Name: specimen preparation, imaging, and analysis protocols for knife-edge scanning microscopy
Uploaded: 2011-12-09T13:00:00
Duration: 10 min 25 s
Description: Watch this Scientific Journal Video about Specimen Preparation, Imaging, and Analysis Protocols for Knife-edge Scanning Microscopy at JoVE.com

This project was funded by NIH/NINDS (#1R01-NS54252), NSF (#0905041,#0079874), the Texas Advanced Technology Program (#ATP-000512-0146-2001, #ATP-000512-0261-2001), Texas A&amp;M Research Foundation, Department of Computer Science and Engineering at Texas A&amp;M University, and 3Scan. We would like to thank Bernard Mesa (Micro Star Technologies) for technical consultation and support for KESM instrumentation. Major parts of the design and implementation of the KESM was done by Bruce H. McCormick, who died in 2007.

1. Specimen preparation: Golgi-Cox
<ol>
	<li>This protocol closely follows the protocol of Mayerich et al.7.</li>
	<li>The C57BL/6J mouse is deeply anesthetized using isoflurane inhalant anesthesia and then decapitated.</li>
	<li>The brain is removed and placed into a Golgi-Cox fixation solution (1% potassium chromate, 1% potassium dichromate, and 1% mercuric chloride in deionized water).</li>
	<li>The brain is left in the Golgi-Cox solution in the dark, at room temperature, for 10 to 16 weeks.</li>
	<li>The brains are rinsed in deionized water overnight in the dark.</li>
	<li>The rinsed brain is immersed in 5% ammonium hydroxide solution in deionized water for 7 to 10 days in the dark at room temperature. This lengthy preparation time was to ensure infiltration of the entire brain, so that the black precipitate had a chance to completely form in the tissue.</li>
	<li>The brain is rinsed again in deionized water at room temperature for 4 hours and then dehydrated through a graded series of ethyl alcohols: 50% and 70% in the refrigerator (4 &deg;C), and 85%, 95% (3 changes), and 100% (4 to 5 changes) in room temperature. The brain is left in each solution for 24 hours.</li>
	<li>The dehydrated brain is then put in acetone (3 to 4 changes, each for one day), followed by an araldite-acetone mixture with araldite-to-acetone ratio of 1:2, 1:1, 2:1, and finally in 100% araldite (3 changes, each time overnight), in the refrigerator (4 &deg;C).</li>
	<li>Finally, the treated brain is embedded in 100% araldite and heated to 60 &deg;C for 3 days (cf. embedding protocol in Abbott and Sotelo2). 
	If specimens are embedded in LR White then the specimens are transferred from the last 100% ethanol solution through three changes of LR White solution that are each kept in the refrigerator overnight, which is a standard procedure prior to polymerization. Three changes are done to ensure full infiltration. The specimen is then transferred to fresh LR White that is polymerized in a closed container at 60 &deg;C for 24 hours, which is the required amount of time and temperature for proper polymerization.</li>
	<li>Fig. 1 shows a Golgi-Cox stained brain embedded in Araldite.</li>
	<li>The cured specimen block is then mounted on the metal specimen ring using epoxy, and the sides of the block trimmed as necessary (see Fig. 2).</li>
</ol>
2. Specimen preparation: Nissl
<ol>
	<li>This protocol closely follows the protocol of Mayerich et al.7</li>
	<li>The mouse is deeply anaesthetized using ketamine and xylazine injected intraperitoneally and then perfused transcardially using 50 mL of room temperature phosphate-buffered saline (pH 7.4), followed by 250 mL of cold 4% phosphate-buffered paraformaldehyde (pH 7.4).</li>
	<li>The mouse is then perfused with 100 mL of a solution of 0.1% thionin dye in deionized water, and the body placed in the refrigerator (4 &deg;C) for 24 hours.</li>
	<li>After 24 hours, the brain is removed from the calvaria and placed in a fresh solution of 0.1% thionin and left at 4 &deg;C for 7 days. The lengthy duration of this procedure was to ensure full infiltration of thionin into the en bloc preparation.</li>
	<li>The brain is then destained and dehydrated through a graded series of ethanols starting with 50% ethanol and water and increasing to 100% ethanol over a time period of 6 weeks. The ethanol changes from 50% to 70% are stored in the refrigerator (4 &deg;C) and all changes of ethanol above 70% are stored at room temperature.</li>
	<li>After three changes of acetone (2-4 days in each solution at room temperature), the brain is then embedded in araldite plastic following the protocol in 1.8-1.9 above. If LR white is the embedding medium then the protocol in 1.10 above is followed.</li>
</ol>
3. Specimen preparation: India ink
<ol>
	<li>The mouse is deeply anaesthetized using ketamine and xylazine (1.7 mg ketamine / 0.26 mg xylazine per 20 grams body weight) injected intraperitoneally and then perfused transcardially using 50 mL of room temperature phosphate-buffered saline (pH 7.4), followed by 250 mL of room temperature 10% neutral buffered formalin (pH 7.4). and finally with 3.0 cc of undiluted India ink.</li>
	<li>Whole body perfusion with saline and fixative is necessary to clear the blood from the cardiovascular system and to fix the tissues. Perfusion with India ink is necessary to completely fill the vasculature of the cardiovascular system.</li>
	<li>The resulting brain is then dehydrated through a series of graded ethyl alcohols (25%-100%) and then embedded in araldite plastic following the protocol in 1.8-1.9 above. If LR white is the embedding medium then the protocol in 1.10 above is followed.</li>
</ol>
4. Specimen preparation: Generic species, generic organs
<ol>
	<li>For the generic case, fixation can be done with either 10% neutral buffered formalin or 4% paraformaldehyde.</li>
	<li>For small tissue volumes, diffusion instead of perfusion is recommended for fixation and staining. In the case of small organisms or organs, staining also can be done during the dehydration process.</li>
	<li>The embedding protocol is the same as above, although the times for solution infiltration can be reduced for organs or organisms that are much smaller than whole mouse brain.</li>
</ol>
5. KESM setup and imaging
<ol>
	<li>Turn off pump, turn on stage, turn on camera, turn on illuminator.</li>
	<li>Raise knife and objective.</li>
	<li>Install specimen ring.</li>
	<li>Measure specimen dimension and create a configuration file for the KESM Stage Controller2 application.</li>
	<li>Start KESM Stage Controller2 and initialize stage.</li>
	<li>Lower knife/objective, turn on pump.</li>
	<li>Focus objective and camera, by turning the focusing knob on the optical train and adjusting the camera's field of view relative to the knife edge.</li>
	<li>Press on [Go] to initiate imaging.</li>
	<li>See Figs. 3-8. See 9, 7 for technical details.</li>
</ol>
6. Image processing and data preparation
<ol>
	<li>Copy the KESM Stack Processor executable into the data folder under the configuration file folder (00000, 00001, etc.).</li>
	<li>Run the KESM Stack Processor to remove lighting artifact and normalize the background intensity in all images. Repeat for all data subfolders.</li>
	<li>Prepare multiscale tiles from the data set and upload and set up the files in the KESM Brain Atlas web server.</li>
	<li>See Fig. 9.</li>
</ol>
7. Data visualization and analysis
<ol>
	<li>Visualization using the KESM Brain Atlas. Go to <a href="http://kesm.cs.tamu.edu">http://kesm.cs.tamu.edu</a> and select the appropriate atlas.
	<ol>
		<li>Navigate as in Google maps.</li>
		<li>Zoom in and out as in Google maps.</li>
		<li>To go to a different depth, select how deep to go at each step from the pull-down menu, and then click on either [+] or [-] to increase or decrease z depth.</li>
		<li>Adjust the overlay number by using the pull-down menu.</li>
		<li>Adjust the overlay interval by using the pull-down menu.</li>
		<li>See Fig. 11.</li>
	</ol>
	</li>
	<li>Visualization using MeVisLab.
	<ol>
		<li>Launch MeVisLab.</li>
		<li>Create a new project.</li>
		<li>In the following, new modules can be creating by typing in the module name in the command box.</li>
		<li>Create module [Compose3Dfrom2DFiles], and go to the desired folder. Enter file string and click in [GetFileList]. Make sure the selected images can fit in memory. Click [Create3D] to create volume.</li>
		<li>Create module [SetWorldMatrix], and set the "Scale" under "Elementary Transforms" x, y, z to the appropriate voxel size (0.6, 0.7, 1.0 for a 10X 0.45NA objective). Link [Compose3Dfrom2DFiles] to [SetWorldMatrix].</li>
		<li>Set Matrix and Transforms under "Matrix Composition" to "Ignore", "Ignore", "Forward".</li>
		<li>Create module [Arithmetic1] and set the "Function" to "Invert(Max-Img)". 
		Link [SetWorldMatrix] to [Arithmetic1].</li>
		<li>Create module [View3D] and connect [Arithmetic1].</li>
		<li>In View3D, while right mouse button is pressed, move the mouse around to adjust the threshold. You can crop regions, change orientation (move mouse while left mouse button is pressed), change illumination (volume rendering or MIP), turn on/off background, etc.</li>
		<li>See Fig. 10.</li>
	</ol>
	</li>
</ol>
8. Representative Results:
Here, we present whole-brain data and details. Figs. 12-15 show whole-brain India Ink, Golgi, and Nissl data sets 1, 4, 3.
<img alt="Figure 1" src="/files/ftp_upload/3248/3248fig1.jpg" /> 
Figure 1. Fixed, stained, and embedded mouse brain
<img alt="Figure 2" src="/files/ftp_upload/3248/3248fig2.jpg" /> 
Figure 2. Embedded mouse brain specimen mounted on the specimen ring.
<img alt="Figure 3" src="/files/ftp_upload/3248/3248fig3.jpg" /> 
Figure 3. The Knife Edge Scanning Microscope (KESM). A photo of the KESM is shown with its major components marked: (1) high-speed line-scan camera, (2) microscope objective, (3) diamond knife assembly and light collimator, (4) specimen tank (for water immersion imaging), (5) three-axis precision air-bearing stage, (6) white-light microscope illuminator, (7) water pump (in the back) for the removal of sectioned tissue, (8) PC server for stage control and image acquisition, (9) granite base, and (10) granite bridge.
<img alt="Figure 4" src="/files/ftp_upload/3248/3248fig4.jpg" /> 
Figure 4. Imaging principles of the KESM. The principal of operation of KESM is illustrated. The objective and the knife is held in place, while the specimen affixed on the positioning stage moves (arrow with solid line) at the resolution of 20 nm and travel speed of 1-5, and gets scraped against the diamond knife (5 mm wide for 10X objective), generating a thin section flowing over the knife (arrow with solid line). Line-scan imaging is done near the very tip of the knife.
<img alt="Figure 5" src="/files/ftp_upload/3248/3248fig5.jpg" /> 
Figure 5. KESM camera and objective focusing controls.
<img alt="Figure 6" src="/files/ftp_upload/3248/3248fig6.jpg" /> 
Figure 6. KESM knife assembly and controls.
<img alt="Figure 7" src="/files/ftp_upload/3248/3248fig7.jpg" /> 
Figure 7. Initial focusing through the observation port.
<img alt="Figure 8" src="/files/ftp_upload/3248/3248fig8.jpg" /> 
Figure 8. KESM Stage Controller 2 application (screenshot).
<img alt="Figure 9" src="/files/ftp_upload/3248/3248fig9.jpg" /> 
Figure 9. KESM stack processor application (screenshot).
<img alt="Figure 10" src="/files/ftp_upload/3248/3248fig10.jpg" /> 
Figure 10. MeVisLab application (screenshot).
<img alt="Figure 11" src="/files/ftp_upload/3248/3248fig11.jpg" /> 
Figure 11. KESM Brain Atlas: Web-interface (screenshot).
<img alt="Figure 12" src="/files/ftp_upload/3248/3248fig12.jpg" /> 
Figure 12. KESM whole-brain India ink data. Volume visualizations of KESM data stacks are shown for the vascular data set. (a) Close-up of the vascular data. Width ~100 um. (b-d) Three standard views of the whole mouse brain vasculature (subsampled from high-resolution data). Width ~10mm.
<img alt="Figure 13" src="/files/ftp_upload/3248/3248fig13.jpg" /> 
Figure 13. KESM whole-brain mouse Golgi data.
<img alt="Figure 14" src="/files/ftp_upload/3248/3248fig14.jpg" /> 
Figure 14. KESM Golgi data details.
<img alt="Figure 15" src="/files/ftp_upload/3248/3248fig15.jpg" /> 
Figure 15. KESM whole-brain Nissl data. (a) Close-up of the Nissl data. ~300 um3. (b-d) Three standard views of the whole mouse brain Nissl data (subsampled from high-resolution data). Cross section shown for a clear view of the internal structures.

<ol>
	<li>Abbott, L. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-throughput+imaging+of+whole+small+animal+brains+with+the+knife-edge+scanning+microscope.">High-throughput imaging of whole small animal brains with the knife-edge scanning microscope.</a> Neuroscience Meeting Planner. Program No. 504.2, (2008).</li><li>Abbott, L. C., Sotelo, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrastructural+analysis+of+catecholaminergic+innervation+in+weaver+and+normal+mouse+cerebellar+cortices.">Ultrastructural analysis of catecholaminergic innervation in weaver and normal mouse cerebellar cortices.</a> Journal of Comparative Neurology. 426, 316-329 (2000).</li><li>Choe, Y., Abbott, L. C., Miller, D. E., Han, D., Yang, H. -. F., Chung, J. R., Sung, C., Mayerich, D., Kwon, J., Micheva, K., Smith, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiscale+imaging,+analysis,+and+integration+of+mouse+brain+networks.">Multiscale imaging, analysis, and integration of mouse brain networks.</a> Neuroscience Meeting Planner. Program No. 516.3, (2010).</li><li>Choe, Y., Han, D., Huang, P. -. S., Keyser, J., Kwon, J., Mayerich, D., Abbott, L. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Complete+submicrometer+scans+of+mouse+brain+microstructure:+Neurons+and+vasculatures.">Complete submicrometer scans of mouse brain microstructure: Neurons and vasculatures.</a> Neuroscience Meeting Planner. Program No. 389.10, (2009).</li><li>Choe, Y., Abbott, L. C., Han, D., Huang, P. S., Keyser, J., Kwon, J., Mayerich, D., Melek, Z., McCormick, B. H., Ravi Rao, A., Cecchi, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Knife-edge+scanning+microscopy:+High-throughput+imaging+and+analysis+of+massive+volumes+of+biological+microstructures.">Knife-edge scanning microscopy: High-throughput imaging and analysis of massive volumes of biological microstructures.</a> High-Throughput Image Reconstruction and Analysis: Intelligent Microscopy Applications. , 11-37 (2008).</li><li>Choe, Y., Abbott, L. C., Ponte, G., Keyser, J., Kwon, J., Mayerich, D., Miller, D., Han, D., Grimaldi, A. M., Fiorito, F., Edelman, D. B., McKinstry, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Charting+out+the+octopus+connectome+at+submicron+resolution+using+the+knife-edge+scanning+microscope.">Charting out the octopus connectome at submicron resolution using the knife-edge scanning microscope.</a> BMC Neuroscience. 11, 136-136 (2010).</li><li>Mayerich, D., Abbott, L. C., McCormick, B. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Knife-edge+scanning+microscopy+for+imaging+and+reconstruction+of+three-dimensional+anatomical+structures+of+the+mouse+brain.">Knife-edge scanning microscopy for imaging and reconstruction of three-dimensional anatomical structures of the mouse brain.</a> Journal of Microscopy. 231, 134-143 (2008).</li><li>Mayerich, D., Kwon, J., Sung, C., Abbott, L. C., Keyser, J., Choe, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+macro-scale+Transmission+Imaging+of+Microvascular+Networks+Using+KESM.">Fast macro-scale Transmission Imaging of Microvascular Networks Using KESM.</a> Biomedical Optics Express. 2, 2888-2896 (2008).</li><li>McCormick, B. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+Brain+Tissue+Scanner.">Development of the Brain Tissue Scanner.</a> Technical Report. , (2002).</li><li>Sporns, O., Tononi, G., K&#246;tter, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+human+connectome:+A+structural+description+of+the+human+brain.">The human connectome: A structural description of the human brain.</a> PLoS Computational Biology. 1, 42-42 (2005).</li></ol>

Todd Huffman is with 3Scan, a commercial company that manufactures and markets the KESM Technology (US patent #6,744,572).

The KESM allows for a submicrometer-level survey of large volumes of biological specimen (~1 cm3). This kind of volume is enough to hold whole small animal organs, such as brains, lungs, hearts, kidneys, etc. Scanned images from such organs can provide unprecedented quantitative information about the structural organization of these organs, and enable computational modeling of various functional aspects of these organs, including circuit and network dynamics, electrical properties, fluid and air flow dynamics, and muscular dynamics. 
The specimen preparation protocol and post-analysis protocol detailed in this article are expected to help outside research groups to have easier access to the KESM and the resulting data. The KESM operation protocol will help these external researchers to appreciate the benefits and limitations of this unique imaging modality.

A correction was made to <a href="http://www.jove.com/video/3248">Specimen Preparation, Imaging, and Analysis Protocols for Knife-edge Scanning Microscopy</a>. Important modifications were made to the Nissl staining protocol.

A correction was made to <a href="http://http://www.jove.com/video/3248">Specimen Preparation, Imaging, and Analysis Protocols for Knife-edge Scanning Microscopy</a>. Important modifications were made to the Nissl staining protocol.

Erratum: Specimen Preparation, Imaging, and Analysis Protocols for Knife-edge Scanning Microscopy

specimen preparation, imaging, and analysis protocols for knife-edge scanning microscopy

Major advances in high-throughput, high-resolution, 3D microscopy techniques have enabled the acquisition of large volumes of neuroanatomical data at submicrometer resolution. One of the first such instruments producing whole-brain-scale data is the Knife-Edge Scanning Microscope (KESM)7, 5, 9, developed and hosted in the authors' lab. KESM has been used to section and image whole mouse brains at submicrometer resolution, revealing the intricate details of the neuronal networks (Golgi)1, 4, 8, vascular networks (India ink)1, 4, and cell body distribution (Nissl)3. The use of KESM is not restricted to the mouse nor the brain. We have successfully imaged the octopus brain6, mouse lung, and rat brain. We are currently working on whole zebra fish embryos. Data like these can greatly contribute to connectomics research10; to microcirculation and hemodynamic research; and to stereology research by providing an exact ground-truth. 
In this article, we will describe the pipeline, including specimen preparation (fixing, staining, and embedding), KESM configuration and setup, sectioning and imaging with the KESM, image processing, data preparation, and data visualization and analysis. The emphasis will be on specimen preparation and visualization/analysis of obtained KESM data. We expect the detailed protocol presented in this article to help broaden the access to KESM and increase its utilization.

Watch this Scientific Journal Video about Specimen Preparation, Imaging, and Analysis Protocols for Knife-edge Scanning Microscopy at JoVE.com

Specimen Preparation, Imaging, and Analysis Protocols for Knife-edge Scanning Microscopy

The full process from brain specimen preparation to serial sectioning imaging using the Knife-Edge Scanning Microscope, to data visualization and analysis is described. This technique is currently used to acquire mouse brain data, but it is applicable to other organs, other species.

Major advances in high-throughput, high-resolution, 3D microscopy techniques have enabled the acquisition of large volumes of neuroanatomical data at ...

specimen-preparation-imaging-analysis-protocols-for-knife-edge

Research

JoVE Journal

Bioengineering

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

Bacterial Immobilization for Imaging by Atomic Force Microscopy

AFM is a high-resolution (nm scale) imaging tool that mechanically probes a surface. It has the ability to image cells and biomolecules, in a liquid environment, without the need to chemically treat the sample. In order to accomplish this goal, the sample must sufficiently adhere to the mounting surface to prevent removal by forces exerted by the scanning AFM cantilever tip. In many instances, successful imaging depends on immobilization of the sample to the mounting surface. Optimally, immobilization should be minimally invasive to the sample such that metabolic processes and functional attributes are not compromised. By coating freshly cleaved mica surfaces with porcine (pig) gelatin, negatively charged bacteria can be immobilized on the surface and imaged in liquid by AFM. Immobilization of bacterial cells on gelatin-coated mica is most likely due to electrostatic interaction between the negatively charged bacteria and the positively charged gelatin. Several factors can interfere with bacterial immobilization, including chemical constituents of the liquid in which the bacteria are suspended, the incubation time of the bacteria on the gelatin coated mica, surface characteristics of the bacterial strain and the medium in which the bacteria are imaged. Overall, the use of gelatin-coated mica is found to be generally applicable for imaging microbial cells.

Live Gram-negative and Gram-positive bacteria can be immobilized on gelatin-coated mica and imaged in liquid using Atomic Force Microscopy (AFM).

AFM is a high-resolution (nm scale) imaging tool that mechanically probes a surface.  It has the ability to image cells and biomolecules, in a liquid ...

Video-rate Scanning Confocal Microscopy and Microendoscopy

Confocal microscopy has become an invaluable tool in biology and the biomedical sciences, enabling rapid, high-sensitivity, and high-resolution optical sectioning of complex systems. Confocal microscopy is routinely used, for example, to study specific cellular targets1, monitor dynamics in living cells2-4, and visualize the three dimensional evolution of entire organisms5,6. Extensions of confocal imaging systems, such as confocal microendoscopes, allow for high-resolution imaging in vivo7 and are currently being applied to disease imaging and diagnosis in clinical settings8,9.
Confocal microscopy provides three-dimensional resolution by creating so-called "optical sections" using straightforward geometrical optics. In a standard wide-field microscope, fluorescence generated from a sample is collected by an objective lens and relayed directly to a detector. While acceptable for imaging thin samples, thick samples become blurred by fluorescence generated above and below the objective focal plane. In contrast, confocal microscopy enables virtual, optical sectioning of samples, rejecting out-of-focus light to build high resolution three-dimensional representations of samples.
Confocal microscopes achieve this feat by using a confocal aperture in the detection beam path. The fluorescence collected from a sample by the objective is relayed back through the scanning mirrors and through the primary dichroic mirror, a mirror carefully selected to reflect shorter wavelengths such as the laser excitation beam while passing the longer, Stokes-shifted fluorescence emission. This long-wavelength fluorescence signal is then passed to a pair of lenses on either side of a pinhole that is positioned at a plane exactly conjugate with the focal plane of the objective lens. Photons collected from the focal volume of the object are collimated by the objective lens and are focused by the confocal lenses through the pinhole. Fluorescence generated above or below the focal plane will therefore not be collimated properly, and will not pass through the confocal pinhole1, creating an optical section in which only light from the microscope focus is visible. (Fig 1). Thus the pinhole effectively acts as a virtual aperture in the focal plane, confining the detected emission to only one limited spatial location.
Modern commercial confocal microscopes offer users fully automated operation, making formerly complex imaging procedures relatively straightforward and accessible. Despite the flexibility and power of these systems, commercial confocal microscopes are not well suited for all confocal imaging tasks, such as many in vivo imaging applications. Without the ability to create customized imaging systems to meet their needs, important experiments can remain out of reach to many scientists.
In this article, we provide a step-by-step method for the complete construction of a custom, video-rate confocal imaging system from basic components. The upright microscope will be constructed using a resonant galvanometric mirror to provide the fast scanning axis, while a standard speed resonant galvanometric mirror will scan the slow axis. To create a precise scanned beam in the objective lens focus, these mirrors will be positioned at the so-called telecentric planes using four relay lenses. Confocal detection will be accomplished using a standard, off-the-shelf photomultiplier tube (PMT), and the images will be captured and displayed using a Matrox framegrabber card and the included software.

The complete construction of a custom, real-time confocal scanning imaging system is described. This system, which can be readily used for video-rate microscopy and microendoscopy, allows for an array of imaging geometries and applications not accessible using standard commercial confocal systems, at a fraction of the cost.

Confocal microscopy has become an invaluable tool in biology and the biomedical sciences, enabling rapid, high-sensitivity, and high-resolution optical ...

Creating Adhesive and Soluble Gradients for Imaging Cell Migration with Fluorescence Microscopy

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as growth factors. Here, we outline a method to create opposing gradients of adhesive and soluble cues in a microfluidic chamber, which is compatible with live cell imaging. A copolymer of poly-L-lysine and polyethylene glycol (PLL-PEG) is employed to passivate glass coverslips and prevent non-specific adsorption of biomolecules and cells. Next, microcontact printing or dip pen lithography are used to create tracks of streptavidin on the passivated surfaces to serve as anchoring points for the biotinylated peptide arginine-glycine-aspartic acid (RGD) as the adhesive cue. A microfluidic device is placed onto the modified surface and used to create the gradient of adhesive cues (100% RGD to 0% RGD) on the streptavidin tracks. Finally, the same microfluidic device is used to create a gradient of a chemoattractant such as fetal bovine serum (FBS), as the soluble cue in the opposite direction of the gradient of adhesive cues.

A method for the assembly of adhesive and soluble gradients in a microscopy chamber for live cell migration studies is described. The engineered environment combines antifouling surfaces and adhesive tracks with solution gradients and therefore allows one to determine the relative importance of guidance cues.

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as ...

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

Preparation of Mica Supported Lipid Bilayers for High Resolution Optical Microscopy Imaging

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction with other compounds, such as drugs or peptides. However SLB characteristics differ depending on the support used. 
Commonly used techniques for SLB imaging and measurements are single molecule fluorescence microscopy, FCS and atomic force microscopy (AFM). Because most optical imaging studies are carried out on a glass support, while AFM requires an extremely flat surface (generally mica), results from these techniques cannot be compared directly, since the charge and smoothness properties of these materials strongly influence diffusion. Unfortunately, the high level of manual dexterity required for the cutting and gluing thin slices of mica to the glass slide presents a hurdle to routine use of mica for SLB preparation. Although this would be the method of choice, such prepared mica surfaces often end up being uneven (wavy) and difficult to image, especially with small working distance, high numerical aperture lenses. Here we present a simple and reproducible method for preparing thin, flat mica surfaces for lipid vesicle deposition and SLB preparation. Additionally, our custom made chamber requires only very small volumes of vesicles for SLB formation. The overall procedure results in the efficient, simple and inexpensive production of high quality lipid bilayer surfaces that are directly comparable to those used in AFM studies.

We present a method of preparing mica supported lipid bilayers for high resolution microscopy. Mica is transparent and flat on an atomic scale, but rarely used in imaging because of handling difficulties; our preparation results in even deposition of the mica sheet, and reduces the material used in bilayer preparation.

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction ...

Hybrid &#181;CT-FMT imaging and image analysis

Fluorescence-mediated tomography (FMT) enables longitudinal and quantitative determination of the fluorescence distribution in vivo and can be used to assess the biodistribution of novel probes and to assess disease progression using established molecular probes or reporter genes. The combination with an anatomical modality, e.g., micro computed tomography (&#181;CT), is beneficial for image analysis and for fluorescence reconstruction. We describe a protocol for multimodal &#181;CT-FMT imaging including the image processing steps necessary to extract quantitative measurements. After preparing the mice and performing the imaging, the multimodal data sets are registered. Subsequently, an improved fluorescence reconstruction is performed, which takes into account the shape of the mouse. For quantitative analysis, organ segmentations are generated based on the anatomical data using our interactive segmentation tool. Finally, the biodistribution curves are generated using a batch-processing feature. We show the applicability of the method by assessing the biodistribution of a well-known probe that binds to bones and joints.

Hybrid  &amp;#181;CT-FMT imaging and image analysis

We describe a protocol for hybrid imaging, combining fluorescence-mediated tomography (FMT) with micro computed tomography (&#181;CT). After fusion and reconstruction, we perform interactive organ segmentation to extract quantitative measurements of the fluorescence distribution.

Fluorescence-mediated tomography (FMT) enables longitudinal and quantitative determination of the fluorescence distribution in vivo and can be used to ...

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

Correlative Light Electron Microscopy (CLEM) for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing virus-infected or transfected cells via EM are the low efficiencies of infection or transfection, hindering the examination of these cells. In order to overcome this difficulty, light microscopy (LM) can be performed first to allocate the subpopulation of infected or transfected cells. Thus, taking advantage of the use of fluorescent proteins (FPs) fused to viral proteins, LM is used here to record the positions of the "positive-transfected" cells, expressing a FP and growing on a support with an alphanumeric pattern. Subsequently, cells are further processed for EM via high pressure freezing (HPF), freeze substitution (FS) and resin embedding. The ultra-rapid freezing step ensures excellent membrane preservation of the selected cells that can then be analyzed at the ultrastructural level by transmission electron microscopy (TEM). Here, a step-by-step correlative light electron microscopy (CLEM) workflow is provided, describing sample preparation, imaging and correlation in detail. The experimental design can be also applied to address many cell biology questions.

Correlative Light Electron Microscopy CLEM for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

A correlative light electron microscopy (CLEM) method is applied to image virus-induced intracellular structures via electron microscopy (EM) in cells that are previously selected by light microscopy (LM). LM and EM are combined as a hybrid imaging approach to achieve an integrated view of virus-host interactions.

Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing ...

Visualizing Adhesion Formation in Cells by Means of Advanced Spinning Disk-Total Internal Reflection Fluorescence Microscopy

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to visualize these highly dynamic events, two complementary light microscopy techniques that allow fast imaging of live samples were combined: spinning disk microscopy (SD) for fast and high-resolution volume recording and total internal reflection fluorescence (TIRF) microscopy for precise localization and visualization of the plasma membrane. A comprehensive and complete imaging protocol will be shown for guiding through sample preparation, microscope calibration, image formation and acquisition, resulting in multi-color SD-TIRF live imaging series with high spatio-temporal resolution. All necessary image post-processing steps to generate multi-dimensional live imaging datasets, i.e. registration and combination of the individual channels, are provided in a self-written macro for the open source software ImageJ. The imaging of fluorescent proteins during initiation and maturation of adhesion complexes, as well as the formation of the actin cytoskeletal network, was used as a proof of principle for this novel approach. The combination of high resolution 3D microscopy and TIRF provided a detailed description of these complex processes within the cellular environment and, at the same time, precise localization of the membrane-associated molecules detected with a high signal-to-background ratio.

An advanced microscope that permit fast and high-resolution imaging of both, the isolated plasma membrane and the surrounding intracellular volume, will be presented. The integration of spinning disk and total internal reflection fluorescence microscopy in one setup allows live imaging experiments at high acquisition rates up to 3.5 s per image stack.

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to ...