Rev 22:151–75View ArticleGoogle Scholar
  33. Wahlen BD, Willis RM, Seefeldt LC (2011) Biodiesel production by simultaneous extraction and conversion of total lipids from microalgae, cyanobacteria, and wild mixed-cultures. Bioresour Technol 102:2724–30View ArticleGoogle Scholar
  34. OECD (2011) Freshwater alga and cyanobacteria, growth inhibition test. In: Test Guideline 201: Organisation for Economic Co-operation and DevelopmentGoogle Scholar
  35. Busscher H, Weerkamp A, van der Mei H, van Pelt A, de Jong H, Arends J (1984) Measurement of the surface free energy of bacterial cell surfaces and its relevance for adhesion. Appl Environ Microbiol 48:980–3Google Scholar
  36. van Oss C (1995) Hydrophobicity of biosurfaces - origin, quantitative determination and interaction energies. Colloids Surf B Biointerfaces 5:91–110View ArticleGoogle Scholar
  37. Ozkan A, Berberoglu H (2013) Cell to substratum and cell to cell interactions of microalgae. Colloids Surf B Biointerfaces 112:302–9View ArticleGoogle Scholar
  38. van Oss CJ (2003) Long-range and short-range mechanisms of hydrophobic attraction and hydrophilic repulsion in specific and aspecific interactions. J Mol Recognit 16:177–90View ArticleGoogle Scholar
  39. Janczuk B, Chibowski E, Bruque J, Kerkeb M, Caballero FG (1993) On the consistency of surface free energy components as calculated from contact angles of different liquids: an application to the cholesterol surface. J Colloid Interface Sci 159:421–8View ArticleGoogle Scholar
  40. Salim S, Bosma R, Vermuë MH, Wijffels RH (2011) Harvesting of microalgae by bio-flocculation. J Appl Phycol 23:849–55View ArticleGoogle Scholar
  41. Gompertz B (1825) On the nature of the function expressive of the law of human mortality, and on a new mode of determining the value of life contingencies. Philos Trans Royal Soc London 115:513–83View ArticleGoogle Scholar
  42. Simões LC, Simões M, Oliveira R, Vieira MJ (2007) Potential of the adhesion of bacteria isolated from drinking water to materials. J Basic Microbiol 47:174–83View ArticleGoogle Scholar
  43. Volpe CD, Siboni S (1997) Some reflections on acid–base solid surface free energy theories. J Colloid Interface Sci 195:121–36View ArticleGoogle Scholar
  44. Ozkan A, Berberoglu H (2013) Physico-chemical surface properties of microalgae. Colloids Surf B Biointerfaces 112:287–93View ArticleGoogle Scholar
  45. Becker E (2007) Microalgae as a source of protein. Biotechnol Adv 25:207–10View ArticleGoogle Scholar
  46. Latała A, Nędzi M, Stepnowski P (2009) Toxicity of imidazolium and pyridinium based ionic liquids towards algae: Chlorella vulgaris, Oocystis submarina (green algae) and Cyclotella meneghiniana, Skeletonema marinoi (diatoms). Green Chem 11:580–8View ArticleGoogle Scholar
  47. Loos E, Meindl D (1982) Composition of the cell wall of Chlorella fusca. Planta 156:270–3View ArticleGoogle Scholar
  48. Northcote D, Goulding K, Horne R (1958) The chemical composition and structure of the cell wall of Chlorella pyrenoidosa. Biochem J 70:391Google Scholar
  49. Bos R, Mei HC, Busscher HJ (1999) Physico-chemistry of initial microbial adhesive interactions - its mechanisms and methods for study. FEMS Microbiol Rev 23:179–230Google Scholar
  50. Ferreira C, Pereira A, Pereira M, Melo L, Simões M (2011) Physiological changes induced by the quaternary ammonium compound benzyldimethyldodecylammonium chloride on Pseudomonas fluorescens. J Antimicrob Chemother 66:1036–43View ArticleGoogle Scholar
  51. González-Fernández C, Ballesteros M (2012) Microalgae autoflocculation: an alternative to high-energy consuming harvesting methods. J Appl Phycol 25:1–9Google Scholar
  52. Hadjoudja S, Deluchat V, Baudu M (2010) Cell surface characterisation of Microcystis aeruginosa and Chlorella vulgaris. J Colloid Interface Sci 342:293–9View ArticleGoogle Scholar
  53. Stumm W, Morgan JJ (2012) Aquatic chemistry: chemical equilibria and rates in natural waters. John Wiley & Sons, New JerseyGoogle Scholar
  54. de Schryver P, Crab R, Defoirdt T, Boon N, Verstraete W (2008) The basics of bio-flocs technology: the added value for aquaculture. Aquaculture 277:125–37View ArticleGoogle Scholar
  55. Zita A, Hermansson M (1994) Effects of ionic strength on bacterial adhesion and stability of flocs in a wastewater activated sludge system. Appl Environ Microbiol 60:3041–8Google Scholar
  56. Manheim D, Nelson Y (2013) Settling and bioflocculation of two species of algae used in wastewater treatment and algae biomass production. Environ Prog Sustain Energy 32:946–54View ArticleGoogle Scholar
  57. Papazi A, Makridis P, Divanach P (2010) Harvesting Chlorella minutissima using cell coagulants. J Appl Phycol 22:349–55View ArticleGoogle Scholar
  58. Garzon-Sanabria AJ, Davis RT, Nikolov ZL (2012) Harvesting Nannochloris oculata by inorganic electrolyte flocculation: effect of initial cell density, ionic strength, coagulant dosage, and media pH. Bioresour Technol 118:418–24View ArticleGoogle Scholar

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© Gonçalves et al.; licensee Springer. 2015

This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly credited.

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