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Influence of exopolysaccharides on the electrophoretic properties of the model cyanobacterium Synechocystis
Colloids and Surfaces B: Biointerfaces 110 (2013) 171–177
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Colloids and Surfaces B: Biointerfaces journal homepage: www.elsevier.com/locate/colsurfb
Influence of exopolysaccharides on the electrophoretic properties of the model cyanobacterium Synechocystis Mariane Planchon a,b , Thichakorn Jittawuttipoka c , Corinne Cassier-Chauvat c , Franc¸ois Guyot d , Franck Chauvat c , Olivier Spalla a,∗ a
CEA Saclay, DSM/IRAMIS/SIS2M/LIONS, UMR CEA-CNRS 3299, 91191 Gif sur Yvette, France Université Paris Diderot, Sorbonne Paris Cité, IPGP, UMR CNRS 7154, Paris, France c CEA Saclay, DSV/IBITEC-S/SB2SM, UMR 8221, 91191 Gif sur Yvette, France d IMPMC, UMR CNRS-Université Pierre et Marie Curie-Université Paris Diderot, 75005 Paris, France b
a r t i c l e
i n f o
Article history: Received 3 January 2013 Received in revised form 26 March 2013 Accepted 29 March 2013 Available online 25 April 2013 Keywords: Extracellular polymeric substance Electrophoretic mobility Soft particle electrophoresis theory Cyanobacteria Synechocystis
a b s t r a c t The influence of extracellular polymeric substances (EPS) on cell electrokinetics was investigated in the model cyanobacterium Synechocystis, in wild-type strains and in ten EPS-depleted mutants. The charge density and the softness of the EPS polyelectrolyte layer were calculated from the dependence of the electrophoretic mobility values of the cells with the ionic strength of the surrounding fluid. Electrophoretic mobility data showed that the eleven Synechocystis strains investigated behave as soft particles and cannot be adequately described by classical electrokinetic models of rigid particles. EPS surrounding the cells, especially those released in the growth medium, significantly increased the softness of the cell surface. Furthermore, the anionic nature of EPS resulted in negative surface charge densities, which appeared to be strongly dependent on the composition of the suspending fluid, as documented by a strong reduction of their absolute values in the presence of calcium cations. These finding stresses the importance of the physicochemical properties of EPS and cell surfaces of cyanobacteria, for both cell-to-medium and cellto-cell communications. In turn, these results emphasize that, whenever possible, natural waters should be used for meaningful ecotoxicological analyses of potential toxics. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Bacteria constitute the most successful form of life on Earth. They colonize most biotopes and produce a huge biomass (5 × 1030 cells and 35–55 × 1016 g of carbon), which represents up to 60–100% of the estimated total carbon in plants [1,2]. The principal reasons for the success of bacteria are their physiological robustness as well as their abilities to rapidly respond and adapt to environmental stimuli [3]. Biofilm is the predominant mode of bacterial growth in most natural, industrial and clinical environments [4]. The biofilm lifestyle is associated with a high tolerance to environmental stresses, including treatments with antibiotics or other biocides. Hence, biofilm formation is a major concern in environmental and biotechnological applications. Biofilms typically consist of densely packed, multispecies populations, encased in a cell-synthesized polymeric matrix attached to a surface [5]. These microbial extracellular polymeric substances (EPS) are heterogeneous in complexity and composition, which varies depending on the microorganisms [6–8]. EPS contain soluble ions and colloidal
∗ Corresponding author. Tel.: +33 1 69 08 57 43. E-mail address: [email protected] (O. Spalla). 0927-7765/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.colsurfb.2013.03.057
insoluble matters and are mainly composed of polysaccharides (neutral and acidic), proteins, nucleic acids, lipids and other biological macromolecules [9]. They provide a highly hydrated gel matrix structure that mediates most of the cell-to-cell and cell-to-surface interactions, which are required for the formation and stabilization of biofilms. In EPS, charged groups may associate or dissociate upon changes in pH or ionic strength of the suspending fluid, or upon the approach of a new charged surface of another cell. Regarding their physicochemical properties, EPS can be assimilated to polyelectrolytes since this matrix is composed of long polymers whose gelation is promoted by multivalent cations [10]. The electrophoretic mobility of an object in a liquid is a macroscopic signature of its surface state in interaction with the surrounding fluid and it can be measured by different means. So, electrophoretic mobility is both a rapid and effective technique to assess the stability and behavior of particles in suspension. This analysis was initially developed for objects having a hard surface and has been very successful in hard colloids science. However, the classical electric double layer model is not appropriate for bacterial cells because their surfaces display complex structures. In hard colloids, the electric charges on the outer surface determine the electrophoretic mobility. However, the permeability of the bacterial surface layer to ions and molecules from the
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surrounding fluid strongly influences the mobility. In addition the mobility of bacteria is affected by the non homogeneous spatial distribution of the charged groups present at the cell surface. This is why Ohshima and co-workers [11] developed an alternative model, called the “soft particle model”, which is better suited to the analysis of bacterial cells, i.e. biological core particles surrounded by a ion-penetrable EPS layer that strongly influences electrophoretic mobility. This “soft particle electrophoresis theory” was successfully used in several studies of the cell surface of the enterobacterium Escherichia coli [12,33] and of other non-photosynthetic bacteria [13–18,34]. However, soft particle electrophoresis has not yet been employed to study the cell surface of cyanobacteria, the cosmopolitan photosynthetic prokaryotes that produce a large part of the terrestrial di-oxygen [19] and biomass for the food chain [20], and which have the potentials for the sustainable production of biofuels [7,21] and bioplastics [22], and for the bioremediation of polluted soils and waters [23]. Many cyanobacteria possess extracellular polymeric substances, mainly of polysaccharidic nature (EPS) [24], which protect cells against environmental stresses (UV, salt, desiccation, heavy metals) and enable them to form biofilms [13]. The cyanobacterial EPS, which are complex [23], can be divided into two main groups: the RPS (released polysaccharides), which are secreted by cells into the surrounding environment, and the CPS (capsular polysaccharides), which are attached to the cell surface [24]. In this study, we investigated the influence of EPS on the electrophoretic mobility of the widely used unicellular model strain Synechocystis PCC6803 (hereafter Synechocystis), which makes up complex anionic EPS that contain 13 different monosaccharides and various uronic acids [9,25]. For this purpose we used, and compared, the wild type strain (WT) and 10 EPS-depleted mutants that were recently constructed [26]. The electrophoretic mobilities of these 11 strains were analyzed in the framework of the Ohshima’s soft particle theory. We also studied the influence on electrophoretic mobility and cell surface parameters of the counter-ions nature in the bulk suspension, emphasizing the strong role of calcium. 2. Materials and methods 2.1. Bacterial strains and culture conditions The round unicellular cyanobacterium Synechocystis PCC6803 strains wild-type (WT) is considered in this work together with and 10 of its EPS-depleted mutants that were recently constructed the details of which are fully reported in [26]. In the present work, the bacteria were grown aerobically under shaking (180 rpm, Infors Multitron II) and white light (2500 lux; 31.25 E m−2 s−1 Mazda TF 16 W lamps) at 30 ◦ C on BG11 medium [27] enriched with 3.78 mM Na2 CO3 [28] thereafter referred to as MM for standard mineral medium (see composition on Supporting information S1). The main composition of Synechocystis PCC6803 EPS is detailed in Supporting Information S2. The EPS-depleted mutants were generated after the single or double deletion of the genes sll0923, sll1581, slr1875 and sll5052 that share sequence homology with EPS-production genes in non-photosynthetic bacteria [29,30]. For commodity, all four single deletion mutants are designated as X− (for instance 0923− stands for the deletion of sll0923) while all six double deletion mutants are noted X− Y− (for instance 0923− 1581− stands for the deletion of both sll0923 and sll1581). All EPS-depleted mutants were grown in the presence of selective antibiotics, which were added at the following concentrations: Streptomycin (Sm, 2.5 g/mL), Spectinomycin (Sp, 2.5 g/mL) and Kanamycin (Km, 50 g/mL) [28]. The growth of the various strains was followed as the timely increase in absorbance (Beckman DU 640 spectrophotometer) at 580 nm (1 OD580 unit corresponding to 2.5 × 107 cells/mL).
2.2. EPS characterization In order to disclose correlations between the content in EPS and the physicochemical properties, the EPS mass quantification data, extracted from a previous study [26] are expressed in mg EPS/m2 of cell surface instead of mg EPS/mg of cell protein mass. This unit, classical in physical chemistry, is more relevant for surface studies than a biological material normalization. The cell being almost spherical with a size of 1 m, the surface per cell is m2 . Moreover, we made the distinction between CPS and RPS mass quantities to better assess the modification of EPS layer structure and nature among the mutants. Indeed, the gene deletion can influence differently the production of CPS or RPS, which cannot be reflected in the total EPS mass quantification. The method of CPS/RPS titration is given in our former paper [26]. Scanning electron microscopy (SEM) was performed to obtain qualitative imaging of the EPS in the WT strain and in the different mutants. 72 h grown cultures of WT and mutant strains (OD580 = 0.7) were chemically fixed overnight with glutaraldehyde (1%) washed twice with UPW (milliQ water) and gradually dehydrated using a CO2 critical point dryer (BAL-TEC CPD030), as described in [31]. Dried samples were mounted on aluminum stubs using double-sided carbon tape and carbon-coated (Leica EM SCD 500). Cells were observed with a Zeiss Ultra 55 FEG SEM microscope operated at 2.0 kV at a working distance of 2.7 mm. Images were acquired in secondary electron mode using an Everhart Thornley detector. 2.3. Measurement of the electrophoretic mobility of the cells Following centrifugation, cells resuspended in the studied media at OD580 = 0.5 (2.5 × 107 cells/mL) were transferred into disposable folded capillary cells with gold covered electrodes (Malvern). The electrophoretic mobility (EPM) of the cells was measured at 25 ◦ C at a 17◦ fixed scattering angle with a Zetasizer Nano ZS instrument (Malvern) equipped with a 633 nm laser, using runs performed at a voltage of 150 V and a frequency of 285 Hz. To investigate the influence of the ionic strength of the medium, EPM measurements were performed with cells suspensions in various dilutions of MM in ultrapure water (100%, 80%, 60%, 40%, 20% and 0%) which lead to an ionic strength range of 1–26.6 mM. One can note that the absence of washing, in order to preserve the EPS structure around cells, leaded to a non-zero ionic strength when cells were resuspended in ultrapure water. All measurements were repeated five times and experiments were carried out in triplicate. 2.4. Modeling using soft-particle theory Electrophoresis is an effective technique to assess the electrical properties of microparticle surfaces but the Smoluchowski mobility formula ( = ·/ε0 ·εr ) is suitable only for hard particles (where electric charges are located only at the ion-impenetrable particle surface of zero thickness) that are large in comparison to the Debye length. In this formula, is the electrophoretic mobility, the zeta potential, ε0 the permittivity of vacuum, εr the relative permittivity of medium and the viscosity of the medium. The zeta potential represents the potential at the stagnant and slip plane which separates the movable part of the ionic double layer and the particle surface (see Supporting Information S3-a). Bacterial cell surfaces cannot be modeled as so-called “rigid particles” because they are often surrounded by a layer of charged polymers (EPS), as occurs in Synechocystis. As described in Supporting Information 3-b, the zeta potential of a soft particle is much more negative than the potential ˚0 at the outside of the soft layer because of the fixed charge density of surface polymers; thus zeta potential cannot reflect the surface potential of bacterial cells.
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The “soft particle theory”, proposed by Ohshima et al. [11,32] describes the EPM of soft particles. It can be applied to microorganisms which contain both membrane fixed charges and a surface charged soft layer. This latter polyelectrolyte layer around microbial cell surfaces deforms upon variations of the fluid ionic strength, and this phenomenon triggers consequent changes in EPM . Thus, from EPM data obtained at various ionic concentrations, the cell surface properties, such as the volumetric charge density and the softness of the polyelectrolyte layer, can be determined based on soft particle theory [12–18,33,34]. Indeed, the general equation for the electrophoretic mobility is derived and based on two characteristics specific to the particle layer. First the softness 1/ (given in length unit) characterizes the degree of resistance to liquid flow penetration in the layer region. It depends on the fluid dynamic viscosity and on the friction coefficient for flow in the polyelectrolyte layer (1/ = (/)1/2 ). The second parameter is the fixed spatial charge density in the layer (expressed in C m−3 or mM since 1 mM = 1.04 × 10−5 C m−3 ), depending on the elementary electric charge e and on the number concentration N and the valence Z of dissociated functional groups in the polyelectrolyte layer ( = NZe). The approximated electrophoretic mobility is defined [11] by: ε0 εr =
(˚0 / m ) + (˚DON /) (1/ m ) + (1/)
+
2
m = 1 +
kT ˚0 = ze
2 1/4
ln
+ 2zen
2zen +
˚DON
kT = ze
(2)
2zen
1−
ln
2zen
2zen
2
2
1/2
+1
1/2 +1
2 2zen
+
2zen
Fig. 1. Quantification of capsular (CPS) and released (RPS) EPS of Synechocystis PCC6803 WT and mutants. The color of data points refers to the two groups identified: without RPS depletion (red), and with RPS depletion (blue). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of the article.)
(1)
with ˚0 the outer surface potential (located at the surface of the ion-penetrable layer of polyelectrolyte), ˚DON the Donnan potential in the polyelectrolyte layer, and m the Debye–Hückel parameter of the polyelectrolyte layer given by:
173
(3)
1/2 +1
(4)
where is the inverse of the Debye length associated with the salt only, k is the Boltzmann constant, T is the temperature (K), and z and n are respectively the valency and concentration of bulk ions. From the plot of experimental mobility data against ionic concentration, curve fitting can be carried out to obtain two unknown parameters and 1/. 3. Results and discussion 3.1. Quantification and visualization of the Synechocystis EPS We found (see Supporting Information S4) that Synechocystis possesses both types of EPS occurring in many, but not all, bacteria: the capsular EPS (CPS), which are linked to the cell surface, and the EPS, which are released in the medium (RPS). CPS and RPS are both abundant in the wild-type strain (WT). In contrast, the amounts of either CPS alone or both CPS and RPS decrease as a result of the deletion of one or two EPS-production genes sll0923, sll1581, slr1875, and sll5052. We report that the EPS-depleted mutants can be divided in two groups. The first group comprises the single deletion mutant 1875− , and the two double deletions
mutants 1875− 5052− and 1581− 5052− (represented with red circles in Fig. 1) which all possess the same high amount of RPS (about 2.6 mg/cm2 ) but less CPS than in the WT strain. We also included in that group the mutant 5052 that − exhibits WT-levels of both RPS and CPS. The second group of mutants contains the two single mutants 0923− and 1581− , and the four double mutants 0923− 5052− , 0923− 1875− , 0923− 1581− and 1581− 1875− (represented with blue squares in Fig. 1), which all have depletions in both RPS and CPS. In this second group of mutants, it is worth noting the clear anti-correlation between RPS and CPS (Fig. 2). Also interestingly, we found that, in most cases, RPS represent only a small fraction of total EPS (about 20–30% for most mutants), except for the most CPS-depleted mutants 1581− 1875− and 0923− 1581− where RPS account for more than 50% of total EPS. The double mutant 1581− 1875− possesses the lowest content of total EPS (CPS + RPS). The SEM observations of Synechocystis cells presented in Fig. 2 confirm the significant presence of EPS around WT, 5052− mutant and 1875− mutant (which belong to “red group”). The distinction between CPS and RPS is not possible by standard electron microscopy but we assume that the observed EPS are mainly CPS because those are usually the most abundant (Fig. 1) and also more RPS are susceptible to be lost during sample preparation, in spite of the supercritical CO2 drying procedure. The frequency of occurrence of extracellular surface structures on those three Synechocystis strains is high and various appendages with different density and lengths up to several m are observed. Indeed, both mutant strain cells exhibit long and sparsely distributed fibrils whereas WT cells are surrounded by a thick slime layer. Regarding 5052− and 1875− mutants, it can thus be concluded that although the total amount of RPS and CPS is close to that of WT, their tridimensional structure, texture and nature are strongly modified by the gene deletions. This can have consequence on physical properties such as electrophoretic mobilities and capacities to form biofilms. These observations of WT and two mutants strains with close total quantities of EPS offer a great perspective to tune EPS properties without altering their quantity which could have applications in biotechnologies [7,22,24,35]. Conversely, no fibrils can be observed at 0923− mutant, 1581− mutant and 1581− 1875− double mutant cell surfaces (which belong to “blue group”). Moreover, the double mutant 1581− 1875− cell surface appears quite bald, in agreement with its very low
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Fig. 2. SEM images of WT and 5 mutant strains of Synechocystis: (up) WT, 5052− mutant and 1875− mutant; (bottom) 0923− , 1581− mutant and 1581− 1875− double mutant. The color on sample name refers to the two groups identified in Fig. 1: without RPS depletion (red), and with RPS depletion (blue). Presented images are typical observations among dozens of cell visualized. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of the article.)
amount of CPS (Fig. 1), whereas EPS, presumably CPS are visible as a fuzzy layer around 0923− and 1581− mutant cell surfaces. SEM imaging following CO2 critical point evaporation protocol allows to observe EPS surrounding bacterial cells and to assess qualitative differences between the EPS microstructures (e.g. compact slime, network of fibrils, thin coverage of the cell surface). These observations correlate well with CPS quantities provided in Fig. 1. The double mutant 1581− 1875− which is almost entirely devoid of CPS shows no visible EPS in SEM images. However, such standard SEM imaging provides no definite clues on the actual physico-chemical effects of the different EPS phenotypes as well as no indication of possible differential effects of RPS and CPS. In the following, we turn to electrophoretic measurements to evaluate the potential of that technique in fast discrimination of the different EPS phenotypes.
The evolution of EPM of the eleven strains of bacterial cells along with the ionic concentration is shown in Fig. 3. The values measured in MM (ionic strength = 26.6 mM) correspond to cells centrifuged and resuspended in MM. The dotted lines represent the best-fit curves to Eq. (1). All strains exhibit negative EPM, indicating a negative surface charge in accordance with the anionic nature of the EPS layer [17]. Indeed, at each fixed ionic strength, the WT strain, as well as the 5052− , 1875− and 1875− 5052− mutant strains, have more negative EPM than the 0923− , 1581− and 1581− 1875− mutant strains. So Synechocystis cells with a significant EPS layer (“red” group in Fig. 1) exhibit a more negative EPM, and so more global surface charge, than EPS-depleted mutant cells (“blue” group in Fig. 1) (see Figure S6 in Supporting Information). The tendency observed for a reduction of the absolute values of EPM correlated with an increase of ionic strength is explained by the compression and suppression of the electrical double layer around the cells which reduces negative potential at the slipping
3.2. Electrophoretic measurements and influence of EPS on surface parameters The reproducibility of EPM measurements has been checked both for a same aliquot (<5% deviation) and for triplicate cultures (<12% deviation) (see Figure S5-a in Supporting Information). The values of EPM used in the determination of surface parameters were calculated through the average of triplicates experiments, each measurement repeated 5 times. The influence of centrifugation on the EPS characteristics, examined by comparing the EPM of growing cells before and after centrifugation and resuspension in MM, gives indications on the cohesion of the EPS layer and its adherence to the surface. Indeed, it appears that centrifugation without washing does not alter the EPS layer of the majority of Synechocystis strains (Figure S5-b) as the EPM does not change (<6% deviation). The difference is more pronounced (20–40% deviation) for four mutants, 0923− , 1581− , 1581− 1875− and 1581− 5052− . This result can be explained by the variability of the EPS layer among the different mutants, which is not just quantitative but also qualitative. These mutants likely possess a weaker cohesive force inside the EPS structure, with RPS layers appearing to be electrostatically linked more loosely to the cell surface than in the others strains.
Fig. 3. Evolution of electrophoretic mobility of WT and 10 mutants strains of Synechocystis as a function of the medium ionic strength (dots). The experiments were repeated three times for each strain and the mean values were plotted. The standard error was 5%. The dotted lines show the best fit to Ohshima’s soft particle model yielding parameters presented in Table 1.
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possibly not imaged by SEM have a strong contribution to softness, as discussed below. The quantity and nature of EPS thus obviously play a role in the softness of the polyelectrolyte layer as the softest strains belong to the red group and the hardest to the blue group as shown in Fig. 4. The obtained results clearly indicate that the RPS surrounding the cell surface increase the particle softness (Fig. 4). Altogether, these results show that, although EPS are complex and heterogeneous, they can be adequately modeled, from the standpoint of electrokinetic analysis, by a polyelectrolyte layer surrounding core bacterial particles. They also show that the RPS component of EPS gives a supplementary contribution to the softness of bacterial particles which explains the relatively small differences in softness between the different strains. An anti-correlated relation between the softness and the charge density can be also extracted from these data. Indeed, the hardest strain has the most negative charge density and the contrary is also true (see Figure S7 in Supporting Information). This phenomenon has been observed in other studies [37] and can be attributed to the density distribution of RPS/CPS ratio on the different strains. We could draw two clear conclusions regarding the RPS amount and less evidence of correlation between EMP and the amount of CPS. This comes from the fact that the RPS effects are dominating the roles of CPS, due to their outer location and also because these bacteria which possess indeed a large amount of RPS. A comparison between estimated values of surface potential of bacterial cells, calculated either with Oshima’s formula based on soft particle theory (˚0 ) or Smoluchowski formula based on conventional double layer theory (zeta potential ), is also shown in Table 1. Magnitudes of the surface potential ˚0 are approximately 1/20th of the zeta potential for each cell, in accordance with description in Figure S3-b. The difference between the two values emphasizes the importance of the consideration of EPS ionic permeability when interpreting global surface charge measurements of bacteria.
Fig. 4. Evolution of softness as function of RPS amount for the Synechocystis WT and mutant strains. (For interpretation of the references to color in the text, the reader is referred to the web version of the article.)
plane. However, the cells mobility converge to non-zero values as the ionic strength increases over 15 mM, which is a characteristic feature of ‘soft particles’ explained by the ion-penetrable polyelectrolyte layer around rigid particle. One can note that the observed EPM values deviate from the theoretical mobility curve at ionic strengths lower than 5 mM (Fig. 4). This phenomenon has already been observed for ionic strength below 15 mM [36] or 50 mM [33] and is generally explained by chemical and physical heterogeneities in the polyelectrolyte layer at the cell surface which trigger a non-uniform distribution of charge density deep inside the surface layer. Moreover, the resistance to liquid flow may change with the distance from the boundary between the surface layer and the surrounding medium because of the multi-sub-layer structure of bacterial cell surface. The two parameters characterizing cell electrokinetic properties of each eleven strains of Synechocystis, the charge density and the softness 1/, are gathered in Table 1. The degree of softness and charge density of the fluid-penetrable layer vary between the different strains but all the studied strains exhibit very soft particle characters since 1/ is comprised between 4 and 7 nm as compared to bacteria with no EPS such as E. coli for instance (1/ found in the 1–2 nm range). These values can be related to the SEM observations of the strains 5052− and 1875− mutants, which exhibiting both similar fibrils, have nevertheless the same high softness (1/ > 6 nm) than the WT strain (1/ = 6.9 nm) whose cells appeared surrounded by a slime layer. The softness of the double mutant 1581− 1875− although lower (1/ = 5 nm) is still high, in spite of its bald-looking cell surface. This suggests that RPS
3.3. Influence of binding ions on cell surface parameters Cyanobacteria inhabiting diverse aquatic biotopes (marine, brackish or freshwaters) are facing various physico-chemical parameters (temperature, ionic strength, ionic composition. . .) depending on their environments. Contrary to MM medium, freshwaters contain a relatively high amount of calcium ions: for example, 2.4 mM on average in the water of the Seine River (Paris, France) that is ten times higher than in MM. The presence of calcium ions Ca2+ in the solution significantly decreases the absolute value of Synechocystis EPM, as shown in Fig. 5a. The influence of medium counter ions, and especially Ca2+ , on the EPS layer characteristics was investigated through the use of MM or ultrapure water both supplemented with dilute ionic solutions of CaCO3 . The influence of Ca2+ on the combination of the charge density and the
Table 1 Best fitted couple of charge density ( ) and softness (1/) for Synechocystis cells based on soft particle analysis; and calculated values of bacterial cell surface potential using Smoluchowski theory (zeta potential ) or soft particle theory (surface potential ˚0 ). The color on sample name refers to the two groups identified in Fig.1: without RPS depletion (red), and with RPS depletion (blue). (For interpretation of the references to color in this table legend, the reader is referred to the web version of the article.). −
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
176
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Fig. 5. (a) Evolution of electrophoretic mobility of Synechocystis cells (WT, 0923− , 1581− , 1875− , 1581− 1875− , 5052− ) as a function of the ionic strength of the medium in absence () or in presence of 2.3 mM calcium ions Ca2+ (♦). The dotted lines show the theoretical curves calculated using Ohshima’s soft particle model with parameters presented in the table (b). The valence z was taken as 1 for data without calcium, or 2 for data with calcium. (b) Combination of charge density ( ) and softness (1/) for Synechocystis cells based on soft particle analysis with and without calcium cations. The red and blue color of strain names refer to the two groups of strains identified in Fig. 1: without RPS depletion (red), and with RPS depletion (blue). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of the article.)
softness of six Synechocystis strains is presented in Fig. 5b. We kept the softness constant and only adjusted the charge density of the polyelectrolyte layer since the interaction of charged ions with the EPS is more likely to influence their electric charge rather than their resistance to fluid flow. The calcium addition turns out to decrease the absolute value of EPS charge density, i.e. the polyelectrolyte layer density of charge is less negative, which is consistent with a complexation of the positively charged ions Ca2+ on the anionic groups of Synechocystis EPS. We demonstrate here that the composition of the surrounding medium can have a significant impact on the EPS electrokinetic properties through complexation processes, which thus can have influence on the capacity of cells to form biofilms or to adsorb onto surfaces such as stones and soils. Further experiments are indeed required for which one can guess contrasted behaviors for the different mutants. This result supports the use of natural waters in ecotoxicological tests since we reported in previous studies that the protective effect of the EPS layer against toxics is linked to its structure. 4. Conclusion In this study, we have investigated the influence of EPS on the cell electrokinetic properties in the model cyanobacterium Synechocystis PCC6803 and ten EPS-affected mutant we recently constructed. All these mutants produced, in a more or less important extent, less capsular and/or released EPS than the wild-type (WT) strain. Furthermore, we report that the nature and structure of the EPS layers were strongly modified in the mutants, even in the less affected 5052− strain. All eleven strains (WT and 10 EPS mutants) presently studied exhibited a negative electrophoretic
mobility, thereby revealing that they display a negative global surface charge. In addition, we found that the anionic character of the EPS layer strongly influenced the electrophoretic mobility value. Indeed, the more EPS the cell is surrounded with, the more negative is its mobility value. Because of their EPS layer, which can be assimilated to a polyelectrolyte layer, bacterial cells cannot be modeled as rigid particles. Consequently, we used the Ohshima’s soft particle theory, rather than the Smoluchowski equation to analyze the electrophoretic mobility data. This strategy enabled us to extract two parameters characterizing the EPS layer of cells: the softness and the charge density of the polyelectrolyte layer. From the electrophoretic mobility measurements, we showed that all eleven strains behaved as very soft particles. Therefore, we found that EPS covering cell surface increased the softness of the cell surface. Indeed, this softness turns out to be mainly correlated with the RPS amount. These findings can lead to important applications, for example an easy and fast screening of EPS-mutant strains produced by high-throughput genetics, since the electrophoretic migration of cells is influenced by the structure of the EPS layer and more precisely the RPS/CPS proportion. Furthermore, the possibility to conservatively design the EPS layer offers a great perspective for biotechnology applications where adhesion on substrate or penetration of solvent in the EPS layer plays a significant role. Finally, as cyanobacteria colonize diverse aquatic biotopes usually characterized by elevated calcium concentrations, we have also studied the influence of calcium ions on electrophoretic properties of Synechocystis cells. Indeed, the concentration of calcium ions in natural waters can vary from 4 mg/L from peat bog waters to 45 mg/L in lake waters and even 430 mg/L in sea waters. The presence of calcium cations in the bulk fluid yields a decrease of the negative charge density by binding, and then neutralizing, the anionic groups of EPS. This phenomenon triggers a physical and phenotypical alteration of the extracellular surface structure, which can lead to modifications of cell behavior (floating, adsorption on surface, flocculation, transport, formation of biofilm) in hard waters. Acknowledgements We thank the French C‘Nano programme (Enseine project) for the postdoc and PhD fellowships to T.J and M.P, respectively. The Scanning Electron Microscope (SEM) facility at the Institut de Minéralogie et de Physique des Milieux Condensés is supported by Région Ile de France grant SESAME 2006 N◦ I-07-593/R, INSUCNRS, INP-CNRS, University Pierre et Marie Curie – Paris 6. Karim Benzerara for insightful advices in sample preparation for SEM. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.colsurfb. 2013.03.057. References [1] W. Whitman, D. Coleman, W. Wiebe, Prokaryotes: the unseen majority, Proc. Natl. Acad. Sci. U.S.A. 95 (1998) 6578–6583. [2] M. Rappe, S. Giovannoni, Their uncultured microbial majority, Annu. Rev. Microbiol. 57 (2003) 369–394. [3] J. Costerton, Z. Lewandowski, Microbial biofilms, Annu. Rev. Microbiol. 49 (1995) 711–745. [4] T. Beveridge, S. Makin, J. Kadurugamuwa, Z. Li, Interactions between biofilms and the environment, FEMS Microbiol. Rev. 20 (1997) 291–303. [5] O. Rendueles, J. Kaplan, J. Ghigo, Antibiofilm polysaccharides, Environ. Microbiol. 15 (2) (2012) 334–336. [6] T. Beveridge, L. Graham, Surface layers of bacteria, Microbiol. Rev. 55 (4) (1991) 684–705.
M. Planchon et al. / Colloids and Surfaces B: Biointerfaces 110 (2013) 171–177 [7] P. Li, S. Harding, Z. Liu, Cyanobacterial exopolysaccharides: their nature and potential biotechnological applications, Biotechnol. Genet. Eng. Rev. 18 (2001) 375–404. [8] H. Liu, H. Fang, Extraction of extracellular polymeric substances (EPS) of sludges, J. Biotechnol. 95 (2002) 249–256. [9] J. Panoff, B. Priem, H. Morvan, F. Joset, Sulphated exopolysaccharides produced by two unicellular strains of cyanobacteria, Synechocystis PCC 6803 and 6714, Arch. Microbiol. 150 (1988) 558–563. [10] M. Delacruz, L. Belloni, M. Delsanti, Precipitation of highly-charged polyelectrolyte solutions in the presence of multivalent salts, J. Chem. Phys. 103 (13) (1995) 5781–5791. [11] H. Ohshima, Electrophoretic mobility of soft particles, J. Colloid Interface Sci. 63 (1994) 474–483. [12] H. Hayashi, H. Seiki, S. Tsuneda, A. Hirata, H. Sasaki, Influence of growth phase on bacterial cell electrokinetic characteristics examined by soft particle electrophoresis theory, J. Colloid Interface Sci. 264 (2003) 565–568. [13] R. Bos, H.H. van der Mei, H. Busscher, ‘Soft-particle’ analysis of the electrophoretic mobility of a fibrillated and non-fibrillated oral streptococcal strain: Streptococcus salivarius, Biophys. Chem. 74 (251) (1998) 255. [14] S. Takashima, H. Morisaki, Surface characteristics of the microbial cell of Pseudomonas syringae and its relevance to cell attachment, Colloids Surf. B: Biointerfaces 9 (1997) 205–212. [15] P. Kiers, R. Bos, H.V. Der, H. Busscher, The electrophoretic softness of the surface of Staphylococcus epidermidis cells grown in a liquid medium and on a solid agar, Microbiology 147 (2001) 757–762. [16] A. Tsuneda, J. Jung, H. Hayashi, H. Aikawa, A. Hirata, H. Sasaki, Influence of extracellular polymers on electrokinetic properties of heterotrophic bacterial cells examined by soft particle electrophoresis theory, Colloids Surf. B: Biointerfaces 29 (29) (2003) 181–188. [17] H. Morisaki, S. Nagai, H. Ohshima, E. Ikemoto, K. Kogure, The effect of motility and cell-surface polymers on bacterial attachment, Microbiology 145 (1999) 2797–2802. [18] E. Donath, A. Budde, E. Knippel, H. Baumler, “Hairy surface layer” concept of electrophoresis combined with local fixed surface charge density isotherms: application to human erythrocyte electrophoretic fingerprinting, Langmuir 12 (1996) 4832–4839. [19] A. Mulkidjanian, E. Koonin, K. Makarova, S. Mekhedov, A. Sorokin, Y. Wolf, et al., The cyanobacterial genome core and the origin of photosynthesis, Proc. Natl. Acad. Sci. U.S.A. 103 (35) (2006) 13126–13131. [20] J.P. Zehr, Nitrogen fixation by marine cyanobacteria, Trends Microbiol. 19 (2011) 162–173. [21] W.J. Ducat, P. Silver, Engineering cyanobacteria to generate high-value products, Trends Biotechnol. 29 (2011) 95–103. [22] R. Abed, S. Dobretsov, K. Sudesh, Applications of cyanobacteria in biotechnology, J. Appl. Microbiol. 106 (2009) 1–12. [23] R. de Philippis, G. Colica, E. Micheletti, Exopolysaccharide-producing cyanobacteria in heavy metal removal from water: molecular basis and practical
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Colloids and Surfaces B: Biointerfaces journal homepage: www.elsevier.com/locate/colsurfb
Influence of exopolysaccharides on the electrophoretic properties of the model cyanobacterium Synechocystis Mariane Planchon a,b , Thichakorn Jittawuttipoka c , Corinne Cassier-Chauvat c , Franc¸ois Guyot d , Franck Chauvat c , Olivier Spalla a,∗ a
CEA Saclay, DSM/IRAMIS/SIS2M/LIONS, UMR CEA-CNRS 3299, 91191 Gif sur Yvette, France Université Paris Diderot, Sorbonne Paris Cité, IPGP, UMR CNRS 7154, Paris, France c CEA Saclay, DSV/IBITEC-S/SB2SM, UMR 8221, 91191 Gif sur Yvette, France d IMPMC, UMR CNRS-Université Pierre et Marie Curie-Université Paris Diderot, 75005 Paris, France b
a r t i c l e
i n f o
Article history: Received 3 January 2013 Received in revised form 26 March 2013 Accepted 29 March 2013 Available online 25 April 2013 Keywords: Extracellular polymeric substance Electrophoretic mobility Soft particle electrophoresis theory Cyanobacteria Synechocystis
a b s t r a c t The influence of extracellular polymeric substances (EPS) on cell electrokinetics was investigated in the model cyanobacterium Synechocystis, in wild-type strains and in ten EPS-depleted mutants. The charge density and the softness of the EPS polyelectrolyte layer were calculated from the dependence of the electrophoretic mobility values of the cells with the ionic strength of the surrounding fluid. Electrophoretic mobility data showed that the eleven Synechocystis strains investigated behave as soft particles and cannot be adequately described by classical electrokinetic models of rigid particles. EPS surrounding the cells, especially those released in the growth medium, significantly increased the softness of the cell surface. Furthermore, the anionic nature of EPS resulted in negative surface charge densities, which appeared to be strongly dependent on the composition of the suspending fluid, as documented by a strong reduction of their absolute values in the presence of calcium cations. These finding stresses the importance of the physicochemical properties of EPS and cell surfaces of cyanobacteria, for both cell-to-medium and cellto-cell communications. In turn, these results emphasize that, whenever possible, natural waters should be used for meaningful ecotoxicological analyses of potential toxics. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Bacteria constitute the most successful form of life on Earth. They colonize most biotopes and produce a huge biomass (5 × 1030 cells and 35–55 × 1016 g of carbon), which represents up to 60–100% of the estimated total carbon in plants [1,2]. The principal reasons for the success of bacteria are their physiological robustness as well as their abilities to rapidly respond and adapt to environmental stimuli [3]. Biofilm is the predominant mode of bacterial growth in most natural, industrial and clinical environments [4]. The biofilm lifestyle is associated with a high tolerance to environmental stresses, including treatments with antibiotics or other biocides. Hence, biofilm formation is a major concern in environmental and biotechnological applications. Biofilms typically consist of densely packed, multispecies populations, encased in a cell-synthesized polymeric matrix attached to a surface [5]. These microbial extracellular polymeric substances (EPS) are heterogeneous in complexity and composition, which varies depending on the microorganisms [6–8]. EPS contain soluble ions and colloidal
∗ Corresponding author. Tel.: +33 1 69 08 57 43. E-mail address: [email protected] (O. Spalla). 0927-7765/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.colsurfb.2013.03.057
insoluble matters and are mainly composed of polysaccharides (neutral and acidic), proteins, nucleic acids, lipids and other biological macromolecules [9]. They provide a highly hydrated gel matrix structure that mediates most of the cell-to-cell and cell-to-surface interactions, which are required for the formation and stabilization of biofilms. In EPS, charged groups may associate or dissociate upon changes in pH or ionic strength of the suspending fluid, or upon the approach of a new charged surface of another cell. Regarding their physicochemical properties, EPS can be assimilated to polyelectrolytes since this matrix is composed of long polymers whose gelation is promoted by multivalent cations [10]. The electrophoretic mobility of an object in a liquid is a macroscopic signature of its surface state in interaction with the surrounding fluid and it can be measured by different means. So, electrophoretic mobility is both a rapid and effective technique to assess the stability and behavior of particles in suspension. This analysis was initially developed for objects having a hard surface and has been very successful in hard colloids science. However, the classical electric double layer model is not appropriate for bacterial cells because their surfaces display complex structures. In hard colloids, the electric charges on the outer surface determine the electrophoretic mobility. However, the permeability of the bacterial surface layer to ions and molecules from the
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surrounding fluid strongly influences the mobility. In addition the mobility of bacteria is affected by the non homogeneous spatial distribution of the charged groups present at the cell surface. This is why Ohshima and co-workers [11] developed an alternative model, called the “soft particle model”, which is better suited to the analysis of bacterial cells, i.e. biological core particles surrounded by a ion-penetrable EPS layer that strongly influences electrophoretic mobility. This “soft particle electrophoresis theory” was successfully used in several studies of the cell surface of the enterobacterium Escherichia coli [12,33] and of other non-photosynthetic bacteria [13–18,34]. However, soft particle electrophoresis has not yet been employed to study the cell surface of cyanobacteria, the cosmopolitan photosynthetic prokaryotes that produce a large part of the terrestrial di-oxygen [19] and biomass for the food chain [20], and which have the potentials for the sustainable production of biofuels [7,21] and bioplastics [22], and for the bioremediation of polluted soils and waters [23]. Many cyanobacteria possess extracellular polymeric substances, mainly of polysaccharidic nature (EPS) [24], which protect cells against environmental stresses (UV, salt, desiccation, heavy metals) and enable them to form biofilms [13]. The cyanobacterial EPS, which are complex [23], can be divided into two main groups: the RPS (released polysaccharides), which are secreted by cells into the surrounding environment, and the CPS (capsular polysaccharides), which are attached to the cell surface [24]. In this study, we investigated the influence of EPS on the electrophoretic mobility of the widely used unicellular model strain Synechocystis PCC6803 (hereafter Synechocystis), which makes up complex anionic EPS that contain 13 different monosaccharides and various uronic acids [9,25]. For this purpose we used, and compared, the wild type strain (WT) and 10 EPS-depleted mutants that were recently constructed [26]. The electrophoretic mobilities of these 11 strains were analyzed in the framework of the Ohshima’s soft particle theory. We also studied the influence on electrophoretic mobility and cell surface parameters of the counter-ions nature in the bulk suspension, emphasizing the strong role of calcium. 2. Materials and methods 2.1. Bacterial strains and culture conditions The round unicellular cyanobacterium Synechocystis PCC6803 strains wild-type (WT) is considered in this work together with and 10 of its EPS-depleted mutants that were recently constructed the details of which are fully reported in [26]. In the present work, the bacteria were grown aerobically under shaking (180 rpm, Infors Multitron II) and white light (2500 lux; 31.25 E m−2 s−1 Mazda TF 16 W lamps) at 30 ◦ C on BG11 medium [27] enriched with 3.78 mM Na2 CO3 [28] thereafter referred to as MM for standard mineral medium (see composition on Supporting information S1). The main composition of Synechocystis PCC6803 EPS is detailed in Supporting Information S2. The EPS-depleted mutants were generated after the single or double deletion of the genes sll0923, sll1581, slr1875 and sll5052 that share sequence homology with EPS-production genes in non-photosynthetic bacteria [29,30]. For commodity, all four single deletion mutants are designated as X− (for instance 0923− stands for the deletion of sll0923) while all six double deletion mutants are noted X− Y− (for instance 0923− 1581− stands for the deletion of both sll0923 and sll1581). All EPS-depleted mutants were grown in the presence of selective antibiotics, which were added at the following concentrations: Streptomycin (Sm, 2.5 g/mL), Spectinomycin (Sp, 2.5 g/mL) and Kanamycin (Km, 50 g/mL) [28]. The growth of the various strains was followed as the timely increase in absorbance (Beckman DU 640 spectrophotometer) at 580 nm (1 OD580 unit corresponding to 2.5 × 107 cells/mL).
2.2. EPS characterization In order to disclose correlations between the content in EPS and the physicochemical properties, the EPS mass quantification data, extracted from a previous study [26] are expressed in mg EPS/m2 of cell surface instead of mg EPS/mg of cell protein mass. This unit, classical in physical chemistry, is more relevant for surface studies than a biological material normalization. The cell being almost spherical with a size of 1 m, the surface per cell is m2 . Moreover, we made the distinction between CPS and RPS mass quantities to better assess the modification of EPS layer structure and nature among the mutants. Indeed, the gene deletion can influence differently the production of CPS or RPS, which cannot be reflected in the total EPS mass quantification. The method of CPS/RPS titration is given in our former paper [26]. Scanning electron microscopy (SEM) was performed to obtain qualitative imaging of the EPS in the WT strain and in the different mutants. 72 h grown cultures of WT and mutant strains (OD580 = 0.7) were chemically fixed overnight with glutaraldehyde (1%) washed twice with UPW (milliQ water) and gradually dehydrated using a CO2 critical point dryer (BAL-TEC CPD030), as described in [31]. Dried samples were mounted on aluminum stubs using double-sided carbon tape and carbon-coated (Leica EM SCD 500). Cells were observed with a Zeiss Ultra 55 FEG SEM microscope operated at 2.0 kV at a working distance of 2.7 mm. Images were acquired in secondary electron mode using an Everhart Thornley detector. 2.3. Measurement of the electrophoretic mobility of the cells Following centrifugation, cells resuspended in the studied media at OD580 = 0.5 (2.5 × 107 cells/mL) were transferred into disposable folded capillary cells with gold covered electrodes (Malvern). The electrophoretic mobility (EPM) of the cells was measured at 25 ◦ C at a 17◦ fixed scattering angle with a Zetasizer Nano ZS instrument (Malvern) equipped with a 633 nm laser, using runs performed at a voltage of 150 V and a frequency of 285 Hz. To investigate the influence of the ionic strength of the medium, EPM measurements were performed with cells suspensions in various dilutions of MM in ultrapure water (100%, 80%, 60%, 40%, 20% and 0%) which lead to an ionic strength range of 1–26.6 mM. One can note that the absence of washing, in order to preserve the EPS structure around cells, leaded to a non-zero ionic strength when cells were resuspended in ultrapure water. All measurements were repeated five times and experiments were carried out in triplicate. 2.4. Modeling using soft-particle theory Electrophoresis is an effective technique to assess the electrical properties of microparticle surfaces but the Smoluchowski mobility formula ( = ·/ε0 ·εr ) is suitable only for hard particles (where electric charges are located only at the ion-impenetrable particle surface of zero thickness) that are large in comparison to the Debye length. In this formula, is the electrophoretic mobility, the zeta potential, ε0 the permittivity of vacuum, εr the relative permittivity of medium and the viscosity of the medium. The zeta potential represents the potential at the stagnant and slip plane which separates the movable part of the ionic double layer and the particle surface (see Supporting Information S3-a). Bacterial cell surfaces cannot be modeled as so-called “rigid particles” because they are often surrounded by a layer of charged polymers (EPS), as occurs in Synechocystis. As described in Supporting Information 3-b, the zeta potential of a soft particle is much more negative than the potential ˚0 at the outside of the soft layer because of the fixed charge density of surface polymers; thus zeta potential cannot reflect the surface potential of bacterial cells.
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The “soft particle theory”, proposed by Ohshima et al. [11,32] describes the EPM of soft particles. It can be applied to microorganisms which contain both membrane fixed charges and a surface charged soft layer. This latter polyelectrolyte layer around microbial cell surfaces deforms upon variations of the fluid ionic strength, and this phenomenon triggers consequent changes in EPM . Thus, from EPM data obtained at various ionic concentrations, the cell surface properties, such as the volumetric charge density and the softness of the polyelectrolyte layer, can be determined based on soft particle theory [12–18,33,34]. Indeed, the general equation for the electrophoretic mobility is derived and based on two characteristics specific to the particle layer. First the softness 1/ (given in length unit) characterizes the degree of resistance to liquid flow penetration in the layer region. It depends on the fluid dynamic viscosity and on the friction coefficient for flow in the polyelectrolyte layer (1/ = (/)1/2 ). The second parameter is the fixed spatial charge density in the layer (expressed in C m−3 or mM since 1 mM = 1.04 × 10−5 C m−3 ), depending on the elementary electric charge e and on the number concentration N and the valence Z of dissociated functional groups in the polyelectrolyte layer ( = NZe). The approximated electrophoretic mobility is defined [11] by: ε0 εr =
(˚0 / m ) + (˚DON /) (1/ m ) + (1/)
+
2
m = 1 +
kT ˚0 = ze
2 1/4
ln
+ 2zen
2zen +
˚DON
kT = ze
(2)
2zen
1−
ln
2zen
2zen
2
2
1/2
+1
1/2 +1
2 2zen
+
2zen
Fig. 1. Quantification of capsular (CPS) and released (RPS) EPS of Synechocystis PCC6803 WT and mutants. The color of data points refers to the two groups identified: without RPS depletion (red), and with RPS depletion (blue). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of the article.)
(1)
with ˚0 the outer surface potential (located at the surface of the ion-penetrable layer of polyelectrolyte), ˚DON the Donnan potential in the polyelectrolyte layer, and m the Debye–Hückel parameter of the polyelectrolyte layer given by:
173
(3)
1/2 +1
(4)
where is the inverse of the Debye length associated with the salt only, k is the Boltzmann constant, T is the temperature (K), and z and n are respectively the valency and concentration of bulk ions. From the plot of experimental mobility data against ionic concentration, curve fitting can be carried out to obtain two unknown parameters and 1/. 3. Results and discussion 3.1. Quantification and visualization of the Synechocystis EPS We found (see Supporting Information S4) that Synechocystis possesses both types of EPS occurring in many, but not all, bacteria: the capsular EPS (CPS), which are linked to the cell surface, and the EPS, which are released in the medium (RPS). CPS and RPS are both abundant in the wild-type strain (WT). In contrast, the amounts of either CPS alone or both CPS and RPS decrease as a result of the deletion of one or two EPS-production genes sll0923, sll1581, slr1875, and sll5052. We report that the EPS-depleted mutants can be divided in two groups. The first group comprises the single deletion mutant 1875− , and the two double deletions
mutants 1875− 5052− and 1581− 5052− (represented with red circles in Fig. 1) which all possess the same high amount of RPS (about 2.6 mg/cm2 ) but less CPS than in the WT strain. We also included in that group the mutant 5052 that − exhibits WT-levels of both RPS and CPS. The second group of mutants contains the two single mutants 0923− and 1581− , and the four double mutants 0923− 5052− , 0923− 1875− , 0923− 1581− and 1581− 1875− (represented with blue squares in Fig. 1), which all have depletions in both RPS and CPS. In this second group of mutants, it is worth noting the clear anti-correlation between RPS and CPS (Fig. 2). Also interestingly, we found that, in most cases, RPS represent only a small fraction of total EPS (about 20–30% for most mutants), except for the most CPS-depleted mutants 1581− 1875− and 0923− 1581− where RPS account for more than 50% of total EPS. The double mutant 1581− 1875− possesses the lowest content of total EPS (CPS + RPS). The SEM observations of Synechocystis cells presented in Fig. 2 confirm the significant presence of EPS around WT, 5052− mutant and 1875− mutant (which belong to “red group”). The distinction between CPS and RPS is not possible by standard electron microscopy but we assume that the observed EPS are mainly CPS because those are usually the most abundant (Fig. 1) and also more RPS are susceptible to be lost during sample preparation, in spite of the supercritical CO2 drying procedure. The frequency of occurrence of extracellular surface structures on those three Synechocystis strains is high and various appendages with different density and lengths up to several m are observed. Indeed, both mutant strain cells exhibit long and sparsely distributed fibrils whereas WT cells are surrounded by a thick slime layer. Regarding 5052− and 1875− mutants, it can thus be concluded that although the total amount of RPS and CPS is close to that of WT, their tridimensional structure, texture and nature are strongly modified by the gene deletions. This can have consequence on physical properties such as electrophoretic mobilities and capacities to form biofilms. These observations of WT and two mutants strains with close total quantities of EPS offer a great perspective to tune EPS properties without altering their quantity which could have applications in biotechnologies [7,22,24,35]. Conversely, no fibrils can be observed at 0923− mutant, 1581− mutant and 1581− 1875− double mutant cell surfaces (which belong to “blue group”). Moreover, the double mutant 1581− 1875− cell surface appears quite bald, in agreement with its very low
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Fig. 2. SEM images of WT and 5 mutant strains of Synechocystis: (up) WT, 5052− mutant and 1875− mutant; (bottom) 0923− , 1581− mutant and 1581− 1875− double mutant. The color on sample name refers to the two groups identified in Fig. 1: without RPS depletion (red), and with RPS depletion (blue). Presented images are typical observations among dozens of cell visualized. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of the article.)
amount of CPS (Fig. 1), whereas EPS, presumably CPS are visible as a fuzzy layer around 0923− and 1581− mutant cell surfaces. SEM imaging following CO2 critical point evaporation protocol allows to observe EPS surrounding bacterial cells and to assess qualitative differences between the EPS microstructures (e.g. compact slime, network of fibrils, thin coverage of the cell surface). These observations correlate well with CPS quantities provided in Fig. 1. The double mutant 1581− 1875− which is almost entirely devoid of CPS shows no visible EPS in SEM images. However, such standard SEM imaging provides no definite clues on the actual physico-chemical effects of the different EPS phenotypes as well as no indication of possible differential effects of RPS and CPS. In the following, we turn to electrophoretic measurements to evaluate the potential of that technique in fast discrimination of the different EPS phenotypes.
The evolution of EPM of the eleven strains of bacterial cells along with the ionic concentration is shown in Fig. 3. The values measured in MM (ionic strength = 26.6 mM) correspond to cells centrifuged and resuspended in MM. The dotted lines represent the best-fit curves to Eq. (1). All strains exhibit negative EPM, indicating a negative surface charge in accordance with the anionic nature of the EPS layer [17]. Indeed, at each fixed ionic strength, the WT strain, as well as the 5052− , 1875− and 1875− 5052− mutant strains, have more negative EPM than the 0923− , 1581− and 1581− 1875− mutant strains. So Synechocystis cells with a significant EPS layer (“red” group in Fig. 1) exhibit a more negative EPM, and so more global surface charge, than EPS-depleted mutant cells (“blue” group in Fig. 1) (see Figure S6 in Supporting Information). The tendency observed for a reduction of the absolute values of EPM correlated with an increase of ionic strength is explained by the compression and suppression of the electrical double layer around the cells which reduces negative potential at the slipping
3.2. Electrophoretic measurements and influence of EPS on surface parameters The reproducibility of EPM measurements has been checked both for a same aliquot (<5% deviation) and for triplicate cultures (<12% deviation) (see Figure S5-a in Supporting Information). The values of EPM used in the determination of surface parameters were calculated through the average of triplicates experiments, each measurement repeated 5 times. The influence of centrifugation on the EPS characteristics, examined by comparing the EPM of growing cells before and after centrifugation and resuspension in MM, gives indications on the cohesion of the EPS layer and its adherence to the surface. Indeed, it appears that centrifugation without washing does not alter the EPS layer of the majority of Synechocystis strains (Figure S5-b) as the EPM does not change (<6% deviation). The difference is more pronounced (20–40% deviation) for four mutants, 0923− , 1581− , 1581− 1875− and 1581− 5052− . This result can be explained by the variability of the EPS layer among the different mutants, which is not just quantitative but also qualitative. These mutants likely possess a weaker cohesive force inside the EPS structure, with RPS layers appearing to be electrostatically linked more loosely to the cell surface than in the others strains.
Fig. 3. Evolution of electrophoretic mobility of WT and 10 mutants strains of Synechocystis as a function of the medium ionic strength (dots). The experiments were repeated three times for each strain and the mean values were plotted. The standard error was 5%. The dotted lines show the best fit to Ohshima’s soft particle model yielding parameters presented in Table 1.
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possibly not imaged by SEM have a strong contribution to softness, as discussed below. The quantity and nature of EPS thus obviously play a role in the softness of the polyelectrolyte layer as the softest strains belong to the red group and the hardest to the blue group as shown in Fig. 4. The obtained results clearly indicate that the RPS surrounding the cell surface increase the particle softness (Fig. 4). Altogether, these results show that, although EPS are complex and heterogeneous, they can be adequately modeled, from the standpoint of electrokinetic analysis, by a polyelectrolyte layer surrounding core bacterial particles. They also show that the RPS component of EPS gives a supplementary contribution to the softness of bacterial particles which explains the relatively small differences in softness between the different strains. An anti-correlated relation between the softness and the charge density can be also extracted from these data. Indeed, the hardest strain has the most negative charge density and the contrary is also true (see Figure S7 in Supporting Information). This phenomenon has been observed in other studies [37] and can be attributed to the density distribution of RPS/CPS ratio on the different strains. We could draw two clear conclusions regarding the RPS amount and less evidence of correlation between EMP and the amount of CPS. This comes from the fact that the RPS effects are dominating the roles of CPS, due to their outer location and also because these bacteria which possess indeed a large amount of RPS. A comparison between estimated values of surface potential of bacterial cells, calculated either with Oshima’s formula based on soft particle theory (˚0 ) or Smoluchowski formula based on conventional double layer theory (zeta potential ), is also shown in Table 1. Magnitudes of the surface potential ˚0 are approximately 1/20th of the zeta potential for each cell, in accordance with description in Figure S3-b. The difference between the two values emphasizes the importance of the consideration of EPS ionic permeability when interpreting global surface charge measurements of bacteria.
Fig. 4. Evolution of softness as function of RPS amount for the Synechocystis WT and mutant strains. (For interpretation of the references to color in the text, the reader is referred to the web version of the article.)
plane. However, the cells mobility converge to non-zero values as the ionic strength increases over 15 mM, which is a characteristic feature of ‘soft particles’ explained by the ion-penetrable polyelectrolyte layer around rigid particle. One can note that the observed EPM values deviate from the theoretical mobility curve at ionic strengths lower than 5 mM (Fig. 4). This phenomenon has already been observed for ionic strength below 15 mM [36] or 50 mM [33] and is generally explained by chemical and physical heterogeneities in the polyelectrolyte layer at the cell surface which trigger a non-uniform distribution of charge density deep inside the surface layer. Moreover, the resistance to liquid flow may change with the distance from the boundary between the surface layer and the surrounding medium because of the multi-sub-layer structure of bacterial cell surface. The two parameters characterizing cell electrokinetic properties of each eleven strains of Synechocystis, the charge density and the softness 1/, are gathered in Table 1. The degree of softness and charge density of the fluid-penetrable layer vary between the different strains but all the studied strains exhibit very soft particle characters since 1/ is comprised between 4 and 7 nm as compared to bacteria with no EPS such as E. coli for instance (1/ found in the 1–2 nm range). These values can be related to the SEM observations of the strains 5052− and 1875− mutants, which exhibiting both similar fibrils, have nevertheless the same high softness (1/ > 6 nm) than the WT strain (1/ = 6.9 nm) whose cells appeared surrounded by a slime layer. The softness of the double mutant 1581− 1875− although lower (1/ = 5 nm) is still high, in spite of its bald-looking cell surface. This suggests that RPS
3.3. Influence of binding ions on cell surface parameters Cyanobacteria inhabiting diverse aquatic biotopes (marine, brackish or freshwaters) are facing various physico-chemical parameters (temperature, ionic strength, ionic composition. . .) depending on their environments. Contrary to MM medium, freshwaters contain a relatively high amount of calcium ions: for example, 2.4 mM on average in the water of the Seine River (Paris, France) that is ten times higher than in MM. The presence of calcium ions Ca2+ in the solution significantly decreases the absolute value of Synechocystis EPM, as shown in Fig. 5a. The influence of medium counter ions, and especially Ca2+ , on the EPS layer characteristics was investigated through the use of MM or ultrapure water both supplemented with dilute ionic solutions of CaCO3 . The influence of Ca2+ on the combination of the charge density and the
Table 1 Best fitted couple of charge density ( ) and softness (1/) for Synechocystis cells based on soft particle analysis; and calculated values of bacterial cell surface potential using Smoluchowski theory (zeta potential ) or soft particle theory (surface potential ˚0 ). The color on sample name refers to the two groups identified in Fig.1: without RPS depletion (red), and with RPS depletion (blue). (For interpretation of the references to color in this table legend, the reader is referred to the web version of the article.). −
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
−
176
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Fig. 5. (a) Evolution of electrophoretic mobility of Synechocystis cells (WT, 0923− , 1581− , 1875− , 1581− 1875− , 5052− ) as a function of the ionic strength of the medium in absence () or in presence of 2.3 mM calcium ions Ca2+ (♦). The dotted lines show the theoretical curves calculated using Ohshima’s soft particle model with parameters presented in the table (b). The valence z was taken as 1 for data without calcium, or 2 for data with calcium. (b) Combination of charge density ( ) and softness (1/) for Synechocystis cells based on soft particle analysis with and without calcium cations. The red and blue color of strain names refer to the two groups of strains identified in Fig. 1: without RPS depletion (red), and with RPS depletion (blue). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of the article.)
softness of six Synechocystis strains is presented in Fig. 5b. We kept the softness constant and only adjusted the charge density of the polyelectrolyte layer since the interaction of charged ions with the EPS is more likely to influence their electric charge rather than their resistance to fluid flow. The calcium addition turns out to decrease the absolute value of EPS charge density, i.e. the polyelectrolyte layer density of charge is less negative, which is consistent with a complexation of the positively charged ions Ca2+ on the anionic groups of Synechocystis EPS. We demonstrate here that the composition of the surrounding medium can have a significant impact on the EPS electrokinetic properties through complexation processes, which thus can have influence on the capacity of cells to form biofilms or to adsorb onto surfaces such as stones and soils. Further experiments are indeed required for which one can guess contrasted behaviors for the different mutants. This result supports the use of natural waters in ecotoxicological tests since we reported in previous studies that the protective effect of the EPS layer against toxics is linked to its structure. 4. Conclusion In this study, we have investigated the influence of EPS on the cell electrokinetic properties in the model cyanobacterium Synechocystis PCC6803 and ten EPS-affected mutant we recently constructed. All these mutants produced, in a more or less important extent, less capsular and/or released EPS than the wild-type (WT) strain. Furthermore, we report that the nature and structure of the EPS layers were strongly modified in the mutants, even in the less affected 5052− strain. All eleven strains (WT and 10 EPS mutants) presently studied exhibited a negative electrophoretic
mobility, thereby revealing that they display a negative global surface charge. In addition, we found that the anionic character of the EPS layer strongly influenced the electrophoretic mobility value. Indeed, the more EPS the cell is surrounded with, the more negative is its mobility value. Because of their EPS layer, which can be assimilated to a polyelectrolyte layer, bacterial cells cannot be modeled as rigid particles. Consequently, we used the Ohshima’s soft particle theory, rather than the Smoluchowski equation to analyze the electrophoretic mobility data. This strategy enabled us to extract two parameters characterizing the EPS layer of cells: the softness and the charge density of the polyelectrolyte layer. From the electrophoretic mobility measurements, we showed that all eleven strains behaved as very soft particles. Therefore, we found that EPS covering cell surface increased the softness of the cell surface. Indeed, this softness turns out to be mainly correlated with the RPS amount. These findings can lead to important applications, for example an easy and fast screening of EPS-mutant strains produced by high-throughput genetics, since the electrophoretic migration of cells is influenced by the structure of the EPS layer and more precisely the RPS/CPS proportion. Furthermore, the possibility to conservatively design the EPS layer offers a great perspective for biotechnology applications where adhesion on substrate or penetration of solvent in the EPS layer plays a significant role. Finally, as cyanobacteria colonize diverse aquatic biotopes usually characterized by elevated calcium concentrations, we have also studied the influence of calcium ions on electrophoretic properties of Synechocystis cells. Indeed, the concentration of calcium ions in natural waters can vary from 4 mg/L from peat bog waters to 45 mg/L in lake waters and even 430 mg/L in sea waters. The presence of calcium cations in the bulk fluid yields a decrease of the negative charge density by binding, and then neutralizing, the anionic groups of EPS. This phenomenon triggers a physical and phenotypical alteration of the extracellular surface structure, which can lead to modifications of cell behavior (floating, adsorption on surface, flocculation, transport, formation of biofilm) in hard waters. Acknowledgements We thank the French C‘Nano programme (Enseine project) for the postdoc and PhD fellowships to T.J and M.P, respectively. The Scanning Electron Microscope (SEM) facility at the Institut de Minéralogie et de Physique des Milieux Condensés is supported by Région Ile de France grant SESAME 2006 N◦ I-07-593/R, INSUCNRS, INP-CNRS, University Pierre et Marie Curie – Paris 6. Karim Benzerara for insightful advices in sample preparation for SEM. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.colsurfb. 2013.03.057. References [1] W. Whitman, D. Coleman, W. Wiebe, Prokaryotes: the unseen majority, Proc. Natl. Acad. Sci. U.S.A. 95 (1998) 6578–6583. [2] M. Rappe, S. Giovannoni, Their uncultured microbial majority, Annu. Rev. Microbiol. 57 (2003) 369–394. [3] J. Costerton, Z. Lewandowski, Microbial biofilms, Annu. Rev. Microbiol. 49 (1995) 711–745. [4] T. Beveridge, S. Makin, J. Kadurugamuwa, Z. Li, Interactions between biofilms and the environment, FEMS Microbiol. Rev. 20 (1997) 291–303. [5] O. Rendueles, J. Kaplan, J. Ghigo, Antibiofilm polysaccharides, Environ. Microbiol. 15 (2) (2012) 334–336. [6] T. Beveridge, L. Graham, Surface layers of bacteria, Microbiol. Rev. 55 (4) (1991) 684–705.
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