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HEPES-Stabilized Encapsulation of Salmonella typhimurium Zhiyong Suo,† Xinghong Yang,‡ Recep Avci,*,† Laura Kellerman,† David W. Pascual,‡ Marc Fries,§ and Andrew Steele§ Imaging and Chemical Analysis Laboratory, Department of Physics, Montana State UniVersity, Bozeman, Montana 59717, Veterinary Molecular Biology, Montana State UniVersity, Bozeman, Montana 59717, and Carnegie Institution of Washington, Washington, DC 20015 ReceiVed July 24, 2006 Most bacteria, planktonic and sessile, are encapsulated inside loosely bound extracellular polymeric substance (EPS) in their physiological environment. Imaging a bacterium with its capsule requires lengthy sample preparation to enhance the capsular contrast. In this study, Salmonella typhimurium was investigated using atomic force microscopy for a practical means of imaging an encapsulated bacterium in air. The investigation further aimed to determine the relation between the buffers used for preparing the bacterium and the preservation of the capsular material surrounding it. It was observed that rinsing bacteria with HEPES buffer could stabilize and promote capsule formation, while rinsing with PBS, Tris, or glycine removes most of the capsular EPS. For bacteria rinsed with HEPES and air-dried, the height images showed only the contour of the capsular material, while the phase and amplitude images presented the detailed structures of the bacterial surface, including the flagella encapsulated inside the capsular EPS. The encapsulation was attributed to the cross-linking of the acidic exopolysaccharides mediated by the piperazine moiety of HEPES through electrostatic attraction. This explanation is supported by encapsulated bacteria observed for samples rinsed with N,N′-bis(2-hydroxyethyl)-piperazine solution and by the presence of entrapped HEPES within the dry capsular EPS suggested by micro-Raman spectroscopy.
Introduction The extracellular polymeric substances (EPS) produced by a bacterial species are a mixture of various components, including polysaccharides, lipopolysaccharides, proteins, peptides, and nucleic acids, in which the polysaccharides constitute the major fraction.1 For most bacterial species, the cells are encapsulated by a certain form and amount of EPS in their planktonic state (sometimes referred to as capsular EPS). The capsular EPS is loosely attached to the bacterium’s outer surface, forming a capsule which moves with the bacterium. Both planktonic and sessile bacteria can form capsules from the very beginning of their existence. The capsule, which usually contains various pathogenic antigens as virulence factors,2 plays a critical role in bacterial adhesion3,4 and can facilitate the evasion of bacteria from host defenses.2 The imaging of bacterial capsules will be an important step toward understanding their structures and biological functions, which may then benefit the development of novel vaccines. Compared to bacterial cells themselves, the capsules surrounding bacterial cells are difficult to observe because of their transparency to visible light.5,6 In order to visualize an EPS capsule by optical microscope or by a traditional electron microscope (EM), it is often necessary to stain the capsules with contrast* To whom correspondence should be addressed. E-mail: avci@ physics.montana.edu. Tel: 406-994-6164. Fax: 406-994-6040. † Department of Physics, Montana State University. ‡ Veterinary Molecular Biology, Montana State University. § Carnegie Institution of Washington. (1) Christensen, B. E.; Characklis, W. G. Physical and chemical properties of biofilms. In Biofilms, Characklis, W. G., Marshall, K. C., Eds.; John Wiley and Sons: New York, 1990; pp 93-130. (2) Cross, A. S. The biological significance of bacterial encapsulation SpringerVerlag: Berlin, 1990; Vol. 150, pp 87-95. (3) Tsuneda, S.; Aikawa, H.; Hayashi, H.; Yuasa, A.; Hirata, A. Extracellular polymeric substances responsible for bacterial adhesion onto solid surface. FEMS Microbiol. Lett. 2003, 223 (2), 287-292. (4) Gubner, R.; Beech, I. B. The effect of extracellular polymeric substances on the attachment of Pseudomonas NCIMB 2021 to AISI 304 and 316 stainless steel. Biofouling 2000, 15 (1-3), 25-36.
enhancing reagents, such as Indian ink particles,7 antibodies,8 cationic proteins,5,9 cationic dyes such as Alcian Blue,10 or heavy metal compounds such as ruthenium red,11,12 alkaline bismuth,13 and/or gold nanoparticles.14 Even then, the quality of images obtained by optical microscope and EM depends largely on the operator’s special skills and the quality of the labor-intensive staining procedure. Furthermore, the traditional EM sample preparation makes it extremely difficult to investigate biological samples in their native physiological environment, and the dehydration and/or nonisotropic collapse of specimens under high vacuum can cause severe artifacts.15 Accordingly, there has been a need for a convenient, practical approach to direct imaging of the bacterial capsule. (5) Bayer, M. E. Visualization of the bacterial polysaccharide capsule. Curr. Top. Microbiol. Immunol. 1990, 150, 129-157. (6) Handley, P. S. Detection of cell surface carbohydrate components. In Microbial cell surface analysis-structural and physicochemical methods; Mozes, N., Handley, P. S., Busscher, H. J., Rouxhet, P. G., Eds.; VCH: New York, 1991; pp 87-108. (7) Duguid, J. P. The demonstration of bacterial capsules and slime. J. Pathol. Bacteriol. 1951, 63, 673-685. (8) Hochkeppel, H. K.; Braun, D. G.; Vischer, W.; Imm, A.; Sutter, S.; Staeubli, U.; Guggenheim, R.; Kaplan, E. L.; Boutonnier, A.; Fournier, J. M. Serotyping and electron-microscopy studies of Staphylococcus-Aureus clinical isolates with monoclonal-antibodies to capsular polysaccharide type-5 and type-8. J. Clin. Microbiol. 1987, 25 (3), 526-530. (9) Ferris, F. G.; Beveridge, T. J. Site specificity of metallic ion binding in Escherichia-Coli K-12 lipopolysaccharide. Can. J. Microbiol. 1986, 32 (1), 52-55. (10) Reuhs, B. L.; Geller, D. P.; Kim, J. S.; Fox, J. E.; Kolli, V. S. K.; Pueppke, S. G. Sinorhizobium fredii and Sinorhizobium meliloti produce structurally conserved lipopolysaccharides and strain-specific K antigens. Appl. EnViron. Microbiol. 1998, 64 (12), 4930-4938. (11) Costerton, J. W.; Irvin, R. T.; Cheng, K. J. The Bacterial Glycocalyx in Nature and Disease. Annu. ReV. Microbiol. 1981, 35, 299-324. (12) Kuno, T.; Naito, S.; Ohta, M.; Kido, N.; Ito, H.; Kato, N. Staining of O-specific polysaccharide chains of lipopolysaccharides with ruthenium red. Microbiol. Immunol. 1986, 30 (8), 743-751. (13) Kuno, T.; Naito, S.; Ito, H.; Ohta, M.; Kido, N.; Kato, N. Staining of the O-specific polysaccharide chains of lipopolysaccharides with alkaline bismuth. Microbiol. Immunol. 1986, 30 (11), 1207-1211. (14) Bayer, M. E.; Bayer, M. H. Effects of Bacteriophage-Fd Infection on Escherichia-Coli Hb11 Envelope - a Morphological and Biochemical-Study. J. Virol. 1986, 57 (1), 258-266.
10.1021/la0621721 CCC: $37.00 © 2007 American Chemical Society Published on Web 12/10/2006
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Atomic force microscopy (AFM) has been used with great success for imaging bacterial species, both Gram positive and Gram negative, in air and in physiological solution.16-18 Due to the high spatial resolution of AFM, the fine structures of bacterial surfaces,18-20 bacterial spores,21,22 native membranes,23 and lipopolysaccharides monolayers24 can be resolved in some detail. At the same time, its high sensitivity to force measurements makes AFM an indispensable technique for the investigation of cellular mechanical properties, such as the elastic modulus of cellulose fibers,25 turgor pressure,26 and the adhesive properties of bacterial cells.27,28 However, only limited attention has been given to the investigation of bacterial capsules using AFM, and most work concerning bacterial capsules has focused on either the morphology of the capsule of distinct bacterial species29-34 or the influence of the production of capsular polysaccharides on bacterial adhesion on biotic and abiotic substrates.27,31,35,36 Little is known about how to enhance the imaging of a bacterium (15) Dubochet, J.; Adrian, M.; Richter, K.; Garces, J.; Wittek, R. Structure of Intracellular Mature Vaccinia Virus Observed by Cryoelectron Microscopy. J. Virol. 1994, 68 (3), 1935-1941. (16) Ubbink, J.; Schar-Zammaretti, P. Probing bacterial interactions: integrated approaches combining atomic force microscopy, electron microscopy and biophysical techniques. Micron 2005, 36 (4), 293-320. (17) Dufrene, Y. F.; Muller, D. J. Microbial surfaces investigated using Atomic Force Microscope. In Microbial Imaging, Metohds in Microbiology; Savidge, T., Pothoulakis, C., Eds.; Elsevier: Amsterdam, 2005; Vol. 34, pp 163-197. (18) Dufrene, Y. F. Using nanotechniques to explore microbial surfaces. Nat. ReV. Microbiol. 2004, 2 (6), 451-460. (19) Sokolov, I. Y.; Firtel, M.; Henderson, G. S. In situ high-resolution atomic force microscope imaging of biological surfaces. J. Vac. Sci. Technol. A. 1996, 14 (3), 674-678. (20) Camesano, T. A.; Natan, M. J.; Logan, B. E. Observation of changes in bacterial cell morphology using tapping mode atomic force microscopy. Langmuir 2000, 16 (10), 4563-4572. (21) Plomp, M.; Leighton, T. J.; Wheeler, K. E.; Pitesky, M. E.; Malkin, A. J. Bacillus atrophaeus outer spore coat assembly and ultrastructure. Langmuir 2005, 21 (23), 10710-10716. (22) Plomp, M.; Leighton, T. J.; Wheeler, K. E.; Malkin, A. J. The highresolution architecture and structural dynamics of Bacillus spores. Biophys. J. 2005, 88 (1), 603-608. (23) Scheuring, S.; Levy, D.; Rigaud, J. L. Watching the components of photosynthetic bacterial membranes and their in situ organisation by atomic force microscopy. Biochim. Biophys. Acta 2005, 1712 (2), 109-127. (24) Roes, S.; Seydel, U.; Gutsmann, T. Probing the properties of lipopolysaccharide monolayers and their interaction with the antimicrobial peptide polymyxin B by atomic force microscopy. Langmuir 2005, 21 (15), 6970-6978. (25) Guhados, G.; Wan, W. K.; Hutter, J. L. Measurement of the elastic modulus of single bacterial cellulose fibers using atomic force microscopy. Langmuir 2005, 21 (14), 6642-6646. (26) Yao, X.; Walter, J.; Burke, S.; Stewart, S.; Jericho, M. H.; Pink, D.; Hunter, R.; Beveridge, T. J. Atomic force microscopy and theoretical considerations of surface properties and turgor pressures of bacteria. Colloids Surf., B 2002, 23 (2-3.), 213-230. (27) Razatos, A.; Ong, Y. L.; Sharma, M. M.; Georgiou, G. Molecular determinants of bacterial adhesion monitored by atomic force microscopy. Proc. Natl. Acad. Sci. U.S.A. 1998, 95 (19), 11059-11064. (28) Touhami, A.; Jericho, M. H.; Boyd, J. M.; Beveridge, T. J. Nanoscale characterization and determination of adhesion forces of Pseudomonas aeruginosa Pili by using atomic force microscopy. J. Bacteriol. 2006, 188 (2), 370-377. (29) Tollersrud, T.; Berge, T.; Andersen, S. R.; Lund, A. Imaging the surface of Staphylococcus aureus by atomic force microscopy. APMIS 2001, 109 (7-8), 541-545. (30) Beech, I. B.; Cheung, C. W. S.; Johnson, D. B.; Smith, J. R. Comparative studies of bacterial biofilms on steel surfaces using atomic force microscopy and environmental scanning electron microscopy. Biofouling 1996, 10 (1-3), 65-70. (31) van der Aa, B. C.; Dufrene, Y. F. In situ characterization of bacterial extracellular polymeric substances by AFM. Colloids Surf., B 2002, 23 (2-3), 173-182. (32) Kalaji, M.; Neal, A. L. IR study of self-assembly of capsular exopolymers from Pseudomonas sp NCIMB 2021 on hydrophilic and hydrophobic surfaces. Biopolymers 2000, 57 (1), 43-50. (33) Teschke, O. Volume of extracellular polymeric substance coverage of individual Acidithiobacillus ferrooxidans bacterium measured by atomic force microscopy. Microscopy Res. Tech. 2005, 67 (6), 312-316. (34) Toikka, J.; Aalto, J.; Hayrinen, J.; Pelliniemi, L. J.; Finne, J. The polysialic acid units of the neural cell adhesion molecule N-CAM form filament bundle networks. J. Biol. Chem. 1998, 273 (44), 28557-28559. (35) Hanna, A.; Berg, M.; Stout, V.; Razatos, A. Role of capsular colanic acid in adhesion of uropathogenic Escherichia coli. Appl. EnViron. Microbiol. 2003, 69 (8), 4474-4481.
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under its capsule or the interaction of capsular EPS with the buffer solutions. Height images, which show the three-dimensional topography of the sample, are the most common image type presented in the literature on the imaging of bacterial species by AFM. However, in height images only the contour of the sample surface is revealed, making it difficult to obtain information about the sample’s internal structure. In tapping mode AFM, the phase image is generated by measuring the delay between the phase angle of the driving oscillator and the phase angle of the tip response at each pixel. This phase signal delay is a result of elastic contribution such as topographical effects and/or inelastic contributions such as energy dissipation and thus is sensitive to variations in the interaction of the tip with the sample due to variations in elasticity, viscoelasticity, or adhesive/repulsive interactions.37-39 This feature makes phase imaging an indispensable method for studying samples with surface and bulk heterogeneity, such as an encapsulated bacterium cell. Although phase imaging in tapping mode has been widely used in the characterization of polymer surfaces, especially block copolymers,40-43 until recently the investigation of bacterial surfaces by means of phase imaging has received limited attention.20,44-46 Salmonella typhimurium is a species of pathogenic bacteria frequently encountered when contaminated food or water is consumed, causing up to four million cases of salmonellosis per year in the United States alone.47 In both planktonic and biofilm form, S. typhimurium can produce extracellular polymers, which contain a rich reservoir of a variety of pathogenic antigens.48 However, until now only limited studies have been concerned with the imaging of the capsule of Salmonella using traditional methods such as optical microscope49 or EM,50 and only one (36) Razatos, A.; Ong, Y. L.; Sharma, M. M.; Georgiou, G. Evaluating the interaction of bacteria with biomaterials using atomic force microscopy. J. Biomater. Sci., Polym. Ed. 1998, 9 (12), 1361-1373. (37) Garcia, R.; Perez, R. Dynamic atomic force microscopy methods. Surf. Sci. Rep. 2002, 47 (6-8), 197-301. (38) Garcia, R.; Tamayo, J.; Calleja, M.; Garcia, F. Phase contrast in tappingmode scanning force microscopy. Appl. Phys. A: Mater. Sci. Process. 1998, 66, S309-S312. (39) Arce, F. T.; Avci, R.; Beech, I. B.; Cooksey, K. E.; Wigglesworth-Cooksey, B. Microelastic properties of minimally adhesive surfaces: A comparative study of RTV11 (TM) and Intersleek elastomers (TM). J. Chem. Phys. 2003, 119 (3), 1671-1682. (40) Bar, G.; Thomann, Y.; Brandsch, R.; Cantow, H. J.; Whangbo, M. H. Factors affecting the height and phase images in tapping mode atomic force microscopy. Study of phase-separated polymer blends of poly(ethene-co-styrene) and poly(2,6-dimethyl-1,4-phenylene oxide). Langmuir 1997, 13 (14), 38073812. (41) Wang, Y.; Song, R.; Li, Y. S.; Shen, J. S. Understanding tapping-mode atomic force microscopy data on the surface of soft block copolymers. Surf. Sci. 2003, 530 (3), 136-148. (42) Knoll, A.; Magerle, R.; Krausch, G. Tapping mode atomic force microscopy on polymers: Where is the true sample surface? Macromolecules 2001, 34 (12), 4159-4165. (43) Magonov, S. N.; Cleveland, J.; Elings, V.; Denley, D.; Whangbo, M. H. Tapping-mode atomic force microscopy study of the near-surface composition of a styrene-butadiene-styrene triblock copolymer film. Surf. Sci. 1997, 389 (13), 201-211. (44) Forsythe, J. H.; Maurice, P. A.; Hersman, L. E. Attachment of a Pseudomonas sp. to Fe(III)-(hydr)oxide surfaces. Geomicrobiol. J. 1998, 15 (4), 293. (45) Korenevsky, A.; Stukalov, O.; Dutcher, J.; Beveridge, T. Preferential adhesion of rough phenotypes to iron oxides from heterogeneous DMRB populations. Geochim. Cosmochim. Acta. 2005, 69 (10), A833-A833. (46) Auerbach, I. D.; Sorensen, C.; Hansma, H. G.; Holden, P. A. Physical morphology and surface properties of unsaturated Pseudomonas putida biofilms. J. Bacteriol. 2000, 182 (13), 3809-3815. (47) Tauxe, R. V.; Pavia, A. T. Salmonellosis: nontyphoidal. Plenum Medical Books: New York, 1998. (48) Ofek, I. H., D.L.; Doyle, R.J. Bacterial adhesion to animal cells and tissues; ASM Press: Washington, DC, 2003. (49) de Rezende, C. E.; Anriany, Y.; Carr, L. E.; Joseph, S. W.; Weiner, R. A. Capsular polysaccharide surrounds smooth and rugose types of Salmonella enterica seroVar typhimurium DT104. Appl. EnViron. Microbiol. 2005, 71 (11), 7345-7351.
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report has documented the use of AFM in the investigation of Salmonella.51 In the imaging of bacterial specimens by AFM, care must be taken to avoid artifacts induced by sample preparation. Certain chemicals with biological activities used during sample preparation can cause the alteration of bacterial surface morphology.19,20,52 Common buffers, such as HEPES and PBS, are widely used in biological experiments and presumed to be inert to the specimens. These buffers are also used in AFM sample preparation protocols to remove loosely attached bacterial cells on a substrate by rinsing before imaging in air or liquid. However, we observed in our experiments that at least one buffer, HEPES, preserved the encapsulation of Gram negative bacterial species such S. typhimurium and Escherichia coli when bacterial cells were exposed to it. This unexpected phenomenon was not observed when substrates were rinsed with other buffers, such as PBS, glycine, or Tris. In this study, we present the observation of the bacterial capsule of S. typhimurium by tapping mode AFM imaging in air. Our results showed the following (1) HEPES stabilizes the capsule formation of S. typhimurium and E. coli strains and (2) EPS capsules can be differentiated from the bacterial cells by means of phase imaging in tapping mode, which reveals the structural details of the bacterial surface and appendages inside the capsule. Experimental Section Bacteria. S. typhimurium H64753 and another Gram negative bacterial strain, E. coli S17-1, were used for capsule observation with AFM. Both species, S. typhimurium and E. coli, were inoculated from frozen bacteria stock at -80 °C onto a Luria-Bertani (LB) plate and incubated at 37 °C overnight. The bacteria were then inoculated into an LB liquid medium without antibiotics and shaken at 125 rpm at 37 °C. The bacterial cells were harvested when the medium’s optical density at 600 nm (OD600) reached about 0.50.6. Depending on the experiment, bacterial samples were either kept in liquid LB medium or resuspended in HEPES (or PBS) after centrifugation of the cells in liquid LB medium at 4000g for 5 min at 25 °C and stored at 4 °C. EPS Purification. Purified EPS was used as a control for the Raman experiments. The EPS of S. typhimurium H647 was purified according to the procedure reported by Bernhard.54 Briefly, S. typhimurium H647 cells grown on LB agar overnight at 37 °C were harvested (∼106 bacteria) and resuspended in 5 mL of PBS. The EPS was separated from the cells by vortexing each sample for 3 min followed by ultracentrifugation at 30 000 rpm (160 000g) for 45 min at 10 °C. The supernatant was removed and dialyzed in H2O for 3 h. The EPS was then precipitated from the supernatant by adding 0.3 M NaCl and 2.5 volumes of ethanol. After 16 h at 4 °C, the precipitated EPS was spooled with a glass rod and left to dry at 37 °C. Rinsing Solutions. Buffers of PBS (0.01 M, pH 7.4), HEPES (0.1 M, pH 7.4 and pH 9), 3-(N-morpholino)propanesulfonic acid (MOPS) (0.1 M, pH 7.4), Tris (0.1 M, pH 7.4), and glycine (0.1 M, pH 6.0) were prepared with solids purchased from Sigma-Aldrich (Milwaukee, WI). N,N′-Bis(2-hydroxyethyl)piperazine (BHEP), (50) Grund, S. Slime capsule and Fimbriae on Salmonella-Typhimurium VarCop - electron-microscopic study. J. Vet. Med. B. 1991, 38 (7), 545-551. (51) Aytac, S. A.; Mercanoglu, B.; Ergun, M. A.; Tan, E. The visualisation of Salmonella enteritidis by atomic force microscopy. Ann. Microbiol. 2003, 53 (3), 337-342. (52) Santamaria, M.; Diaz-Marrero, A. R.; Hernandez, J.; Gutierrez-Navarro, A. M.; Corzo, J. Effect of thorium on the growth and capsule morphology of Bradyrhizobium. EnViron. Microbiol. 2003, 5 (10), 916-924. (53) Ascon, M. A.; Hone, D. M.; Walters, N.; Pascual, D. W. Oral immunization with a Salmonella typhimurium vaccine vector expressing recombinant enterotoxigenic Escherichia coli K99 fimbriae elicits elevated antibody titers for protective immunity. Infect. Immun. 1998, 66 (11), 5470-5476. (54) Wehland, M.; Bernhard, F. The RcsAB box - Characterization of a new operator essential for the regulation of exopolysaccharide biosynthesis in enteric bacteria. J. Bio. Chem. 2000, 275 (10), 7013-7020.
Langmuir, Vol. 23, No. 3, 2007 1367 CaCl2, Na2SO3, and Na2SO4 were purchased from Acros Organics (Morris Plains, NJ), and the 0.1 M solutions were prepared with deionized water. The pH value of the BHEP solution alone was adjusted to pH 6.2; the pH values of solutions of CaCl2, Na2SO3, and Na2SO4 were kept unadjusted. Water purified by Milli-Q ultrapure water purification systems (Billerica, MA) was used in all the experiments. Sample Preparation for AFM Imaging. A 100 µL turbid suspension of bacteria in LB medium was brought to room temperature, placed on a freshly cleaved muscovite mica disk, and kept at ambient conditions for 10 min before being rinsed with one of the preselected rinsing solutions (4 × 200 µL) and then dried with a stream of dry nitrogen. Some control experiments were conducted with bacteria resuspended in PBS or HEPES buffer (see the Bacteria section) in order to verify the presence of capsular shells around the bacteria. The sample preparation for these experiments was identical to that used for bacteria in LB medium except that the rinsing buffer was the same as the resuspension buffer, i.e., if the bacteria were resuspended in HEPES buffer, we used HEPES buffer to rinse the excess bacteria after 10 min of incubation. A similar rule applied to the cells resuspended in PBS. AFM Experiments. All measurements were carried out with a Nanoscope IIIa Extended Multimode AFM from Veeco (Santa Barbara, CA) with a J-type scanner. Imaging in air was performed in tapping mode to reduce the tip-sample interaction and the lateral forces. The tapping mode silicon probes (RTESPW, Veeco, Santa Barbara, CA) used for imaging have a nominal spring constant of 40 N/m, nominal tip radius of ∼10 nm, and nominal resonance frequency of ∼300 kHz. For each mica disk containing immobilized bacteria, about 20 individual bacterial cells were imaged. In addition, a force volume measurement was conducted on one of these cells after the imaging. The details of force volume experiments have been described in our previous publication,39,55 and similar procedures were employed in the force-volume experiments reported here. Raman Microscopy. These experiments were conducted at the Carnegie Institution in Washington, D.C., using a WITec CRM 200 Raman microscope to acquire Raman spectra and images of HEPESrinsed and dried encapsulated S. typhimurium in air. Spectra of purified EPS from S. typhimurium and HEPES solids were also collected under identical conditions as the controls. Data were acquired with a 543 nm excitation laser (10 mW) at a scan speed of 0.1 s per pixel to minimize sample heating and fluorescence while achieving a diffraction-limited spatial resolution of ∼360 nm.
Results Immobilization of Bacterial Cells. Most bacterial species have a negatively charged cell surface in the physiological pH and ionic environment,5 so it is desirable to immobilize the bacteria on positively charged surfaces such as poly-L-lysine-modified glass or amino-terminated silicon wafers. However, our experience showed that S. typhimurium and E. coli can be easily immobilized on freshly cleaved mica that is negatively charged by incubating the bacterial suspension at ambient conditions. Pretreatment of the mica surface with poly-L-lysine did not significantly enhance the attachment of bacterial cells, suggesting that the adhesion of these bacterial species may be dominated by EPS rather than by a direct electrostatic attraction between cell surface and mica, as speculated in the Discussion section below. Rinsing with deionized water did remove EPS efficiently, probably because of the large aqueous solubility of EPS. However, S. typhimurium H647 is sensitive to osmotic pressure changes, and rinsing with water for even a few seconds resulted in the lysing of most of the bacterial cells, which compromised the cellular integrity. The cellular plasma of lysed cells formed sticky (55) Suo, Z. Y.; Arce, F. T.; Avci, R.; Thieltges, K.; Spangler, B. Dendritic structures of poly(ethylene glycol) on silicon nitride and gold surfaces. Lagmuir 2006, 22, 3844-3850.
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Figure 1. Structures of some compounds used for preparing the buffers and rinsing solutions.
solids with irregular morphology on and around the cell surfaces, making imaging difficult because of the strong adhesion between the AFM tip and the exposed endocellular materials. Therefore, all the bacterial samples were rinsed with buffers or rinsing solutions to reduce osmotic pressure differences between the interiors and the exteriors of the immobilized cells. The chemical structures of some compounds used in our experiments to prepare buffers and rinsing solutions are shown in Figure 1. The attached bacterial cells can be imaged easily using a low-magnification (∼200×) optical microscope incorporated into our AFM. In this setup, the bacterial cells appear as small black dots uniformly distributed over the mica surface. Generally, samples rinsed with water, PBS, MOPS, Tris, or glycine buffers or Na2SO4 solution showed much lower densities of bacterial cells on mica than those rinsed with CaCl2 or BHEP solution or HEPES buffer. High-resolution imaging of individual bacteria and clusters of bacteria revealed that there was a definite correlation between the rinsing solution and the observation of capsular shells around the bacteria. For example, rinsing with HEPES always resulted in at least half of the cells being covered by capsular shells while rinsing with PBS resulted in minimal capsular material and no capsular shells around the bacteria, as described in the next section. Typically, the cell density on mica substrate is higher for samples showing encapsulated bacteria than for those showing only nonencapsulated bacteria except for one special case: the samples rinsed with Na2SO3 solution had the largest number of bacterial cells on mica surface with no capsular shells surrounding the bacteria. In short, not only does the choice of rinsing solution affect capsule formation, it also dictates the bacterial adhesion on mica. Imaging of S. typhimurium Using Tapping Mode. It is desirable to investigate the bacterial capsule in its physiological environment where the native states of the bacterial surface and the capsule are well preserved. However, imaging in liquid suffers from a loss of both lateral and perpendicular resolution,55 not to mention the difficulty and challenges of immobilizing bacteria in an aqueous environment.17,56 Furthermore, bacterial cells, in particular the capsular polysaccharides of Gram negative bacteria, are very soft in their native state and can deform easily upon contact with an AFM probe. The turgor pressure for a live Gram negative bacterial cell is on the order of ∼105 Pa;57 hence, the images obtained from bacterial cells in liquid merely show the compressed contours of the cell walls.26,56 Added to all of these challenges, the fundamental limitations associated with the low Q-value of the oscillating cantilever in liquid58,59 hinder the (56) Doktycz, M. J.; Sullivan, C. J.; Hoyt, P. R.; Pelletier, D. A.; Wu, S.; Allison, D. P. AFM imaging of bacteria in liquid media immobilized on gelatin coated mica surfaces. Ultramicroscopy 2003, 97 (1-4), 209-216. (57) Arnoldi, M.; Fritz, M.; Bauerlein, E.; Radmacher, M.; Sackmann, E.; Boulbitch, A. Bacterial turgor pressure can be measured by atomic force microscopy. Phys. ReV. E 2000, 62 (1), 1034-1044. (58) Chen, G. Y.; Warmack, R. J.; Thundat, T.; Allison, D. P.; Huang, A. Resonance Response of Scanning Force Microscopy Cantilevers. ReV. Sci. Instrum. 1994, 65 (8), 2532-2537.
Figure 2. S. typhimurium rinsed with PBS buffer: (a) height image; (b) section profile along the black line in (a). The height of the capsular EPS ring (∼28 nm) is marked by two black arrow heads; (c) phase image; (d) amplitude image. Location on the capsular EPS ring where the height measurement is made is marked with a white arrow in the height, phase and amplitude images. The scale bars shown in (a), (c), and (d) correspond to 1 µm.
sensitivity of the tip to small variations in soft tissues, hence causing serious degradation in image quality. Therefore, in our experiments, imaging in air was chosen as the practical method for studying the bacterial capsule. Artifacts due to tip contamination20 or tip geometry60 have been reported for bacteria imaging. To avoid these artifacts in our experiments, the tip was replaced on a regular basis or whenever any sign of image degradation was encountered. As briefly described previously, the observation of capsules on S. typhimurium is related to the choice of the rinsing solution. Samples rinsed with PBS, Tris, or glycine buffer or a solution of Na2SO3 or Na2SO4 showed only nonencapsulated cells stuck on the mica surface. Only a very small fraction of the total attached cells were encapsulated when the samples were rinsed with MOPS buffer, and the majority of total cells were nonencapsulated, similar to the results with PBS buffer. As an example, images of a sample rinsed with PBS buffer are shown in Figure 2 in which no capsules except for thin rings surrounding the cells are observed. The bacterial surface and the flagella can be clearly resolved in phase and amplitude images. The texture of the bacterial surface is presumably caused by the dehydration of the cell. Two of the three cells appear to be dividing, as observed in the phase and amplitude images in Figure 2. It is notable that all the cells are surrounded by a thin layer of crust (ring), ∼2030 nm thick, as marked by the white arrows in Figure 2, which is similar to the EPS observed by Beech et al. for sulfate-reducing bacteria.30 These rings are attributed to the remnant capsular EPS. The small particulates, with sizes around 100-200 nm, scattered on and around the cells are attributed to nanocrystals of the buffer salts formed during the drying of the sample subjected (59) Kokavecz, J.; Horvath, Z.; Mechler, A. Dynamical properties of the Q-controlled atomic force microscope. Appl. Phys. Lett. 2004, 85 (15), 32323234. (60) Velegol, S. B.; Pardi, S.; Li, X.; Velegol, D.; Logan, B. E. AFM imaging artifacts due to bacterial cell height and AFM tip geometry. Langmuir 2003, 19 (3), 851-857.
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Figure 4. S. typhimurium rinsed with HEPES buffer (pH 7.4): (a) height image; (b) section profile corresponding to the black line in (a); (c) phase image; (d) amplitude image. Note that all the flagella are broken and part of the capsular EPS is confined by the flagella, as shown on the right side of (c). The scale bars shown in (a), (c), and (d) correspond to 1 µm.
Figure 3. Phase images of S. typhimurium cells rinsed with different solutions: (a) with Tris buffer; (b) with MOPS buffer; (c) with glycine buffer; (d) with Na2SO4 solution; (e) with Na2SO3 solution; and (f) zoomed out image of bacteria rinsed with Na2SO3 solution, showing the hydrophobic areas circling around the cells (see text for details). The scale bars shown in (a-f) correspond to 1 µm.
to the nitrogen gas flow. Similar particulates were observed on mica when a drop of LB medium without bacteria was rinsed with PBS as a negative control. The samples rinsed with a Tris or glycine buffer or a Na2SO3 or Na2SO4 solution resulted in bacterial images similar to those observed with PBS buffer in that there were no capsules around the bacteria, as shown in Figure 3. Those cells rinsed with a Na2SO3 or Na2SO4 solution appeared to lose more EPS than those rinsed with PBS, Tris, or MOPS, leaving behind a discontinuous ring of dots surrounding the cell, as seen for example in Figure 3e,f. In Figure 3f, each bacterium appears to be surrounded by a relatively clean substrate surface containing no buffer particulates. A similar feature is observed for cells rinsed with PBS (Figure 2d). Inside this clean area no evidence of nanocrystals or particulate matter associated with the rinsing salt is observed; at the same time, there is a layer of densely packed crystals or particulate matter formed outside this area. One possible explanation for this observation is that a small amount of the hydrophobic components of the capsular EPS was absorbed on the mica surface during the rinsing procedure, making these areas hydrophobic. This hypothesis is supported by the size of these particle-free areas, which is comparable to the size of the observed capsules around the bacteria (see Figures 4 and 5). These hydrophobic areas then repelled water, preventing the condensation of crystals near the bacteria.
Figure 5. Aggregate of three S. typhimurium cells covered by an EPS capsule. This sample was also rinsed with HEPES buffer (pH 7.4), and almost all the flagella are intact: (a) height image, (b) phase image, (c) amplitude image. Note part of the EPS is confined by the flagella, as shown in the upper right-hand side of (b) and (c). The scale bars shown in (a-c) correspond to 2 µm.
Capsules were observed around S. typhimurium cells when they were rinsed with HEPES buffer (pH 7.4). AFM images of such a cell taken in tapping mode are shown in Figure 4a,c,d. The height image (Figure 4a) shows clearly the contour of the cell plus the capsule without many details of the bacterial surface or the bacterial appendages inside the capsule. The cross-section profile shown in Figure 4b corresponds to the diagonal black line in Figure 4a. The phase image shown in Figure 4c clearly
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resolves the sub-micrometer details associated with the cell surface and the bacterial flagella, as well as the boundaries associated with the bacterial capsule. The wrinkles on the cell surface shown in Figure 4c are clearly visible and have a different pattern than those shown in Figure 2c for cells rinsed with PBS. This difference is most likely due to the slower dehydration of the cell covered by the capsule. The sharp contrast between the capsular material (dark area in Figure 4c) and the rest of the bacterial features is attributed to viscoelastic differences between them. The contrast observed in the phase image is a function of the dehydration conditions of the sample and the amplitude setpoint during tapping mode imaging. For light tapping, i.e., Aset / Ao > 0.8, where Ao is the amplitude of the freely oscillating cantilever and Aset is the amplitude setpoint, the phase image starts to lose the sharp contrast observed under normal experimental conditions. In our experiments the best contrast was observed when the samples were dried under ambient conditions for about 12 h. Images taken immediately after sample preparation were dominated by surface features of capsular EPS. The amplitude image shown in Figure 4d, on the other hand, does not show as much contrast between the capsular material and the rest of the bacterium; however, it differentiates clearly the details of the bacterium and its surroundings, hence supporting and complementing the height and phase images shown in Figure 4a,c. Another interesting observation for HEPES-rinsed cells is the capsular EPS becomes confined within the areas defined by the flagella, as shown in Figure 4c,d. The so-called hydrophobic areas (Figure 3f) observed for samples rinsed with Na2SO3 or another solution that removes EPS were not found for HEPES-rinsed samples. Figures 4c,d also show a number of broken pieces of the flagellum associated with this cell that are scattered on the right-hand side of the cell. These are most likely due to rough handling of the sample during preparation, including rinsing. Figure 5 shows another sample also rinsed with HEPES. Note that almost all the flagella associated with the cells appear to be intact and can be traced to the anchoring points on the cell surfaces. Consistency could be observed for Figures 4 and 5 in the sense that the height images show merely the morphology of the encapsulated microorganisms, while the phase and amplitude images show highly resolved surface features of the cells under the protective coating of the capsular material. In particular, the phase image reveals clear boundaries for the capsular EPS relative to the rest of the bacterial surface features and appendages. Both encapsulated and nonencapsulated cells could be observed for S. typhimurium rinsed with HEPES buffer (pH 7.4), roughly in a ratio of 1:1. However, no encapsulated cell was found for the sample rinsed with the same buffer but with a higher pH value (pH 9.0), as shown in Figure 6a,b. Similarly, if the LB growth medium was removed by centrifugation for 5 min at 5000g and the S. typhimurium cells were resuspended in PBS buffer for 30 min, no capsular features were observed; only nonencapsulated cells similar to those shown in Figure 2 were imaged. On the other hand, when S. typhimurium cells were resuspended in HEPES buffer (pH 7.4) (using a procedure similar to that described for PBS) and observed under AFM, the majority of the cells immobilized on the substrate were encapsulated. However, if the LB growth medium, without bacteria, was used as a control and subjected to the same sample preparation, no capsular features were observed, indicating that the capsular features are directly related to the presence of bacteria. The encapsulation of S. typhimurium cells is not limited to HEPES-exposed samples. Similar encapsulated cells were also observed for samples rinsed with CaCl2 solution and BHEP solution, as shown, respectively, in Figure 6c,d and Figure 6e,f.
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Figure 6. Height and phase images of S. typhimurium rinsed with HEPES buffer (pH 9.0) (a,b), with CaCl2 solution (c,d) and with BHEP solution (e,f). Images (a), (c), and (e) are 3-D height images, and those marked as (b), (d), and (f) correspond to phase images. The scale bars shown in (b), (d), and (f) correspond to 1 µm.
Similar to the HEPES-rinsed samples, these bacterial cells were covered by EPS capsules. These observations help explain the encapsulation phenomenon as elaborated in the Discussion section below. HEPES-stabilized encapsulation is not limited to S. typhimurium. Another Gram negative bacterial species, E. coli, also showed encapsulation when rinsed with HEPES (pH 7.4), as shown in Figure 7a,b. Compared with the encapsulated S. typhimurium (Figures 4c and 5b), the capsule for HEPES-rinsed E. coli contains a smaller amount of capsular EPS. However, it is clearly shown in Figure 7a,b that the E. coli cells were covered with a layer of EPS which extends beyond the edges of cells. On the other hand, the E. coli cells rinsed with PBS showed no encapsulation (Figure 7c,d) and displayed a hydrophobic area surrounding the cell, similar to PBS-rinsed S. typhimurium (Figure 2d). Force-Volume Measurement. In order to shed light on the phase contrast shown in Figures 4c and 5b, the elasticity of the three encapsulated cells shown in the phase image in Figure 8a was studied by the force-volume method: a pair of force vs displacement curves was recorded at each of the 32 × 32 pixels over an ∼5 µm2 area. The corresponding force-volume image generated for the piezo displacement (tip displacement) kept at ∼30 nm is shown in Figure 8b. Three pairs of force vs displacement curves, representing the elastic response of mica, capsular EPS, and encapsulated bacteria, are shown in Figure 8c.
HEPES-Stabilized Encapsulation of S. typhimurium
Figure 7. Height and phase images of E. coli rinsed with HEPES buffer (pH 7.4) (a,b) and with PBS buffer (c,d). Images (a) and (c) are 3-D height images, and (b) and (d) are phase images. Note the EPS capsule in (b) and the hydrophobic area in (d) are similar to those shown in Figures 4c and 5b and Figure 3f, respectively. The scale bars shown in (b) and (d) correspond to 1 µm.
The corresponding indentation curves extracted from these pairs of force vs displacement curves, by subtracting the cantilever deflection from the piezo displacement, are shown in Figure 8d. The maximum cantilever deflection was 30 nm, and the corresponding maximum load was about ∼1200 nN. The force vs displacement curves associated with the capsular EPS and encapsulated cells presented in Figure 8c show hystereses between the loading and unloading curves, suggesting a plastic deformation. In fact, under the loading conditions used in our study, no elastic recovery was observed. This can be seen from the indentation curves extracted from the unloading curves (the dotted lines in Figure 8c) resulting in vertical lines similar to those associated with the mica surface. Mica, being a hard material, shows no visible indentation in the load and indentation regime used in our studies. As the tip presses on the mica surface, the load on the surface increases without a discernible indentation, giving rise to a vertical line for both the loading and unloading processes. Similar vertical lines are observed for the indentation curves corresponding to the unloading force vs distance curves associated with the compressed EPS and with the encapsulated cell. Basically, there is no elastic recovery once the capsular material is compressed under the maximum load. These vertical lines are placed at the ends of the indentation curves shown in Figure 8d for clarity. Notice also that the stiffness (slope of the indentation curve) of the capsular EPS increases rapidly toward the end of its maximum indentation (at ∼46 nm), while the stiffness of cell is slightly smaller at the end of its indentation (at ∼76 nm) than that of the capsular EPS. The size of the indentation is always larger on the cells than on the capsule outside the cells, implying that during tapping mode the AFM tip penetrates deeper into the encapsulated cell than it does into the capsular EPS surrounding the cells. These differences are translated into the contrast observed in the phase images, as described in the Discussion section. Raman Microscopy Results. The encapsulated S. typhimurium (Figures 4 and 5) obtained by rinsing with HEPES buffer (pH 7.4) was also studied with Raman microscopy and spectroscopy.
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Figure 8. Elasticity measurement on encapsulated S. typhimurium in air: (a) tapping mode phase image showing three encapsulated cells; (b) force-volume image of the same cells generated for the piezo displacement at ∼30 nm; (c) three pairs of force vs displacement curves, each pair corresponding to the loading (full line) and unloading (dotted line) trace of the AFM tip. These curves are representative of the elastic responses of a spot on the mica, on the capsular EPS, and on the encapsulated bacterial cell; (d) three pairs of indentation curves, obtained from the corresponding pairs of force vs distance curves shown in (c). Hysteresis is observed between the loading and unloading curves shown in (c) for both the encapsulated cell and the capsular EPS. The hystereses are attributed to plastic deformation of the dehydrated encapsulated cells and capsular EPS. On mica, no such deformation is expected. Note that the indentation curves associated with mica are superimposed vertical lines, as expected because mica is a hard mineral and shows no indentation in the load and indentation regime of interest. Similarly, the indentation curves associated with the unloading curves of cell and capsule are vertical lines, placed at the ends of the indentation curves for clarity. The vertical indentation profile means there is no elastic recovery of compressed capsule or encapsulated cell. The scale bars shown in (a) and (b) correspond to 1 µm.
In Figure 9a, a reflective optical image of a sample also used in our AFM experiments is presented. The dashed square in the image is where we conducted the micro-Raman studies presented in Figure 9b-f. The capsular EPS covering the cells can be visualized in the laser reflection image (Figure 9b) and the total Raman spectrum image (Figure 9c) with moderate resolution. In the image (Figure 9e) corresponding to the C-H stretch (28323031 cm-1), strong Raman signals can be observed for both the encapsulated cells and the capsular EPS (Figure 9f); only the contour of the whole capsule is visible without resolving the individual cells (Figure 9e) because both the encapsulated cells and the capsular EPS undergo strong Raman scattering associated with C-H stretches, as evidenced by their Raman spectra in Figure 9f. In the image shown in Figure 9d, which corresponds to the SdO stretch (1015-1054 cm-1), the bacterial cells under the capsular EPS show the strongest Raman signal, while the capsular EPS outside the cells show a weaker signal; at the same time, most of the mica substrate appears as a dark background with minimal or no emission, hence differentiating the encapsulated bacteria from the capsular EPS. Figure 9f shows four Raman spectra; the lower two spectra were taken on the mica substrate and on the encapsulated cell surface corresponding to the locations marked by the black and white arrows in Figure 9c. The spectrum acquired on the encapsulated cell shows Raman features centered at 1043, 2891, 2944, and 2973 cm-1, which are
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capsular EPS is stabilized by the HEPES molecules via crosslinking the exopolymeric material as discussed below.
Discussion
Figure 9. (a) Reflective optical image of S. typhimurium rinsed with HEPES (pH 7.4). The area inside the square of the white dashed line was imaged with a Raman microscope. (b) Laser reflection image; (c) entire Raman spectrum image; (d) SdO stretch image; (e) C-H stretch image. (f) Raman spectra of HEPES and EPS as controls, and Raman spectra obtained from the sample spots marked by white and black arrows in (c). The scale bars shown in (b), (c), (d), and (e) correspond to 1 µm.
characteristic of the HEPES Raman features, seen in the spectrum taken from a pure HEPES control sample, also shown in Figure 9f. Spectra taken of a second control sample of dried capsular EPS (also presented in Figure 9f) showed somewhat distinct spectral features easily distinguishable from those of HEPES. The point of all of these spectra is that capsular EPS shows spectral features very similar to those of HEPES, indicating that
High-resolution AFM imaging of bacterial cells requires a proper immobilization of these cells on a flat substrate without damaging the integrity of the cellular structure. Bacterial cells are usually attached onto positively charged surfaces, such as poly-L-lysine-covered glass or amino-terminated silicon wafer, via the electrostatic interaction between the negatively charged bacterial surface and the positively charged substrate surface.56,61 However, between the bacterial cell and the substrate surface stands the EPS capsule that also makes a significant contribution to the attachment of bacterial cells to an abiotic substrate.3 In our experiments, the modification of the mica surface with polyL-lysine did not enhance the cell attachment significantly. Considering the sizable EPS capsules surrounding S. typhimurium cells, as shown in Figures 4c and 5b, it is possible that the EPS modifies the mica surface and the adhesion between the bacterial cells and the EPS-modified mica surface overcomes the repulsive interaction between the bacterial cell and the mica surface (both were initially negatively charged). Therefore, when considering the electrostatic charges on opposing surfaces, the role of capsular EPS must be taken into account. As presented above, the presence of capsular EPS around bacteria totally negates the use of polyL-lysine precoating. The rinsing of samples is a key sample preparation step that has notable influence on the stabilization or the removal of the capsular EPS surrounding the bacteria. For samples rinsed with HEPES buffer or CaCl2 or BHEP solution, the bacterial attachment on mica involves encapsulated (together with nonencapsulated) cells. In some cases, a majority of the cells are covered with capsular EPS. Rinsing with a PBS, Tris, or glycine buffer or a Na2SO4 solution resulted in a low cell density on mica, and no encapsulated cell could be observed. These results suggested that capsular EPS plays a positive role in the attachment of S. typhimurium cells on mica. One exception to this conclusion is the samples rinsed with Na2SO3 solution which showed the largest amount of bacterial cell immobilization on mica surface among all the samples studied, even though these cells were all nonencapsulated. The mechanism by which Na2SO3 mediates bacterial attachment onto mica is not clear at this time. It is worthwhile to note that due to the limited number of bacterial cells imaged for each sample, ca. 20 cells, a very small fraction of cells associated with samples rinsed with Tris, glycine, Na2SO3, or Na2SO4 were possibly encapsulated but escaped our detection. For example, for samples rinsed with MOPS buffer, in one case two encapsulated bacterial aggregates were observed even though the majority of the cells were nonencapsulated. However, samples rinsed with HEPES and PBS were prepared many times, and the results were very consistent among different preparations. Bacterial cells exposed to certain chemicals can undergo considerable modifications in their morphology.20 In order to understand the interaction of HEPES with capsular EPS, some idea of the composition of the EPS is imperative. Although this has not yet been thoroughly described for S. typhimurium, it has been reported that colanic acid, an acidic exopolysaccharide, is one of the components of the capsular EPS of both S. typhimurium (61) Arnoldi, M.; Kacher, C. M.; Ba¨uerlein, E.; Radmacher, M.; Fritz, M. Elastic properties of the cell wall of Magnetospirillumgryphiswaldense investigated by atomic forcemicroscopy. Appl. Phys. A 1998, 66, S613-S617.
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and E. coli.62,63 Taking into account the fact that HEPES-stabilized encapsulation has been observed for both S. typhimurium and E. coli, we propose that the encapsulation is a result of crosslinking of the acidic exopolysaccharides of capsular EPS mediated by HEPES molecules via the electrostatic attraction. Furthermore, we assume that the piperazine moiety (the ring structure) of HEPES, with its two nitrogen atoms, serves as the positively charged cross-linker for these acidic exopolysaccharides through electrostatic attraction. One consequence of the cross-linking is assumed to be a considerable reduction in the aqueous solubility of the capsular EPS, possibly due to the increase in the molecular weight and size of the exopolysaccharides, and thus facilitate the formation of the EPS capsule. For bacterial cells rinsed with other buffers, such as PBS or Tris, no cross-linking of EPS was induced because of the lack of piperazine or other linkage moiety, and hence, the majority of EPS was removed by the rinsing processes, resulting in nonencapsulation. In order to confirm the role of the piperazine moiety of HEPES in the cross-linking process, a control experiment was conducted with BHEP, similar to HEPES but without the SO3-, as shown in Figure 1. As anticipated, a large amount of encapsulated S. typhimurium cells was observed by AFM imaging, proving the crucial role of the piperazine moiety in the encapsulation. Furthermore, the S. typhimurium cells were centrifuged to remove the LB growth medium and resuspended in either HEPES or PBS buffer. Similar to the samples rinsed with PBS buffer, no encapsulated cells were found for bacteria resuspended in PBS. On the other hand, the population of the encapsulated cells was enhanced remarkably for S. typhimurium resuspended in HEPES medium, and compared with the bacteria in LB medium rinsed with HEPES, almost all the resuspended cells in HEPES were encapsulated. The extended incubation time in HEPES buffer, ∼30 min, presumably enhanced the cross-linking of EPS and further facilitated the formation of EPS capsules. The resuspension experiments furthermore excluded the possibility that the encapsulation was a result of the reaction between HEPES buffer and components in the LB growth medium. The HEPES-mediated cross-linking of EPS is sensitive to the pH value of the buffer. At pH 9.0, the positively charged nitrogen atom of HEPES is neutralized and the HEPES molecules become negatively charged.64 This negatively charged HEPES moiety is unable to cross-link the acidic exopolysaccharides because they, too, are negatively charged; hence, no encapsulated S. typhimurium were observed when the cells were rinsed with HEPES at pH 9.0 (Figure 6a,b). Similarly, SO32- and SO42- had only negative charges and hence were not able to induce cross-linking of EPS. The question of whether the zwitterionic nature of HEPES contributed to the cross-linking of EPS was tested using two zwitterionic buffers, glycine and MOPS, and neither of them led to any encapsulation. In brief, the piperazine moiety of HEPES is the most likely candidate to mediate the cross-linking process. The existence of HEPES in the dried capsule is confirmed by the results of Raman microscopy. As shown in Figure 9f, the Raman spectra of the encapsulated cells showed characteristic frequencies associated with the HEPES molecule. More importantly, the SdO stretch image could be related to both the capsule and cells. The SdO stretch at 1015-1054 cm-1 was not found
in the purified EPS polymers and was unique to HEPES, so the SdO stretch was strong evidence for the presence of HEPES inside the capsule. The sampling depth of Raman microscopy for transparent samples, such as EPS, is about 1 µm, which is larger than the height of the encapsulated cells. Because of the fast decrease of the capsular EPS thickness (Figure 4b), the edges of the capsule have a smaller effective sampling depth than the encapsulated cells. This might explain why a strong SdO stretch signal was observed at the locations of the cells and a weaker signal was observed where the capsular EPS is located (Figure 9d). Similar to the piperazine moiety in HEPES, calcium ions could also serve as the cross-linker for acidic exopolysaccharides. It is well known that Ca2+ can induce the gelation of another acidic polysaccharide, alginate, by forming a stable well-defined helix structure with poly-R-L-guluronate.65 The elasticity of this Ca2+alginate gel depends on the amount of D-mannuronic acid and L-guluronic acid present in the alginate.66 For bacterial polysaccharides, Korstgens and others report that Ca2+ induces a large increase in the Young’s modulus of the biofilm of P. aeruginosa, which is believed to be due to the cross-linking of alginate by calcium ions.67 In our experiments, the observation of Ca2+induced capsulation (Figure 6c,d) proved the validity of divalent cation Ca2+ as a cross-linker to induce the formation of EPS capsules for S. typhimurium, and this further lends support to the role of HEPES as a cross-linker for the acidic exopolysaccharides during the capsulation. Our results also demonstrated that phase imaging is an effective means of revealing the structural details inside an EPS capsule. In imaging encapsulated bacterial cells with AFM, one concern is to resolve the detailed features inside the EPS capsules. Usually the height image gives only a three-dimensional topographic contour of the encapsulated cell surface and is not sensitive to the small features underneath the surface, such as the flagella and/or fimbriae inside the bacterial capsule, especially when these are combined with sharp variations in the surface morphology (Figures 4a and 5a). In this case, phase imaging raced to the rescue, resolving the features inside the EPS capsule because of its sensitivity to the viscoelastic heterogeneity of the sample. As shown in Figures 4c, 5b, and 6d,f, the EPS capsules of cells could be differentiated in the phase images, where they were in sharp contrast to the encapsulated cell and its appendages. Phase imaging also clearly revealed the edge between the cell and the EPS polymer. It has been observed that for a given sample the amplitude setpoint has significant influence on phase contrast. With fairly hard tapping, the phase signal delay is largely related to the viscoelastic properties of the surface.40 In our experiment, the amplitude setpoint was set in such a way as to exert fairly hard tapping (Aset/Ao ≈ 0.6) on the samples, and a large contrast could be observed between the capsule and the cell, which implies that the dissipative processes associated with the tip-surface interactions were different for the EPS capsule, the encapsulated cell surface, and the flagellum. During tapping mode imaging the speed of the AFM tip in the medium is ∼104 µm/s, which probes the dynamic viscoelastic properties of the medium; on the other hand, during force-volume imaging the speed is reduced by 4 orders of magnitude, which then reveals
(62) Sutherland, I. W. Structural studies on colanic acid, common exopolysaccharide found in enterobacteriaceae, by partial acid hydrolysis - oligosaccharides from colanic acid. Biochem. J. 1969, 115 (5), 935-945. (63) Grant, W. D.; Sutherland, I. W.; Wilkinson, J. F. Exopolysaccharide colanic acid and its occurrence in the Enterobacteriaceae. J. Bacteriol. 1969, 100 (3), 1187-1193. (64) Good, N. E.; Winget, G. D.; Winter, W.; Connolly, T. N.; Izawa, S.; Singh, R. M. M. Hydrogen ion buffers for biological research. Biochemistry 1966, 5 (2), 467-477.
(65) Draget, K. I.; Stokke, B. T.; Yuguchi, Y.; Urakawa, H.; Kajiwara, K. Small-angle x-ray scattering and rheological characterization of alginate gels. 3. Alginic acid gels. Biomacromolecules 2003, 4 (6), 1661-1668. (66) Donati, I.; Holtan, S.; Morch, Y. A.; Borgogna, M.; Dentini, M.; SkjakBraek, G. New hypothesis on the role of alternating sequences in calcium-alginate gels. Biomacromolecules 2005, 6 (2), 1031-1040. (67) Korstgens, V.; Flemming, H. C.; Wingender, J.; Borchard, W. Influence of calcium ions on the mechanical properties of a model biofilm of mucoid Pseudomonas aeruginosa. Water Sci. Technol. 2001, 43 (6), 49-57.
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the velocity-independent elastic properties of the system.68 During phase imaging the dissipative processes are the major contribution to the phase contrast.69 In this sense the penetration depth of the tip into the viscoelastic medium and hence the time the tip spends in the dissipative medium and the interaction of the tip with the medium all contribute to the phase contrast. Typically, the higher the dehydration, the stiffer the medium becomes. In our force-volume experiments on encapsulated S. typhimurium cells, there was a discernible difference between the stiffness (slope of the indentation curve) of the capsular EPS and that of the encapsulated cells (see Figure 8d). This difference is partly due to the effective depth of indentation and partly to the differential dehydration of the polysaccharides surrounding the encapsulated cells and the cells themselves.70,71 The differential dehydration is expected to be a function of how long the encapsulated cells are exposed to the ambient conditions. This explains why waiting under ambient conditions for ∼12 h improved the phase contrast. It is also clear that the penetration depth of the AFM tip into the encapsulated cells is much larger than that into the capsular EPS around the edges, as is clearly demonstrated in the indentation curves presented in Figure 8d. Furthermore, the hystereses shown in Figure 8c suggest a larger deformation energy for the tip-cell interactions than for the tip-capsule interactions because the size of the area confined between the loading and unloading curves is larger for force vs displacement curves obtained on cell than that on capsule. All (68) Tsukruk, V. V.; Gorbunov, V. V.; Huang, Z.; Chizhik, S. A. Dynamic microprobing of viscoelastic polymer properties. Polym. Int. 2000, 49 (5), 441444. (69) Aime, J. P.; Boisgard, R.; Nony, L.; Couturier, G. Influence of noncontact dissipation in the tapping mode: Attempt to extract quantitative information on the surface properties with the local force probe method. J. Chem. Phys. 2001, 114 (11), 4945-4954. (70) Wilkinson, K. J.; Balnois, E.; Leppard, G. G.; Buffle, J. Characteristic features of the major components of freshwater colloidal organic matter revealed by transmission electron and atomic force microscopy. Colloids Surf., A 1999, 155 (2-3), 287-310. (71) Santschi, P. H.; Balnois, E.; Wilkinson, K. J.; Zhang, J. W.; Buffle, J.; Guo, L. D. Fibrillar polysaccharides in marine macromolecular organic matter as imaged by atomic force microscopy and transmission electron microscopy. Limnol. Oceanogr. 1998, 43 (5), 896-908.
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of these nanoscale differences in viscoelastic properties, effective penetration depth of the AFM tip, and interactions of the tip with the viscoelastic medium are expected to give rise to the contrast at nanoscale in the phase imaging as presented here.
Summary We studied the HEPES-stabilized encapsulation of S. typhimurium and E. coli by imaging encapsulated bacterial cells with AFM in tapping mode. The phase and amplitude images taken in air revealed structural details of the cells buried under the capsular EPS surrounding the cell and the phase image taken differentiated the capsular EPS from the features of the encapsulated cell with remarkable contrast. Rinsing is a key step in sample preparation to obtain encapsulated bacteria. In the case of S. typhimurium cells rinsed with HEPES buffer, BHEP, or CaCl2 solution, CaCl2 facilitated the formation of capsules covering the cells, while no encapsulation of the cells was observed if the bacteria were rinsed with PBS, Tris, glycine buffers, or a solution of Na2SO4, Na2SO3, or deionized water. Although the number of encapsulated cells was significantly enhanced when S. typhimurium cells were resuspended in HEPES (pH 7.4), no encapsulation was observed if a HEPES buffer with increased pH value (pH 9.0) was used. The encapsulation process was hypothesized to be the result of a cross-linking of the acidic exopolysaccharides of S. typhimurium mediated by the positively charged piperazine moiety of HEPES and BHEP molecules or a divalent cation such as Ca2+. In contrast, negatively charged ions such as SO32-, SO42-, and HPO42-, and monovalent cations, including Na+ and Tris+ did not cross-link the acidic exopolysaccharides of EPS or promote the encapsulation of cells. Acknowledgment. This work is funded by NASA-EPSCOR under Grant No. NCC5-579 and in part by U.S. Public Service Grant AI-41123, and in part by Montana Agricultural Station and USDA Formula Funds. The authors acknowledge helpful discussions with Prof. I. B. Beech on bacterial EPS. LA0621721
