In situ imaging of Sulfobacillus thermosulfidooxidans on pyrite under conditions of variable pH using tapping mode atomic force microscopy

Process Biochemistry 46 (2011) 966–976

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In situ imaging of Sulfobacillus thermosulfidooxidans on pyrite under conditions of variable pH using tapping mode atomic force microscopy T. Becker a,∗ , N. Gorham a , D.W. Shiers b , H.R. Watling b a

Parker Cooperative Research Centre for Integrated Hydrometallurgy Solutions, Nanochemistry Research Institute, Curtin University, Bentley, WA 6845, Australia Parker Cooperative Research Centre for Integrated Hydrometallurgy Solutions, CSIRO Minerals Down Under Flagship, CSIRO Process Science and Engineering, Karawara, WA 6152, Australia b

a r t i c l e

i n f o

Article history: Received 3 October 2010 Received in revised form 21 December 2010 Accepted 10 January 2011 Keywords: Atomic force microscopy (AFM) Bacterial attachment Sulfide Bioleaching Sulfobacillus

a b s t r a c t Sulfobacillus thermosulfidooxidans cells attached to pyrite surfaces were imaged using in situ tapping mode atomic force microscopy (AFM). The attached bacteria were dispersed in small groups without preferred orientation or site specific preference. The importance of extracellular polymeric substances (EPS) production by cells upon changing pH was examined. Bacteria detached or were dislodged and no EPS was detected for a rapid decrease from pH 2.2 to pH 1.0 in the solution. In contrast, a layer of EPS with increasing thickness was observed for a decrease from pH 2.2 to pH 1.4. Upon further acidification (pH 0.9) the cells were not dislodged but the thickness of the EPS layer decreased immediately, which is attributed to structural re-arrangements of the polymeric chains within the EPS network. This suggests that the bacteria respond to stress caused by increased acidity by expressing EPS, which acts as a buffer against unfavourable conditions in the environment. Secondary mineral formation (‘scale’) on the pyrite surface was found to form only in the presence of microorganisms and eventually covered the pyrite surface. Bacterial cells did not attach to the scale surface. Upon scale removal, etch pits with similar shape to bacterial cells were observed in the underlying pyrite surface. © 2011 Elsevier Ltd. All rights reserved.

1. Introduction Iron- and sulfur-oxidising acidophilic bacteria and archaea naturally colonise sulfide-rich ores, as evidenced by numerous examples of acidic, iron-rich drainage water containing diverse microbial populations [1]. The natural processes of iron- and sulfur oxidation and growth on organic compounds are exploited in commercial-scale heap and dump ‘bioreactors’ for the extraction of metals from low-grade sulfide ores [2]. Heaps are constructed of relatively large ore particles (tens of millimetres in diameter) containing a diverse mineral assemblage which exhibits various physical and chemical properties during leaching. In some ores, the sulfide minerals, such as secondary copper sulfides, are concentrated on particle surfaces and readily available for microbial colonisation. In others, the sulfides occur as discrete grains encapsulated within gangue minerals with limited exposure on particle surfaces. Another peculiarity of heaps is that they are dynamic environments in which (i) the sulfide content diminishes with time, (ii) acid is consumed through gangue mineral dissolution but periodi-

∗ Corresponding author at: Curtin University, Nanochemistry Research Institute, Dept. Chemistry, GPO Box U 1987, Perth, WA 6845, Australia. Tel.: +61 8 9266 7806; fax: +61 8 9266 4699. E-mail address: [email protected] (T. Becker). 1359-5113/$ – see front matter © 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.procbio.2011.01.014

cally augmented to maintain the leaching environment, giving rise to variations in solution pH, and (iii) secondary minerals may be precipitated on particle surfaces. For the most part, the sampling of ore heaps, particularly at depth, during mining operations is viewed unfavourably because compaction of heap surfaces and inadvertent disturbance of irrigation lines or other heap equipment impacts on metals solubilisation and/or heap management and thus affects productivity. Therefore an indirect approach has been adopted in our laboratories through which to describe and, where possible, quantify microbial processes in the various micro-environments that exist in heaps. The broad research program covers studies on microbial responses to changed conditions of temperature, pH, ionic strength, harmful cations and anions and organic compounds [3]. Wherever possible, studies are conducted using simulated heap leach conditions and monitored using in situ probes. It is anticipated that the knowledge gained will assist in optimising heap leaching conditions for better metals extraction as well as inform operators of the consequences of excursions outside those optimum operating conditions. The particular focus of this paper is bacterial attachment to sulfide surfaces and the impact of changed pH as well as the effects of secondary scale formation. It exploits the unique capabilities of tapping mode atomic force microscopy (AFM) applied in situ to visualise bacterial responses.

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Since its invention in 1986 [4] AFM has been employed in a vast variety of experiments in many fields of science, including the imaging of bacteria. The ability to operate the microscope in situ in most fluids and over a wide temperature range makes AFM a valuable research tool. Tapping mode AFM in liquids, also referred to as intermittent-contact or dynamic-force mode AFM, uses a cantilever (typical spring constant 0.01–0.5 N/m) that oscillates close to the sample, intermittently touching the surface. It has the advantages that lateral forces such as drag are greatly reduced and lateral resolution on soft surfaces is improved. As a consequence, bacterial cells are less likely to be damaged by the AFM tip. Two examples from many published data illustrate the power of high-resolution tapping mode AFM: Camesano [5] using ex situ tapping mode AFM to compare bacterial cell morphologies (Pseudomonas putida and Burkholderia cepacia), showed that cell surface roughness was increased after exposure to selected surface-active chemicals and that some of the chemicals caused flattening of cells; Camesano and Logan [6] used a combination of in situ tapping mode AFM and force measurements to show that the equilibrium length of bacterial surface polymers increased significantly with increased pH for both their test species (B. cepacia G4 and P. putida KT2442). Regarding AFM applications on microorganisms commonly found in bioleaching systems, our study of bacterial attachment on sulfides is based on research by Sand and colleagues at the ‘Biofilm Centre’ [7–14] complemented by studies at several other institutions [15–20]. Studies on six of the known bioleaching bacteria have been undertaken [7,8] using either contact mode [12,19] or tapping mode AFM [19], in air (ex situ) [7,19] or in solution (in situ) [12,17], sometimes in combination with a second technique [11,15]. Collectively, the results indicate that: • Cell morphology differs between planktonic cells and sessile cells; • Substrate and crystal orientation may influence attachment and cell morphology, and EPS mediated attachment of cells; • Bacterial coverage of the surface is often of low density, but there is evidence of communal activity in construction of biofilms around even two or three cells; • Stressful conditions, such as dehydration, induce greater EPS production but the amount of EPS produced is also influenced by the nature of the surface on which the cells have been dried; hydrophilic (more EPS) or hydrophobic (less EPS). Some problems have been encountered during in situ imaging of bacteria, such as tip-induced movement and/or detachment from the substrate and in situ tapping mode AFM generates images of bacteria that lack surface detail, compared with ex situ, contact mode images [12]. Nevertheless this technique was chosen to study the responses of the bacterium Sulfobacillus thermosulfidooxidans to controlled but varied conditions relevant to heap bioleaching, allowing the real time observation of the behaviour of the bacteria in an environment as close as possible to the conditions in a heap. The selected surface for attachment was pyrite. In this study, AFM measurements were carried out in situ using a pyrite coupon previously exposed to a bacterial suspension to facilitate cell attachment immersed in a flow cell through which nutrient medium was pumped very slowly to avoid disturbing the imaging process. The coupon surface was never exposed to air during transfer from the bacterial suspension to the flow cell during the experiment. Further, with careful choice of operating conditions, it was possible to image bacteria in situ using tapping mode AFM over periods of up to several days without causing bacteria to detach. Following the verification of stable imaging conditions, the responses of attached bacteria subjected to increased acidity were examined.

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2. Materials and methods 2.1. Bacterial culture Basal salts medium (BSM) was prepared as follows: (NH4 )2 SO4 (1.5 g L−1 ), KH2 PO4 (0.25 g L−1 ) and MgSO4 (0.25 g L−1 ) were dissolved in deionised water (1 L), the pH adjusted to 1.8 by dropwise-addition of concentrated H2 SO4 and the solution sterilized at 121 ◦ C, 100 kPa for 20 min. Where required, FeSO4 ·7H2 O (10 g) was dissolved in 25 mL BSM. This solution was filter sterilized (Pall P/N60301 membrane filter, pore size 0.2 ␮m) back into the BSM, followed by the addition of 1 mL of 10% sterile yeast extract solution (BSM-YE). The final concentration of iron(II) was 2.01 g L−1 (0.036 M). The moderately thermophilic mixotroph S. thermosulfidooxidans DSM9293T was grown at 45 ◦ C in BSM-YE medium containing FeSO4 ·7H2 O (10 g L−1 ) and was subcultured into fresh medium, with or without added FeSO4 ·7H2 O, 24–48 h prior to each experiment and then filtered through a 0.2 ␮m pore size membrane. Cells were resuspended into 30 mL BSM-YE to yield a cell concentration of approximately 2.0 × 108 cells mL−1 . 2.2. Ferrous ion oxidation tests Sterile solutions of BSM-YE supplemented with FeSO4 ·7H2 O (10 g L−1 ) were prepared in the pH range 1.0–3.0, in increments of 0.2 pH units. The solutions were sterilized by passage through a 0.2 ␮m pore-size filter. Conical flasks containing 100 mL of the pH-adjusted media were inoculated with 10 mL of the selected bacterial culture to yield an initial cell density of ∼1 × 107 cells mL−1 . Control flasks were not inoculated and all tests were incubated at 45 ◦ C. Ferrous ion concentrations were determined periodically after 16 h in biotic and abiotic tests using a spectrophotometric method based on Wilson [21]. Spectrophotometry was chosen because the total iron concentration in some solutions changed during the experiment due to the precipitation of iron(III) species at pH > 2. The difference between abiotic and biotic tests measured at the same time was attributed to bacterial activity. Rate data were plotted against the final solution pH (16 h). 2.3. Preparation of pyrite coupons The pyrite coupons were prepared from pyrite (1 cm3 ) embedded in an epoxy resin and cut into slices of approximately 3 mm thickness. These slices were ground manually and polished (three grinding steps: #600, #1000, and #1200, two polishing steps: 3 ␮m diamond suspension and 1 ␮m diamond suspension). The sample disc for the in situ experiments was cut to size and two holes were drilled into the slice to fit it into the flow cell holder. All samples were cleaned in ethanol in an ultrasonic bath and with an UV/ozone cleaner prior to exposure to a bacterial suspension of S. thermosulfidooxidans cells. 2.4. In situ atomic force microscopy A Molecular Imaging PicoPlus AFM system (Agilent, Texas) with a fluid flowthrough cell mounted on a heating stage was used to investigate the behaviour of bacteria attached to pyrite surfaces. All pyrite samples were freshly polished and cleaned prior to immersion in the S. thermosulfidooxidans culture for 24 h to facilitate bacterial attachment on the surface. The coupon was removed from the bacterial suspension and mounted on an AFM sample stage with integrated heating element (T = 45 ◦ C) together with the flow cell. During this process the sample surface was constantly covered with a film of liquid. The bacterial suspension was recirculated between the reservoir and the flow cell using a peristaltic pump. Both, reservoir and AFM sample stage were kept at a constant temperature of 45 ◦ C. A small pump-rate was used to avoid large temperature gradients in the flow cell and to keep the pulsing caused by the pump to a minimum. The system was then left for at least 1 h before scanning the sample to allow thermal equilibration. The in situ tapping mode images were acquired using very soft silicon nitride cantilevers with a spring constant of 0.06 N m−1 (type DNP, Veeco, California). The amplitude set point was kept as high as possible (typical engage setpoint: 92–95% of free cantilever oscillation) to avoid tip-influenced distortion or detachment of the bacteria during scanning. With the appropriate imaging parameters and the system fully equilibrated it was possible to scan the surface of the sample over a period of at least 24 h without detaching bacteria.

3. Results and discussion The purpose of imaging S. thermosulfidooxidans cells in situ was to obtain a real-time response to deliberately modified conditions. It was anticipated that observable responses would include one or more of: attachment, detachment, cell replication, changed cell morphology, EPS production or spore formation when stressful growth conditions were introduced.

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and cell division. Further, bacterial cells might not attach within the area of the scan if they were disturbed during their initial settling phase. Fig. 3 shows a typical phase image for bacteria attached to the pyrite substrate. The brighter features correspond to bacteria, whereas the less-bright, disc-shaped features are secondary mineral phases crystallising on the pyrite surface. Whereas Gehrke et al. [9] report attachment of the cells specifically to dislocation sites on pyrite such as cracks and grain boundaries, our experiments show relatively evenly distributed bacteria over the scanned area and no preference for pyrite crystal orientation or specific surface features such as scratches or voids. Our observations are consistent with the results of Pace et al. [23]. 3.3. Rapid increase in acidity

Fig. 1. Effect of pH on bacterial growth. The growth of bacteria such as S. thermosulfidooxidans can be related to the formation of ferric ion from the oxidation of ferrous iron. The graph shows the effect of the pH of the growth medium on the growth of the bacteria and highlights optimum pH of 1.6 (initial cell density ∼1 × 107 cells mL−1 ; replicate experiments indicated by , ).

3.1. pH and growth (ferrous ion oxidation) The rate of growth of bioleaching bacteria approximates the rate of product formation; thus, for iron(II)-oxidising bacteria, such as S. thermosulfidooxidans, growth can be related to the formation of ferric ion from the oxidation of ferrous ion [22]. Using this approximation, the effects on ferrous ion biooxidation of changing the growth-medium pH were tested (Fig. 1). The data (two experiments under similar conditions indicated by different symbols) highlight not only the relatively narrow range of pH conditions for S. thermosulfidooxidans growth but also the significant reduction in ferric ion generation rates as conditions deviate from the estimated optimum pH value of pH 1.6. These data served to guide the choice of conditions for subsequent experiments on the effect of increased acidity, such as may be experienced when acid contents in heap irrigation solutions are replenished. 3.2. Cell attachment to pyrite The method of establishing an initial population of bacterial cells attached to a pyrite coupon was to immerse the coupon in a vertical position in a bacterial suspension for 24 h. Subsequent in situ images obtained using the flow-cell with BSM-YE over 64 h showed that attachment was successful and that bacterial cells could be scanned repeatedly in tapping mode without being dislodged from the pyrite surface. In topographical images (Fig. 2a–c), the brighter colour indicates a higher feature. Phase images (Fig. 2d–f), collected simultaneously with the topographical data allow features with different physical properties such as visco-elastic response, adhesion or friction to be distinguished. In these images, brighter colour indicates a softer surface such as a bacterial cell, compared with a mineral crystal. These data are discussed further in Section 3.4. Attempts to observe in situ the attachment of new bacteria to the substrate or the cell division of attached bacteria were unsuccessful. A possible explanation is that the process of attachment of the bacteria was disturbed by the scanning AFM probe. A typical scan took approximately 5.7 min; thus the probe would return to the same site on the sample every 5.7 min. Depending on the image size, the probe would scan over a single bacterium for up to 1 min. Tapping could sufficiently disturb bacterial cells to inhibit growth

As shown in Fig. 1, bacteria are sensitive to changes in pH. Their activity, in this case ferrous ion oxidation to form ferric ion, diminishes as the solution pH deviates from the optimum pH for bacterial growth. Solution pH within a heap varies significantly, spatially and with depth, as (i) the acid is consumed through gangue mineral dissolution or (ii) the acid is replenished and applied through the irrigation system to maintain the desired leaching environment. In the following experiment, the response of bacteria attached to a pyrite coupon subsequent to a rapid increase in acidity was investigated. The pyrite sample was prepared as described above. The surface was imaged in situ at 45 ◦ C with tapping mode AFM using a scan rate of 1 Hz (approximately 8.5 min per image). Fig. 4 shows successive topography images (a–d) and corresponding phase images (e–h) of the pyrite sample with attached bacteria (e.g. white outlined area). After sufficient time to achieve stable imaging, the pH in the solution was decreased from pH 2.2 to pH 1.0 by the addition of sulfuric acid into the reservoir towards the end of a scan (Fig. 4a, indicated by the white arrow). This drop in pH represented a change from conditions conducive to bacterial iron(II) oxidation to conditions where iron(II) oxidation would be minimal (Fig. 1). Due to the very slow setting of the peristaltic pump, which recirculates the fluid between the AFM fluid cell and the reservoir, it takes several minutes until the condition of pH 1.0 is established in the fluid cell. Comparing the two images taken subsequent to acidification (Fig. 4b and c), Fig. 4c showed the rapid loss of many bacteria. At the same time, features of the underlying pyrite surface became more distinct. The image conveyed the impression that the surface had been “wiped clean”. Interestingly, the corresponding phase image (Fig. 4g) still showed evidence of the bacteria or possibly of remnant EPS which was still evident in the image acquired 25 min after acidification (Fig. 4h). The phase images showed a residual footprint of organic matter, even though the corresponding topographic images at identical locations appeared to represent clean, smooth pyrite surfaces. From the experimental data it was not possible to determine whether dead or viable bacteria were dislodged from the surface. For a more detailed investigation of the response of the bacteria to decreased pH, the experiment was repeated with a staged change in acidity. 3.4. Staged change in acidity The images presented in Fig. 2 show topography (a–c) and phase images (d–f) at three different times. The clean pyrite coupon was immersed in an iron-free bacterial suspension (pH 2.2) for 24 h to facilitate bacterial attachment and then transferred to freshly prepared iron-free medium (BSM-YE, pH 1.4) in the AFM flow cell. The sample was imaged continuously for the next 39 h. Comparison of a topography image acquired 0.5 h after the pH was changed from pH

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Fig. 2. Time series of AFM topography (a–c) and corresponding phase images (d–f) for a staged change in acidity (pH 2.2–pH 1.4, later pH 1.4–pH 0.9 (not shown)). The in situ tapping-mode images show S. thermosulfidooxidans on a pyrite coupon in the flow-cell with BSM-YE at three different times after acidification to change from pH 2.2 to pH 1.4 (0.42 h, 9.42 h and 24.42 h). Each imaged area was scanned over a period of approximately 4 min (topography images: height range: 280 nm; phase images: phase angle range: 70◦ ).

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more acidic environment. With a sufficient amount of EPS, the bacteria remain on the surface even when the pH of the solution is decreased further to pH 0.9. Conversely, a rapid decrease in pH of the leaching solution may hinder production of sufficient amounts of EPS with the consequence that bacteria are exposed directly to a too high concentration of acid. Cells may die, lyse or be dislodged, leaving behind a small amount of organic matter which is clearly visible in the phase signal. We assume that this is remaining EPS, which may dehydrate and shrink with time and become partially dislodged and removed by the scanning AFM probe. However, even at the end of the experiments, the topography images of the sample did not show the same level of detail as was observed in the initial scans. This suggests that the EPS layer is not completely removed from the surface. 3.5. pH and EPS production

Fig. 3. Bacterial attachment on pyrite surface. The AFM phase image reveals different interactions of the probe with the sample and allow here the identification of the bacteria (bright features). The bacteria do not show an orientation preference with respect to pyrite surface features and exhibit a disperse attachment to the pyrite surface (phase angle range: 70◦ ).

2.2 to pH 1.4, with an image at 31.8 h showed a significant difference in the appearance of the sample surface. The bacteria were still on the surface, but all surface features appeared blurred in the topography images (Fig. 2a–c). Further, the phase image at 25 min (Fig. 2d) showed a strong contrast, revealing many surface details such as the bacteria and polishing marks on the pyrite surface, whereas there was less contrast in the phase image at 31.8 h (Fig. 2f). The reduced phase contrast later in the experiment suggested that the surface had become covered with a nearly homogeneous layer. This assumption is supported by a comparison of cross sections through the topography images at different times during the experiment. Fig. 5 shows two cross sections from the same location on the sample surface at 0.5 h and 31.8 h, respectively. Polishing marks and grooves were responsible for the background structure of the sample profile and were clearly present in both sections, but only the section at 0.5 h showed the fine detail of the surface. The cross section at 31.8 h was significantly smoother. Moreover, detailed comparison of several cross sections from corresponding locations at 0.5 h and 31.8 h showed that features protruding from the surface broadened and that the widths of grooves and scratches decreased with time. These observations are attributed to the formation of a near-coherent EPS layer over the pyrite surface, consistent with the observations of Teschke et al. [19] of EPS materials encapsulating groups of bacteria (termed an EPS ‘corral’ by those authors). After 39 h at pH 1.4 the solution in the AFM flow cell was acidified to pH 0.9 and further images collected. Remarkably, no further significant differences in the appearance of the surface compared with the measurements at 39 h were observed. Until the end of the experiment at 63 h, the topography images appeared slightly blurred and the phase signal showed only small variations. Clearly the behaviour observed for a staged decrease in pH differed significantly from that observed with a rapid change in acidity, described in the previous section. The increasing thickness of the EPS layer observed for the staged change (Fig. 2) was not seen in the case of the rapid pH change (Fig. 4). It is assumed that this increased layer of EPS acts as a protective barrier against the

The observed change in the appearance of the sample surface during in situ AFM imaging (Fig. 2) indicated a significant influence of the solution pH on the production of EPS. Similar results were reported recently, where a pyrite surface with attached bacteria (S. thermosulfidooxidans) exposed to higher acidity became “sticky” and difficult to scan with ex situ AFM, due to the strong adhesion of the tip to the sample surface Gorham et al. [24]. In the present study it was possible to determine the change in thickness or the ‘growth rate’ of the EPS layer from the in situ AFM topography images collected as a function of time. Due to the formation of this EPS layer over the whole sample area, it was not possible to take advantage of the high resolution capability of the AFM along the z-axis to measure the thickness of the EPS layer. However, by measuring the width of protruding surface features at different times, the change in thickness can be determined and the ‘growth rate’ of the EPS layer estimated. The change in thickness EPS of the EPS layer is represented by: EPS =

t − t0 2

where t0 is the width of a feature at the beginning of the experiment and t is the width of the same feature at time t. In Fig. 6 the broadening effect due to the coverage with EPS is shown with topographical cross section at two different sample locations at three different times during one experiment. The pH of the solution for all of the presented cross sections in Fig. 6 was pH 1.4. To extract the increase of the amount of EPS covering the respective feature, the full width at half maximum was determined. In case of the profile shown in Fig. 6a and b, the EPS film increases by 48 nm (35 nm) between 0.42 h and 9.42 h after increasing the acidity of the solution to pH 1.4, and increases by a further 57 nm (75 nm) between 9.42 h and 24.42 h after acidification. Fig. 7 shows the changes in the EPS layer versus time for two separate experiments by considering the cross sections of the same surface feature taken at different times. The open diamonds represent the changes for the experiment described above (Fig. 2), and the solid squares show the results for a control sample subjected to solution of pH 1.4. The values shown in Fig. 7 are measured with reference to the width of protruding features at time t = 0, immediately following the first decrease from pH 2.2 to pH 1.4. The thickness of the EPS layer on the pyrite surface increased in an approximately linear fashion at a rate of 15.6 nm h−1 , and achieved a maximum change in thickness of 215 nm in the experiment. Whereas, during continuous imaging, the appearance of the surface did not appear to change significantly upon further acidification to pH 0.9 at t = 39 h, the thickness of the EPS layer decreased by approximately 100 nm as an immediate response to the increased acidity. After the initial decrease in EPS thickness due to increased acidity, the EPS layer remained relatively constant.

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Fig. 4. Time series of AFM topography (a–d) and corresponding phase images (e–h) for a rapid change in acidity (pH 2.2–pH 1.0). The white arrow in (a) indicates the time when the acid was added to the system. The majority of attached bacteria (as for example within the white outlined area) disappeared over the course of the next 2 images, whereas the phase images still show a contrast at the attachment site, indicating evidence of bacteria or more likely a residual EPS footprint of the previously attached bacteria (topography images: height range: 300 nm; phase angle range: 6◦ ).

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Fig. 5. Coverage of pyrite surface. Cross sections of the sample topography at 0.5 h and 31.8 h after acidification at identical locations within the scan area. The smoother profile indicates the presence of a layer of EPS covering the surface.

Similar values for the thickness of the EPS layer determined with ex situ contact mode AFM have been reported previously [17,18]. Teschke et al. [18] found evidence of a continuous layer of EPS covering the substrate with a thickness varying between 20 nm and 600 nm; the average thickness of the EPS film covering individual bacteria was reported to be 250 ± 150 nm. Our in situ measurements are in good agreement with these results. Composition analysis has shown that EPS form a highly hydrated gel-like structure consisting of sugars and hydrogen-bonded water [19,25]. SAXS investigations of the EPS of a marine bacterial isolate revealed major rearrangements of the EPS structure in response to variation of the pH [26]. EPS can be described as networks of polymers with regions of unequal densities, for example due to the presence of coiled structures [26,27]. Dogsa et al. [26] reported a shrinking EPS network with decreasing pH–pH 0.7 with an increase of denser areas of polymeric chains. A similar process may explain the decrease of EPS film thickness upon acidification observed in this study. In a separate experiment the thickness of the EPS coating was monitored without further acidification of the environment. These data are shown as solid squares in Fig. 7. The growth rate was comparable to the experiment described above. After about 40 h the EPS thickness reached a maximum and remained relatively constant. However, after a further 20 h, the thickness of the EPS began to decrease. This decrease might possibly be related to a small but not insignificant amount of solution evaporated from the AFM flowcell which could cause structural rearrangement within the EPS network and result in a measurable decrease in EPS film thickness. Within the scope of this study it was not possible to determine the exact cause for the shrinkage of the EPS layer in this case, but this will be considered in future investigations.

Fig. 6. Broadening of protruding surface features. Topography cross sections at identical locations for different times on two different surface features show the broadening of the features due to coverage of the sample surface with EPS (EPS thickness increase: (a) 0.42 h → 9.42 h: 48 nm, 9.42 h → 24.42 h: 57 nm, (b) 0.42 h → 9.42 h: 35 nm, 9.42 h → 24.42 h: 70 nm).

media (e.g. BSM). Some previous studies regarding precipitates formed in the presence of micro-organisms, including At. ferrooxidans, showed these precipitates to be potassium and ammonium jarosites [28–31]. Jarosite formation on the pyrite surface poses two questions that relate to the efficiency of bacterial attachment and subsequent catalytic action in enhancing sulfide dissolution: Do the bacteria compete with the jarosite for surface area on the pyrite and do the bacteria also attach to the jarosite surface when the sulfide has become covered?

3.6. pH and secondary mineral formation In addition to the attachment of bacterial cells on the pyrite surface, formation of deposits on the surface was observed on most of the samples. The deposits formed by a process of nucleation and subsequent crystal growth and the crystals with disc-like morphology eventually covered the pyrite surface. The precipitation of similar disc like structures on pyrite surfaces in the presence of different micro-organisms has been reported [28]; the precipitates appeared in stages, starting with sub-micron sized residues later developing into disc-like structures with sizes of 1–2 ␮m. In our experiments, disc-shaped features of similar size to those of Mikkelsen et al. [28] were imaged using AFM and SEM (Fig. 8). Raman spectroscopy (Fig. 9) and energy dispersive X-ray spectroscopy (EDS) of the discs showed them to be a mixture of potassium and hydronium jarosites. The formation of these types of jarosites is plausible taking into account that potassium and ammonium salts are components of bacterial nutrient

Fig. 7. Thickness of EPS layer. Change of thickness of EPS layer for two separate experiments: control sample at pH 1.4 (); sample with pH 1.4 for t = 0 → 39 h, then addition of acid to change to pH 0.9 for t = 39 → 64 h (♦). The thickness of the EPS layer was estimated by comparing the width of protruding surface features at different times with the width of the same features at the beginning of the experiment.

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Fig. 8. Secondary mineral formation. (a) AFM and (b) SEM view of the layer formed by secondary minerals on the pyrite surface in the presence of microbes in the iron(III)-rich suspension. (AFM topography image: height range: 1.3 ␮m).

For the investigation of the effect of jarosite formation, pyrite coupons were exposed to different solutions. In one experiment a freshly polished and cleaned sample was mounted in the AFM flow cell and exposed to ferric sulfate solution (concentration 1 g L−1 , pH 1.8, T = 24 ◦ C, abiotic). The sample was imaged continuously for ∼45 h. During the experiment, some surface dissolution was apparent, but the formation of jarosite discs as seen in biooxidation experiments was not observed. The absence of jarosite was

Fig. 9. Raman spectrum of secondary minerals. Comparison of the Raman spectrum taken of the pyrite coupon covered with a layer of secondary minerals after 94 h of exposure to iron(III)-rich bacterial suspension with a jarosite spectrum (taken from RRUF Project Raman spectroscopy database).

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Fig. 10. Secondary minerals. AFM amplitude signal of surface with scale (a) and topography cross section of disc-like scale (b and c) to determine the step height of grown crystals as an attempt to determine one parameter of the unit cell.

confirmed by Raman spectroscopy, where spectra acquired from the sample surface were identical before and after exposure to the ferric sulfate solution. In a second experiment, a freshly polished and cleaned pyrite sample was exposed for 24 h to a bacterial suspension prepared by harvesting cells from their iron(II)/iron(III) growth medium and resuspending them in iron-free BSM-YE (pH 2.2). Here, the formation of scale on the surface was not observed, in particular no disc shaped features were found. The absence of scale in this case is explained by the fact that iron in solution could only originate from the pyrite sample as a consequence of oxidation. Consequently, the concentration of dissolved iron in the solution, which is required to facilitate the scale growth, is significantly smaller than for a bacterial suspension prepared with iron. At this stage we do not exclude the possible growth of scale, but within the timeframe and on the scale of the AFM images of this study no significant amounts of scale were observed. These results are also confirmed by a comparison of abiotic and biotic pyrite leaching experiments conducted by Mikkelsen et al. [28]. In their abiotic test some secondary products formed on pyrite particles after 35 days, but there were no significant changes to the pyrite particles. In contrast, significant amounts of secondary products in the shapes of discs formed in their biotic test after just 10 days. In a third experiment, a freshly polished and cleaned pyrite sample was exposed to a bacterial suspension prepared with iron (initially 10 g L−1 FeSO4 ). In this case, the bacteria had oxidised the iron(II) to iron(III) and were at the end of the exponential growth

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Fig. 11. Thickness of secondary mineral layer. (a) AFM topography image of the sample covered sample covered with secondary minerals after 94 h exposure to iron(III)-rich bacterial suspension. The surface has been scratched with a needle to reveal the underlying pyrite surface. The dotted line indicates the location where the cross section shown in (b) has been taken to measure the thickness of the secondary minerals layer (AFM topography image: height scale: 2.8 ␮m).

phase prior to their exposure to the pyrite coupon. In contrast to the second experiment in which the pyrite sample was exposed to an iron-free bacterial suspension, the pyrite coupon was now immersed in the iron(III)-rich bacterial suspension for 24 h before being mounted in the AFM flow cell. Initial AFM images showed that the surface was already covered with a significant amount of jarosite (Fig. 8). The jarosite discs appeared to be randomly oriented presenting a continuous and uniform coverage of the pyrite surface. In previous experiments, bacteria were detected interspersed among the discs on surfaces of similar appearance. However, in this experiment (Fig. 8) no bacteria were found in between the discs or on the surface of the jarosite. The total exposure time of this pyrite coupon to the bacterial suspension was 94 h. After this time, the sample was rinsed with MilliQ water, dried with nitrogen gas and prepared for Raman spectroscopy and SEM/EDS analysis. Both techniques confirmed that the material covering the pyrite surface was comprised of potassium and hydronium jarosite and the SEM images confirmed that no bacteria were attached to the surface of the jarosite.

Fig. 12. Etch pits caused by bacteria. (a) The AFM topography image of the pyrite surface after removal of the secondary minerals layer revealed etched pits caused by the oxidation of the surface by the bacteria. (b) The hump in the centre of the etch pit as shown in the cross section of the topography image (as indicated by the white line in a) suggests the presence of the bacterial cell, which might be encapsulated in secondary minerals. The measurements were taken 94 h after inoculation and a maximum depth of the pits of ∼50 nm was measured (AFM topography image: height scale: 400 nm).

The amplitude signal of the AFM images showed the presence of crystal steps (Fig. 10) on the surfaces of most jarosite discs. Cross sections taken from the topography images allowed the height of these steps to be measured and the results compared with the ˚ crystallographic dimensions of the jarosite unit cell (a = 7.304 A; c = 17.268 A˚ and Z = 3). The majority of the profiles extracted from the AFM images showed significantly larger step sizes on the jarosite discs suggesting multiple steps. The smallest step height measured on a disc was h = 3.4 nm, corresponding to twice the height of a unit cell. More detailed and accurate analysis of the step height of the discs could have been obtained with higher lateral and vertical resolution but these conditions were not suited to the main goals of this work. The thickness of the jarosite layer was investigated by scratching with a needle to reveal the underlying pyrite surface. This scratch method is commonly used in the AFM community to determine the thickness of, for example, coatings or polymer films. Fig. 11 shows the AFM topography image of a region at the edge of the

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scratch and a profile (indicated by the dotted line) taken from the image. The thickness of the scale was estimated to be 1.27 ± 0.2 ␮m. An additional image of the blank pyrite showed that scratching had not damaged the pyrite surface and also revealed evidence of bacterial attachment in the appearance of etched pits with similar dimensions to bacterial cells. These etched pits are thought to be the result of pyrite oxidation below the attached bacterial cell [32,33]. Fig. 12a shows a scan of an area where the jarosite has been removed. Whereas the sample after polishing and cleaning only showed some minor scratch marks from polishing and cavities due to the polycrystalline nature of the sample, some dark features with the shape and dimension of the bacteria were visible, in particular along the diagonal from the top right to the bottom left of the image within the white outlined area. A cross section through one of these features is shown in Fig. 12b, revealing a trench along the outline of the location where the bacterium once had attached to the surface. From the size and appearance of these etched pits visible in the AFM image Fig. 12a, it is assumed that the bacterial cell is still present in the etched pit but possibly encapsulated by secondary minerals. This result indicated that a certain amount of mineral had been leached due to the presence of the bacterial cell on the surface. However, since it was not possible to monitor the behaviour of the attached bacterial cells once they had been covered with jarosite, the degree of dissolution might provide information about the efficiency and lifetime of the cells on the surface. In the literature, the depth of these etched pits has been reported to be on average approximately 320 nm for samples with bacteria growing for four months [23]. Our measurements were of significantly shorter duration and thus the depth of the etched pits was smaller (∼50 nm). A further interesting aspect is how long the cells are capable of maintaining their own micro-environment. Pace et al. [23] reported the presence of some secondary minerals surrounding or partially encapsulating the attached bacterial cells, but their pyrite surface was not covered with secondary mineral scale, as in our experiments. The coverage of the surface with secondary minerals, which separates the bacterial cells from the bulk leaching solution might have an effect on the efficiency of bacterial oxidation. This aspect may be investigated with future experiments.

4. Summary In this study, bacteria attached to a pyrite surface were dispersed in small groups rather than forming a large localised population. No evidence was found for preferential attachment sites or preferred orientation of the bacteria with respect to discontinuities of the substrate surface. The results show a significant reduction in ferric ion generation rates as conditions deviate from the estimated optimum pH value of pH 1.6 and at the same time an increase of thickness of the EPS on the sample surface. The results highlighted the importance of EPS expressed by the bacteria. When the solution pH was decreased rapidly from pH 2.2 to pH 1.0, the bacteria detached or were dislodged from the pyrite surface and no layer of EPS was detected. In contrast, a layer of EPS with increasing thickness was detected with a staged change in acidity from pH 2.2 to pH 1.4. Upon further addition of acid to pH 0.9, the thickness of the EPS layer decreased immediately. The mechanism responsible for the decreased film thickness was attributed to the formation of areas with denser arrangements of the polymeric chains within the EPS network. The observed behaviour suggested that the bacteria expressed EPS as a response to the stress caused by the increased acidity of the leaching solution. The EPS formed a layer covering the cells and the pyrite surface and acted as a buffer against the unfavourable conditions of the surrounding solution. This behaviour is of particular interest for heap leaching, as the pH

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in the leaching solution is affected by both irrigation with acid and consumption of acid by dissolution of the gangue material. Secondary minerals formed on the pyrite surface only when bacteria were present and eventually covered the entire sulfide surface. Bacterial cells did not attach to the surface covered with this scale. After careful removal of the scale from the pyrite surface, etch pits in the pyrite surface in the shape of the bacteria were observed. The depth of these etch pits may be used in future investigations to determine how long the bacteria remain active after presumably being isolated from the leaching solution by the formation of secondary minerals on the pyrite surface. Acknowledgements The financial support of the Australian Government through CSIRO Minerals Down Under Flagship and the Parker Cooperative Research Centre for Integrated Hydrometallurgy Solutions is gratefully acknowledged. References [1] Johnson DB, Hallberg KB. The microbiology of acidic mine waters. Res Microbiol 2003;154:466–73. [2] Watling HR. The bioleaching of sulphide minerals with emphasis on copper sulphides – a review. Hydrometallurgy 2006;84:81–108. [3] Watling HR, Watkin ELJ, Ralph DE. The resilience and versatility of acidophiles that contribute to the bio-assisted extraction of metals from mineral sulphides. Environ Technol 2010;31:915–33. [4] Binnig G, Quate CF, Gerber C. Atomic force microscope. Phys Rev Lett 1986;56:930–3. [5] Camesano TA, Natan MJ, Logan BE. Observation of changes in bacterial cell morphology using tapping mode atomic force microscopy. Langmuir 2000;16:4563–72. [6] Camesano TA, Logan BE. Probing bacterial electrostatic interactions using atomic force microscopy. Environ Sci Technol 2000;34:3354–62. [7] Bellenberg S, Florian BM, Vera MA, Rohwerder T, Sand W. Comparative study of planktonic and sessile cells from pure and mixed cultures of Acidithiobacillus ferrooxidans and Acidiphilium cryptum growing on pyrite. In: Donati ER, Viera MR, Tavani EL, Giaveno MA, Lavalle TL, Chiacchiarini PA, editors. Biohydrometallurgy: a meeting point between microbial ecology, metal recovery processes and environmental remediation. Zurich: Trans Tech Publications; 2009. p. 333–6. [8] Florian B, Noël N, Bellenberg S, Huergo J, Rohwerder T, Sand W. Attachment behaviour of leaching bacteria to metal sulfides elucidated by combined atomic force and epifluorescence microscopy. In: Donati ER, Viera MR, Tavani EL, Giaveno MA, Lavalle TL, Chiacchiarini PA, editors. Biohydrometallurgy: a meeting point between microbial ecology, metal recovery processes and environmental remediation. Zurich: Trans Tech Publications; 2009. p. 337–40. [9] Gehrke T, Telegdi J, Thierry D, Sand W. Importance of extracellular polymeric substances from Thiobacillus ferrooxidans for bioleaching. Appl Environ Microbiol 1998;64:2743–7. [10] Harneit K, Goksel A, Kock D, Klock J-H, Gehrke T, Sand W. Adhesion to metal sulfide surfaces by cells of Acidithiobacillus ferrooxidans, Acidithiobacillus thiooxidans and Leptospirillum ferrooxidans. Hydrometallurgy 2006;83:245–54. [11] Mangold S, Harneit K, Rohwerder T, Claus G, Sand W. Novel combination of atomic force microscopy and epifluorescence microscopy for visualization of leaching bacteria on pyrite. Appl Environ Microbiol 2008;74:410–5. [12] Mangold S, Laxander M, Harneit K, Rohwerder T, Claus G, Sand W. Visualization of Acidithiobacillus ferrooxidans biofilms on pyrite by atomic force and epifluorescence microscopy under various experimental conditions. Hydrometallurgy 2008;74:410–5. [13] Sand W, Florian B, Noël N. Mechanisms of bioleaching and the visualization of these by combined AFM & EFM. In: Donati ER, Viera MR, Tavani EL, Giaveno MA, Lavalle TL, Chiacchiarini PA, editors. Biohydrometallurgy: a meeting point between microbial ecology, metal recovery processes and environmental remediation. Zurich: Trans Tech Publications; 2009. p. 297–302. [14] Telegdi J, Keresztes Z, Pálinkás G, Kálmán E, Sand W. Microbially influenced corrosion visualized by atomic force microscopy. Appl Phys A 1998;66:S639–42. [15] Bevilaqua D, Diez-Perez I, Fugivara CS, Sanz F, Benedetti AV, Garcia O. Oxidative dissolution of chalcopyrite by Acidithiobacillus ferrooxidans analysed by electrochemical impedance spectroscopy and atomic force microscopy. Bioelectrochem 2004;64:79–84. [16] Pisapia C, Humbert B, Chaussidon M, Mustin C. Perforative corrosion of pyrite enhanced by direct attachment of Acidithiobacillus ferrooxidans. Geomicrobiol J 2008;25:261–73. [17] Taylor ES, Lower SK. Thickness and surface density of extracellular polymers on Acidithiobacillus ferrooxidans. Appl Environ Microbiol 2008;74:309–11. [18] Teschke O. Volume of extracellular polymeric substance coverage of individual Acidithiobacillus ferrooxidans bacterium measured by atomic force microscopy. Microsc Res Tech 2005;67:312–6.

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[19] Teschke O, De Souza EF, Silva-Stenico ME, Fiore M, de F, Etchegaray A. Quorum sensing detected by atomic force microscopy imaging of corrals surrounding multicellular arrangement of bacteria. Microsc Res Tech 2008;71: 112–8. [20] Liu H-L, Chen B-Y, Lan Y-W, Cheng Y-C. SEM and AFM images of pyrite surfaces after bioleaching by the indigenous Thiobacillus thiooxidans. Appl Microbiol Biotechnol 2003;62:414–20. [21] Wilson AD. The micro-determination of ferrous iron and silicate minerals by a volumetric and colorimetric method. Analyst 1960;85:823–7. [22] DiSpirito AA, Tuovinen OH. Uranous ion oxidation and carbon dioxide fixation by Thiobacillus ferrooxidans. Arch Microbiol 1982;133:28–32. [23] Pace DL, Mielke RE, Southam G, Porter TL. Scanning Force Microscopy studies of the colonization and growth of A. ferrooxidans in the surface of pyrite minerals. Scanning 2005;27:136–40. [24] Gorham N, Becker T, Shiers DW, Watling HR. Visualisation of bacterial behaviour using tapping-mode atomic force microscopy. In: Donati ER, Viera MR, Tavani EL, Giaveno MA, Lavalle TL, Chiacchiarini PA, editors. Biohydrometallurgy: a meeting point between microbial ecology, metal recovery processes and environmental remediation. Zurich: Trans Tech Publications; 2009. p. 341–4. [25] Roberson EB, Firestone MK. Relationship between desiccation and exopolysaccharide production in a soil Pseudomonas sp. Appl Environ Microbiol 1992;58:1284–91.

[26] Dogsa I, Kriechbaum M, Stopar D, Laggner P. Structure of bacterial polymeric substances at different pH values as determined by SAXS. Biophys J 2005;89:2711–20. [27] Ding Q, LaBelle M, Yang BY, Montgomery R. Physiochemical studies of extracellular polysaccharides of Erwinia chrysanthemia spp. Carbohydr Polym 2003;51:333–46. [28] Mikkelsen D, Kappler U, Webb RI, Rasch R, McEwan AG, Sly LI. Visualisation of pyrite leaching by selected thermophilic archaea: nature of microorganism–ore interactions during bioleaching. Hydrometallurgy 2007;88:143–53. [29] Sasaki K, Konno H. Morphology of jarosite-group compounds precipitated from biologically and chemically oxidized Fe ions. Can Mineral 2000;38:45–56. [30] Sasaki K, Nakamuta Y, Hirajima T, Tuovinen OH. Raman characterization of secondary minerals formed during chalcopyrite leaching with Acidithiobacillus ferrooxidans. Hydrometallurgy 2009;95:153–8. [31] Gramp JP, Jones FS, Bigham JM, Tuovinen OH. Monovalent cation concentrations determine the types of Fe(III) hydroxysulfate precipitates formed in bioleach solutions. Hydrometallurgy 2008;94:29–33. [32] Kinzler K, Gehrke T, Telegdi J, Sand W. Bioleaching – a result of interfacial processes caused by extracellular polymeric substances (EPS). Hydrometallurgy 2003;71:83–8. [33] Li Y-Q, Wan D-S, Huang S-S, Leng F-F, Yan L, Ni Y-Q, et al. Type IV pili of Acidithiobacillus ferrooxidans are necessary for sliding, twitching motility, and adherence. Curr Microbiol 2010;60:17–24.