J. Phys. Chem. B 1998, 102, 5947-5950
5947
Silver Colloids Impregnating or Coating Bacteria S. Efrima*,†,‡ and B. V. Bronk‡ Department of Chemistry, Ben Gurion UniVersity, P.O. Box 653, Beer-SheVa, Israel 84105, and B. V. Bronk, U.S. Air Force Research Laboratory at US ERDEC, SCBRD-RT, E5951, APG, Maryland 21010-5423 ReceiVed: March 6, 1998; In Final Form: June 2, 1998
We produce silver nanocolloids selectively within and on Escherichia coli bacteria. Silver preferentially concentrates in the bacteria, distributes uniformly within them, or, alternatively, forms a rough coating over them. Oscillations in polarized light scattering vs scattering angle are more pronounced and shift to larger angles, compared to a control that has no silver. Very intense surface-enhanced Raman signals are observed for E. coli with a “wall colloid”. The main bands can be associated with peptides and polysaccharides in the cell wall and its membrane. This work was extended to Gram-positive Bacillus megaterium.
Introduction In this communication we demonstrate that deposits of metallic silver may be generated selectively inside or on bacteria in a controlled manner and discuss several aspects of the behavior of the combined metal-bacteria system. The methods developed here are potentially of wide general use, for example, in the study of the biochemistry and biophysics of bacteria or in material science. Specifically, the methods can be applied to the identification of bacteria, on one hand, as well as to the control of the production of colloids in materials processing, on the other hand, in line with the directions outlined recently by Mann.1 Deposition of inorganic materials in biological organisms is essential for a variety of natural phenomena such as biomineralization,2 which is also important geologically,3 and accumulation of minerals by bacteria,4 which has industrial implications. It is also important for laboratory applications (e.g., staining techniques5). Under the proper circumstances silver colloids provide extremely large enhancements of Raman spectra of adsorbed molecules, yielding highly detailed information concerning the molecular environments near the solid surface.6 Such spectra might also be used as fingerprints for categorizing and identifying bacteria. Here we describe production of silver colloids selectively in bacteria or on their surface and discuss the behavior of the combined system with respect to Raman scattering and polarized light scattering. Results and Discussion Electron Microscopy. Figure 1 shows an electron micrograph of Escherichia coli with a silver colloid deposited mainly on the bacterium wall (“wall colloid”). The TEM measurements are done to determine the location of the silver deposits when compared to various controls that show the bacterial image as much less dense. Thus the micrographs shown here are made without any of the usual preparations to enhance the image. They utilize only the silver from the experiment to add density to the bacterial image. The silver colloid-bacteria combination is achieved by soaking washed bacteria freshly grown to the * To whom correspondence should be addressed. E-mail: efrima@ bgumail.bgu.ac.il. † Ben Gurion University. ‡ U.S. Air Force Research Laboratory.
Figure 1. TEM of E. coli with a wall colloid. The colloid was produced using 5 × 10-4 M AgNO3.
late logarithmic phase in 0.05 M sodium borohydride, followed by spinning them down (at 2200g for 10 min) and resuspension in distilled water to get rid of the excess reagent in the solution. After an additional centrifugation, the bacteria are resuspended in a 5 × 10-4 M silver nitrate solution. The reductant within the cells reacts with the incoming silver nitrate and forms a colloid and a subsequent deposit predominately where the two diffusion fronts meet, at the bacterial wall. Evidently, as Figure 1 shows, this process deposits a rough film, coating the bacteria and causes some sticking together of the bacteria. As the shapes of the bacteria are still partly discernible, we infer that the thickness of the silver coating is usually smaller than ∼0.05 µm. In contrast, when an “internal colloid” is formed, by first saturating washed bacteria with a 0.05 M silver nitrate solution, spinning down twice, resuspending in water to wash away the free solution silver ions, and only then adding the reductant to the suspension of the bacteria, a uniform colloid forms predominately inside the bacteria, with the bacterial shape being rather well preserved (Figure 2). The bacteria remain mostly separated, and there is no evidence for bridging between them. In this case the silver ions in the cell are not mobile, probably being attached to various nitrogen-, sulfur-, or carboxylate-rich bonding sites, as is known from silver colloidal chemistry. This is evidenced by the enrichment of the cell with silver ions and
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5948 J. Phys. Chem. B, Vol. 102, No. 31, 1998
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Figure 4. UV-vis extinction of E. coli with an internal colloid and a wall colloid. The background due to scattering was subtracted. Silver concentration in the parent reagent solution is 0.05 M. Figure 2. TEM of E. coli with an internal colloid. The colloid was produced using 0.05 M AgNO3.
Figure 3. TEM of a silver colloid produced in E. coli as an internal colloid and then released into solution from damaged cells. Shown is a region of Formvar on the Lacey Formvar/carbon 300 mesh copper grids. The colloid was produced using 0.05 M AgNO3.
the fact that even following the soaking of the bacteria in highly dionized water the silver is still retained in it. Also silver ion toxicity at very low concentrations attests to the same conclusion. These sites also are expected to serve as convenient nucleation sites for the colloid. A quantitative analysis of the intensity profile along a bacterium in the latter case shows it to be consistent with a uniform silver distribution within undeformed, essentially cylindrical bacteria. Typical bacterial length from the electron micrographs in the case of an internal colloid is about 2 µm, and the diameter of the bacteria is about 1 µm. Both results are in good agreement with literature values for E. coli K12 in the stationary phase.7 The apparent diameter is about 2-fold smaller for the bacteria with the wall colloid. It will take further experimentation to determine the cause of the difference. Figure 3 shows a micrograph of a silver colloid produced as an internal colloid, released into solution from damaged cells, filtered through a 0.22 µm filter, and deposited on the Formvar regions of Lacey Formvar/carbon-coated copper grids (Pelco International 01883-F 300 mesh grids). The average radius is 1 ( 0.6 nm. Larger particles (radius 2-5 nm) are observed to preferentially deposit onto the carbon fibers on the grids. UV-Visible Absorption. Figure 4 shows a UV-visible absorption spectrum (scattering background subtracted) from a
suspension of E. coli cells in which an internal colloid or a wall colloid formed. In both cases a signature of a silver nanocolloid with maximum at 415-430 nm is observed. Comparison of the two spectra indicates that the internal colloid most probably consists of well-isolated particles, as indicated by the symmetric, featureless, and relatively narrow extinction band. Similar spectra are seen often but at much lower silver concentrations (∼10-4 M).8 The wall colloid is aggregated, giving a wider, less symmetric band. The structure seen in the band is characteristic of chainlike aggregates. A comparison of Figure 4 to the absorption spectrum of a cell-free colloid obtained from the supernatant (not shown) indicates that the particles that escape to the supernatant are similar to those inside the cells. TEM shows that the internal colloid consists of predominately spherical particles with diameters in the range 1-4 nm. In the absence of the bacteria and using the same silver and reductant concentrations, one obtains only a black silver precipitate. Thus the bacteria have a profound effect on the production of the colloid, its size distribution, and its stability. Elemental Analysis. Elemental analysis giving total organic carbon (by a Durham DC-80 Total Organic Carbon System) and silver (using a Perkin-Elmer Plasma II Inductively Coupled Plasma) shows a significant enrichment of the cells by silver, compared to the ambient solution from which the silver was extracted in each case. For an “internal colloid” we typically find in the cell silver/carbon atom ratios of 0.02-0.05 and concentration enrichment factors of ∼10 and ∼1600 for 0.05 and 0.0005 M AgNO3, respectively (assuming that the dry weight is ∼16% of the total cell weight). This falls in line with data of the uptake of silver ions by E. coli K12 reported in the context of toxicity studies9 and silver resistant bacteria.4,10 In the case of the wall colloid, the silver/carbon atom ratios are in the range 0.06-0.1. For ∼16% dry weight in a 0.5 µm diameter cylindrical cell, this corresponds to an average silver coating of 1-2 nm. As the TEM (Figure 1) clearly shows surface roughness of 50 nm, the coating probably is not continuous but similar to the well-known island film deposits of silver.11 Furthermore, the UV-visible spectrum (Figure 4) shows features characteristic of aggregated and single silver particles, in line with the average thickness of 1-2 nm we estimated from the elemental analysis. Polarized Light Scattering. The S34/S11 ratio in the polarized elastic light scattering, where Sij is the ij component of the scattering, “Mueller” matrix,12,13 has been shown to depend on bacterial size, in addition to its optical coefficients.7 Figure 5 shows the results for bacteria impregnated with a silver colloid (internal colloid) compared to a control of E. coli that was treated similarly, except that water was used instead of a silver
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Figure 5. Light scattering: E. coli impregnated with a silver colloid, control, and cyanide-treated. The initial silver concentration in the reagent solution is 0.05 M.
Figure 6. Raman spectrum of E. coli with a wall colloid. The background has been subtracted out. The initial silver concentration in the reagent solution is 0.05 M.
nitrate solution, and another control where a cyanide treatment dissolved the colloid from the impregnated bacteria. The oscillations as a function of the scattering angle are much more pronounced when the bacteria are impregnated with the colloid, and a shift to higher angles occurs. After the silver is dissolved, the scattering profile reverts almost to that of the untreated control. Similar features were obtained with E. coli treated to obtain a wall colloid. The silver colloid increases the difference between the optical constant of the bacteria and that of the aqueous solution in which they are suspended. Therefore it is plausible to expect changes in the scattering profile. The shift to larger angles, however, does not result from a possible change of the refractive index, as shown by a preliminary calculation based on Mie scattering for a composite particle made up of cell material and silver nanoparticles. Thus, it appears that size changes are responsible, but this will require further investigation. In any event, impregnating bacteria with silver colloids clearly enhances the intensity of the polarized scattering signature and can provide a system for a further study of the relationship between scattering and small biological particle attributes. Surface-Enhanced Raman Scattering. Figure 6 shows the Raman spectrum obtained from E. coli coated with a silver colloid. The system consists of a Coherent Innova 90/5 laser, a Jarrell Ash MonoSpec 27 single stage monochromator (Model 82499) with a ruled grating of 600 gr/mm, one or two Kaiser Holographic SuperNotch filters, HSNF-514.5, and a PI 512 element intensified diode array. This spectrum is very intense, orders of magnitude stronger than that obtained from untreated bacteria. Typically it takes just a few seconds to obtain this spectrum using 1-2 mW argon laser light power (at 514.5 nm),
J. Phys. Chem. B, Vol. 102, No. 31, 1998 5949 while untreated cells give much weaker spectra using even ∼100 mW laser power and 30-60 min exposures. The large intensities are due to surface-enhanced Raman scattering (SERS): i.e., the spectra were enhanced by the presence of the silver colloid, as has been seen for many other molecules and biomolecules in the past.6,14 To our knowledge this is the first report of SERS of bacterial cells. The main vibrational bands are seen at 1612(s), 1558(w), 1500-1520(s), 1442(w), 1390(sh), 1332(vs), 1274(s), 1146(s), 1080(s), 808(m), 732(m), 612(m), 530(s), 450(w), 290(m), and 210(m) cm-1. The intensities are denoted in parentheses by w, weak; m, medium; sh, shoulder; s, strong; vs, very strong. A noticeable resemblance to the UV resonance Raman of E. coli reported by Nelson’s group15 is observed. However, the intensity ratios are quite different, as expected from the different mechanisms, and some of the bands are different. They assign the various bands to groups associated either with aromatic amino acids or to the nucleotide bases in the nucleic acids (mostly RNA), as these moieties are expected to be the active chromophores. They also discuss the difficulty involved in the assignments. It is noteworthy that considering the number and diversity of biomolecules in the bacteria cell one would expect a highly congested Raman spectrum. However, SERS is selective, as is UV resonance Raman spectroscopy, the latter showing bands associated with the chromophores, while the former shows predominately those molecules and functional groups that are in the immediate proximity of the metal surface. It is expected that in general the colloid will form mainly at sites that are rich in thiol, amino, or carboxylate groups (i.e. amino acids, nucleic bases, etc.) that may serve as convenient bonding and nucleation sites for the emerging colloid. Furthermore, a wall colloid will favor the molecular components of the wall and membrane (such as peptidoglycan) over nucleic acids that are located internally. Indeed, some of the major bands we observe can be associated with amino acids15 (1612, 1558, 1500-1520 cm-1). The 1486 band, the strongest peak usually associated with A + G (adenine and guanine), is absent in our spectra, demonstrating the preferential enhancement of the wall components rather than the internal RNA. Similarly, a 1242 cm-1 band associated with U (uracyl) is not observed by us at all. We do see an intense band at 1332 cm-1 close to 1335 cm-1, which Britton et al.15 associate with A + G. In fact, it is the most intense band in our spectra. However medium or strong bands at about this position were seen in the Raman spectra and the SERS of several proteins, like albumin, globulin and lysozyme,16-18 peptides19,20 and amino acids such as cystine.21 Thus the 1332 cm-1 band we observe can be attributed to proteins and protein components. Britton et al.15 also observed a weakening of the 1486, 1335, and 1242 cm-1 signals when using the shorter wavelength excitations (at 231 and 222 nm). They proposed that at these wavelengths “the outer cell wall, which contains materialss especially aromatic amino acidsswhich absorb” this light, “attenuates the incident beam to an extent sufficient to cause a major reduction in scattering from regions within the cytoplasm”. This explanation is consistent with our findings with a wall colloid as described above. We observe additional bands at 1442, 1390, 1274, 1080, 808, 732, 612, 540, 450, 290, and 210 cm-1, many of which (especially the first three bands in the list) are observed in the SERS spectra of proteins, peptides, and amino acids.17-19,21 Thus the SERS spectrum is particularly rich. Some tentative assignments are (symmetric) carboxylate stretch at 1400-1430 cm-1,
5950 J. Phys. Chem. B, Vol. 102, No. 31, 1998 amide III band, alkane CH2 twist and rock modes and C-O-C modes of pyranose rings around at 1270 cm-1, alkane C-C stretch at 1080 cm-1, and skeletal and ring breathing modes at the lower Raman shifts.22 These assignments are in line with the cell wall providing the major contribution to the spectra we measure. Note that the 1008 cm-1 band associated with the aromatic ring breathing mode is missing in our spectra. The excitation wavelength of 514.5 nm is too long to resonate with excitations of these rings. Unlike in the case of UV resonance Raman, their scattering is not significantly enhanced above that of most other amino acids, which are the majority. It is more difficult to obtain SERS from bacteria impregnated with the internal colloid. It seems that, unlike on the wall, the silver particles within the cell are isolated and do not produce the appropriate conditions for enhancement. It is well-known that major enhancements are obtained in colloidal systems only when they are allowed to aggregate to some extent.23 The UVvisible spectra of the bacteria-colloid systems corroborate this conclusion. The spectrum of an internal colloid is characteristic of an ensemble of well-isolated particles, while that from the wall colloid shows significant aggregation. It stands to reason that within the cell, when there is a limited supply of silver ions, only separated particles form. In the case of a wall colloid, there is practically an inexhaustible supply of mobile silver ions, which react at the surface of the bacteria. With this situation in mind, very recently we did manage to obtain SERS from E. coli impregnated with an internal colloid, by adding silver ions to bacteria that were already impregnated with a silver colloid and then repeating the reduction. A weak but definitely enhanced Raman spectrum appeared that is quite different from the one obtained with the wall colloid. We are now trying to optimize the preparation protocol to achieve better spectra that can be easily analyzed. Other Experiments. We repeated most of this study for a nonsporulating strain of the Gram-positive bacterium Bacillus megaterium. We find essentially the same results as with E. coli. We can form internal colloids and wall colloids. Electron microscopy again shows striking differences in the results of the two methods of colloid preparation. However, owing to the larger size of this bacterium, we cannot see directly whether the silver colloid is indeed uniformly distributed in the cell for the internal colloid. Silver impregnated or coated B. megaterium also show shifts in the polarized light scattering to larger angles and large intensities, similarly to E. coli. Also SERS spectra are obtained for a wall colloid. The spectra are essentially the same as for E. coli, reflecting the basic similarity of cell and cell-wall chemical composition for even very different bacterial species. There are definite differences, most probably related to the difference between the wall thickness of the Gram-positive versus the Gram-negative bacteria. It is hoped that such spectra will offer a novel opportunity to study the molecular environment within a bacterium in a detailed and intimate way. The differences between the spectra might be used to distinguish between bacterial species and provide a convenient means of identification. Summary In summary, we showed here that metal silver colloids can form within and on bacteria. On one hand, the bacteria modify the properties of the colloid, which is important for material science. Specifically, stable nanocolloidal dispersions are obtained under conditions that yield only powdery deposits in
Letters the absence of the bacteria. On the other hand, the colloid modifies the optical and spectroscopic response of the bacteria and offers new opportunities for studying details of bacterial structure. This approach seems to be general, as it applies equally well to E. coli and B. megaterium, which are very different. We believe that other colloids (other metals, semiconductors, etc.) can be used in a similar fashion. The intense SERS spectra can be very useful in the study of bacteria at various phases of their life cycle, or of their response to a variety of stimulants. They might also be useful in the study of metalstaining techniques and metal toxicity, as well as of colloid immunological tagging and therapy. Acknowledgment. This work was conducted in the framework of the National Research Council associateship program, in the USAF Armstrong laboratory at Aberdeen, MD. We thank J. Cze’ge’, N. Feay, R. Herd, Z. Z. Li, O. I. Sindoni, M. Milham, T. Coleano, and J. Slunt for assistance and expertise in various aspects of these studies. References and Notes (1) Mann, S. Nature 1993, 365, 499. (2) Addadi, L.; Mazur, S. Angew. Chem., Int. Ed. Engl. 1992, 31, 153. (3) Hess, P. P. Mar. Geol. 1994, 117, 1. See also preface to: Mann, S., Webb, J., Williams, R. J. P., Eds. Biomineralization, Chemical and Biochemical PerspectiVes; VCH: New York, 1989. Lowenstam, H.; Weiner, S. On Biomineralization; Oxford University Press: New York, 1989. (4) Belly, R. T.; Kydd, G. C. In Developments in Industrial Microbiology, Vol. 23. Proceedings of the 38th Meeting of the Society of Industrial Microbiology, August 9-14, 1981, Richmond, VA; Chapter 55, p 567. Pooley, F. D. Nature 1982, 296, 642. Goddard, P. A.; Bull, A. T. Appl. Microbiol. Biotechnol. 1989, 31, 308. Slawson, R. M.; Van Dyke, M. I.; Lee, H.; Trevors, J. T. Plasmid 1992, 27, 72. Holmes, J. D.; Richardson, D. J.; Saed, S.; Evansgowing, R.; Russell, D. A.; Sodeau, J. R. Microbiology 1997, 143, 2521 and references therein. (5) Hayat, M. A. Stains and Cytochemical Methods; Plenum Press: New York, 1993. (6) Efrima, S. In Modern Aspects of Electrochemistry; Conway, B. E., White, R., Bockris, J. O’M., Eds.; Plenum Press: New York, 1985; Vol. 16, p 253. (7) Bronk, B. V.; Van De Merwe, W. P.; Stanley, M. Cytometry 1992, 13, 155. (8) Mulvaney, P.; Linnert, T.; Henglein, A. J. Phys. Chem. 1991, 95, 7843. Ershov, B. G.; Janata, E.; Henglein, A. J. Phys. Chem. 1993, 97, 339. Gutierrez, M.; Henglein, A. J. Phys. Chem. 1993, 97, 11368. (9) Ghandour, W.; Hubbard, J. A.; Deistung, J.; Hughes, M. N.; Poole, R. K. Appl. Microbiol. Biotechnol. 1988, 28, 559. (10) Bridges, K.; Kidson, A.; Lowbury, J. L.; Wilkins, M. D. Brit. Med. J. 1979, 1, 446. Pumpel, T.; Schinner, F. Appl. Microbiol. Biotechnol. 1986, 24, 244. (11) Rouard, R. P.; Messen, A. Prog. Opt. 1977, XV, 79. (12) Bohren, C. F.; Huffman, D. R. Absorption and Scaterring of Light by Small Particles; Wiley: New York, 1983. Bronk, B. V.; Van De Merwe, W. P.; Huffman, D. R. In Modern Techniques for Rapid Microbiological Analysis; Nelson, W., Ed.; VCH Publishers: New York, 1992; p 171. (13) Van De Merwe, W. P.; Li, Z. Z.; Bronk, B. V.; Cze’ge’, J. Biophys. J. 1997, 73, 500. (14) Cotton, T. M.; Kim, J. H.; Chumanov, G. D. J. Raman Spectrosc. 1991, 22, 729. (15) Britton, K. A.; Dalterio, R. A.; Nelson, W. H.; Britt, D.; Sperry, J. F. Appl. Spectrosc. 1988, 42, 782. (16) Ahern, A. M.; Garrell, R. L, Langmuir 1991, 7, 254. (17) Grabbe, E. S.; Buck, R. P. J. Am. Chem. Soc. 1989, 111, 8362. (18) Chumanov, G. D.; Efremov, R. G.; Nabiev, I. R. J. Raman Spectrosc. 1990, 21, 43. (19) Herne, T. M.; Ahern, A. M.; Garrell, R. L. J. Am. Chem. Soc. 1989, 113, 846. (20) Lee, H. I.; Suh, S. W.; Kim, M. S. J. Raman Spectrosc. 1988, 19, 491. (21) Watanabe, T.; Maeda, H. J. Phys. Chem. 1989, 93, 3258. (22) Lin-Vien, D.; Colthup, N. B.; Fateley, W. G.; Grasselli, J. G. The Handbook of Infrared and Raman Characteristic Frequencies of Organic Molecules; Academic Press: San Diego, CA, 1991. Scharder, B. Raman/ Infrared Atlas of Organic Compounds, 2nd ed.; VCH: Weinheim, 1989. (23) Blatchford, C. G.; Campbell, J. R.; Creighton, A. J. Surf. Sci. 1982, 120, 435.
