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Probing the surface ultrastructure of Brevibacillus laterosporus using atomic force microscopy
Micron 131 (2020) 102827
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Probing the surface ultrastructure of Brevibacillus laterosporus using atomic force microscopy
T
Khalid Alzahrania,b, Arun Kumar Shuklab,*, Javed Alamb, Abdurahman A. Niazyc, Abdullah M. Alsouwailehd, Mansour Alhoshanb,e, Jamal Khalidf, Hamdan S. Alghamadic a
Department of Physics and Astronomy, King Abdullah Institute for Nanotechnology, King Saud University, Riyadh 11451, Saudi Arabia King Abdullah Institute for Nanotechnology, King Saud University, Riyadh 11451, Saudi Arabia c Molecular and Cell Biology Laboratory, College of Dentistry, King Saud University, Riyadh 11545, Saudi Arabia d Department of Chemistry, College of Science, King Saud University, Riyadh 11451, Saudi Arabia e Department of Chemical Engineering, College of Engineering, King Saud University, P.O. Box 800, Riyadh 11421, Saudi Arabia f Department of Botany and Microbiology, College of Science, King Saud University, Riyadh 11451, Saudi Arabia b
A R T I C LE I N FO
A B S T R A C T
Keywords: Cell immobilization Atomic force microscopy Imaging live bacteria Brevibacillus laterosporus Surface ultrastructure
One of the main obstacles to studying the surface ultrastructure of microbial cells by atomic force microscopy (AFM) is determining how to immobilize live cells on the AFM substrates. Each method has its own advantages and disadvantages. The aim of this study was to characterize a new simple and inexpensive method using two types of polyethersulfone (PES) membrane filters that differ in pore size (micropore and nanopore) to immobilize live and dead Brevibacillus laterosporus for AFM imaging. B. laterosporus was easily trapped by the microporous PES membrane, facilitating the successful AFM scanning of the bacterial surface ultrastructure. In addition, B. laterosporus strongly attached to the nanoporous membranes and withstood the pulling forces exerted by the AFM tip during scanning. These methods of immobilization did not affect the cell viability. The nanostructure and roughness of the bacterial surface were also observed for live, fixed, and air-dried cells. Live and dead bacteria displayed similar morphologies at low resolution, while at high resolution, live bacteria displayed a more convoluted surface ("brain-like structure").
1. Introduction Atomic force microscopy (AFM) has attracted widespread attention as a powerful tool for imaging and mapping several properties of biological and non-biological surfaces (Kailas et al., 2009; Dorobantu et al., 2012; Ozkan et al., 2016; Alzahrani and Swedan, 2017; Gomand et al., 2018). In comparison to other high-resolution imaging techniques, such as transmission and scanning electron microscopy, AFM is favorable through its capability to image live bacteria under physiological conditions. Imaging the surface ultrastructure of live bacteria is fundamental for understanding the correlation between the structural properties and functions of bacteria, and for exploring the behaviors and responses of bacterial cells to different stimuli and changes in their environment. However, the imaging of living microbial cells remains a challenge. In general, the contact force between microbial cells and the AFM substrate is weak, often leading to the detachment of the adherent cells by lateral forces exerted by the AFM tip during scanning. Thus, a novel approach is needed to immobilize living cells on the AFM
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substrate, while leaving the structure and cell viability unaffected. Different strategies have been developed for immobilizing living microbial cells on AFM substrates. In general, these strategies can be classified as chemistry-based or involving physical entrapment. Chemistry-based approaches rely on the use of substrates functionalized by a substance that can enhance cell adhesion. These approaches are feasible for cells of different shapes and sizes. The most common approach is the adsorption of cells onto a polymer surface, such as gelatin or poly-L-lysine (PLL) (Lonergan et al., 2014; Allison, 2010; Doktycz et al., 2003; Kuyukina et al., 2014). Immobilization on PLL results from the electrostatic interaction force created between the positively charged PLL substrate and negatively charged cell membrane. However, the use of PLL may not be applicable to high ionic solutions (Kuyukina et al., 2014; Meyer et al., 2010); moreover, PLL can affect the chemistry and morphology of the surface (Lonergan et al., 2014). For bacterial cells of different shapes and sizes, gelatin is a suitable surface. A gelatin layer is extremely thin, such that bacteria cannot be pushed into the layer during scanning (Allison, 2010). The main
Corresponding author. E-mail address: [email protected] (A.K. Shukla).
https://doi.org/10.1016/j.micron.2020.102827 Received 10 November 2019; Received in revised form 12 January 2020; Accepted 12 January 2020 Available online 13 January 2020 0968-4328/ © 2020 Elsevier Ltd. All rights reserved.
Micron 131 (2020) 102827
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activity of the bacteria. From the resulting culture, 10 ml was added to 100 ml of fresh nutrient broth and cultivated under the same conditions for 18 h. Bacteria were harvested in mid-logarithmic phase by centrifugation at 8000 × g for 5 min. The optical density was 0.8 ± 0.2 at 550 nm using a visible spectrophotometer. The bacterial cells were washed three times using sterile normal saline (0.89 % NaCl).
disadvantage of using gelatin in AFM imaging is the possibility of contaminating the AFM tip during scanning as a result of gelatin desorption from the substrate. Physical entrapment approaches are occasionally used and are relatively simple compared with other methods. One example is the employment of porous membrane filters with lithographically patterned surfaces as mechanical traps for bacteria (Kailas et al., 2009; Dufrêne, 2008; Vadillo-Rodríguez et al., 2004). However, physical entrapment is not applicable for microbial cells that vary in shape and size. For example, an isopore polycarbonate membrane filter can trap coccoid bacteria but not rod-shaped bacteria (Kasas and Ikai, 1995). Mechanical entrapment methods are also likely to induce the deformation of growing cells (Kailas et al., 2009), which may affect their ultrastructural properties. Presently, we developed a cost-effective method to image living Brevibacillus laterosporus. The porous polyethersulfone (PES) membrane used was prepared in our laboratory. The pore sizes were controlled to produce a microporous PES membrane (micro-PES) and a nanoporous PES membrane (nano-PES). The use of micro-PES allowed bacteria of diverse shapes and sizes to be mechanically entrapped, owing to the high heterogeneity of pore sizes and shapes. Bacteria trapped in the pores of micro-PES were imaged under liquid. Meanwhile, the use of nano-PES allowed the imaging of live bacteria in air. The attachment of bacteria was sufficiently strong to withstand the detachment forces exerted by the AFM tip during scanning. Consequently, the morphology and fine structure of the B. laterosporus surface were resolved using the two types of membranes.
2.4. Immobilization of bacteria for AFM The bacterial suspension was divided into four portions. One portion was directly applied to the micro-PES membrane, as the suspension was filtered through the membrane to trap the bacteria in the pores. The membrane was rinsed gently with phosphate buffered saline to remove untrapped cells and attached to a steel disk. The disk was immediately placed on the AFM stage and submerged in PBS. Imaging was performed in liquid with a silicon AFM tip (SNL, k =0.3 N/m, ƒ =65 kHz, Bruker) using the AFM tapping mode. The second portion was centrifuged at 3000 rpm for 2 min. The pellet was washed in phosphate buffered saline (PBS) and resuspended in ultrapure water. A droplet of the suspension was pipetted onto the nano-porous membrane and allowed to partially air-dry partially for 10−20 min. The third portion was treated as previously described (Chao and Zhang, 2011). Briefly, the bacteria were washed twice in PBS, fixed using 2.5 % glutaraldehyde in PBS for 2 h, and then washed twice in ultrapure water to remove any traces of glutaraldehyde. The fixed cells were then resuspended in ultrapure water. A droplet of suspension was filtered through a micro-PES membrane. The membrane was then washed with ultrapure water to remove the untrapped cells, adhered to a steel support, and left to dry. The fourth portion was processed as described for the second portion. A drop of the solution was deposited on a round glass coverslip and left to air-dry. All four surfaces were glued to a steel disk and immediately mounted onto the AFM stage. AFM images were acquired in air in AFM tapping mode using a cantilever (TESPA, Bruker) with a nominal spring constant of 42 N/m and resonance frequency of 320 kHz. All experiments were conducted in triplicate.
2. Materials and methods 2.1. Fabrication of PES membrane The micro-PES and nano-PES membranes were fabricated using the immersion precipitation phase inversion technique. Vacuum-dried PES granules (15 and 17 wt%) and polyethylene glycol 600, serving as the pore-forming agent (10 wt%), were dissolved in N-methyl pyrrolidone and mixed completely by magnetic stirring at 70 ± 2 °C until a homogeneous and transparent casting solution was obtained. The solution was degassed in a vacuum at 25 ± 1 °C for another 6 h to remove air bubbles. The casting solution was dispersed onto a smooth clean glass plate through a film applicator (DeltaE Srl, Italy) from the gap of 70 ± 2 μm and 115 ± 2 μm between the casting knife and glass plate surface. Contact with air was maintained for 30 s for nano-PES and 2 min for micro-PES. Subsequently, the cast liquid membrane was horizontally immersed in a water coagulation bath at 30 ± 2 °C (microPES) and 15 ± 1 °C (nano-PES). The obtained micro-PES and nano-PES membranes were washed several times with Milli-Q water and preserved until further study.
2.5. Viability test The membrane sheet was cut into 5 mm-diameter disks, sterilized in 70 % ethanol for 2 h, and washed several times by using sterile deionized (DI) water to wash off the ethanol. The disks were then placed into 96 well plates. Each well containing a disk received a 100 μl lysogeny broth (LB) broth. Three wells containing only LB broth and membrane disks served the negative control. Bacteria (5 × 105) were added to each test well. The plate was then incubated at 37 °C for 24, 48, and 72 h. At the appropriate times, each disk was removed from the well and dipped into sterile DI water three times to remove all unattached bacteria. The disks were placed in sterile 1 ml conical tubes, and 1000 μl sterile DI water was added to each tube. Each tube was vortexed at 3000 rpm for 1 min to detach the bacteria from the membrane. The suspension was serially diluted, and the aliquots were plated on LB agar. Plates were incubated overnight, and colony forming units (CFUs) were counted. The average CFU/ml was 5.5 × 104, 6.2 × 104, and 8.0 × 104 at 24, 48, and 72 h, respectively. The nano-PES membranes with immobilized bacteria have been prepared as mentioned in Section 2.4. The bacterial cells have been recovered from the membranes; the cells were left for 5 h on the membrane, and then washed out using sterile normal saline. The suspension was serially diluted and aliquots were plated on LB agar. Plates were incubated overnight and the viability was proved using colony forming units (CFUs) were counted.
2.2. Membrane zeta potential measurement The surface charge of the PES membranes was calculated by the zeta potential measurement using the tangential streaming potential method with a SurPASS electrokinetic analyzer (Anton Paar, GmbH, Austria). Zeta potential measurements were carried out in an adjustable clamping cell with 1 mM KCl electrolyte solution as a function of pH (pH 5.6–2.5). The electrolyte solution pH was adjusted with 0.05 M HCl via titration, and values was calculated by the Attract® software from the streaming potential. 2.3. Bacterial growth conditions B. laterosporus was isolated from the digestive tract of honey bees, as previously described (Khaled et al., 2018) and grown in a nutrient broth (Oxoid, UK) at 37 °C for 24 h at 180 rpm. The sub-cultivation was repeated at least three times using these conditions to obtain optimum 2
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Fig. 1. (a) SEM top surface image of micro-PES membrane. (b and c) AFM top surface image of micro-PES membrane. (d) Cross-sectional view of micro-PES membrane obtained by SEM. (e) AFM top surface image of nano-PES membrane. Scale bars represent (a and b) 2 μm, (d) 10 μm, and (e) 150 nm. Channel-like structures within the membrane are depicted with a white arrow and a black arrow, indicating the membrane top surface.
3. Results
laterosporus was mechanically trapped in the micro-PES membrane (Fig. 3a), on which living B. laterosporus were directly deposited from the nutrient broth and immediately imaged under PBS by AFM. These immobilized B. laterosporus were stably imaged, and high-resolution images were obtained. The bacteria showed no signs of damage, and the cell wall appeared to be intact. The bacterial viability was further checked by the viability assay. The main morphological feature of the cells was the aforementioned canoe-like shape with a slight indentation in the middle, whereas a ripple- or wave-like texture could be observed on the bacterial cell wall. Fig. 3d shows an AFM image of B. laterosporus fixed using glutaraldehyde and physically entrapped into a pore of the micro-PES membrane. Although the bacterium was slightly squeezed into the pore and was affected by the chemical treatment, the main features of the bacterial morphology remained intact. AFM revealed bacteria anchored on the nano-PES membrane displaying the canoe-like shape with a slightly rippled surface texture (Fig. 3e). Although the bacteria prepared in the latter manner were imaged in air in a humid environment, there were no indications of cell disruption nor dehydration, and the viability was not detrimentally affected.
3.1. SEM and AFM evaluation of PES membranes Representative SEM/AFM images for the micro-PES and nano-PES membranes are shown in Fig. 1a–e. The SEM micrograph in Fig. 1d shows a cross-sectional view taken along the membrane. Channel-like structures that become wider from the top to the bottom of the membrane are visible, allowing the fast draining of the applied solution. Representative images of micro-PES membrane shown the average pore size approximately 1.3 μm and presented in Fig. 1a–d, and a representative image of the nano-PES is displayed in Fig. 1e. The average size of the nanopores was estimated to be 40 nm, while the root mean square (RMS) of the membrane roughness was less than 1 nm.
3.2. SEM and AFM evaluation of B. laterosporus onto micro- and nano-PES membranes The SEM images of B. laterosporus revealed a canoe-like shape with a smooth surface (Fig. 2a). A representative AFM image of B. laterosporus cells is illustrated in Fig. 2b. The bacteria were deposited on the glass coverslip and air-dried without further treatment. The majority of the bacteria are shown to exhibit a rod-like shape with a rough surface. The bacteria were aggregated closely together, probably because of the tendency of single cells to approach each other during drying. A clear variation in the morphology between that examined by SEM and that examined by AFM was evident and could reflect the different preparation methods. Fig. 3a–e shows the AFM images of B. laterosporus on PES. B.
3.3. AFM of the surface ultrastructure of B. laterosporus At high magnification (Fig. 4a–f,h), the ultrastructure of the cell surface appears convoluted with a "brain-like shape". Bacteria immobilized on nano-PES membranes (Fig. 4c and d) displayed nearly the same ultrastructure as those mechanically entrapped in the pores of the micro-PES membrane (Fig. 4a and b). In contrast, the glutaraldehydefixed (Fig. 4e and f) and air-dried (Fig. 4g and h) bacteria appeared to be damaged, exhibiting a highly modified surface. 3
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Fig. 2. (a) SEM image of B. laterosporus. (b) AFM image of B. laterosporus deposited on glass coverslip and air-dried without further treatment. The scale bar is 1 μm.
deleterious effect of the glutaraldehyde on the bacterial surface.
Finally, Fig. 5 presents the RMS of the bacterial roughness quantified using AFM. A slight increase in the roughness of bacteria attached to the nano-PES membranes, compared with bacteria trapped in the pores of the micro-PES membranes, was evident, with a small variation in cell size. The observations could reflect imaging under different ambient conditions. In comparison, the surface of the glutaraldehydefixed cells was rougher and irregular, as the RMS roughness of the fixed cells was noticeably higher and widely distributed, implying the
4. Discussion The bacterial surface ultrastructure is highly complex and primarily consists of an inner cytoplasmic membrane and cell wall. In gram-positive bacteria, the cell wall mainly consists of a thick layer of peptidoglycan, whereas in gram-negative bacteria a thin layer of
Fig. 3. (a) Schematic diagram of B. laterosporus mechanically entrapped in pores of a micro-PES membrane. (b) Schematic diagram of B. laterosporus mechanically anchored to a nano-PES membrane. (c) AFM image of live B. laterosporus entrapped into a micro-PES membrane. (d) AFM image of B. laterosporus fixed using glutaraldehyde and physically entrapped into a pore of a micro-PES membrane. (e) AFM image of a live B. laterosporus anchored to a nano-PES membrane. Scale bars are 1 μm. 4
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Fig. 4. High-resolution AFM images of B. laterosporus: height images (a, c, e, and g) and the corresponding phase images (b, d, f, and h) using different treatments. (a and b) Untreated bacteria trapped in a micro-PES membrane. (c and d) Untreated bacteria immobilized on a nano-PES membrane. (e and f) Glutaraldehyde-fixed bacteria trapped in a micro-PES membrane. (g and h) Air-dried bacteria on a glass plate. Scan size 500 nm × 500 nm.
Thus, electrostatic interactions, hydrophobic interactions, glue, and chemical fixation are commonly used to attach cells to the AFM cantilever. Meyer et al. (2010) also demonstrated that the bacteria were easily immobilised by physisorption to chemically modified glass surfaces. This observation was presumably due to electrostatic and hydrophobic interactions. Whereas imaging of cells immobilised to surfaces was relatively straight forward and detached. However, the use of chemicals can alter the cell surface, which can make the experiment unreliable. To avoid these difficulties, a physical-entrapment-based technique was employed in the present study using micro- and nanoPES membranes. We anticipate that this method will be helpful in applying AFM to probe the surface ultrastructure of living bacteria. The use of a porous PES membrane with a wide variety of pore sizes and shapes is an alternative approach to immobilizing bacteria. In the micro-PES membrane, irregularity in the pore geometry allows multiple surface contacts between the pore and the bacterial surface. Therefore, a pore with a size larger than the bacterium can still be capable of immobilizing the cell, while leaving ample space for bacterial growth and division. In contrast, the immobilization of bacterial cells onto the nano-PES membrane does not require entrapment in pores. The surface of a nano-PES membrane contains several nanosize pores (approximately 40 nm), exhibiting a roughness of less than 1 nm. The nature of the adhesive forces involved in the traction of the bacterial surface to the membrane is not clear. However, we hypothesize that the nanoscale pores are responsible for anchoring the bacteria to the surface via capillary forces, as illustrated schematically in Fig. 6. Water is trapped in the nanopores, forming a nanolayer of water on top of each pore. At that location, a nano-contact point will develop between the bacterial surface and the membrane. We hypothesize that the presence of the water contact points provides adhesive forces at the nanoscale between the cell surface and the membrane, as well as prevents cell dehydration. The configuration of the force on the surface depends on the scale and distribution of the pores. The possibility of electrostatic force can be rejected, owing to the negative charge of both surfaces (bacteria and membrane, as determined by zeta potential measurements (result not shown)), leading to an electrostatic repulsion between the surfaces. At the nano-contact, the capillary forces dominate over the other forces,
Fig. 5. Roughness of bacterial surface measured by AFM.
peptidoglycan is sandwiched between the inner cytoplasmic membrane and outer membrane. Knowledge of the morphology and physiochemical properties of the bacteria are essential to understanding the mechanisms that govern the bacterial interactions with other surfaces and their functions. SEM and TEM are microscopic methods widely used to study bacterial surface morphology. The use of both requires relatively extensive and tedious preparation processes that may affect the integrity of the cell surface, leading to artifacts or surface damage. Accordingly, highresolution AFM has emerged as a powerful tool in cell biology, with the capability to visualize cells in their native states. Despite the advantages of AFM, it has been limited by the lack of an appropriate method to secure a force sufficient to maintain the attachment of a single bacterial cell onto the AFM substrate during scanning. Kailas et al., 2009 was monitored the activity of the whole cell under different conditions using a AFM and the results showed that the cells were not cramped inside the holes because of shape and size in lithographically patterned substrates provided the cells freedom to carry on with their natural behavior. A weak bond between the cell and substrate can lead to cell detachment. 5
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Fig. 6. Schematic diagram of the possible mechanism of bacterial anchorage to the nano-PES membrane surface.
pattern. However, the remarkable similarity in the structural pattern between the gram-positive B. laterosporus and gram-negative bacteria, such as E. coli, indicates that LPS may play little to no role in forming the observed patterned. Gram-positive bacteria are surrounded by a thick layer of peptidoglycan, whereas gram-negative bacteria are surrounded by a thin layer of peptidoglycan and the LPS-containing outer membrane. One possible explanation of this pattern observed for E. coli and B. laterosporus, which are structurally different, is that the preparation method used for E. coli might have led to the degradation of the outer membrane and exposure of the peptidoglycan layer. To demonstrate that the observed pattern is naturally occurring and not an artifact, the experiment was repeated several times using different treatments and approaches. Comparing AFM images obtained with the nano-PES membrane to those of living bacteria revealed similar patterns. However, the width of the filaments was slightly smaller than those apparent on the living cells, owing to a partial loss of water. Thus, the imaging of nano-PES membrane does not influence the cell morphological properties. For glutaraldehyde-fixed bacteria, the surface pattern was nearly identical to that of the living bacteria. Fixation and drying approaches caused the pattern to be less pronounced and partially induced damage to the organization and structure of the pattern; however, the pattern was still evident. Consequently, the observed pattern is not an artifact due to the imaging or treatment approach. This pattern is not random, and further specific experiments are required to verify its origin.
such as electrostatic forces (Yoon et al., 2003). The influence of the capillary forces was presently investigated by repeating the experiment under liquid. Samples imaged under liquid displayed a sharply deteriorated image quality, indicating that the stability of the bacterial cells during AFM imaging was more likely due to capillary forces. Further investigation is necessary to verify the nature and significance of this force. Images obtained on either micro-PES or nano-PES membranes revealed the unique canoe-shaped morphology of B. laterosporus. The bacterial viability did not appear to be affected by the immobilization method, as demonstrated by the results of the growth-based enumeration of colonies. AFM images of the B. laterosporus surface using the nano-PES membrane were almost identical to those of the micro-PES membrane under liquid. Therefore, although the imaging of the nanoPES membrane was performed in air, the structural properties of the bacteria appeared to remain unchanged, indicating that the bacteria were still hydrated and viable. The use of the nano-PES membrane allowed AFM scanning in air without the complexities that could affect the stability and quality of the image obtained in liquid. However, the main disadvantage of using this method is the time restriction, as the sample needs to be imaged approximately 3 h before the image starts to deteriorate. At nanoscale resolution, the surface ultrastructure of B. laterosporus displayed a consistent structural pattern of nanodimensional filaments with a wave- or “brain-like” shape. The width of the filaments varied between 30 and 60 nm, and the height varied between 3 and 10 nm. A high degree of consistency between the phase images and the corresponding topography was evident. The contrast (bright and dark areas) observed in the phase images indicated a variation in the energy dissipation during imaging that could be attributed to the heterogeneity of the surface composition or different molecular organization on the surface (Gammoudi et al., 2016). The brighter areas (filaments) are more likely to be composed of rigid materials, whereas the darker areas (void spaces between the filaments) most likely correspond to soft or viscous materials. It is possible that the bright areas represent peptidoglycan, while the darker areas represent the periplasmic space located beneath the peptidoglycan. Similar surface patterns were reported for the gram-negative bacterium Escherichia coli (Gammoudi et al., 2016; Mathelié-Guinlet et al., 2016). Greif et al. (2010) demonstrated that the ultrastructural properties of the surfaces of fixed and dried Sinorhizobium meliloti, another gram-negative bacterium, imaged in air by AFM and SEM, displayed a pattern that was nearly identical to those of B. laterosporus and E. coli. However, when imaged under liquid, the bacterial surface was covered with a pattern mostly composed of round protrusions. The lipopolysaccharide (LPS) found in the outer membrane of gram-negative bacteria, including E. coli and S. meliloti, was proposed as the origin of this
5. Conclusions Two types of PES membranes were fabricated (micro-PES and nanoPES), and used to facilitate optimal AFM imaging of the surface ultrastructure of living as well as dead B. laterosporus. Owing to the high heterogeneity of the pore sizes and shapes within the microporous membranes, bacteria with different shapes and sizes could be mechanically entrapped. We anticipate that nano-PES membranes will permit cell immobilization via nano-capillary forces created between water-filled pores and the outer bacterial surface. The morphology and fine structure of the surface of live B. laterosporus were resolved using the two types of membranes and compared with those imaged by SEM. The ultrastructure of the bacterial surfaces exhibited a particular pattern, which we describe as a "brain-like structure", which was more pronounced for non-treated cells. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. 6
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Acknowledgments
Gomand, F., et al., 2018. High-throughput screening approach to evaluate the adhesive properties of bacteria to milk biomolecules. Food Hydrocoll. 84, 537–544. Greif, D., et al., 2010. High resolution imaging of surface patterns of single bacterial cells. Ultramicroscopy 110, 1290–1296. Kailas, L., et al., 2009. Immobilizing live bacteria for AFM imaging of cellular processes. Ultramicroscopy 109, 775–780. Kasas, S., Ikai, A., 1995. A method for anchoring round shaped cells for atomic force microscope imaging. Biophys. J. 68, 1678–1680. Khaled, J.M., et al., 2018. Brevibacillus laterosporus isolated from the digestive tract of honeybees has high antimicrobial activity and promotes growth and productivity of honeybee’s colonies. Environ. Sci. Pollut. Res. 25, 10447–10455. Kuyukina, M., et al., 2014. Methods of microorganism immobilization for dynamic atomic-force studies. Appl. Biochem. Microbiol. 50, 1–9. Lonergan, N., Britt, L., Sullivan, C., 2014. Immobilizing live Escherichia coli for AFM studies of surface dynamics. Ultramicroscopy 137, 30–39. Mathelié-Guinlet, M., et al., 2016. Silica nanoparticles assisted electrochemical biosensor for the detection and degradation of Escherichia coli bacteria. Procedia Eng. 168, 1048–1051. Meyer, R.L., et al., 2010. Immobilisation of living bacteria for AFM imaging under physiological conditions. Ultramicroscopy 110, 1349–1357. Ozkan, A.D., et al., 2016. Atomic force microscopy for the investigation of molecular and cellular behavior. Micron 89, 60–76. Vadillo-Rodríguez, V., et al., 2004. Comparison of atomic force microscopy interaction forces between bacteria and silicon nitride substrata for three commonly used immobilization methods. Appl. Environ. Microbiol. 70, 5441–5446. Yoon, E.-S., et al., 2003. An experimental study on the adhesion at a nano-contact. Wear 254, 974–980.
The authors thank Deanship of Scientific Research at King Saud University for funding this work through the research project no. (RG1439-026). The authors would like to extend their appreciation to Researchers Support Services Unit for their technical support. References Allison, D.P., 2010. Atomic force microscopy of biological samples. Wiley Interdiscip. Rev. Nanomed. Nanobiotechnol. 2, 618–634. Alzahrani, K., Swedan, H., 2017. Nanostructural changes in the cell membrane of gammairradiated red blood cells. Indian J. Hematol. Blood Transfus. 33109–33115. Chao, Y., Zhang, T., 2011. Optimization of fixation methods for observation of bacterial cell morphology and surface ultrastructures by atomic force microscopy. Appl. Microbiol. Biotechnol. 92, 381. Doktycz, M., et al., 2003. AFM imaging of bacteria in liquid media immobilized on gelatin coated mica surfaces. Ultramicroscopy 97, 209–216. Dorobantu, L.S., Goss, G.G., Burrell, R.E., 2012. Atomic force microscopy: a nanoscopic view of microbial cell surfaces. Micron 43, 1312–1322. Dufrêne, Y.F., 2008. Atomic force microscopy and chemical force microscopy of microbial cells. Nat. Protoc. 3, 1132. Gammoudi, I., et al., 2016. Morphological and nanostructural surface changes in Escherichia coli over time, monitored by atomic force microscopy. Colloids Surf. B 141, 355–364.
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Probing the surface ultrastructure of Brevibacillus laterosporus using atomic force microscopy
T
Khalid Alzahrania,b, Arun Kumar Shuklab,*, Javed Alamb, Abdurahman A. Niazyc, Abdullah M. Alsouwailehd, Mansour Alhoshanb,e, Jamal Khalidf, Hamdan S. Alghamadic a
Department of Physics and Astronomy, King Abdullah Institute for Nanotechnology, King Saud University, Riyadh 11451, Saudi Arabia King Abdullah Institute for Nanotechnology, King Saud University, Riyadh 11451, Saudi Arabia c Molecular and Cell Biology Laboratory, College of Dentistry, King Saud University, Riyadh 11545, Saudi Arabia d Department of Chemistry, College of Science, King Saud University, Riyadh 11451, Saudi Arabia e Department of Chemical Engineering, College of Engineering, King Saud University, P.O. Box 800, Riyadh 11421, Saudi Arabia f Department of Botany and Microbiology, College of Science, King Saud University, Riyadh 11451, Saudi Arabia b
A R T I C LE I N FO
A B S T R A C T
Keywords: Cell immobilization Atomic force microscopy Imaging live bacteria Brevibacillus laterosporus Surface ultrastructure
One of the main obstacles to studying the surface ultrastructure of microbial cells by atomic force microscopy (AFM) is determining how to immobilize live cells on the AFM substrates. Each method has its own advantages and disadvantages. The aim of this study was to characterize a new simple and inexpensive method using two types of polyethersulfone (PES) membrane filters that differ in pore size (micropore and nanopore) to immobilize live and dead Brevibacillus laterosporus for AFM imaging. B. laterosporus was easily trapped by the microporous PES membrane, facilitating the successful AFM scanning of the bacterial surface ultrastructure. In addition, B. laterosporus strongly attached to the nanoporous membranes and withstood the pulling forces exerted by the AFM tip during scanning. These methods of immobilization did not affect the cell viability. The nanostructure and roughness of the bacterial surface were also observed for live, fixed, and air-dried cells. Live and dead bacteria displayed similar morphologies at low resolution, while at high resolution, live bacteria displayed a more convoluted surface ("brain-like structure").
1. Introduction Atomic force microscopy (AFM) has attracted widespread attention as a powerful tool for imaging and mapping several properties of biological and non-biological surfaces (Kailas et al., 2009; Dorobantu et al., 2012; Ozkan et al., 2016; Alzahrani and Swedan, 2017; Gomand et al., 2018). In comparison to other high-resolution imaging techniques, such as transmission and scanning electron microscopy, AFM is favorable through its capability to image live bacteria under physiological conditions. Imaging the surface ultrastructure of live bacteria is fundamental for understanding the correlation between the structural properties and functions of bacteria, and for exploring the behaviors and responses of bacterial cells to different stimuli and changes in their environment. However, the imaging of living microbial cells remains a challenge. In general, the contact force between microbial cells and the AFM substrate is weak, often leading to the detachment of the adherent cells by lateral forces exerted by the AFM tip during scanning. Thus, a novel approach is needed to immobilize living cells on the AFM
⁎
substrate, while leaving the structure and cell viability unaffected. Different strategies have been developed for immobilizing living microbial cells on AFM substrates. In general, these strategies can be classified as chemistry-based or involving physical entrapment. Chemistry-based approaches rely on the use of substrates functionalized by a substance that can enhance cell adhesion. These approaches are feasible for cells of different shapes and sizes. The most common approach is the adsorption of cells onto a polymer surface, such as gelatin or poly-L-lysine (PLL) (Lonergan et al., 2014; Allison, 2010; Doktycz et al., 2003; Kuyukina et al., 2014). Immobilization on PLL results from the electrostatic interaction force created between the positively charged PLL substrate and negatively charged cell membrane. However, the use of PLL may not be applicable to high ionic solutions (Kuyukina et al., 2014; Meyer et al., 2010); moreover, PLL can affect the chemistry and morphology of the surface (Lonergan et al., 2014). For bacterial cells of different shapes and sizes, gelatin is a suitable surface. A gelatin layer is extremely thin, such that bacteria cannot be pushed into the layer during scanning (Allison, 2010). The main
Corresponding author. E-mail address: [email protected] (A.K. Shukla).
https://doi.org/10.1016/j.micron.2020.102827 Received 10 November 2019; Received in revised form 12 January 2020; Accepted 12 January 2020 Available online 13 January 2020 0968-4328/ © 2020 Elsevier Ltd. All rights reserved.
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activity of the bacteria. From the resulting culture, 10 ml was added to 100 ml of fresh nutrient broth and cultivated under the same conditions for 18 h. Bacteria were harvested in mid-logarithmic phase by centrifugation at 8000 × g for 5 min. The optical density was 0.8 ± 0.2 at 550 nm using a visible spectrophotometer. The bacterial cells were washed three times using sterile normal saline (0.89 % NaCl).
disadvantage of using gelatin in AFM imaging is the possibility of contaminating the AFM tip during scanning as a result of gelatin desorption from the substrate. Physical entrapment approaches are occasionally used and are relatively simple compared with other methods. One example is the employment of porous membrane filters with lithographically patterned surfaces as mechanical traps for bacteria (Kailas et al., 2009; Dufrêne, 2008; Vadillo-Rodríguez et al., 2004). However, physical entrapment is not applicable for microbial cells that vary in shape and size. For example, an isopore polycarbonate membrane filter can trap coccoid bacteria but not rod-shaped bacteria (Kasas and Ikai, 1995). Mechanical entrapment methods are also likely to induce the deformation of growing cells (Kailas et al., 2009), which may affect their ultrastructural properties. Presently, we developed a cost-effective method to image living Brevibacillus laterosporus. The porous polyethersulfone (PES) membrane used was prepared in our laboratory. The pore sizes were controlled to produce a microporous PES membrane (micro-PES) and a nanoporous PES membrane (nano-PES). The use of micro-PES allowed bacteria of diverse shapes and sizes to be mechanically entrapped, owing to the high heterogeneity of pore sizes and shapes. Bacteria trapped in the pores of micro-PES were imaged under liquid. Meanwhile, the use of nano-PES allowed the imaging of live bacteria in air. The attachment of bacteria was sufficiently strong to withstand the detachment forces exerted by the AFM tip during scanning. Consequently, the morphology and fine structure of the B. laterosporus surface were resolved using the two types of membranes.
2.4. Immobilization of bacteria for AFM The bacterial suspension was divided into four portions. One portion was directly applied to the micro-PES membrane, as the suspension was filtered through the membrane to trap the bacteria in the pores. The membrane was rinsed gently with phosphate buffered saline to remove untrapped cells and attached to a steel disk. The disk was immediately placed on the AFM stage and submerged in PBS. Imaging was performed in liquid with a silicon AFM tip (SNL, k =0.3 N/m, ƒ =65 kHz, Bruker) using the AFM tapping mode. The second portion was centrifuged at 3000 rpm for 2 min. The pellet was washed in phosphate buffered saline (PBS) and resuspended in ultrapure water. A droplet of the suspension was pipetted onto the nano-porous membrane and allowed to partially air-dry partially for 10−20 min. The third portion was treated as previously described (Chao and Zhang, 2011). Briefly, the bacteria were washed twice in PBS, fixed using 2.5 % glutaraldehyde in PBS for 2 h, and then washed twice in ultrapure water to remove any traces of glutaraldehyde. The fixed cells were then resuspended in ultrapure water. A droplet of suspension was filtered through a micro-PES membrane. The membrane was then washed with ultrapure water to remove the untrapped cells, adhered to a steel support, and left to dry. The fourth portion was processed as described for the second portion. A drop of the solution was deposited on a round glass coverslip and left to air-dry. All four surfaces were glued to a steel disk and immediately mounted onto the AFM stage. AFM images were acquired in air in AFM tapping mode using a cantilever (TESPA, Bruker) with a nominal spring constant of 42 N/m and resonance frequency of 320 kHz. All experiments were conducted in triplicate.
2. Materials and methods 2.1. Fabrication of PES membrane The micro-PES and nano-PES membranes were fabricated using the immersion precipitation phase inversion technique. Vacuum-dried PES granules (15 and 17 wt%) and polyethylene glycol 600, serving as the pore-forming agent (10 wt%), were dissolved in N-methyl pyrrolidone and mixed completely by magnetic stirring at 70 ± 2 °C until a homogeneous and transparent casting solution was obtained. The solution was degassed in a vacuum at 25 ± 1 °C for another 6 h to remove air bubbles. The casting solution was dispersed onto a smooth clean glass plate through a film applicator (DeltaE Srl, Italy) from the gap of 70 ± 2 μm and 115 ± 2 μm between the casting knife and glass plate surface. Contact with air was maintained for 30 s for nano-PES and 2 min for micro-PES. Subsequently, the cast liquid membrane was horizontally immersed in a water coagulation bath at 30 ± 2 °C (microPES) and 15 ± 1 °C (nano-PES). The obtained micro-PES and nano-PES membranes were washed several times with Milli-Q water and preserved until further study.
2.5. Viability test The membrane sheet was cut into 5 mm-diameter disks, sterilized in 70 % ethanol for 2 h, and washed several times by using sterile deionized (DI) water to wash off the ethanol. The disks were then placed into 96 well plates. Each well containing a disk received a 100 μl lysogeny broth (LB) broth. Three wells containing only LB broth and membrane disks served the negative control. Bacteria (5 × 105) were added to each test well. The plate was then incubated at 37 °C for 24, 48, and 72 h. At the appropriate times, each disk was removed from the well and dipped into sterile DI water three times to remove all unattached bacteria. The disks were placed in sterile 1 ml conical tubes, and 1000 μl sterile DI water was added to each tube. Each tube was vortexed at 3000 rpm for 1 min to detach the bacteria from the membrane. The suspension was serially diluted, and the aliquots were plated on LB agar. Plates were incubated overnight, and colony forming units (CFUs) were counted. The average CFU/ml was 5.5 × 104, 6.2 × 104, and 8.0 × 104 at 24, 48, and 72 h, respectively. The nano-PES membranes with immobilized bacteria have been prepared as mentioned in Section 2.4. The bacterial cells have been recovered from the membranes; the cells were left for 5 h on the membrane, and then washed out using sterile normal saline. The suspension was serially diluted and aliquots were plated on LB agar. Plates were incubated overnight and the viability was proved using colony forming units (CFUs) were counted.
2.2. Membrane zeta potential measurement The surface charge of the PES membranes was calculated by the zeta potential measurement using the tangential streaming potential method with a SurPASS electrokinetic analyzer (Anton Paar, GmbH, Austria). Zeta potential measurements were carried out in an adjustable clamping cell with 1 mM KCl electrolyte solution as a function of pH (pH 5.6–2.5). The electrolyte solution pH was adjusted with 0.05 M HCl via titration, and values was calculated by the Attract® software from the streaming potential. 2.3. Bacterial growth conditions B. laterosporus was isolated from the digestive tract of honey bees, as previously described (Khaled et al., 2018) and grown in a nutrient broth (Oxoid, UK) at 37 °C for 24 h at 180 rpm. The sub-cultivation was repeated at least three times using these conditions to obtain optimum 2
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Fig. 1. (a) SEM top surface image of micro-PES membrane. (b and c) AFM top surface image of micro-PES membrane. (d) Cross-sectional view of micro-PES membrane obtained by SEM. (e) AFM top surface image of nano-PES membrane. Scale bars represent (a and b) 2 μm, (d) 10 μm, and (e) 150 nm. Channel-like structures within the membrane are depicted with a white arrow and a black arrow, indicating the membrane top surface.
3. Results
laterosporus was mechanically trapped in the micro-PES membrane (Fig. 3a), on which living B. laterosporus were directly deposited from the nutrient broth and immediately imaged under PBS by AFM. These immobilized B. laterosporus were stably imaged, and high-resolution images were obtained. The bacteria showed no signs of damage, and the cell wall appeared to be intact. The bacterial viability was further checked by the viability assay. The main morphological feature of the cells was the aforementioned canoe-like shape with a slight indentation in the middle, whereas a ripple- or wave-like texture could be observed on the bacterial cell wall. Fig. 3d shows an AFM image of B. laterosporus fixed using glutaraldehyde and physically entrapped into a pore of the micro-PES membrane. Although the bacterium was slightly squeezed into the pore and was affected by the chemical treatment, the main features of the bacterial morphology remained intact. AFM revealed bacteria anchored on the nano-PES membrane displaying the canoe-like shape with a slightly rippled surface texture (Fig. 3e). Although the bacteria prepared in the latter manner were imaged in air in a humid environment, there were no indications of cell disruption nor dehydration, and the viability was not detrimentally affected.
3.1. SEM and AFM evaluation of PES membranes Representative SEM/AFM images for the micro-PES and nano-PES membranes are shown in Fig. 1a–e. The SEM micrograph in Fig. 1d shows a cross-sectional view taken along the membrane. Channel-like structures that become wider from the top to the bottom of the membrane are visible, allowing the fast draining of the applied solution. Representative images of micro-PES membrane shown the average pore size approximately 1.3 μm and presented in Fig. 1a–d, and a representative image of the nano-PES is displayed in Fig. 1e. The average size of the nanopores was estimated to be 40 nm, while the root mean square (RMS) of the membrane roughness was less than 1 nm.
3.2. SEM and AFM evaluation of B. laterosporus onto micro- and nano-PES membranes The SEM images of B. laterosporus revealed a canoe-like shape with a smooth surface (Fig. 2a). A representative AFM image of B. laterosporus cells is illustrated in Fig. 2b. The bacteria were deposited on the glass coverslip and air-dried without further treatment. The majority of the bacteria are shown to exhibit a rod-like shape with a rough surface. The bacteria were aggregated closely together, probably because of the tendency of single cells to approach each other during drying. A clear variation in the morphology between that examined by SEM and that examined by AFM was evident and could reflect the different preparation methods. Fig. 3a–e shows the AFM images of B. laterosporus on PES. B.
3.3. AFM of the surface ultrastructure of B. laterosporus At high magnification (Fig. 4a–f,h), the ultrastructure of the cell surface appears convoluted with a "brain-like shape". Bacteria immobilized on nano-PES membranes (Fig. 4c and d) displayed nearly the same ultrastructure as those mechanically entrapped in the pores of the micro-PES membrane (Fig. 4a and b). In contrast, the glutaraldehydefixed (Fig. 4e and f) and air-dried (Fig. 4g and h) bacteria appeared to be damaged, exhibiting a highly modified surface. 3
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Fig. 2. (a) SEM image of B. laterosporus. (b) AFM image of B. laterosporus deposited on glass coverslip and air-dried without further treatment. The scale bar is 1 μm.
deleterious effect of the glutaraldehyde on the bacterial surface.
Finally, Fig. 5 presents the RMS of the bacterial roughness quantified using AFM. A slight increase in the roughness of bacteria attached to the nano-PES membranes, compared with bacteria trapped in the pores of the micro-PES membranes, was evident, with a small variation in cell size. The observations could reflect imaging under different ambient conditions. In comparison, the surface of the glutaraldehydefixed cells was rougher and irregular, as the RMS roughness of the fixed cells was noticeably higher and widely distributed, implying the
4. Discussion The bacterial surface ultrastructure is highly complex and primarily consists of an inner cytoplasmic membrane and cell wall. In gram-positive bacteria, the cell wall mainly consists of a thick layer of peptidoglycan, whereas in gram-negative bacteria a thin layer of
Fig. 3. (a) Schematic diagram of B. laterosporus mechanically entrapped in pores of a micro-PES membrane. (b) Schematic diagram of B. laterosporus mechanically anchored to a nano-PES membrane. (c) AFM image of live B. laterosporus entrapped into a micro-PES membrane. (d) AFM image of B. laterosporus fixed using glutaraldehyde and physically entrapped into a pore of a micro-PES membrane. (e) AFM image of a live B. laterosporus anchored to a nano-PES membrane. Scale bars are 1 μm. 4
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Fig. 4. High-resolution AFM images of B. laterosporus: height images (a, c, e, and g) and the corresponding phase images (b, d, f, and h) using different treatments. (a and b) Untreated bacteria trapped in a micro-PES membrane. (c and d) Untreated bacteria immobilized on a nano-PES membrane. (e and f) Glutaraldehyde-fixed bacteria trapped in a micro-PES membrane. (g and h) Air-dried bacteria on a glass plate. Scan size 500 nm × 500 nm.
Thus, electrostatic interactions, hydrophobic interactions, glue, and chemical fixation are commonly used to attach cells to the AFM cantilever. Meyer et al. (2010) also demonstrated that the bacteria were easily immobilised by physisorption to chemically modified glass surfaces. This observation was presumably due to electrostatic and hydrophobic interactions. Whereas imaging of cells immobilised to surfaces was relatively straight forward and detached. However, the use of chemicals can alter the cell surface, which can make the experiment unreliable. To avoid these difficulties, a physical-entrapment-based technique was employed in the present study using micro- and nanoPES membranes. We anticipate that this method will be helpful in applying AFM to probe the surface ultrastructure of living bacteria. The use of a porous PES membrane with a wide variety of pore sizes and shapes is an alternative approach to immobilizing bacteria. In the micro-PES membrane, irregularity in the pore geometry allows multiple surface contacts between the pore and the bacterial surface. Therefore, a pore with a size larger than the bacterium can still be capable of immobilizing the cell, while leaving ample space for bacterial growth and division. In contrast, the immobilization of bacterial cells onto the nano-PES membrane does not require entrapment in pores. The surface of a nano-PES membrane contains several nanosize pores (approximately 40 nm), exhibiting a roughness of less than 1 nm. The nature of the adhesive forces involved in the traction of the bacterial surface to the membrane is not clear. However, we hypothesize that the nanoscale pores are responsible for anchoring the bacteria to the surface via capillary forces, as illustrated schematically in Fig. 6. Water is trapped in the nanopores, forming a nanolayer of water on top of each pore. At that location, a nano-contact point will develop between the bacterial surface and the membrane. We hypothesize that the presence of the water contact points provides adhesive forces at the nanoscale between the cell surface and the membrane, as well as prevents cell dehydration. The configuration of the force on the surface depends on the scale and distribution of the pores. The possibility of electrostatic force can be rejected, owing to the negative charge of both surfaces (bacteria and membrane, as determined by zeta potential measurements (result not shown)), leading to an electrostatic repulsion between the surfaces. At the nano-contact, the capillary forces dominate over the other forces,
Fig. 5. Roughness of bacterial surface measured by AFM.
peptidoglycan is sandwiched between the inner cytoplasmic membrane and outer membrane. Knowledge of the morphology and physiochemical properties of the bacteria are essential to understanding the mechanisms that govern the bacterial interactions with other surfaces and their functions. SEM and TEM are microscopic methods widely used to study bacterial surface morphology. The use of both requires relatively extensive and tedious preparation processes that may affect the integrity of the cell surface, leading to artifacts or surface damage. Accordingly, highresolution AFM has emerged as a powerful tool in cell biology, with the capability to visualize cells in their native states. Despite the advantages of AFM, it has been limited by the lack of an appropriate method to secure a force sufficient to maintain the attachment of a single bacterial cell onto the AFM substrate during scanning. Kailas et al., 2009 was monitored the activity of the whole cell under different conditions using a AFM and the results showed that the cells were not cramped inside the holes because of shape and size in lithographically patterned substrates provided the cells freedom to carry on with their natural behavior. A weak bond between the cell and substrate can lead to cell detachment. 5
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Fig. 6. Schematic diagram of the possible mechanism of bacterial anchorage to the nano-PES membrane surface.
pattern. However, the remarkable similarity in the structural pattern between the gram-positive B. laterosporus and gram-negative bacteria, such as E. coli, indicates that LPS may play little to no role in forming the observed patterned. Gram-positive bacteria are surrounded by a thick layer of peptidoglycan, whereas gram-negative bacteria are surrounded by a thin layer of peptidoglycan and the LPS-containing outer membrane. One possible explanation of this pattern observed for E. coli and B. laterosporus, which are structurally different, is that the preparation method used for E. coli might have led to the degradation of the outer membrane and exposure of the peptidoglycan layer. To demonstrate that the observed pattern is naturally occurring and not an artifact, the experiment was repeated several times using different treatments and approaches. Comparing AFM images obtained with the nano-PES membrane to those of living bacteria revealed similar patterns. However, the width of the filaments was slightly smaller than those apparent on the living cells, owing to a partial loss of water. Thus, the imaging of nano-PES membrane does not influence the cell morphological properties. For glutaraldehyde-fixed bacteria, the surface pattern was nearly identical to that of the living bacteria. Fixation and drying approaches caused the pattern to be less pronounced and partially induced damage to the organization and structure of the pattern; however, the pattern was still evident. Consequently, the observed pattern is not an artifact due to the imaging or treatment approach. This pattern is not random, and further specific experiments are required to verify its origin.
such as electrostatic forces (Yoon et al., 2003). The influence of the capillary forces was presently investigated by repeating the experiment under liquid. Samples imaged under liquid displayed a sharply deteriorated image quality, indicating that the stability of the bacterial cells during AFM imaging was more likely due to capillary forces. Further investigation is necessary to verify the nature and significance of this force. Images obtained on either micro-PES or nano-PES membranes revealed the unique canoe-shaped morphology of B. laterosporus. The bacterial viability did not appear to be affected by the immobilization method, as demonstrated by the results of the growth-based enumeration of colonies. AFM images of the B. laterosporus surface using the nano-PES membrane were almost identical to those of the micro-PES membrane under liquid. Therefore, although the imaging of the nanoPES membrane was performed in air, the structural properties of the bacteria appeared to remain unchanged, indicating that the bacteria were still hydrated and viable. The use of the nano-PES membrane allowed AFM scanning in air without the complexities that could affect the stability and quality of the image obtained in liquid. However, the main disadvantage of using this method is the time restriction, as the sample needs to be imaged approximately 3 h before the image starts to deteriorate. At nanoscale resolution, the surface ultrastructure of B. laterosporus displayed a consistent structural pattern of nanodimensional filaments with a wave- or “brain-like” shape. The width of the filaments varied between 30 and 60 nm, and the height varied between 3 and 10 nm. A high degree of consistency between the phase images and the corresponding topography was evident. The contrast (bright and dark areas) observed in the phase images indicated a variation in the energy dissipation during imaging that could be attributed to the heterogeneity of the surface composition or different molecular organization on the surface (Gammoudi et al., 2016). The brighter areas (filaments) are more likely to be composed of rigid materials, whereas the darker areas (void spaces between the filaments) most likely correspond to soft or viscous materials. It is possible that the bright areas represent peptidoglycan, while the darker areas represent the periplasmic space located beneath the peptidoglycan. Similar surface patterns were reported for the gram-negative bacterium Escherichia coli (Gammoudi et al., 2016; Mathelié-Guinlet et al., 2016). Greif et al. (2010) demonstrated that the ultrastructural properties of the surfaces of fixed and dried Sinorhizobium meliloti, another gram-negative bacterium, imaged in air by AFM and SEM, displayed a pattern that was nearly identical to those of B. laterosporus and E. coli. However, when imaged under liquid, the bacterial surface was covered with a pattern mostly composed of round protrusions. The lipopolysaccharide (LPS) found in the outer membrane of gram-negative bacteria, including E. coli and S. meliloti, was proposed as the origin of this
5. Conclusions Two types of PES membranes were fabricated (micro-PES and nanoPES), and used to facilitate optimal AFM imaging of the surface ultrastructure of living as well as dead B. laterosporus. Owing to the high heterogeneity of the pore sizes and shapes within the microporous membranes, bacteria with different shapes and sizes could be mechanically entrapped. We anticipate that nano-PES membranes will permit cell immobilization via nano-capillary forces created between water-filled pores and the outer bacterial surface. The morphology and fine structure of the surface of live B. laterosporus were resolved using the two types of membranes and compared with those imaged by SEM. The ultrastructure of the bacterial surfaces exhibited a particular pattern, which we describe as a "brain-like structure", which was more pronounced for non-treated cells. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. 6
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Acknowledgments
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