Effects of antibacterial mineral leachates on the cellular ultrastructure, morphology, and membrane integrity of Escherichia coli and methicillin-resistant Staphylococcus aureus
© Otto et al; licensee BioMed Central Ltd. 2010
Received: 27 July 2010
Accepted: 16 September 2010
Published: 16 September 2010
We have previously identified two mineral mixtures, CB07 and BY07, and their respective aqueous leachates that exhibit in vitro antibacterial activity against a broad spectrum of pathogens. The present study assesses cellular ultrastructure and membrane integrity of methicillin-resistant Staphylococcus aureus (MRSA) and Escherichia coli after exposure to CB07 and BY07 aqueous leachates.
We used scanning and transmission electron microscopy to evaluate E. coli and MRSA ultrastructure and morphology following exposure to antibacterial leachates. Additionally, we employed Bac light LIVE/DEAD staining and flow cytometry to investigate the cellular membrane as a possible target for antibacterial activity.
Scanning electron microscopy (SEM) and transmission electron microscopy (TEM) imaging of E. coli and MRSA revealed intact cells following exposure to antibacterial mineral leachates. TEM images of MRSA showed disruption of the cytoplasmic contents, distorted cell shape, irregular membranes, and distorted septa of dividing cells. TEM images of E. coli exposed to leachates exhibited different patterns of cytoplasmic condensation with respect to the controls and no apparent change in cell envelope structure. Although bactericidal activity of the leachates occurs more rapidly in E. coli than in MRSA, LIVE/DEAD staining demonstrated that the membrane of E. coli remains intact, while the MRSA membrane is permeabilized following exposure to the leachates.
These data suggest that the leachate antibacterial mechanism of action differs for Gram-positive and Gram-negative organisms. Upon antibacterial mineral leachate exposure, structural integrity is retained, however, compromised membrane integrity accounts for bactericidal activity in Gram-positive, but not in Gram-negative cells.
With the advent of antibiotics in the early 20th century, morbidity and mortality from bacterial infections were dramatically reduced in the industrialized world. In recent decades, these advances have been tempered by the rapid, widespread emergence of microorganisms that are resistant to multiple, commonly used antibiotics . As our arsenal of effective antibiotics is diminishing, the pursuit of novel therapeutic agents is becoming progressively more urgent.
Minerals have been utilized in traditional medicine for centuries as topical treatments for cutaneous wounds, digestive treatments for gastrointestinal ailments, nutritional supplements, and for removal of toxins from the body [2–4]. Traditionally, the mechanism of mineral-based healing activities has been attributed to physical properties, such as the expansive surface area and resulting highly adsorptive properties of clays present in the mixtures .
Recently, various mineral products marketed for their health benefits have been investigated for their potential antimicrobial properties [5–8]. However, only a small number of clay products have been shown to be antibacterial and the mechanism of antibacterial activity has been elucidated for very few of these products . Falkinham et al.  attributed the antibacterial effects of Jordan's red soils to bacteriocins produced by bacteria present in the clays. It was hypothesized that application of the red soil to an infected area of the skin allowed the inherent organisms to proliferate, produce bacteriocins, and thus kill the infectious pathogens . Mpuchane et al. [7, 9] tested a total of 102 clays from South Africa and determined that only nine of these clay samples had antibacterial activity. The antibacterial properties of these South African medicinal clays were attributed to the low pH environment of the hydrated mineral suspensions (pH < 4), and it was further postulated that metal cations could contribute to toxicity [7, 9]. While Mpuchane et al.  determined that nine clays had antibacterial properties, none of the clays specifically sold for use against bacterial infections had antibacterial activity. Therefore, it is essential to scientifically validate the efficacy of these mineral products prior to use in a clinical setting.
Clay minerals are excellent adsorbent materials due to their small particle size (< 2 μm), stable layered structure, and high cation exchange capacity . In a pH-dependent manner, exchangeable cations can bind to the clay surface, balancing the negative charge of the clay structure. In hydrated suspensions, the adsorbate can then be released into the aqueous solution, varying the cationic composition of the solution [10, 11]. These released metal ions are known to have toxic effects on bacteria by competing with essential enzyme cofactors, irreversibly binding biological molecules to inhibit function, replacing ions essential to membrane stabilization, and inducing DNA mutations [12–15]. For example, metal cations, such as iron, copper, and chromium, have been implicated in production of elevated levels of reactive oxygen species which can lead to DNA damage, lipid peroxidation, protein oxidation, and eventual cell death [16–18]. Metal ion toxicity varies with pH and appears to be related to changes in ion species that occur as the pH is adjusted [12, 15, 19]. These alterations in toxicity are due to the relative abilities of the ion species to bind cell surfaces and exert their effects .
In a prior study, we identified two mineral mixtures, arbitrarily designated BY07 and CB07, that exhibit antibacterial activity . From these mineral mixtures, we prepared aqueous leachates that contain metal ions released from the clay minerals, but are absent of all solid particles. These leachates retain antibacterial activity, establishing that the mechanism of action is dependent on chemical, not physical interactions . Further investigations revealed that the antibacterial activity of BY07 and CB07 mineral mixtures is related to the pH-dependent bioavailability of toxic metal ions in a low pH environment . While we have discovered that pH-dependent ion toxicity mediates CB07 and BY07 antibacterial activity, further investigations must be performed to fully understand the precise mechanism of action. In this study, we assessed whether cell lysis occurs in E. coli and MRSA cells during leachate exposure and investigated cellular membrane integrity as possible mechanisms of action of the aqueous leachates.
Bacterial strains and growth conditions
E. coli ATCC 25922, obtained from the American Type Culture Collection, and MRSA, obtained from Sonora Quest Laboratories (Tempe, AZ, USA), were used for all studies as previously described . E. coli was grown on Luria-Bertani (LB) agar or in LB broth, and MRSA was grown on trypticase soy agar (TSA) or in trypticase soy broth (TSB). Both bacterial strains were grown at 37°C with gentle rotary mixing.
Mineral leachate preparation
Mineralogical and major chemical characterization of the CB07 and BY07 mineral mixtures has been previously described . Briefly, the CB07 mineral is primarily composed of quartz (45.5%), illite (19.8%), and calcium smectite (17.2%), while the BY07 mineral primarily consists of calcium smectite (37.3%), anorthoclase feldspar (23.0%), and quartz (13.7%) . Major oxide chemical analyses reveal that CB07 and BY07 are primarily composed of silicon, aluminum, iron, calcium, sodium, potassium, and sulfur . Leachates of CB07 and BY07 mineral samples were prepared as previously described . Briefly, 1 g of autoclaved minerals was vigorously agitated in 20 mL of UV-irradiated, ultrapure, deionized H2O (dH2O) for 18 - 24 hours at room temperature. The suspension was centrifuged at 31,000 × g for 3 h to remove the remaining insoluble minerals and then sterilized by passage through a 0.22 μm filter.
Antibacterial susceptibility testing of mineral leachates
E. coli and MRSA exponential phase cultures were prepared by diluting overnight cultures into fresh growth medium to a concentration of 107 CFU/mL and continuing growth at 37°C with gentle rotary mixing until the cultures reached mid-logarithmic phase of growth. Bacterial cells were collected by centrifugation, washed once in phosphate-buffered saline (PBS), and suspended in the appropriate leachate solution or sterile dH2O at an initial concentration of 107 CFU/mL. Initial CFU concentrations were confirmed by plating the control bacterial population and enumerating colonies after 24 h incubation at 37°C. Due to experimental sample processing, the 0 h experimental exposure times were ~ 3 min. Experimental samples were incubated at 37°C with gentle rotary mixing for a specified time, and cell survival was determined by plating duplicate 10-fold serial dilutions for each sample at appropriate time points and enumerating colonies after 24 h incubation at 37°C.
Scanning electron microscopy (SEM)
Bacterial cells were prepared for SEM as described above, with the exceptions that cultures were collected at late logarithmic phase of growth and initial concentrations were 108 CFU/ml. Following 24 h exposure to the leachates, washed cells were inoculated onto a poly-L-lysine-coated coverglass slide and allowed to adhere for 5 min at room temperature. After washing the slide in 50 mM sodium phosphate buffer, pH 7, the cells were chemically crosslinked onto the slide in 2% gluteraldehyde (buffered in 50 mM sodium phosphate, pH 7). The immobilized cells were then fixed in 2% osmium tetroxide for 15 min at room temperature, washed three times in 50 mM sodium phosphate buffer, and dehydrated in 5 min washes in a sequential acetone series (20%, 40%, 60%, 80%, 3× 100%). The samples were critical point dried in a Balzers 020 critical point dryer, attached to aluminum mounting stubs, sputter coated with gold-palladium, and imaged with an XL30 Environmental SEM equipped with a field emission gun. A minimum of 200 cells was counted from each of three independent replicates.
Transmission electron microscopy (TEM)
E. coli and MRSA exponential phase cultures were prepared as described above for SEM with an initial concentration of 108 CFU/mL. Following 24 h exposure to the leachates, cells were fixed in 2% gluteraldehyde buffered in 50 mM phosphate, pH 7, for 2 h at room temperature. The cells were then washed in 50 mM phosphate and resuspended in 1% agarose (final concentration). The agarose-embedded cell pellets were fixed in 2% osmium tetroxide (buffered in 50 mM phosphate) for 2 h at room temperature, washed three times in 50 mM phosphate buffer, washed three times in dH2O, and en bloc stained in 0.5% uranyl acetate overnight at 4°C. The pellets were dehydrated in 10 min washes with a sequential acetone series (20%, 40%, 60%, 80%, 3× 100%) and infiltrated with Spurr's resin. Thin sections (70 nm) were cut using an Ultracut R ultramicrotome (Leica Microsystems, Vienna, Austria). Sections were captured on formvar-coated, 300-mesh copper grids, post-stained in uranyl acetate and Sato's lead citrate, and observed on a Philips CM12 TEM at 80 kV. A minimum of 60 cells was counted from each of three independent replicates.
Flow cytometric measurements
To evaluate the membrane integrity of E. coli and MRSA following exposure to the leachates, the Bac Light LIVE/DEAD membrane permeability kit (Invitrogen, Carlsbad, CA, USA) was used following the manufacturer guidelines. E. coli and MRSA mid-logartithmic phase cultures were prepared as described above and harvested at an initial concentration of 108 CFU/mL. A standard curve was prepared by mixing live (0.85% saline-exposed) cells and dead (40% isopropanol-exposed) cells together at various proportions of live:dead cells (100%, 75%, 50%, 25%, 0% alive). Following exposure to the leachates or control conditions, cells were incubated in 5 μM SYTO9 and 30 μM propidium iodide (PI) for 15 min in the dark and then immediately subjected to flow cytometric analysis. E. coli cells were analyzed following 1 h exposure to CB07 leachate (CB07-L) and 6 h exposure to BY07 leachate (BY07-L), while MRSA cells were analyzed following 15 h exposure to either CB07-L or BY07-L. These time points represent the exposure time required for bactericidal activity (≥ 99.9% killing) of the different leachates against the two cell types. A Cytomics FC 500 flow cytometer (Beckman Coulter, Inc., Brea, CA, USA) fitted with a 488 nm excitation laser was used for membrane permeability analyses. Green fluorescence was detected on channel FL1 with a 525 nm bandpass filter. Red fluorescence was detected on channel FL3 with a 620 bandpass filter. Since the SYTO9 dye emits a strong signal at a wavelength of 600 nm, it overlaps with the PI emission . Therefore, membrane permeabilization is determined by a horizontal population shift that occurs down the green fluorescent intensity axis. For each series of flow cytometric measurements, 50,000 cells were counted and analyzed.
Antibacterial mineral leachates
Scanning electron microscopy
As shown in Figure 1b, MRSA was completely killed after 24 h exposure to the leachates. SEM images of MRSA demonstrated intact cells following exposure to CB07-L and BY07-L (Figures 2h and 2j), with damaged or lysed cells observed in 2.5% and 6.8% of cells, respectively (Figure 3b). MRSA cells incubated in water for 24 h showed flattened and distorted cells with bleb-like structures or deposits (Figure 2d), while cells in broth and low pH buffer exhibited a smooth cell surface appearance (Figures 2b and 2f). In contrast, following exposure to the leachates, the MRSA cell surface appeared rough, showed the appearance of bleb-like structures, and had an increased abundance of extracellular debris (Figures 2h and 2j). Further, 95.4% and 94.5%, respectively, of CB07-L- and BY07-L-treated cells exhibited bleb-like structures, while only 11.1% of cells grown in TSB showed blebs (Figure 3b).
Transmission electron microscopy
TEM images of MRSA confirmed that the cells remain intact following exposure to the leachates (Figures 4h and 4j), indicating that cell lysis is not the antibacterial mechanism of action in Gram-positive cells. Enumeration of triplicate TEM sample images verified that 100% of leachate-exposed cells remain intact (Figure 5b). Cytoplasmic disruption of MRSA (Figure 4h and 4j; black arrows) following exposure to CB07-L and BY07-L was observed in 96.7% and 92% of cells, respectively (Figure 5b). Possible disruptions in the cytoplasmic membrane (Figures 4h and 4j; white arrows) were observed in 67.3% and 89.0% of CB07-L- and BY07-L-treated cells, respectively (Figure 5b). Cells with a distorted shape were also observed in triplicate samples of MRSA exposed to the mineral leachates. When compared to the broth control, a decreased frequency of dividing cells was observed in leachate-exposed MRSA cells (Figure 5b). Moreover, many of these leachate-exposed dividing cells showed evidence of a distorted septum (Figure 5b). Following 24 h exposure to dH2O, MRSA viability decreased by 1-log10 unit (Figure 1b). Water-exposed MRSA cells exhibited evidence of hypo-osmotic environmental stress through cell lysis, wavy cell envelope structures, and separation of cytoplasmic contents from the membrane (Figure 4d). Notably, these ultrastructural alterations were not evident in the broth-exposed or leachate-exposed cells (Figures 4b,h, and 4j), thus indicating that leachate-induced toxicity differs significantly from prolonged cell incubation in water.
CB07-L and BY07-L generate low pH environments ranging between 3.3 - 3.7 . To evaluate the effects of the low pH on cellular ultrastructure, we exposed E. coli and MRSA to a 100 mM phosphate buffer at pH 3.6 to mimic the low pH environment of the leachates. After 24 h exposure to low pH buffer, E. coli viability was reduced by 3-log10 units  as compared to the complete loss of viability observed after 24 h exposure to CB07-L and BY07-L (Figure 1a). This maintenance of viability was expected since E. coli exhibits an inducible acid tolerance in order to facilitate passage through the low pH environment of the digestive tract . Low pH buffer-exposed E. coli displayed different patterns of cytoplasmic condensation from that of the leachate-exposed cells (Figures 4e,g, and 4i). In contrast to E. coli, MRSA cells exposed to the low pH buffer for 24 h were killed completely (data not shown). TEM images of low pH buffer-exposed MRSA cells revealed an even distribution of cytoplasmic contents (Figure 4f), similar to cells grown in broth (Figure 4b), demonstrating that the toxic effects induced by a low pH environment differ from that of the antibacterial leachates. Images of low pH buffer-exposed MRSA cells exhibited mesosome-like structures (Figure 4f; striped arrow), however, this effect was only observed in the low pH buffer-exposed cells and was likely a processing artifact due to exposure to the low pH phosphate buffer.
While electron microscopy (EM) provides useful insight into the mechanism of action of antibacterial agents, the resulting images are observational only. Other techniques must, therefore, be used in tandem to verify the observations generated from the EM images. Accordingly, we investigated the effects of BY07-L and CB07-L on the membrane permeability of E. coli and MRSA by using the Bac Light LIVE/DEAD bacterial viability kit. This assay uses two DNA intercalating dyes: green fluorescent SYTO9, which penetrates all membranes and red fluorescent propidium iodide (PI), which can only penetrate permeabilized membranes due to its large size and negative charge . Red fluorescence is produced in the membrane-permeabilized cell by combined displacement of SYTO9 by PI and quenching of SYTO9 emission by fluorescence resonance energy transfer (FRET) .
Summary of E. coli survival and membrane permeability following incubation in LB, water, pH 3.6 phosphate buffer, and mineral leachates
percent intact membranes
(avg ± SD)a
Plate count viability,
CFU enumeration (%)b
99.9 (99.9 ± 0.1)
0.5-log10 Increase (n/a)c
99.5 (99.0 ± 0.6)
< 0.5-log10 Decrease (50.0)
pH 3.6 Buffer
95.4 (92.3 ± 5.4)
< 0.5-log10 Decrease (50.0)
96.4 (93.0 ± 6.6)
1.5-log10 Decrease (5.0)
99.3 (99.9 ± 0.3)
2.5-log10 Decrease (0.5)
Summary of MRSA survival and membrane permeability following incubation in TSB, water, pH 3.6 phosphate buffer, and mineral leachates
percent intact membranes
(avg ± SD)a
(% viability reduction)b
96.6 (95.8 ± 2.6)
2-log10 Increase (n/a)c
61.9 (78.0 ± 16.0)
1-log10 Decrease (90.0)
pH 3.6 Buffer
6.1 (2.6 ± 3.0)
5-log10 Decrease (99.999)
3.2 (3.3 ± 2.4)
4-log10 Decrease (99.99)
1.7 (1.7 ± 1.0)
4.5-log10 Decrease (99.995)
The abundance of antibiotic resistant pathogens has incrementally increased since the introduction of penicillin in the 1940s. In the United States alone, more than 70% of hospital-acquired infections are antibiotic-resistant, and community-acquired exposure to antibiotic-resistant pathogens is becoming increasingly prevalent [25, 26]. This problem is further exacerbated by waning research and development of novel antibacterial agents. Historically, new antibiotics were developed by recapitulating a small set of molecular scaffolds, thus allowing opportunities for further antibiotic resistance to develop [27, 28]. These alarming trends highlight the urgent need to develop novel and alternative antibacterial agents.
In the past 25 years, ~70% of commercially available antibiotics have been derived from natural sources [28, 29]. However, many unexplored natural resources remain promising for the discovery of new antibacterial agents. Clay minerals have been used historically for cosmetic purposes and to treat ailments of the digestive tract, but more recently have been investigated for their potential antibacterial properties [2, 30]. The research presented here documents an understanding into the mechanism of action of mineral leachates, CB07-L and BY07-L, and scientifically validates the antibacterial efficacy of these clay mineral mixtures as promising new antibacterial agents. Further, an understanding of the antibacterial mechanism of action of these natural products will allow development of a chemically-derived, synthetic alternative, thus guaranteeing consistent efficacy.
Previously, we showed that the BY07 and CB07 mineral mixtures are composed of 37.3% and 21.4% smectite, respectively . Smectite clays have a layered structure with an expandable interlayer and a high cation exchange capacity attributed to their overall net negative charge. In a hydrated suspension, these cations can be exchanged with ions in the external solution, provided charge balance is maintained . In a low pH environment, the abundant protons saturate metal binding sites in the solution, maximizing the concentration of soluble metal ions. Consequently, metal ions become more bioavailable, and possibly more toxic, as the pH of a solution decreases . Some metal ions, such as iron, copper, nickel, magnesium, manganese, and zinc, have specific biological functions as enzyme cofactors or to stabilize proteins and bacterial cell walls [15, 32]. Alternatively, ions, such as aluminum, arsenic, lead, and mercury, have no biological function. Such metals can exert toxic effects by irreversibly binding sulfhydryl groups in proteins or enzyme metal binding sites [19, 33]. Regardless of their function, all metals can exert toxic effects at high concentrations due to non-specific binding . As a consequence, microorganisms have adapted several mechanisms, such as active transport, sequestration, and enzymatic detoxification to exclude heavy metals and regulate intracellular concentrations of essential metals [34–36]. Moreover, when metal ions are present in combination, toxicity can be magnified due to synergistic effects. For example, when lead and mercury are present together, the toxic effects are amplified 100-fold . It is likely that the toxic effects of these mineral mixtures are due to the synergistic effects mediated by a combination of ions present in the solution . However, further research is needed to determine which elements are mediating toxicity and their specific molecular targets.
Many groups have used EM to visualize cellular ultrastructure following exposure to silver ions [22, 38, 39], however, minimal literature exists on the use of EM to assess antibacterial activity and ultrastructural influences of other metal ions. Following exposure to silver ions, E. coli and MRSA cells exhibit condensed DNA and electron-dense granules bound to the cell envelope [22, 38], in a similar manner as the CB07 and BY07 leachates (Figures 2 and 4). To our knowledge, we report the first EM ultrastructural analysis of E. coli and MRSA following exposure to antibacterial mineral leachates, demonstrating that neither E. coli nor MRSA cells lysed following exposure to the mineral leachates. Furthermore, changes in cell shape observed in MRSA cells following exposure to the leachates were not evident in E. coli. Condensation of the cytoplasm was observed in E. coli, and small (~ 10 nm), electron-dense deposits were visualized on the surface of the cells.
CB07-L and BY07-L exhibit bactericidal activity against E. coli and MRSA as determined by CFU enumeration. Overall, a phenomenon occurs whereby E. coli is rapidly killed by CB07-L and BY07-L without permeabilization to the membrane, while MRSA is killed at a slower rate with membrane permeabilization. Tables 1 and 2 summarize the membrane permeability data determined by flow cytometry and cell viability determined by CFU enumeration. Notably, following a 6 h exposure to BY07-L, E. coli viability decreased by 2.5 log10 units (99.5% decrease), while LIVE/DEAD staining determined that only 0.7% of the cell membranes were permeabilized. These data demonstrate the careful consideration needed when using LIVE/DEAD staining as a direct indicator of cellular viability.
It is known that bacterial cell walls interact strongly with metal cations, maintaining control over the type and amount of ions that gain access to the cytoplasm . Gram-positive cell walls specifically have been shown to have a higher charge capacity, allowing containment of a larger volume of cations . For example, it has been demonstrated that Bacillus subtilis binds 28 to 33 times more Cu2+ than E. coli. As a potent metal ion chelator, the thick peptidoglycan layer present in S. aureus likely contributes to the delayed toxicity seen with MRSA exposure to the leachates. Moreover, while these ions are trapped in the peptidoglycan, they can propagate oxidative damage to the membrane . Because E. coli is killed without loss of membrane integrity, it is possible that membrane permeabilization observed in MRSA cells is not the principle mechanism of action, but rather a secondary consequence due to the slow passage through the thick peptidoglycan.
MRSA exposure to the leachates resulted in a distinct and reproducible fluorescence staining pattern as observed with flow cytometry. Interestingly, the dot plots of leachate-exposed MRSA cells show two red-fluorescent populations. The first population moved in a distinct curve-shaped manner, transitioning from higher red fluorescence intensity to lower red fluorescence intensity (Figures 7h and 7j; black arrow). This feature is likely due to intermediate states, characterized by different concentrations of SYTO9 and PI dyes within the cells . The second population, although still red, occurred at much lower fluorescence intensity (Figures 7h and 7j; white arrow). This phenomenon could possibly be due to an overall lower abundance of nucleic acids in these cells. Alternatively, due to FRET, PI fluoresces with greater intensity when in the presence of SYTO9 and at a lower intensity when present alone . Therefore, the separation of these two populations may be due to a greater abundance of SYTO9 in the upper population and a decreased abundance of SYTO9 in the lower population.
Natural sources have historically played an important role in the discovery of novel antibacterial agents . CB07 and BY07 mineral mixtures and their leachate derivatives could offer an additional complementary treatment option against topical bacterial infections. However, efficacy of these minerals can vary widely despite having a common source. It is therefore essential to characterize their specific antibacterial mechanism of action in order to improve quality control, guarantee consistent efficacy, and maximize their performance as an antibacterial agent.
In summary, these data suggest that the mineral leachate antibacterial killing activity differs for Gram-positive and Gram-negative organisms and have guided us in our understanding of the leachate antibacterial mechanism of action. Upon antibacterial mineral leachate exposure, structural integrity is retained, however, compromised membrane integrity accounts for bactericidal activity in Gram-positive, but not in Gram-negative cells.
We thank D. Lowry and R. Roberson in the ASU School of Life Sciences for their assistance with the electron microscopy. Additionally, we gratefully acknowledge the use of facilities within the LeRoy Eyring Center for Solid State Science at Arizona State University. This research was supported by Public Health Service grants AT004690 and AT003618 awarded to S.E.H. from the National Center for Complementary and Alternative Medicine at the National Institutes of Health.
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