Effect of pH on the Electrophoretic Mobility of Spores ...

Effect of pH on the Electrophoretic Mobility of Spores of Bacillus anthracis and Its Surrogates in Aqueous Solutions Colin P. White,a Jonathan Popovici,a Darren A. Lytle,b Noreen J. Adcock,b and Eugene W. Ricec Department of Biological Sciences, University of Cincinnati, Cincinnati, Ohio, USAa; ORD, NRMRL, WSWRD, U.S. Environmental Protection Agency, Cincinnati, Ohio, USAb; and ORD, NHSRC, WIPD, U.S. Environmental Protection Agency, Cincinnati, Ohio, USAc

The electrophoretic mobility (EPM) of endospores of Bacillus anthracis and surrogates was measured in aqueous solution across a broad pH range and several ionic strengths. EPM values trended around phylogenetic clustering based on the 16S rRNA gene. Measurements reported here provide new insight for Bacillus anthracis surrogate selection and for attachment/detachment and transport studies.

iven the national security and public health concerns associated with Bacillus anthracis, research on its mobility in the environment, treatment effectiveness, and decontamination approaches is necessary. However, because of its limited availability, surrogate organisms are often relied upon for research. The properties of a surrogate for any species must be carefully selected prior to experimentation. Optimally, the surrogate will have similar characteristics, including cell electrophoretic mobility (EPM), and therefore behave similarly in the test environment, to the original species (1, 13, 18). Bacterial endospores are of particular interest, as they are generally more resistant to disinfection than vegetative cells, thus allowing for persistence in the environment, and in the case of B. anthracis, they have been identified as both a bioterrorism agent and an agent of public health concern (3, 18, 26, 36, 37). Although studies have reported the hydrophobicity of vegetative cells and their endospores, few have made systematic measurements of the cell’s electrokinetic properties (e.g., electrophoretic mobility [EPM]) (1, 10, 11, 16, 17, 23, 24, 26–29). Determining the EPM of microorganisms is an integral part of elucidating cell physiology and mobility in the environment. The cell’s surface charge has been shown to be directly related to coagulation, disinfection, environmental transport, adhesion to surfaces, and uptake of chemicals (2, 8, 9, 12, 14, 33). As cell surface charge is difficult to measure directly, EPM, used in this study, is often used as a proxy measurement (27–29, 43). The fate and transport of microorganisms and spores in soil, ground and surface waters, and surfaces used in food preparation are of interest. Reports have shown that the surface properties (hydrophobicity and EPM) of microorganisms are fundamental in attachment and detachment mechanisms, thus controlling movement (3, 26, 32, 38, 42). The electrokinetic properties of microorganisms in drinking water are of particular interest, as they have been linked to degraded drinking water quality and waterborne disease outbreaks and related to water treatment effectiveness (25, 30, 34–36, 40, 41). In addition to indigenous bacteria, select agents are also of concern from a biosecurity perspective (31, 36, 37, 40). Although the EPMs of some microorganisms have been reported, interpretation and correlation of results to date have been confounded by the array of growth and sporulation conditions, culture purification techniques, and suspension test buffers, all of which can significantly impact EPM. There have been a limited

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number of published studies on the surface charge of B. anthracis (7, 11). Recently, Da Silva et al. (11) reported zeta potential values for spores of B. anthracis Sterne(pAFp8gfp) in three solutions, but no comparisons were made with other surrogate species. Zeta potential rather than electrophoretic mobility presents a challenge when data are being interpreted; the zeta potential calculation requires the use of the Smoluchowski approximation to convert the directly measured EPM value to zeta potential. Conversion of EPM to zeta potential requires the experimental conditions to fall with the limits of the Smoluchowski model. This calculation is often performed by the instrument whose exact method of calculation is often elusive. Herein, we present electrophoretic mobility measurements of endospores of attenuated Bacillus anthracis and related species using microelectrophoretic-phase light scattering under comparable conditions. EPMs were measured in several buffers with different ionic strengths across a pH range of 2 to 10. Bacteria used in this study were B. anthracis Sterne 34F2 (received from Laura Rose at the Centers for Disease Control and Prevention [CDC]), B. cereus ATCC 14579, B. atrophaeus subsp. globigii (received from Dugway Proving Grounds), B. megaterium ATCC 14581, B. subtilis subsp. subtilis ATCC 6051, and B. thuringiensis var. israelensis ATCC 35646. Species-level identification was confirmed using biochemical or molecular assays (data not shown). All organisms were cultured in generic sporulation broth (8.0 g nutrient broth, 0.040 g MnSO4, and 0.100 g CaCl2 per liter of deionized water) with agitation at a temperature sufficient to produce spores, as determined by phase-contrast microscopy. Spores were then washed through a series of centrifugations (relative centrifugal force [RCF] of 5,900 for 15 min each time) in cold, sterile deionized water. Purified spore preparations were then stored at 4°C in sterile deionized water until use, typically within 24 h. Scanning electron microscopy was used to determine the presence of exosporia. Briefly, washed spores were air dried on a glass slide,

Received 29 May 2012 Accepted 12 September 2012 Published ahead of print 21 September 2012 Address correspondence to Colin P. White, [email protected]. Copyright © 2012, American Society for Microbiology. All Rights Reserved. doi:10.1128/AEM.01337-12

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KH2PO4 and dechlorinated tap water (D). , B. anthracis Sterne; Œ, B. subtilis; , B. cereus; }, B. megaterium; , B. atrophaeus subsp. globigii; , B. thuringiensis. Error bars represent 1 standard deviation.

heat fixed, and sputter coated with gold-palladium and imaged using a JEOL 6490LV microscope (JEOL, Peabody, MA). EPM measurements were made using a Malvern Zetasizer Nano ZS90 (Malvern, Worcestershire, United Kingdom) that was calibrated daily using a manufacturer-provided standard. Two milliliters (approximately 107 spores ml⫺1) of purified spores were added to 100 ml of 9.15 mM, 91.5 mM, and 150 mM KH2PO4 in deionized water and to municipal tap water that had been dechlorinated with granular activated carbon (DTW) (Cincinnati, OH). The suspension of spores in buffer was then computer titrated using either 0.6 N HCl or 0.6 N NaOH, with mixing, in single-pH-unit increments from pH 2 to 10. Care was taken to add only HCl or NaOH, depending on the starting pH of the buffer. One milliliter of sample was collected at each pH increment and immediately analyzed using the ZS90 at 25 ⫾ 1°C.

The results show differences in EPM between the species tested as well as a clear delineation of three groups for a given condition (Fig. 1). In the weakest phosphate buffer (9.15 mM), two EPM grouping trends formed within the genetically related B. cereuslike clade (B. anthracis, B. cereus, B. thuringiensis). B. anthracis Sterne had the lowest (most positive) EPM across the entire pH range tested, with values from 0.045 to ⫺0.72 ␮m · cm V⫺1 s⫺1, forming a unique group within the clade, averaging ⫺0.42 ␮m · cm V⫺1 s⫺1 (Fig. 1; Table 1). Forming the second group, B. thuringiensis and B. cereus were closely related, with average EPMs of ⫺1.20 ␮m · cm V⫺1 s⫺1 and ⫺1.07 ␮m · cm V⫺1 s⫺1, respectively. The final group was comprised of B. atrophaeus subsp. globigii, B. megaterium, and B. subtilis, which averaged ⫺2.84 ␮m · cm V⫺1 s⫺1, ⫺2.20 ␮m · cm V⫺1 s⫺1, and ⫺2.88 ␮m · cm V⫺1 s⫺1, respectively. Isoelectric points of spores were found to generally fall be-

TABLE 1 Summary of experimental measurements of Bacillus sporesa EPM (␮m · cm V⫺1 s⫺1) at pH 7 Condition

B. anthracis Sterne (ND)b

B. subtilis (⫺)

B. megaterium (⫹)

B. thuringiensis (⫹)

B. cereus (⫹)

B. atrophaeus subsp. globigii (⫺)

Conductivity (mS/cm)

KH2PO4 9.15 mM 91.5 mM 150 mM

⫺0.6 ⫾ 0.03 ⫺0.24 ⫾ 0.04 ⫺0.27 ⫾ 0.12

⫺3.22 ⫾ 0.03 ⫺2.38 ⫾ 0.01 ⫺2.31 ⫾ 0.14

⫺3.02 ⫾ 0.2 ⫺2.64 ⫾ 0.16 ⫺2.27 ⫾ 0.3

⫺1.68 ⫾ 0.08 ⫺1.02 ⫾ 0.26 ⫺0.74 ⫾ 0.16

⫺1.6 ⫾ 0.09 ⫺1.04 ⫾ 0.09 ⫺0.9 ⫾ 0.09

⫺3.5 ⫾ 0.08 ⫺2.31 ⫾ 0.24 ⫺2.04 ⫾ 0.17

1.47 ⫾ 0.08 10.81 ⫾ 0.78 15.27 ⫾ 1.58

DTW

⫺0.64 ⫾ 0.05

⫺1.85 ⫾ 0.06

⫺1.96 ⫾ 0.14

⫺1.23 ⫾ 0.08

⫺1.05 ⫾ 0.02

⫺1.87 ⫾ 0.06

0.51 ⫾ 0.13

n ⫽ 6 for all assays. All values are means ⫾ standard deviations. b ⫹ or ⫺, presence or absence of an exosporium; ND, not determined. a


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FIG 1 Effect of pH on electrophoretic mobilities of endospores in various buffers. Endospores were suspended in 9.15 mM (A), 91.5 mM (B), and 150 mM (C)

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surrogate” for all types of experimentation on B. anthracis may not exist, a well-characterized group of surrogates will help researchers choose the appropriate surrogate species for their particular study. The data presented here should help researchers interpret results obtained in studies requiring the use of surrogates as well as aiding in the selection of appropriate surrogates. ACKNOWLEDGMENTS We acknowledge Amanda Jennings for assistance in culture and preparation of spores and Emily Nauman of Pegasus Technical Services for editorial comments. Any opinions expressed in this paper are those of the authors and do not necessarily reflect the official position and policies of the U.S. EPA. Any mention of products or trade names does not constitute recommendation for use by the U.S. EPA.

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low pH 3 (Fig. 1). EPMs exhibited pH dependence below pH 6, steeply increasing in the positive direction with decreasing pH. Above pH 6, only a slight change in the positive direction was observed. The trends were toward a consistent relative charge but an increase in EPM in the positive direction with increasing buffer strength. Suspending spores in DTW affected EPM. Specifically, the range of EPM was decreased, presumably due to a higher ionic strength and complex mixture of ions in solution. The organic load and alkalinity may have also had an effect via surface sorption to the spores. Care must be taken in evaluating the EPMs measured in drinking water, as water chemistry can change. For instance, corrosion and seasonal changes can have drastic effects on water quality from tap to tap, changing the ionic species present in the water. It is important to note that the data presented here form groups that mirror phylogenetic clustering. It has been reported that B. cereus, B. anthracis, and B. thuringiensis are closely related based on 16S rRNA phylogenetic analysis (19). The data here present a phenotypic property that supports this hypothesis, with these three species forming a EPM group distinct from the more distantly related B. atrophaeus subsp. globigii, B. subtilis, and B. megaterium. Spore morphology, including features such as size, density, and the presence or absence of exosporia, must be considered in surrogate selection (5, 18). The EPM groupings observed in this study may be the result of the presence or absence of an exosporium and nap-like appendages exterior to the spore coat (Table 1). B. cereus, B. anthracis Sterne, and B. thuringiensis have all been shown to produce either an exosporium, appendages, or a combination of the two, and these properties have been correlated with increased hydrophobicity (4, 6, 15, 20, 22, 24, 39). Further research is needed to elucidate the effect of natural organic matter on EPM as well as matrix interaction studies to evaluate the interaction of spores and substrate in the environment. Such studies could assist in environmental attachment/detachment and transport models. For example, the data presented here, in the context of spore persistence, support the findings of Hugh-Jones and Blackburn (21). Spores of B. anthracis survive best in humus-rich, slightly alkaline, calciferous soils; the observed negative charge in the higher pH range supports the idea that spores would attract positively charged cations, which would favor maintaining calcium in the spore core matrix and extend viability and germinative capacity. Conversely, as the magnitude of the negative charge decreases with decreasing pH, this could disrupt the equilibrium, causing cations to leach out of the spore, resulting in loss of viability and also weakening adherence to humus particles. The results presented here add new evidence supporting the selection of B. thuringiensis (18) as the most appropriate surrogate for B. anthracis in fate and transport studies (18). These findings represent a comprehensive set of electrophoretic mobilities of Bacillus endospores with practical applications in ecology, food science, physiology, and engineering. The data also show that EPM can be impacted by water chemistry, and care must be taken to ensure that these variables are constant when data are collected and compared. Measurements of endospores of virulent strains of B. anthracis are needed to fully evaluate the data presented here in the context of surrogate selection for virulent strains. Though a “universal

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