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Correlative Fluorescence- and Scanning, Transmission Electron Microscopy for Biomolecular Investigation Kristina Jahn , Deborah Barton , and Filip Braet Australian Key Centre for Microscopy & Microanalysis, The University of Sydney, NSW 2006, Australia These days, cellular and molecular biology are closely integrated due to the development of novel correlative biomolecular microscopy platforms, even though this avant-garde vision of combining different imaging tools has been around since the late 1960s. The current boom and successful application of correlative imaging methods to dissect the cell’s internal structure is progressing faster than ever as witnessed by the growth in the number of papers dealing with correlative microscopy. In this chapter we outline in-depth two straightforward methods to combine fluorescent and electron microscopic information of a single cell by using confocal laser-, scanning electron- and transmission electron microscopy, and discuss how these methods contribute to our understanding of biomolecular cell processes, such as cell movement and cytoskeleton dynamics. Key words: Combined Microscopic Imaging, CeLLocate Coverslips, Electron microscopy, Fluorescence Microscopy, Finder Grids, Confocal Imaging, Image Analysis, Immunocytochemistry, Immunogold Labelling 1. Introduction Correlative microscopy, by using combined light-, probe-, laser- and electron microscope techniques has become increasingly important for the analysis of the structure and function of cells and tissues. New concepts and progress in structural and molecular cell biology have been discovered thanks to improving correlative microscopy techniques that continue to rely predominantly on advances in new, three- dimensional (X, Y and Z) visualization techniques [1]. It is clear from the literature that the development of correlative imaging methods has not come to an end and that soon the time dimension (X, Y, Z and t) will be added by bridging the time resolution gap through the use of rapid-sample transfer systems [2]. Ideally, correlative microscopy can be defined as an imaging platform that aims to capture exactly the same structures within a single cell using two or more different microscopy techniques, preferably with different resolution limits [3, 4]. Consequently, correlative imaging allows the researcher to gain additional novel morphological information about their sample and this provides a degree of confidence about the structures of interest, as information obtained with one method can be compared to that seen with the other methods [3, 5, 6]. Nowadays, these combined microscopy methods have a significant impact in the fields of biology, biotechnology and biomedical sciences in which gathering correlative structural information is a fundamental requirement at both tissue, cellular and molecular levels [7]. For excellent reviews dealing with this matter we refer to references [8, 9]. In this book chapter, we discuss with the aid of practical examples two straightforward methods to combine fluorescent and electron microscopic information of actin microfilaments within the same cell using confocal laser-, scanning electron- or transmission electron microscopy. We also compare two different ways to fluorescently label the filaments and outline the specific specimen preparation techniques necessary to achieve these. These two techniques in combination provide complementary information about the structure of actin filament arrays within cells. Similar correlative imaging methods have been applied previously in our quest to gather new structural information about the fine Corresponding author: E-mail: [email protected], Phone: + 61 2 93517619 K.J. and D.B contributed equally to this work Modern Research and Educational Topics in Microscopy. A. Méndez-Vilas and J. Díaz (Eds.) ©FORMATEX 2007 _______________________________________________________________________________________________ 203

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Correlative Fluorescence- and Scanning, Transmission Electron Microscopy for Biomolecular Investigation

Kristina Jahn†, Deborah Barton†, and Filip Braet∗ Australian Key Centre for Microscopy & Microanalysis, The University of Sydney, NSW 2006, Australia

These days, cellular and molecular biology are closely integrated due to the development of novel correlative biomolecular microscopy platforms, even though this avant-garde vision of combining different imaging tools has been around since the late 1960s. The current boom and successful application of correlative imaging methods to dissect the cell’s internal structure is progressing faster than ever as witnessed by the growth in the number of papers dealing with correlative microscopy. In this chapter we outline in-depth two straightforward methods to combine fluorescent and electron microscopic information of a single cell by using confocal laser-, scanning electron- and transmission electron microscopy, and discuss how these methods contribute to our understanding of biomolecular cell processes, such as cell movement and cytoskeleton dynamics.

Key words: Combined Microscopic Imaging, CeLLocate Coverslips, Electron microscopy, Fluorescence Microscopy, Finder Grids, Confocal Imaging, Image Analysis, Immunocytochemistry, Immunogold Labelling

1. Introduction

Correlative microscopy, by using combined light-, probe-, laser- and electron microscope techniques has become increasingly important for the analysis of the structure and function of cells and tissues. New concepts and progress in structural and molecular cell biology have been discovered thanks to improving correlative microscopy techniques that continue to rely predominantly on advances in new, three-dimensional (X, Y and Z) visualization techniques [1]. It is clear from the literature that the development of correlative imaging methods has not come to an end and that soon the time dimension (X, Y, Z and t) will be added by bridging the time resolution gap through the use of rapid-sample transfer systems [2]. Ideally, correlative microscopy can be defined as an imaging platform that aims to capture exactly the same structures within a single cell using two or more different microscopy techniques, preferably with different resolution limits [3, 4]. Consequently, correlative imaging allows the researcher to gain additional novel morphological information about their sample and this provides a degree of confidence about the structures of interest, as information obtained with one method can be compared to that seen with the other methods [3, 5, 6]. Nowadays, these combined microscopy methods have a significant impact in the fields of biology, biotechnology and biomedical sciences in which gathering correlative structural information is a fundamental requirement at both tissue, cellular and molecular levels [7]. For excellent reviews dealing with this matter we refer to references [8, 9]. In this book chapter, we discuss with the aid of practical examples two straightforward methods to combine fluorescent and electron microscopic information of actin microfilaments within the same cell using confocal laser-, scanning electron- or transmission electron microscopy. We also compare two different ways to fluorescently label the filaments and outline the specific specimen preparation techniques necessary to achieve these. These two techniques in combination provide complementary information about the structure of actin filament arrays within cells. Similar correlative imaging methods have been applied previously in our quest to gather new structural information about the fine

∗ Corresponding author: E-mail: [email protected], Phone: + 61 2 93517619 †K.J. and D.B contributed equally to this work

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organization of peculiar actin structures around transendothelial pores in liver endothelial cells [4, 10] and the microtubular organization in plant cells [11].

2. Materials and Methods

Cultured cancer cells were prepared for correlative microscopic examination essentially through two major steps: (i) they were extracted and fluorescently labelled for confocal laser scanning microscopy investigation; and (ii) the same cells were then postfixed, stained and dried for high-resolution imaging using scanning or transmission electron microscopy. Actin filaments within the cells were labelled either with a fluorescently tagged phallotoxin known to bind to actin filaments [12] (Section 2.1) or using anti-actin antibodies [13, 14] (i.e., immunocytochemistry) (Section 2.2). For both methods, cells were cultured on nickel finder grids (Proscitech, Australia) that had been coated with a thin film of formvar and pretreated with 20 µg / mL collagen (Sigma-Aldrich, Australia). The formvar film provided a surface on which the cells could grow, and the nickel finder grids were used to locate the precise position and orientation of cells using the different microscopes.

2.1. Combined confocal laser scanning- and transmission electron microscopy

In this section, a rapid sample preparation protocol for the labelling of filamentous actin in cultured cells using fluorescent phalloidin and subsequent correlative imaging with confocal laser scanning microscopy & transmission electron microscopy is outlined. To image other components of a cell’s interior, probes such as fluorescent taxol that labels microtubules [12], can be substituted for the fluorescent phalloidin. The following protocol in vide infra is used routinely for actin labelling: - Culture medium was removed and cells were immediately extracted for 5 min at room temperature with cytoskeleton extraction buffer composed of PEM-buffer (100 mM piperazine-N,N'-bis [2-ethanesulfonic acid], 1 mM ethylene glycol bis [2-aminoethylether]-N,N,N',N' tetra-acetic acid, 1 mM MgCl2), containing 4% polyethylene glycol 20.000 (PEG), 1% Triton X-100 (BDH Chemicals Ltd., Poole, England) and 5 µM phalloidin (Sigma Aldrich, P-2141) at pH 7.1. - The samples were then extensively rinsed with PEM-buffer and grids were recovered by a pair of magnetic tweezers to preserve the integrity of the EM supports and placed on parafilm. - Samples were stained for filamentous actin with 165 nM Alexa Fluor® 594 labelled phalloidin (Invitrogen, Cat. No. A12381) and with 0.5 µg DAPI per mL for nuclear identification (Sigma Aldrich, Cat. No. D9564). Both stains were mixed in PEM-buffer and 10 µl of this solution was applied to each grid for 20 min at room temperature under humidified conditions. - Then the samples were washed twice in PEM-buffer. - Grids were mounted onto microscopic glass slides in a 1:1 anti-fading mixture composed of Citifluor® (Leica, Cat. No. R1321 AF2) and PEM-buffer containing 3 mg / mL ascorbic acid. - The grids were covered with glass coverslips that were sealed to the slides with nail polish. - The cells were examined immediately with a confocal laser scanning microscope and those containing structures of interest were imaged. The locations of these cells of interest were documented using the finder grid marks in transmission light optical mode. - Next the grids were recovered from the slides and rinsed twice in PEM-buffer. - The cells were then fixed in 2% glutaraldehyde in 0.1 M sodium cacodylate buffer for 20 min at room temperature. - The samples were incubated in 0.1% aqueous tannic acid solution for 20 min, then rinsed twice in distilled water followed by incubation for 5 min in distilled water. - Following this, the samples were treated with 0.16% uranyl acetate dissolved in distilled water for 20 min. - Samples were then rinsed twice with distilled water followed by replacing the water phase by 10% ethanol before further dehydration in a graded ethanol series.

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- After two rinses with 100% ethanol the samples were transferred to 0.16% uranyl acetate in absolute ethanol solution for 20 min. - The grids were rinsed several times in 100% ethanol, stored for 5 min in 100% ethanol and subsequently transferred to 100% hexamethyldisilazane (HMDS) (Sigma Aldrich, 999-97-3) for 3 minutes [15]. - The grids were transferred to a desiccator for at least 30 min before rotary shadowing with platinum (10 sec at α 45°) followed by carbon coating for 20 sec at an angle of 90°. - Samples were examined with a transmission electron microscope at a voltage of 120 kV. - Cells previously imaged with the confocal laser scanning microscope were relocated by using the α numerical marks. Digital TEM images were captured at the same end magnification using a CCD camera. - The digital data obtained from both microscopies were transferred to Adobe Photoshop® or ImageJ software [16] for colour adjustment and figure assembly by using the replace colour and duplicate layer/merge options [10].

Fig. 1. Cells cultured on electron microscopy supports were stained for filamentous actin and subsequently imaged using (A) confocal laser scanning microscopy and (B) transmission electron microscopy. Image C shows the merged image information. Note, large arrows denote the cortical actin bands at the rim of the cells; the small arrows show the cytoplasmic actin fibres. The asterisk shows a cell that was initially visualized with the confocal microscope, but was lost in the subsequent preparation steps for transmission electron microscopy. Scale bars A-C, 20 µm. (D) Figure D is reproduced at a higher magnification by analysing the cytoplasmic area (see, dotted box in image C for

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comparative reasons) using ImageJ, revealing the fine individual labelled actin fibres (arrows) at high resolution. Scale bar D, 2 µm.

2.2. Combined confocal laser scanning- and scanning electron microscopy

The sample preparation protocol as outlined in Section 2.1 is not recommended for antibody labelling of the actin cytoskeleton of cultured cells. Instead a second method that allows successful imaging of immunofluorescent labelling both with confocal laser scanning microscopy and high-resolution scanning electron microscopy is outlined below, and is partially based on a modification of the method by Svitkina & Borisy [17]: - Cells cultured on nickel finder grids were rinsed twice briefly in warm culture medium, then extracted in PEM-buffer including 4% PEG 20,000 and 1% Triton-X-100 (without the addition of phalloidin) for 4 min. - The cells were immediately rinsed in PEM-buffer, with three rinses over approximately 5 min. - They were immediately fixed in 1% paraformaldehyde (16% stock solution from Proscitech, Australia) and 0.5% glutaraldehyde (25% stock solution from Proscitech, Australia), diluted in phosphate buffered saline (PBS: 2.68 mM KCl, 1.47 mM KH2PO4, 137 mM NaCl and 4.5 mM Na2HPO4). - After rinsing in PBS the cells were further extracted by incubating them with 33% methanol for 15 min at 37ºC. - The cells were rehydrated in PBS over 20 min, then blocked for non-specific antibody binding with 3% bovine serum albumen (BSA) dissolved in PBS. - They were then incubated with the primary antibody, anti-actin (raised in mouse, Clone C4, MP Biomedicals, USA), diluted to 1/100 in PSB containing 1% BSA, for one hour at 37ºC. - Following rinsing in PBS, the cells were incubated with FluoroNanogold® conjugated anti-mouse (Nanoprobes, Yaphank, USA) diluted to 1/25 in PBS, for one hour at 37ºC. - After rinsing in PBS, the grids were mounted onto glass slides in a 1:1 mixture of Citifluor® diluted in PBS including 3 mg / mL ascorbic acid, covered with glass coverslips and sealed with nailpolish. - The cells were imaged with a confocal laser scanning microscope using a 100× oil immersion lens. Series of images collected in the z-plane were combined as single projected images. The positions of imaged cells on the finder grid were recorded for later use. - The grids were rescued from the slides and rinsed thoroughly with PBS. - The 1.4 nm Nanogold particles of the FluoroNanoGold® secondary antibody were then enhanced in size using a Goldenhace kit (Nanoprobes, Yaphank, USA). In brief, the cells were firstly rinsed with 50 mM glycine, then with PBS containing 0.05% Tween-20 (polyoxyethylenesorbitan monolaurate; Sigma-Aldrich, Australia) and 3% BSA, followed by a thorough rinsing in distilled water. - The gold-enhance solution was made by adding 1:5:1:1 solutions A:B:C:D (substitute comprising 0.05 M NaH2PO4 and 0.1 M NaCl) and was next applied to the cells for 5 min. Following which the reaction was stopped by the addition of 3% sodium thiosulfate for 30 seconds. The cells were rinsed in 10% acetic acid and 10% glucose, followed by rinsing thoroughly in distilled water. They were rinsed also in PBS. - The cells were postfixed in 0.5% osmium tetroxide for 10 minutes at 4ºC, rinsed in PBS, then stained with 0.1% aqueous tannic acid for 10 minutes. They were rinsed in distilled water, following which they were stained with 0.16% aqueous uranyl acetate and again rinsed in distilled water. - The cells were dehydrated through a series of ethanol concentrations. Following two rinses with pure 100% ethanol the cells were transferred to 100% HMDS for 3 min. - They were then transferred to a dessicator to dry for at least 30 min before being coated with approximately 2-3 nm platinum in a sputter coater. - The cells were imaged using an in-lens field-emission scanning electron microscope (FESEM) at 10 kV. Cells imaged with confocal microscopy were relocated following directions recorded previously. Images of a single area were collected with both the secondary electron and backscattered electron detectors.

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- Images of the same cell taken with confocal microscopy and scanning electron microscopy were overlaid using Adobe Photoshop®.

3. Results and Discussion

3.1. Combined confocal laser scanning- and transmission electron microscopy

In the past three decades, fluorescent phalloidin has been used extensively as a probe for actin filaments [18]. Phalloidin itself is a phallotoxin that works by binding to single actin filaments and can be used on both live and fixed samples. However, when individual actin filaments (with a diameter in the order of 6 nm) are labelled with fluorescent phalloidin, they cannot be resolved using confocal laser scanning microcopy due to its resolution limits (~ 0.225 µm). It is necessary therefore, to use electron microscopy to study the structure and organisation of actin filaments within cells. But by combining confocal laser scanning microscopy and electron microscopy analysis of a single cell, this can provide complementary information about both the wider distribution of filaments (confocal microscopy) and the finer structural details (electron microscopy) of the arrays. Therefore, to correlate fluorescently-labelled actin structures with electron microscopic information, we designed a protocol to compare fluorescent and transmission electron microscopy images of an actin array within a single cell. This enabled us to correlate the overall actin distribution as well as the fine structure of the labelled cytoskeletal proteins down to the nanometer scale.

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Fig. 2. Cells grown on Formvar coated nickel finder grids were extracted, chemically fixed then immunolabelled with an antibody against actin. (A) Using a FluoroNanogold® secondary antibody, actin filaments were imaged using confocal laser scanning microscopy. Notable features are the finger-like protrusions, or filopodia, and the bright leading edge of actin filaments at the cell periphery. (B) The cell was processed and subsequently imaged with high resolution scanning electron microscopy. Note that although the filopodia are easily resolved at low magnification, individual actin filaments are not. (C) Overlaying the confocal image over the high resolution scanning electron microscopy image allows a more detailed analysis of the precise location of actin bundles throughout the cell. Scale bars A-C, 10 µm. (D) At higher magnification of the boxed area in (B), individual actin filaments are resolved (arrows) and the leading edge is seen as a conglomeration of filaments (double ended arrow). (E) The immunogold label of the FluoroNanogold® secondary antibody was enhanced in size and is seen as bright globules in the backscattered electron image. Scale bars D-E, 500 nm.

Examination of Alexa Fluor® 594 phalloidin stained cells revealed intense circular bundles lining the cell periphery and straight bundles traversing the cytoplasm (Fig. 1A). These same cells were also imaged using a transmission electron microscope (Fig. 1B). Projecting the electron micrograph on top of the corresponding confocal image clearly illustrates the matching structural relationship between filamentous actin and the entire interior of the whole mount cell (Fig. 1C). Close examination by post processing with ImageJ revealed fine staining of actin fibres at the rim of the cell which co-localizes with fine individual finger-like protrusions (Fig. 1D). By applying fluorescent-labelled phalloidin, or other probes, after detergent extraction of whole-mount cells, difficult and complex procedures that demand highly skilled expertise such as serial sectioning, fine structure immunogold labelling or microinjection of fluorescent-labelled anti-actin antibodies are avoided. This leads overall to a shorter experimentation time and consequently bridges the time gap between specimen preparation and observation by both microscopies [2]. We advise therefore the use of simple and uncomplicated preparation protocols as outlined under Section 2.1, since this direct approach reduces the number of possible specimen preparation artefacts. Nevertless, you can take as many precautions as you want, artefacts are always there, it is just a matter of discovering them. Fig. 1C for example nicely illustrates that point in which a cell was lost during the preparation for transmission electron microscopy.

3.2. Combined confocal laser scanning- and scanning electron microscopy

Immunofluorescence cytochemistry, while a little more time consuming in specimen preparation, can provide just as much information about the localisation of actin filaments within the arrays of cultured cancer cells as using conventional probes (Fig. 2). Features such as filopodia and the bright leading edge of actin filaments at the periphery of lamellipodia are visible, as well as thicker bundles around the nucleus (Fig. 2A). These correspond to features seen in the high resolution scanning electron micrograph (Fig. 2B). When the corresponding images are overlaid (Fig. 2C) the position of the large filpodia and lamellipodia edges correlate well to the finger-like protrusions and cell edge as seen in the high resolution scanning electron micrograph. At higher magnification, individual actin filaments composing the leading edge of a small lamellipodium are visible (Fig. 2D), as is the immunogold label (Fig. 2E). This antibody labelling method is a powerful and useful technique that can be used to locate precisely the position of actin filaments within arrays at both the optical and electron microscope level. Also, providing that there are antibodies available, it can be used to locate the precise location of other proteins associated with actin filaments throughout an array. This method also has the advantage of not using phalloidin to stabilise the actin filaments prior to fixation, so although some filaments may be lost during the initial extraction step, the structure of the arrays has not been affected by any drug treatment. Indeed, the affect of drug application to the individual actin filaments can be investigated using this technique.

3.3. Sample preparation tips for successful correlative imaging

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An important contributor to the success of these methods for correlative fluorescence- and electron microscopy observations was the use of a special mounting medium that allowed us to observe the samples for prolonged periods. Photobleaching or fading is a universal problem encountered during fluorescence or confocal laser microscopy observation and results in underestimating and poor imaging of the labelled structures or in the worse case in no detection at all. To limit this, ascorbic acid was found to be an essential additive and was included in the commercial mounting media available. Furthermore, glycerol / phosphate buffered saline-based anti-fading solutions were preferred over other mounting media, because the samples could be easily recovered for subsequent transmission electron microscopy processing. Hexamethyldisilazane (HMDS) is an organic, low surface tension having-, and at room temperature quickly evaporating compound that has been applied by many others for drying samples for electron microscopy [19]. Critical shrinkage induced by critical point drying is a well known artefact in electron microscopy and accounts for about 15% of the overall dimensional changes during sample preparation. In addition, it is generally known that the thick, complex nuclear area is prone to distortion during alcohol dehydration and subsequent drying, whereas the flat cytoplasmic areas are not affected to the same extent. We proved previously that hexamethyldisilazane is extremely useful for drying whole-mount cells for electron microscopy and that it results in less perinculear tension and damage during the drying process [15]. Therefore, we advise the use of this organic compound in specimen preparation because not only does it save a considerable amount of time, it is also proven to be a reproducible drying method.

4. Concluding Remarks

From these findings it is clear that the application of two different, but complementary with respect to sample preparation, high-resolution correlative microscope methods facilitates the collection of new structural data that cannot be resolved by classical or standard microscopy methods. In other words, you get a very powerful platform when you start combining the strengths of different microscopy instruments and sample preparation methods (Fig. 3) [20]. One would imagine that there are still numerous combinations of imaging methods versus sample preparation protocols that are yet to be defined. So, we might expect in the forthcoming years a second boom in the development of correlative imaging methods. This will undoubtedly result in even better, faster and user friendly protocols for combined biomolecular imaging. In the future, ground-breaking developments in correlative microscopy can be expected in this exciting era of digital technology [21], and will certainly speed up scientific breakthrough on cellular nanostructures over a wide range of length and time scales. High-speed cameras, predictive computer models, new data-processing methods and multifunctional microscopes are a few examples that will be all important contributors in the availability of new correlative microscopy imaging techniques for the broader scientific community. Ideally, this technology, in combination with fast sample transfer systems [2], and the development of novel probes for labelling cells [22, 23], will contribute directly to identifying and characterizing the complex molecular machinery of cells in the third (and fourth) dimension.

Acknowledgements. The authors acknowledge the facilities as well as technical assistance from staff at the Australian Key Centre for Microscopy and Microanalysis (AKCMM) of The University of Sydney. The authors are grateful also to Ms. E. Korkmaz for excellent assistance in preparing the samples for microscopy and Dr. L. Soon for stimulating discussions. This work was supported by the Cancer Research Fund of the University of Sydney (2006) and partially by the ARC/NHMRC FABLS Research Project Network (RN0460002 [D.B. & F.B]). K. Jahn. and D. Barton contributed equally to this work.

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Fig. 3. Schematic diagram depicting the key sample preparation steps for the two different correlative imaging methods as illustrated in Figures 1 and 2. The dotted areas (- - -) show the common preparation steps in processing the samples for combined confocal laser scanning- and electron microscopy. The solid line box () illustrates the point were image analysis becomes the main activity in combining the structural and fluorescent information. Briefly, digital images are calibrated using magnification calibration standards, processed for colour adjustment and figure assembly by using the replace colour and duplicate layer/merge options for comparative reasons. Note, the

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definition “correlative sample preparation & imaging platform”, means that the application of two different electron microscopy imaging methods (i.e., scanning- and transmission electron microscopy), also result in additional and different information as surface and internal structural data are collected, respectively.

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