JOURNAL OF CLINICAL MICROBIOLOGY, Nov. 2002, p. 4372–4374 0095-1137/02/$04.00⫹0 DOI: 10.1128/JCM.40.11.4372–4374.2002 Copyright © 2002, American Society for Microbiology. All Rights Reserved.
Vol. 40, No. 11
Comparison of Two Silver Staining Techniques for Detecting Lipopolysaccharides in Polyacrylamide Gels Li Tan and Parwinder S. Grewal* Department of Entomology, The Ohio State University, Wooster, Ohio 44691 Received 7 June 2002/Returned for modification 8 July 2002/Accepted 27 August 2002
nella enterica serovar Typhimurium were purchased from Sigma Chemical Company, St. Louis, Mo. LPS preparations were treated for 5 min at 100°C in 0.05 M Tris-HCl buffer (pH 6.8) containing 2% (wt/vol) SDS, 10% (wt/vol) sucrose, and 0.01% bromophenol blue. Ten microliters of each sample was then loaded on precast Ready Gel Tris-HCl polyacrylamide gels (86 by 68 by 1.0 mm) containing 4 and 15% acrylamide in the stacking and separating gels, respectively (Bio-Rad Laboratories, Inc., Hercules, Calif.). Electrophoresis was performed at 12 mA in the stacking gels and 25 mA in the separating gels until the bromophenol blue had run about 6.7 cm. LPSs in the gels were visualized by either the classic method (11) or the modified method (2). The sensitivities of the two methods were compared by using from 50 ng to 5 g of M. osloensis LPS (Fig. 1). The LPS was revealed to be a rough-type LPS, because only one main band was detected in the gels by both methods. The band patterns obtained at 50 ng by the classic method were equivalent to or better than those obtained at 1 g by the modified method. Therefore, the classic method is at least 20 times more sensitive than the modified method for detecting M. osloensis LPS. The sensitivities of the two methods were also compared by
The classic silver staining method for detecting bacterial lipopolysaccharides (LPSs) in polyacrylamide gels (C. Tsai and C. E. Frasch, Anal. Biochem. 119:115-119, 1982) was at least 20 times more sensitive than the modified silver staining method (A. Fomsgaard, M. A. Freudenberg, and C. Galanos, J. Clin. Microbiol. 28:2627-2631, 1990) for detecting LPS from the bacterium Moraxella osloensis. However, the classic method is only about three to four times more sensitive than the modified method for detecting LPSs from Escherichia coli J5, EH100, and O111:B4 or Salmonella enterica serovar Typhimurium. The reduction of sensitivity is due to omission of the initial fixing step in the modified method. The retention of LPS fractions in the gels during fixing and/or oxidation may depend on the structures of their lipid A moieties. Lipopolysaccharides (LPSs) play a major role in the pathogenesis of gram-negative infections (1). Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) followed by silver staining has been used extensively to characterize LPS (3, 11, 12). Tsai and Frasch (11) first reported a highly sensitive classic silver staining method for detecting LPS in polyacrylamide gels. This classic method can detect even less than 5 ng of the rough type of LPS. However, Fomsgaard et al. (2) revealed that the classic method did not stain certain LPS preparations containing a low number of fatty acids, which were washed out of the gels during the initial fixing step (40% ethanol–5% acetic acid, overnight). Thus, they developed a modified silver staining method by omitting the fixing step and increasing the LPS periodic acid oxidation (the second step) time from 5 min to 20 min to restore the ability to detect all LPSs. We purified LPS from Moraxella osloensis, a bacterium associated with a slug-parasitic nematode, Phasmarhabditis hermaphrodita (8, 9). M. osloensis LPS is an active endotoxin against the slug Deroceras reticulatum (10). Analysis of different quantities of M. osloensis LPS or four commercially available LPSs from other bacteria by SDS-PAGE followed by each of the two methods revealed that the modified method was less sensitive than the classic method. M. osloensis LPS was purified by classical phenol-water extraction (13), with modification as described by Gu et al. (3), from 3-day pure cultures of M. osloensis supplied by MicroBio, Ltd., Cambridge, United Kingdom. LPS preparations from Escherichia coli J5, EH100, and O111:B4 strains and Salmo-
FIG. 1. Comparison of sensitivities of two silver staining methods for detecting M. osloensis LPS. (A) Modified method. (B) Classic method. The lanes contain the following amounts of M. osloensis LPS: 1, 50 ng; 2, 100 ng; 3, 1 g; 4, 5 g. The figure was created with Adobe Photoshop 5.5 software (Adobe Systems Inc., 1999).
* Corresponding author. Mailing address: Department of Entomology, Ohio State University, OARDC, Wooster, Ohio 44691. Phone: (330) 263-3963. Fax: (330) 263-3686. E-mail:
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FIG. 2. Comparison of sensitivities of two silver staining methods for detecting E. coli J5 or EH100 LPS. (A) Modified method. (B) Classic method. Lanes 1 to 4 contain the following amounts of E. coli J5 LPS: 1, 50 ng; 2, 100 ng; 3, 1 g; 4, 5 g. Lanes 5 to 8 contain the following amounts of E. coli EH100 LPS: 5, 50 ng; 6, 100 ng; 7, 1 g; 8, 5 g. The figure was created with Adobe Photoshop 5.5 software (Adobe Systems, Inc., 1999).
using the same quantities of rough-type LPS from E. coli J5 or EH100 (Fig. 2). The band patterns obtained at 50 ng by the classic method were better than those obtained at 100 ng by the modified method. However, the band patterns obtained at 1 g by the classic method were less clear than those obtained at 5 g by the modified method. Thus, the classic method is about three to four times more sensitive than the modified method for detecting the two rough-type LPSs. A similar level (three to four times) of difference in LPS detection sensitivities between the two methods was also observed for the smooth-type LPS from E. coli O111:B4 or S. enterica serovar Typhimurium (Fig. 3). M. osloensis LPS in the gels was also visualized by the modified method, except that the periodic acid oxidation time was increased from 20 min to 100 min (Fig. 4). With the increase in oxidation time, the band patterns obtained at 1 or 5 g of LPS became better and clearer. It is estimated that the sensitivity of the modified method for detecting M. osloensis LPS increases about one to two times with the increase in the oxidation time from 20 min to 100 min. Thus, increasing the oxidation time of LPS from 5 min to 20 min in the modified method at least should not reduce LPS detection sensitivity. Therefore, the reduction in sensitivity is due to the omission of the initial fixing step in the modified method. It is not fully clear why the detection sensitivities of the two methods differ for M. osloensis LPS and the other LPSs tested. The four purchased LPSs have different structures of polysaccharide moiety, but had similar levels (three to four times) of difference in LPS detection sensitivity by the two methods. This could be due to the same structure of lipid A moiety being present in the four LPSs (5, 7). The structure of M. osloensis
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FIG. 3. Comparison of sensitivities of two silver staining methods for detecting E. coli O111:B4 or S. enterica serovar Typhimurium LPS. (A) Modified method. (Note that the background was overstained.) (B) Classic method. (Note that a small piece of the gel was lost in lane 8.) Lanes 1 to 4 contain the following amounts of E. coli O111:B4 LPS: 1, 50 ng; 2, 100 ng; 3, 1 g; 4, 5 g. Lanes 5 to 8 contain the following amounts of S. enterica serovar Typhimurium LPS: 5, 50 ng; 6, 100 ng; 7, 1 g; 8, 5 g. The figure was created with Adobe Photoshop 5.5 software (Adobe Systems, Inc., 1999).
LPS is unknown. Since the structure of lipid A for a given bacterial genus usually exhibits constant characteristics (6), the structure of lipid A from M. osloensis LPS can be forecast from the known lipid A structure for Moraxella catarrhalis LPS (4). Thus, it is forecast that lipid A from M. osloensis LPS contains seven fatty acids, including four 3-hydroxydodecanoic acids, two decanoic acids, and one dodecanoic acid. In contrast, lipid A from the purchased LPS only contains six fatty acids, including five 3-hydroxytetradecanoic acids and one dodecanoic acid (5, 7). Fomsgaard et al. (2) suggested that the retention of LPS fractions in the gels during fixing and/or oxidization may be a property of the number of fatty acids present in their lipid A moiety. However, since the lipid A from the purchased LPS
FIG. 4. Effects of 20 (A) and 100 (B) min of periodic acid oxidation on sensitivity of the modified silver staining method for detecting M. osloensis LPS. Lanes 1 and 2 contain 1 and 5 g, respectively, of M. osloensis LPS. The figure was created with Adobe Photoshop 5.5 software (Adobe Systems, Inc., 1999).
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contains the same number of fatty acids as M. osloensis lipid A, the retention of LPS fractions in the gels during fixing and/or oxidation may depend on the structure of their lipid A moiety (e.g., fatty acid pattern or conformation of lipid A), but not the number of the fatty acids. It is possible that those LPSs containing a low number of fatty acids are fixed weakly or rarely in the gels, whereas M. osloensis LPSs are fixed much more slowly than the other LPSs tested, thus requiring more time (e.g., overnight) to be fixed mostly or completely in the gels. We conclude that each of the two methods has its advantages and disadvantages. The classic method is more sensitive, but time-consuming. Furthermore, it does not detect those LPSs containing a low number of fatty acids. In contrast, the modified method is simpler and faster and detects LPS that would not be stained by the classic method. However, the present results reveal that the modified method has lower LPS detection sensitivity than the classic method for all LPSs tested, especially that from M. osloensis. Therefore, it is suggested that unknown bacterial LPS preparations in the gels be visualized by both methods. This work was supported by a Graduate Research Competitive Grant from the Ohio Agricultural Research and Development Center and by a Presidential Fellowship to L. Tan. REFERENCES 1. Enright, M. C., and H. McKenzie. 1997. Moraxella (Branhamella) catarrhalis: clinical and molecular aspects of a rediscovered pathogen. J. Med. Microbiol. 46:360–371.
2. Fomsgaard, A., M. A. Freudenberg, and C. Galanos. 1990. Modification of the silver staining technique to detect lipopolysaccharide in polyacrylamide gels. J. Clin. Microbiol. 28:2627–2631. 3. Gu, X.-X., C.-M. Tsai, M. A. Apicella, and D. J. Lim. 1995. Quantitation and biological properties of released and cell-bound lipooligosaccharides from nontypeable Haemophilus influenzae. Infect. Immun. 63:4115–4120. 4. Holme, T., M. Rahman, P. E. Jansson, and G. Widmalm. 1999. The lipopolysaccharide of Moraxella catarrhalis: structural relationships and antigenic properties. Eur. J. Biochem. 265:524–529. 5. Holst, O., S. Muller-Loennies, B. Lindner, and H. Brade. 1993. Chemical structure of the lipid A of Escherichia coli J-5. Eur. J. Biochem. 214:695– 701. 6. Rietschel, E. T., and O. Luderitz. 1980. Struktur von Lipopolysakkarid und Taxonomie Gram-negativer Bakterien. Forum Mikrobiol. 1:12–20. 7. Takayama, K., N. Qureshi, and P. Mascagni. 1983. Complete structure of lipid A obtained from the lipopolysaccharides of the heptoseless mutant of Salmonella typhimurium. J. Biol. Chem. 258:12801–12803. 8. Tan, L., and P. S. Grewal. 2001. Infection behavior of the rhabditid nematode Phasmarhabditis hermaphrodita to the grey garden slug Deroceras reticulatum. J. Parasitol. 87:1349–1354. 9. Tan, L., and P. S. Grewal. 2001. Pathogenicity of Moraxella osloensis, a bacterium associated with the nematode Phasmarhabditis hermaphrodita, to the slug Deroceras reticulatum. Appl. Environ. Microbiol. 67:5010– 5016. 10. Tan, L., and P. S. Grewal. 2002. Endotoxin activity of Moraxella osloensis against the grey garden slug, Deroceras reticulatum. Appl. Environ. Microbiol. 68:3943–3947. 11. Tsai, C., and C. E. Frasch. 1982. A sensitive silver stain for detecting lipopolysaccharides in polyacrylamide gels. Anal. Biochem. 119:115–119. 12. Vaneechoutte, M., G. Verschraegen, G. Claeys, and A.-M. Van Den Abeele. 1990. Serological typing of Branhamella catarrhalis strains on the basis of lipopolysaccharide antigens. J. Clin. Microbiol. 28:182–187. 13. Westphal, O., and K. Jann. 1965. Bacterial lipopolysaccharides: extraction with phenol-water and future applications of the procedure. Methods Carbohydr. Chem. 5:83–91.
