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Applied and Environmental Microbiology, January 2008, p. 410-415, Vol. 74, No. 2
0099-2240/08/$08.00+0     doi:10.1128/AEM.01812-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Novel Combination of Atomic Force Microscopy and Epifluorescence Microscopy for Visualization of Leaching Bacteria on Pyrite

Stefanie Mangold,^1,2 Kerstin Harneit,¹ Thore Rohwerder,¹ Günter Claus,² and Wolfgang Sand^1*

University of Duisburg-Essen, Biofilm Centre, Aquatic Biotechnology, Geibelstr. 41, 47057 Duisburg, Germany,¹ Mannheim University of Applied Sciences, Institute of Technical Microbiology, Windeckstrasse 110, 68163 Mannheim, Germany²

Received 3 August 2007/ Accepted 24 October 2007

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Bioleaching of metal sulfides is an interfacial process comprising the interactions of attached bacterial cells and bacterial extracellular polymeric substances with the surface of a mineral sulfide. Such processes and the associated biofilms can be investigated at high spatial resolution using atomic force microscopy (AFM). Therefore, we visualized biofilms of the meso-acidophilic leaching bacterium Acidithiobacillus ferrooxidans strain A2 on the metal sulfide pyrite with a newly developed combination of AFM with epifluorescence microscopy (EFM). This novel system allowed the imaging of the same sample location with both instruments. The pyrite sample, as fixed on a shuttle stage, was transferred between AFM and EFM devices. By staining the bacterial DNA with a specific fluorescence dye, bacterial cells were labeled and could easily be distinguished from other topographic features occurring in the AFM image. AFM scanning in liquid caused deformation and detachment of cells, but scanning in air had no effect on cell integrity. In summary, we successfully demonstrate that the new microscopic system was applicable for visualizing bioleaching samples. Moreover, the combination of AFM and EFM in general seems to be a powerful tool for investigations of biofilms on opaque materials and will help to advance our knowledge of biological interfacial processes. In principle, the shuttle stage can be transferred to additional instruments, and combinations of AFM and EFM with other surface-analyzing devices can be proposed.

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By combining atomic force microscopy (AFM) and fluorescence microscopy, two techniques with complementary strengths and weaknesses are joined to yield a powerful tool for the investigation of biological samples. The atomic force microscope (1, 2, 5) is a mechanical imaging device that requires minimal sample preparation and creates three-dimensional images with high spatial resolution. Since its introduction in 1986 (2), AFM has widely been used in biology owing to its ability to visualize samples under physiological conditions. Consequently, this gives a great advantage over electron microscopy where extensive sample preparation and measurement in high vacuum disallow investigations of native samples. Subnanometer resolution with AFM, for example, has been achieved in the visualization of bacterial cell surface structures consisting of two-dimensional arrays of protein or glycoprotein subunits under physiological conditions (25, 26, 35). The topographic information for AFM images is acquired by scanning a surface with a sharp probe. Since the underlying tip-sample interaction forces are nonspecific, AFM is applicable to a wide range of samples. However, at the same time this limits the capacity of AFM to discern different structures of a sample unless they have a particular shape and size, which have to be known a priori. In contrast, optical imaging techniques such as fluorescence microscopy are routinely used for the identification of various species comprising a sample. By fluorescence labeling of DNA, RNA, proteins, polysaccharides, or lipids, these biomolecules can be tracked in complex samples such as biofilms, single cells, or membranes. However, the resolving power of optical imaging is fundamentally limited by diffraction to about 250 nm or more precisely to half the wavelengths of the light used (17). Consequently, both techniques complement each other in that AFM has excellent spatial resolution but lacks identification capabilities, whereas fluorescence microscopy offers poor resolution but excellent identification properties. Hence, the combination of both microscopy techniques provides AFM with the desirable identification ability. Therefore, the combined microscopy technique has been established for translucent samples within the last decade (9, 15-17, 19, 21, 23, 28, 39). Indeed, the integration of both AFM and transmission fluorescence microscopy in one device is straightforward since an inverted optical microscope can be used for imaging a sample from the back side and AFM measurements can be performed upon the sample. Hence, combinations of AFM with confocal microscopy (9, 16, 17, 19) or with total internal reflection fluorescence microscopy (23, 28) have been reported. However, for the investigation of natural biofilms or other biotic, as well as abiotic, structures on opaque substrata, this combination of both microscopes in one device is not applicable. Nevertheless, considering the outstanding role of opaque materials in many fields, e.g., in biocorrosion and biodeterioration, there is an essential demand for joint imaging with AFM and epifluorescence microscopy (EFM).

In particular, investigations of biofilms on opaque substrata have a high relevance for bioleaching research since specific microbe-mineral surface interactions are involved in the microbially mediated release of heavy metal ions from insoluble compounds. In case of metal sulfides, the metal recovery is based on the action of aerobic, extremely acidophilic iron(II)- and sulfur compound-oxidizing bacteria and archaea (31, 36). By employing these prokaryotes, bioleaching has become an established biotechnological method for the mining of heavy metals such as copper, zinc, cobalt, and gold (31, 33). The same microbial activity, on the other hand, is responsible for the formation of acidic, heavy metal-contaminated waters in natural or anthropogenic leaching biotopes, which are generally termed acid mine drainage (AMD) or acid rock drainage (29). For both optimization of metal bio-winning processes and mitigation of AMD, the underlying bioleaching mechanisms have to be fully understood. It is already known that leaching prokaryotes attach to their solid substrate, i.e., the metal sulfides, and form a monolayered biofilm (7). For doing this, extracellular polymeric substances (EPS) mediate the contact between the prokaryotic cell and the mineral sulfide and play a crucial role in interfacial interactions (10, 14). In the current model, a reaction space is proposed to be formed between mineral and microbial cell surface filled with EPS (32, 34). In this space, EPS-complexed iron(III) ions oxidize the metal sulfide and, subsequently, metal ions, as well as intermediary sulfur compounds, are released. Finally, the reduced iron ions and sulfur species are oxidized by prokaryotic enzyme systems. Consequently, microbial attachment to the metal sulfide surface and the bioleaching mechanism are tightly connected. Hence, the fundamentals of both processes can only be understood by considering the interactions of the prokaryotic cell with the mineral surface. Possibly, the application of new microscopic methods, such as the combination of AFM and EFM, will help to elucidate these specific interactions.

In the present study, we introduce a novel microscopic system allowing investigations on the same location on an opaque sample using both AFM and EFM. The system is based on two separate microscopes and shuttling of the sample between the two devices. As a first example for demonstrating the feasibility of the new system, we studied attached cells of the leaching bacterium Acidithiobacillus ferrooxidans on coupons of the metal sulfide pyrite (FeS₂). For imaging with EFM, DNA of the bacterial cells was stained with DAPI (4',6'-diamidino-2-phenylindole). Subsequently, samples were studied with AFM in liquid or in air.

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Bacterial strain and cultivation.
A. ferrooxidans strain A2 was used, which was previously isolated from the disused sulfidic ore mine Rammelsberg near Goslar, Germany (3). Bacteria were grown aerobically at 28°C in mineral salt solution according to the method of Mackintosh (22) containing 2 g of ferrous iron per liter of medium. Bacterial cells were harvested in their exponential growth phase by vacuum filtration so cells retained their EPS. Strain A2 is maintained on the ferrous iron medium in our strain collection at the University of Duisburg-Essen.

Substratum and biofilm formation.
Coupons with a final size of about 10 by 10 by 1 mm were cut from each face of single cubic pyrite crystals (museum-grade) originating from Navajum, Spain. The coupons were immersed in boiling 6 M HCl for 30 min to remove iron ions and washed with acetone to remove sulfur compounds as described elsewhere (14). Next, three to five sterile pyrite coupons were incubated aerobically at 28°C in a bacterial suspension with 5 x 10⁸ cells per ml of mineral salt solution in volumes of 40 to 50 ml.

AFM and EFM instrumentation.
A NanoWizardII atomic force microscope (JPK Instruments, Germany) and an upright epifluorescence microscope (AxioImager A1m; Zeiss, Germany) with a x100 water-immersible objective (Achroplan; numerical aperture 1.0; working distance, 0.97 mm) without using coverslips were combined using the BioMaterialWorkstation (JPK Instruments). Throughout the present study the prototype of this new system was used. The key feature of the BioMaterialWorkstation was a shuttle stage that carried the actual sample precisely fixed on a glass slide. This shuttle stage could be transferred between the atomic force microscope and the epifluorescence microscope, giving a precise positioning of the stage on both microscopes. Furthermore, a precision sample clamp guaranteed a tight and accurate fixation of the sample to the shuttle stage, thereby allowing the retrieval of the same sample location with AFM and EFM with an error of no more than 3 to 5 µm. For sequential investigations, this shuttling could be repeated as often as required without losing position. For a successful combination of both microscopes, meaning the visualization of the same sample location, the variable position of the AFM cantilever had to be aligned to the static optical axis of the epifluorescence microscope in order to match the AFM scan region with the epifluorescence microscope's field of view. For AFM imaging, silicon cantilever CSC37 A (Mikromasch, Estonia) with the following features was used: typical length, 250 µm; width, 35 µm; thickness, 2 µm; resonance frequency, 41 kHz; and nominal force/spring constant, 0.65 N/m. Each AFM image consists of 512 by 512 pixels. The AFM image acquisition details are given in the legends of the figures. Mineral salt solution was used as immersion medium for EFM.

Biofilm preparation and imaging conditions.
Biofilms and attached cells on pyrite coupons were investigated with combined AFM and EFM. Coupons were incubated in bacterial suspension for 4 days to allow attachment and biofilm formation. Subsequently, they were stained with 0.01% (wt/vol) DAPI for 10 min and either kept hydrated with mineral salt solution or air dried at ambient temperature for 1 h prior to imaging. After staining, the pyrite coupons were glued to a glass slide using a two-component adhesive on epoxy resin base (Uhu Plus Sofortfest; Uhu, Germany), because the shuttle stage was designed to hold a glass slide. AFM imaging was performed by contact mode in air, by contact mode in mineral salt solution (fluid), or by intermittent contact mode in mineral salt solution using cantilever CSC37 A.

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Combined imaging by EFM and AFM in air.
An example of combined AFM and EFM imaging of a DAPI-stained biofilm sample is presented in Fig. 1. Figure 1A shows the vertical deflection image of an AFM scan acquired by contact mode in air. Several rod-shaped structures of 1 to 2 µm length are visible, representing mostly single attached cells of A. ferrooxidans. Figure 1B displays the EFM image of the same sample location showing as many DNA-containing fluorescent spots as rod-shaped structures are evident in the corresponding AFM image. The unambiguous pattern of the bacterium-like structures in both images confirms that the same sample location was imaged. Consequently, with the EFM image, the topographic features in the AFM image can be clearly identified as bacterial cells. Figure 2 displays high-resolution images of cells of A. ferrooxidans on pyrite in air. The surface structure is visible in detail. In addition, EPS surrounding the cell and granular precipitates about 100 nm in length are visible. In the course of AFM imaging of A. ferrooxidans biofilms on pyrite in air, the bacterial cells remained always at their location, even for repeated AFM scans.

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FIG. 1. Combined AFM and EFM images of a biofilm sample of cells of A. ferrooxidans A2 on pyrite stained with DAPI. (A) Vertical deflection image of an AFM scan acquired by contact mode in air (20 by 20 µm; line rate, 2 Hz for both trace and retrace; overscan time, 0.031 s; total time, 4.87 min). (B) EFM image of same sample location. Scale bar, 5 µm.

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FIG. 2. High-resolution images of cells of A. ferrooxidans on pyrite. Vertical deflection images of AFM scans acquired by contact mode in air. (A) Imaging of a single cell (2 by 2 µm; line rate, 0.7 Hz; overscan time, 0.077 s; total time, 13.50 min). (B) Two neighboring cells (1.5 by 1.5 µm; line rate, 0.8 Hz; overscan time, 0.068 s; total time, 11.83 min). 1, Surface structure of a cell of A. ferrooxidans; 2, bacterial EPS surrounding the cell; 3, precipitations.

Combined imaging by EFM and AFM in mineral salt solution.
In Fig. 3A, bacteria attached to a pyrite surface have been visualized using AFM imaging in mineral salt solution. Generally, due to the hydration the cells seemed to be larger and their surfaces were smoother than those observed by contact mode imaging in air (data not shown). However, in contrast to imaging by AFM in air, AFM scanning in mineral salt solution caused detachment of some cells from the pyrite surface. When the sample was constantly kept hydrated during sample preparation, bacteria were detached by contact mode, as well as by intermittent contact mode. In Fig. 3, the detachment of some cells due to scanning in contact mode in mineral salt solution is documented. In detail, Fig. 3A to C display a series of vertical deflection images of sequential AFM scans at the same sample location. Figure 3D and E are the corresponding EFM images, where Fig. 3D was recorded prior to AFM scanning, and Fig. 3E was acquired after AFM scanning. In the first AFM image (Fig. 3A), all cells which are present in the corresponding EFM image within the AFM scan area (white inset frame in Fig. 3D) are visible. In the following AFM images (Fig. 3B and C), some cells are missing. Apparently, they were detached due to the interaction with the AFM probe. The white streaks in these images are scanning disturbances, which can be of various origins. Generally, the tip surface interaction is obstructed by some sample constituent in such a way that true surface information is not collected and image noise is displayed instead. The obstructions can be due to EPS residues adhering to the AFM tip until the tip breaks free of this interaction as it moves along the sample. In the AFM scans presented here, the image noise close to partly imaged bacterial cells (Fig. 3A to C) was caused by the loosened cells themselves. The white streaks correspond to the cell as it rapidly moves out of the scan area. In addition, the EFM image taken after AFM scanning (Fig. 3E) demonstrates that the cells were actually relocated on the surface. Only one bacterial cell resisted the AFM scanning forces and remained at the same position throughout the imaging process. After relocation of the cells, sometimes microbial footprints become visible in the AFM images (Fig. 3B and C) at those locations where formerly the cells had been attached. These footprints, indicated by the number 3 in Fig. 3C, are circular and have the size and shape of bacterial cells. Additional AFM images with microbial footprints acquired by contact mode in fluid are presented in Fig. 4, showing the footprints of several cells at high resolution. Air drying of the sample for 1 h prior to an AFM scan in liquid prevented a detachment of cells by intermittent contact mode but not by contact mode (data not shown).

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FIG. 3. Relocation of cells by AFM scans visualized by AFM and EFM. Images of a biofilm sample, i.e., cells of A. ferrooxidans A2 on pyrite stained with DAPI are shown. (A to C) Vertical deflection images of a series of three AFM scans (20 by 20 µm; the line rate, overscan time, and total time were, respectively, 2 Hz, 0.031 s, and 4.87 min [A], 1 Hz, 0,056 s, and 9.49 min [B], and for 1.8 Hz, 0,033 s, and 5.30 min [C]). The "3" in panel C indicates microbial footprints. (D) Corresponding EFM image prior to AFM scans. (E) EFM image after AFM scans. White inset frame indicates the AFM scan area. 1, Attached bacterial cells outside the scan area remained unaffected from scanning; 2, bacterial cells relocated due to AFM scanning. The white arrow in panels A to E indicates a bacterial cell not relocated during scanning. Scale bar, 5 µm.

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FIG. 4. Microbial footprints of cells of A. ferrooxidans on pyrite after detachment of bacteria. Vertical deflection images of AFM scans recorded by contact mode in fluid. (A) Footprints of three cells (5 by 5 µm; line rate, 0.9 Hz; overscan time, 0.061 s; total time, 10.52 min). (B) Footprint of a single cell (2.5 by 2.5 µm; line rate, 1 Hz; overscan time, 0.056 s; total time, 9.49 min). The "1" indicates microbial footprints.

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In this study, a combination of AFM with EFM for opaque materials was successfully demonstrated. Imaging of the same sample location by both techniques is thus possible. Imaging with AFM of cells of A. ferrooxidans A2 on pyrite was possible by scanning in contact mode in air or by intermittent contact mode in fluid after the sample was air dried. In previous studies, it has been noted that planktonic bacteria that are not sufficiently immobilized to the surface might be moved by the AFM probe during imaging in fluid (5, 40). In the case of planktonic bacteria, simple physical adsorption of whole bacterial cells to a substratum is often not sufficiently strong as the contact area between cell and surface is small. To circumvent this problem, air drying, chemical fixation, or mechanical immobilization using membrane filters were suggested (5). In contrast, the problem of a relocation of sessile bacterial cells, meaning not attached planktonic cells but the ones grown in a biofilm environment, by the interaction with the AFM probe has not yet been addressed (1). However, the important role of bacterial EPS for attachment and the natural immobilization of bacteria within biofilms are pointed out. In conclusion, the investigations of native biofilms without the need of additional fixation are preferable.

Thus far, biofilms of A. ferrooxidans on metal sulfides have not been visualized under physiological conditions, since AFM investigations have been carried out in air only (14, 24, 30). However, fully hydrated biofilms of other organisms have been visualized by using AFM (4, 18). In the study of Bremer et al. (4), 7-day-old freshwater biofilms on copper surfaces were examined by scanning in culture medium. These AFM images revealed a heterogeneous multilayered biofilm and bacterial cells covered with EPS. Detachment or dislocation of bacterial cells within these biofilms is not reported. In contrast, the work of Kolari et al. (18) demonstrated that cells of Deinococcus geothermalis in 1-day-old, hydrated biofilms are dislocated but not detached by the AFM probe. These authors attribute this behavior to flexible, adhesive polymers that provide bacterial cells with a strong but flexible gluing to the surface. As shown in our study the attachment of A. ferrooxidans after 4 days of biofilm formation is not sufficiently strong to withstand the lateral forces caused by the AFM probe under some experimental conditions. As bacterial attachment grows stronger with time, imaging of unfixed natural biofilms of A. ferrooxidans might be possible with older samples.

After bacteria were dislocated by AFM scanning, microbial footprints were visible at the former attachment location on the pyrite. In line with this observation are previous reports indicating that bacteria are able to label surfaces with various substances, mainly EPS (27). Most probably, the microbial footprints of A. ferrooxidans A2 visualized here are such EPS, since these play a pivotal role in attachment and the so-called contact mechanisms in bioleaching (10, 12, 14, 32, 34). Using the new system, it becomes now possible to validate the role of EPS and their composition by staining them with fluorescently labeled lectins (20, 38), followed by a combined visualization with AFM and EFM.

In the present study, we show that the new system for combined imaging with AFM and EFM on opaque samples is feasible for the application to A. ferrooxidans on pyrite. Most materials, whether they are manufactured or natural, are not translucent. For this great variety of materials, the novel system described here offers new opportunities for combined investigation by AFM and EFM. As demonstrated here there will be a great potential for combined AFM and EFM in the investigation of biofilms on opaque substrata. Biofilms are the most important mode of growth for prokaryotes, and they have implications in industry, infectious disease, and the natural environment (1, 13, 37). Thus, it is important to learn more about this stage of prokaryotic life. In this connection, the novel system can help to elucidate many aspects of biofilms formed on their natural or synthetic opaque substrata. AFM images of hydrated biofilms can provide information about their spatial arrangement and thus give indications about mass transfer from bulk solution to single cells. In combination with EFM, various constituents of the biofilm and its matrix can be identified. Possible applications are microbially influenced corrosion, biofilms formed on medical implants, or biofouling in industrial plants. In addition, combined AFM and EFM can now be applied to the investigation of opaque plant material. Moreover, not only are biological applications possible but also investigations in surface chemistry involving fluorescent polymers or coatings can profit from the combination of AFM and optical microscopy. Thus, the combination of AFM and EFM is a powerful tool for the research on surface related phenomena and offers new opportunities for the investigation of all kinds of opaque samples. In principle, the employed shuttle stage can also be transferred to additional devices. Thus, a combination with other instruments, e.g., for measuring charge differences on the surface of materials by a Kelvin probe (10) or the quantitative mapping of elemental distributions by synchrotron X-ray fluorescence microscopy (6, 8, 11), may be achieved in the future.

 
This study was carried out in the frame of COST D33 and BioMinE (European project contract NMP1-CT-500329-1). We acknowledge the financial support given to this project by the European Commission under the Sixth Framework Programme for Research and Development.

We also thank JPK Instruments for their cooperation.

 
* Corresponding author. Mailing address: University of Duisburg-Essen, Biofilm Centre, Aquatic Biotechnology, Geibelstr. 41, 47057 Duisburg, Germany. Phone: 49 203 379 4475. Fax: 49 203 379 4495. E-mail: wolfgang.sand{at}uni-due.de

Published ahead of print on 26 November 2007.

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Applied and Environmental Microbiology, January 2008, p. 410-415, Vol. 74, No. 2
0099-2240/08/$08.00+0     doi:10.1128/AEM.01812-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

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