The authors acknowledge Stefan Vogt for his assistance in the fitting of the representative data shown in this paper, and helpful discussions. The authors also acknowledge Chris Jacobsen for his support to Q. J. 
Use of the Advanced Photon Source, beamlines 2-ID-E and 8-BM-B, at Argonne National Laboratory was supported by the U. S. Department of Energy, Office of Science, Office of Basic Energy Sciences, under Contract No. DE-AC02-06CH11357.

1. Preparation of Instruments, Substrates, Culture Media and Dishes
<ol>
	<li>Handling of Silicon Nitride (Si3N4) windows.
	<ol>
		<li>Open the capsule slightly by squeezing one end containing the window in such a way as to not squeeze the window itself, while rotating the other end of the capsule with the other hand.</li>
		<li>Check a pair of reverse, fine-tipped tweezers under stereomicroscope to make sure there are no adhesive, gaps or bends at the tip. Otherwise, the windows may be broken by the tweezers or fly off of them during the subsequent steps.</li>
		<li>Remove the window from the capsule by carefully grasping the frame of the window with tweezers, while gently squeezing the capsule near the open end applying pressure in a direction so as to make the capsule wider where the edges of the window need to come out.</li>
	</ol>
	</li>
	<li>Window pre-wash (optional).
	<ol>
		<li>If desired, pre-wash windows by gently dipping them into distilled water for 5 sec, followed by 2 min wash in 70% ethanol and 2 min wash in 100% ethanol.</li>
	</ol>
	</li>
	<li>Affixing and sterilizing windows on the culture dish.
	<ol>
		<li>To affix the windows, set the window with flat side up at the bottom of the culture dish, place a piece of tape on a clean glass slide. Cut about a quarter inch strip off down the long side of the tape. Take a slice of tape off the short side (at &#8539;&#8221; by &#190;&#8221;) with a clean razor blade.</li>
		<li>Use tweezers to apply the slice of tape to the edge of the grid or window. Use enough to securely adhere it without covering the window opening itself. Manipulating the adhered tape with tweezers, set the window down onto the bottom of the culture dish. Use the tip of the tweezers to adhere tape to the bottom of the culture dish firmly.</li>
		<li>Apply another tape strip to the other edge and adhere this to the bottom firmly with the tip of the tweezers.</li>
		<li>Once all windows are taped, sterilize windows in culture dishes with ultraviolet (UV) radiation. Accomplish this by placing the dish under the UV lamp in a laminar flow hood for about 1 hr.</li>
		<li>(Optional) If windows are to be pre-coated with poly-lysine, do so following window sterilization. Coat the window with 10 &#956;l of sterile 0.01% poly-L-lysine solution for 1 hr in a 37 &#176;C incubator, and then rinse the remaining solution off with the culture media.</li>
	</ol>
	</li>
	<li>Prepare buffers for final wash before XFM.
	<ol>
		<li>Prepare either a Tris-glucose or piperazine-N,N&#8217;-bis(ethanesulfonic acid) (PIPES) -sucrose buffer free of any trace metals with osmolality and pH equivalent to Dulbecco&#39;s phosphate buffer saline (D-PBS) without calcium and magnesium. Use one of these buffers to rinse cells after cells are chemically fixed.
		<ol>
			<li>To make the Tris-glucose solution (10 mM Tris, 260 mM glucose, 9 mM acetic acid,) add 1.4 g of glucose, 36 mg of Tris base and 16 &#181;l of acetic acid to 20 ml of ultrapure water. Then, adjust final volume to 30 ml with ultrapure water. No pH adjustment is needed. Store at RT up to one week.</li>
			<li>To make the PIPES-sucrose (20 mM PIPES, 200 mM sucrose) add 0.18 g of PIPES and 2.0 g of sucrose to 20 ml of ultrapure water and adjust the pH to 7.4 with small amounts of concentrated acetic acid. Then, adjust final volume to 30 ml with ultrapure water. Store at RT up to one week. 
			NOTE: The reason to avoid using PBS in the final rinse is that the concentrations of phosphate, chloride and potassium in PBS are so high that it can interfere with XFM if it is not completely removed before air drying.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Prepare buffers for cell plating.
	<ol>
		<li>Prepare or take out all media, such as RPMI medium 1640, 1x trypsin-EDTA solution, D-PBS, and any other reagents as needed for the particular adherent cell line as would normally be done for passaging the adherent cells of interest, and warm them to 37 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Plating of Cells
<ol>
	<li>To the sterilized culture dish containing the windows, in the laminar flow hood, gently add media (10 ml for a 100 mm culture dish), along the side of the culture dish with it tilted at an angle. Avoid creating unnecessary surface tension or dropping media directly onto surface of silicon nitride windows. As media is added, slowly relieve the tilt angle to coat the grid and window with media.</li>
	<li>Prepare the adherent cell line of interest appropriately, trypsinizing or scraping the cells off of the plates as usual for passaging the cell line. Perform any cell counting, or other preparation necessary before applying the cells to the fresh plate.</li>
	<li>Add cells to the culture dish with windows at a cell density that is needed to typically reach 50%-70% confluence within 24-72 hr, and incubate at 37 &#176;C in a humidified incubator with 5% carbon dioxide.</li>
	<li>Observe the cells&#8217; growth occasionally, using an inverted light microscope, until the desired cell density (usually 50%-70% confluence) is reached.</li>
</ol>
3. Fixation of Cells
<ol>
	<li>Prepare fresh 4% paraformaldehyde in D-PBS by diluting a purchased stock (such as 25% paraformaldehyde) and adjusting the pH to 7.4 with acetic acid (to avoid adding excess sodium to the sample).</li>
	<li>When cells have been grown as desired, and any vital stains (such as Hoescht) have been applied and imaged, remove the media by gentle aspiration, tilting the dish at a 45&#176; angle with one hand, and aspirating by hand pipette with the other. 
	NOTE: Instead of pipetting spent media, an alternative is to pick the window up, dip the window into microcentrifuge tubes with D-PBS twice and then place it into a new culture vessel with fixative solutions for 20 min followed by 2 times of PBS wash to remove the residual fixatives. If this is chosen, go directly to step 3.5.</li>
	<li>Rinse the dish gently with D-PBS and replace the removed liquid immediately by gently adding the fresh 4% PFA/PBS while holding the dish at a 45&#176; angle, pipetting towards the side of the dish, and slowly relaxing the tilt angle to allow the fixative solution to cover the windows. Keep the cells covered with the fixative for 20 min at RT.</li>
	<li>Remove the liquid by gentle aspiration, again as above, tilting the dish slightly and aspirating by hand.</li>
	<li>Replace the removed liquid by gently adding some of the PIPES/Sucrose solution, using the tilting and pipetting method described in Step 3.3.</li>
	<li>Repeat Steps 3.4 and 3.5.</li>
	<li>Prepare the materials for drying.
	<ol>
		<li>Gather lint-free towels such as Kimwipes, and pull one out of the box so it is handy very quickly and easily.</li>
		<li>Prepare a clean, non-level place to set the window to dry. Set out a rubber grid mat of the kind used in electron microscopy, which has ridges so the window can be propped on one end while it dries. This will keep the window from getting stuck to the drying surface as it dries, and drain any remaining buffer to the edge of the sample.</li>
	</ol>
	</li>
	<li>Carefully tweeze the tape attached to window to hold the window up out of the liquid. Retrieve windows that come off of the tape by applying a small amount of PIPES/sucrose to get them to float and then picking them up with the reverse tweezers.</li>
	<li>Carefully (without touching the window itself) and thoroughly daub any excess PIPES/sucrose from the edge of the window with a Kimwipe. Also daub the back indented area using a rounded fold of a Kimwipe. 
	NOTE: The goal is to have as little crystallization of salts or buffers on the surface of the window as possible.</li>
	<li>Place the window onto the grid mat. Make sure that the window is not lying flat on the grid mat, but propped up on an edge (using the ridges of the grid mat) and touching as little of the grid mat as possible. Let the window dry.</li>
</ol>
4. Sample Mounting and Imaging
<ol>
	<li>Once the sample has dried, verify the presence of cells on the windows using a light microscope. Perform this verification, before preparing the samples for transport to a synchrotron.</li>
	<li>Package the windows for transportation to the synchrotron beamline hosting your experiment. Gently affix the windows with tape on two sides to a glass slide, then affix the glass slide with tape to the bottom of a plastic Petri dish. Tape the Petri dish shut. 
	NOTE: Prior to this point, these steps may be carried out in any typical lab. The following steps would best be done at a synchrotron beamline.</li>
	<li>Remove the tape from the window gently with tweezers. Use nail polish, in a thin even coating along the edge of the window, to secure the windows to an aluminum holder provided by the beamline collaborators at the synchrotron.</li>
	<li>Insert the aluminum holder into a kinematic mount, and then place it into position in the X-ray microscope. Mount the sample on a bracket connecting it to motorized stages, inside the sample chamber at the beamline.</li>
	<li>After exiting the sample X-ray microscope instrument area (a hutch made of lead-filled walls), setup the scan. Using beamline-specific software, choose the appropriate area of the sample to scan, while viewing the position of the X-ray beam on the sample using a camera equipped with a video cross-hair and pre-aligned with a downstream scintillator camera.</li>
	<li>Choose the appropriate resolution for the sample, based on an estimate of the size of the smallest feature of interest. For imaging single mammalian cells (~20-40 &#181;m in size), use a resolution of 0.25-0.5 &#956;m. For imaging areas of tissues, and distinguishing cell layers from each other, use a resolution of 2-20 &#956;m.</li>
	<li>Choose the appropriate dwell time for the sample, based on an estimate of the quantity of the metal of interest present. For a quick overview scan, or if high quantities of metals are present, use fly scans with dwell times of 30-300 msec per pixel. If little material is present, or to measure metals of low relative abundance, use step scans with dwell times of 1-2 sec per pixel.</li>
	<li>Program the scans into the beamline-specific software. Acquire a raster-scan image. Work together with the beamline collaborators to analyze the data with software that will identify the characteristic energy peaks for each element.</li>
</ol>

The authors have nothing to disclose.

X-ray fluorescence imaging is useful in many fields, including geosciences, materials science, and chemical biology26-34. Advances in synchrotron X-rays, and their focusing, have produced very high-intensity beams. Focused X-ray beams sufficient to excite the endogenous metals in cells now exist, producing signal that can be measured with currently available silicon drift detector technology. And studying the chemical biology of metals in the cell is one application in particular that makes use of many of the ...

X-ray fluorescence imaging allows for both the identity and quantity of elements present in a sample to be spatially resolved. Incident X-rays, of an energy selected to be greater than the electron binding energy of the heaviest element of interest, overcome the binding energy of inner-shell electrons to the nucleus1. This creates a &#8216;hole&#8217; in the electron shell. As higher-energy electrons fall down into these holes, fluorescent X-rays are emitted whose wavelength is dependent on the energy separation of those orbitals. Since the energy spacing of the orbitals is characteristic of a given element, the X-ray fluorescence emission also has characteristic wavelengths, dependent on the element. It is this emission at a characteristic wavelength that allows the identification of the elements present. Calibration of the fluorescence intensity allows the quantitation of the elements present.
X-ray fluorescence microscopy (XFM) has become increasingly utilized, partly due to the development of very brilliant X-ray synchrotron sources, such as those at Spring-8 in Japan, the European Radiation Synchrotron Facility (ESRF) in France, and the Advanced Photon Source (APS) in the US2. These sources provide very high intensity X-ray beams. At the same time, improvements in X-ray optics, such as zone plate technology, allowed the focusing of these beams to sub-micron spots, albeit rather inefficiently3. With very high-intensity beams, even a relatively small amount of light that can be focused is sufficient to excite the endogenous metals in cells, producing signal that can be measured with currently available detector technology. Thus, studying the chemical biology of metals in the cell is one application in particular that makes use of many of the recent developments in this technique4-10.
There are many critical factors to be considered while applying XFM to investigate the elemental distribution and quantification of cultured mammalian cells or other biological samples. Firstly, the sample needs to be kept intact, both structurally and with respect to its elemental composition, in order for the measurement to be meaningful. Secondly, the sample must also be preserved in some way so that it is hardy to the radiation damage that can be caused by a focused X-ray beam. One way that a sample can meet both of these criteria at once is to be rapidly frozen into a vitreous, amorphous ice11,12. Rapid freezing is often achieved through various cryopreservation techniques such as plunge freezing or high pressure freezing13-16. It is generally accepted that cryopreservation preserves overall cellular architecture and chemical compositions in biological samples as close to native state as possible. Chemical fixation, on the other hand, due to the slow and selective penetration of fixatives into cells and tissues as well as subsequent changes in membrane permeability, may allow various cellular ions especially the diffusible ions such as Cl, Ca and K to be leached, lost or relocated, thus rendering investigation of these elements suboptimal17-19. Despite the clear advantage of cryo-fixation over chemical fixation in general, for adherent mammalian cells in particular, cryopreservation has various limitations20-23. The most obvious one is that not every research lab has easy access to cryopreservation instruments. Most current high pressure freezers or even plunge freezers are costly and owned only by a subset of cryo facilities, which may be far from where cells are incubated. The benefit of cryopreservation might be traded for the disadvantage of travel stress placed on the cells. Thus, while cryopreservation is surely the most rigorous way to preserve samples for X-ray fluorescence analysis, it is certainly not the most accessible to all researchers under all circumstances; nor is it always essential&#160;&#8212; if the metals of interest are tightly bound to fixable macromolecules, and the resolution at which the sample will be imaged is greater than the damage to the ultra-microstructure that might occur during drying. Mindful of the caveats24, chemical fixation and drying may be a suitable choice.
Other factors in a successful X-ray fluorescence imaging experiment include proper analysis. X-ray fluorescence imaging is fundamentally X-ray fluorescence emission spectroscopy combined with raster-scanning to provide spatial resolution. The X-ray fluorescence emission spectra collected contain a combination of overlapping emission peaks, background, and the elastic and inelastic scattering peaks of the incident beam. Software that enables the de-convolution of these contributions, and the fitting of the emission peaks, has been a critical development to this field25. Also, the development and commercial distribution of thin-film standards of known composition, used to calibrate fluorescence intensity relative to material quantity, has also been very important.
This protocol provides a description of the preparation of adherent cells by chemical fixation and air drying. A vital step in this process is the growth of the cells on the silicon nitride windows, which often do not adhere well, making gentle rinsing in a particular fashion key to success.

<table><tbody><tr><td>Silicon nitride windows</td><td>Silson Ltd/J B J Business Park/Northampton Rd, Northampton NN7 3DW, United Kingdom</td><td>SiRN-5.0-200-2.0-200. Order by size. Membrane size: 1.5 mm x 1.5 mm. Thickness: 200&amp;nbsp;nm. Frame size: 5&amp;nbsp;mm x 5 mm. Frame thickness: 200 &amp;micro;m.</td><td>Alternate source: SPI Supplies / Structure Probe, Inc. West Chester, PA.</td></tr><tr><td>Reverse tweezers</td><td>Electron Microscopy Sciences, P.O. Box 550, 1560 Industry Road, Hatfield, PA 19440, Tel: 215-412-8400, Toll Free: 800-523-5874, Fax: 215-412-8450</td><td>78520-5X</td><td>EMS 5X, NC &amp;ndash; Ultra Fine Tweezers</td></tr><tr><td>Rubber grid mat</td><td>Electron Microscopy Sciences, P.O. Box 550, 1560 Industry Road, Hatfield, PA 19440, Tel: 215-412-8400, Toll Free: 800-523-5874, Fax: 215-412-8450</td><td>71170</td><td>Round Grid Mat</td></tr><tr><td>Acetic acid</td><td>Sigma-Aldrich, 3050 Spruce St., St. Louis, MO 63103, Tel: 800-325-3010, Fax: 800-325-5052</td><td>338826</td><td>trace metals grade concentrated acetic acid</td></tr><tr><td>PIPES buffer</td><td>Sigma-Aldrich, 3050 Spruce St., St. Louis, MO 63103, Tel: 800-325-3010, Fax: 800-325-5052</td><td>P6757</td><td>solid PIPES buffer</td></tr><tr><td>Formaldehyde stock solution</td><td>Electron Microscopy Sciences, P.O. Box 550, 1560 Industry Road, Hatfield, PA 19440, Tel: 215-412-8400, Toll Free: 800-523-5874, Fax: 215-412-8450</td><td>RT 17113</td><td>10 x 10 ml ampules of 20% aqueous paraformaldehyde</td></tr></tbody></table>

preparing adherent cells for x-ray fluorescence imaging by chemical fixation

X-ray fluorescence imaging allows us to non-destructively measure the spatial distribution and concentration of multiple elements simultaneously over large or small sample areas. It has been applied in many areas of science, including materials science, geoscience, studying works of cultural heritage, and in chemical biology. In the case of chemical biology, for example, visualizing the metal distributions within cells allows us to study both naturally-occurring metal ions in the cells, as well as exogenously-introduced metals such as drugs and nanoparticles. Due to the fully hydrated nature of nearly all biological samples, cryo-fixation followed by imaging under cryogenic temperature represents the ideal imaging modality currently available. However, under the circumstances that such a combination is not easily accessible or practical, aldehyde based chemical fixation remains useful and sometimes inevitable. This article describes in as much detail as possible in the preparation of adherent mammalian cells by chemical fixation for X-ray fluorescent imaging.

The ability of X-ray fluorescence imaging to provide information about biological samples is contingent upon these samples being prepared in such a way that they are robust to radiation damage on the time-scale of the experiment, and yet their chemical and structural features are well preserved. In viewing the result of a sample that has been prepared as described above and imaged, it is possible to see that there is variation in the elements present &#8212; indicating that fixation preserved these aspects of the cell (<...

Watch this Scientific Journal Video about Preparing Adherent Cells for X-ray Fluorescence Imaging by Chemical Fixation at JoVE.com

Preparing Adherent Cells for X-ray Fluorescence Imaging by Chemical Fixation

Here, we present a protocol on how to determine the quantity and distribution of metals in a sample using synchrotron X-ray fluorescence. We focus on adherent cells, and describe the chemical fixation method to prepare this sample. We then describe how to mount and image the sample using synchrotron X-rays.

X-ray fluorescence imaging allows us to non-destructively measure the spatial distribution and concentration of multiple elements simultaneously over ...

preparing-adherent-cells-for-x-ray-fluorescence-imaging-chemical

Research

JoVE Journal

Chemistry

9.4K Views.  Argonne National Laboratory. Here, we present a protocol on how to determine the quantity and distribution of metals in a sample using synchrotron X-ray fluorescence. We focus on adherent cells, and describe the chemical fixation method to prepare this sample. We then describe how to mount and image the sample using synchrotron X-rays. 

Video: Preparing Adherent Cells for X-ray Fluorescence Imaging by Chemical Fixation

In Situ Detection and Single Cell Quantification of Metal Oxide Nanoparticles Using Nuclear Microprobe Analysis

Micro-analytical techniques based on chemical element imaging enable the localization and quantification of chemical composition at the cellular level. They offer new possibilities for the characterization of living systems and are particularly appropriate for detecting, localizing and quantifying the presence of metal oxide nanoparticles both in biological specimens and the environment. Indeed, these techniques all meet relevant requirements in terms of (i) sensitivity (from 1 up to 10 &#181;g.g-1 of dry mass), (ii) micrometer range spatial resolution, and (iii) multi-element detection. Given these characteristics, microbeam chemical element imaging can powerfully complement routine imaging techniques such as optical and fluorescence microscopy. This protocol describes how to perform a nuclear microprobe analysis on cultured cells (U2OS) exposed to titanium dioxide nanoparticles. Cells must grow on and be exposed directly in a specially designed sample holder used on the optical microscope and in the nuclear microprobe analysis stages. Plunge-freeze cryogenic fixation of the samples preserves both the cellular organization and the chemical element distribution. Simultaneous nuclear microprobe analysis (scanning transmission ion microscopy, Rutherford backscattering spectrometry and particle induced X-ray emission) performed on the sample provides information about the cellular density, the local distribution of the chemical elements, as well as the cellular content of nanoparticles. There is a growing need for such analytical tools within biology, especially in the emerging context of Nanotoxicology and Nanomedicine for which our comprehension of the interactions between nanoparticles and biological samples must be deepened. In particular, as nuclear microprobe analysis does not require nanoparticles to be labelled, nanoparticle abundances are quantifiable down to the individual cell level in a cell population, independently of their surface state.

In Situ Detection and Single Cell Quantification of Metal Oxide Nanoparticles Using Nuclear Microprobe Analysis

We describe a procedure for the detection of chemical elements present in situ in human cells as well as their in vitro quantification. The method is well-suited to any cell type and is particularly useful for quantitative chemical analyses in single cells following in vitro metal oxide nanoparticles exposure.

Micro-analytical techniques based on chemical element imaging enable the localization and quantification of chemical composition at the cellular level. ...

Processing Embryo, Eggshell, and Fungal Culture for Scanning Electron Microscopy

Although scanning electron microscopy (SEM) is being widely used for the ultra-structural analysis of various biological and non-biological samples, methods involved in processing different biological samples involve unique practices. All conventional practices described in the literature for processing samples still find useful applications, but subtle changes in the sample preparation can alter image quality, as well as, introduce artifacts. Hence, using a unique sample preparation technique specific to the type of tissue analyzed is required to obtain a good quality image with ultrastructural resolution. The focus of this study is to provide the optimal sample preparation protocols for imaging embryos, rigid eggshells, and fungal cultures using SEM. The following optimizations were recommended to yield good results for the three different delicate biological samples studied. Use of milder fixatives like 4% paraformaldehyde or 3% glutaraldehyde followed by dehydration with ethanol series is mandatory. Fungal mycelium on agar blocks obtained by slide cultures yields a better ultrastructural integrity compared to cultures taken directly from agar plates. Chemical drying of embryos with HMDS provides drying without introducing surface tension artifacts compared to critical point drying. HMDS prevents cracking caused by shrinkage as samples are less brittle during drying. However, for fungal culture, critical point drying provides acceptable image quality compared to chemical drying. Eggshells can be imaged with no special preparation steps except for thorough washing and air drying prior to mounting. Preparation methodologies were standardized based on acceptable image quality obtained with each trial.&#160;

Here, we present detailed processing protocols for imaging delicate tissue samples using scanning electron microscopy (SEM). Three different processing methods, namely, hexamethyl disilazana (HMDS) chemical drying, simple air drying, and critical point drying are described for preparing rigid eggshells, embryos at early developmental stages, and fungal cultures respectively.

Although scanning electron microscopy (SEM) is being widely used for the ultra-structural analysis of various biological and non-biological samples, ...

Environment

Cell Culture on Silicon Nitride Membranes and Cryopreparation for Synchrotron X-ray Fluorescence Nano-analysis

Very little is known about the distribution of metal ions at the subcellular level. However, those chemical elements have essential regulatory functions and their disturbed homeostasis is involved in various diseases. State-of-the-art synchrotron X-ray fluorescence nanoprobes provide the required sensitivity and spatial resolution to elucidate the two-dimensional (2D) and three-dimensional (3D) distribution and concentration of metals inside entire cells at the organelle level. This opens new exciting scientific fields of investigation on the role of metals in the physiopathology of the cell. The cellular preparation is a key and often complex procedure, particularly for basic analysis. Although X-ray fluorescence techniques are now widespread and various preparation methods have been used, very few studies have investigated the preservation of the elemental content of cells at best, and no stepwise detailed protocol for the cryopreparation of adherent cells for X-ray fluorescence nanoprobes has been released so far. This is a description of a protocol that provides the stepwise cellular preparation for fast cryofixation to enable synchrotron X-ray fluorescence nano-analysis of cells in a frozen hydrated state when a cryogenic environment and transfer is available. In case nano-analysis has to be performed at room temperature, an additional procedure for freeze-drying the cryofixed adherent cellular preparation is provided. The proposed protocols have been successfully used in previous works, most recently in studying the 2D and 3D intracellular distribution of an organometallic compound in breast cancer cells.

Presented here is a protocol for cell culture on silicon nitride membranes and plunge-freezing prior to X-ray fluorescence imaging with a synchrotron cryogenic X-ray nanoprobe. When only room temperature nano-analysis is provided, the frozen samples can be further freeze-dried. These are critical steps to obtain information on the intracellular elemental composition.

Very little is known about the distribution of metal ions at the subcellular level. However, those chemical elements have essential regulatory functions ...

Biochemistry

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 ...

Bioengineering

Bioorthogonal Chemical Imaging of Cell Metabolism Regulated by Aromatic Amino Acids

Essential aromatic amino acids (AAAs) are building blocks for synthesizing new biomasses in cells and sustaining normal biological functions. For example, an abundant supply of AAAs is important for cancer cells to maintain their rapid growth and division. With this, there is a rising demand for a highly specific, noninvasive imaging approach with minimal sample preparation to directly visualize how cells harness AAAs for their metabolism in situ. Here, we develop an optical imaging platform that combines deuterium oxide (D2O) probing with stimulated Raman scattering (DO-SRS) and integrates DO-SRS with two-photon excitation fluorescence (2PEF) into a single microscope to directly visualize the metabolic activities of HeLa cells under AAA regulation. Collectively, the DO-SRS platform provides high spatial resolution and specificity of newly synthesized proteins and lipids in single HeLa cell units. In addition, the 2PEF modality can detect autofluorescence signals of nicotinamide adenine dinucleotide (NADH) and Flavin in a label-free manner. The imaging system described here is compatible with both in vitro and in vivo models, which is flexible for various experiments. The general workflow of this protocol includes cell culture, culture media preparation, cell synchronization, cell fixation, and sample imaging with DO-SRS and 2PEF modalities.

We present a protocol to directly visualize metabolic activities in cells regulated by amino acids using deuterium-oxide (heavy water D2O) probed stimulated Raman scattering (DO-SRS) microscopy, which is integrated with two-photon excitation fluorescence microscopy (2PEF).

Essential aromatic amino acids (AAAs) are building blocks for synthesizing new biomasses in cells and sustaining normal biological functions. For example, ...

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...

Biology

Imaging Intermediate Filaments and Microtubules with 2-dimensional Direct Stochastic Optical Reconstruction Microscopy

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, polarity, and motility. The organization of the actin and microtubule networks has been extensively studied but that of IFs is not yet fully characterized. IFs have an average diameter of 10 nm and form a network extending throughout the cell cytoplasm. They are physically associated with actin and microtubules through molecular motors and cytoskeletal linkers. This tight association is at the heart of the regulatory mechanisms that ensure the coordinated regulation of the three cytoskeletal networks required for most cell functions. It is therefore crucial to visualize IFs alone and also together with each of the other cytoskeletal networks. However, IF networks are extremely dense in most cell types, especially in glial cells, which makes its resolution very difficult to achieve with standard fluorescence microscopy (lateral resolution of ~250 nm). Direct STochastic Optical Reconstruction Microscopy (dSTORM) is a technique allowing a gain in lateral resolution of one order of magnitude. Here, we show that lateral dSTORM resolution is sufficient to resolve the dense organization of the IF networks and, in particular, of IF bundles surrounding microtubules. Such tight association is likely to participate in the coordinated regulation of these two networks and may, explain how vimentin IFs template and stabilize microtubule organization as well as could influence microtubule dependent vesicular trafficking. More generally, we show how the observation of two cytoskeletal components with dual-color dSTORM technique brings new insight into their mutual interaction.

The overall goal of this methodology is to give the optimal experimental conditions from sample preparation to image acquisition and reconstruction in order to perform 2D dual color dSTORM images of microtubules and intermediate filaments in fixed cells

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, ...

Serial Block-Face Scanning Electron Microscopy (SBF-SEM) of Biological Tissue Samples

Serial block-face scanning electron microscopy (SBF-SEM) allows for the collection of hundreds to thousands of serially-registered ultrastructural images, offering an unprecedented three-dimensional view of tissue microanatomy. While SBF-SEM has seen an exponential increase in use in recent years, technical aspects such as proper tissue preparation and imaging parameters are paramount for the success of this imaging modality. This imaging system benefits from the automated nature of the device, allowing one to leave the microscope unattended during the imaging process, with the automated collection of hundreds of images possible in a single day. However, without appropriate tissue preparation cellular ultrastructure can be altered in such a way that incorrect or misleading conclusions might be drawn. Additionally, images are generated by scanning the block-face of a resin-embedded biological sample and this often presents challenges and considerations that must be addressed. The accumulation of electrons within the block during imaging, known as "tissue charging," can lead to a loss of contrast and an inability to appreciate cellular structure. Moreover, while increasing electron beam intensity/voltage or decreasing beam-scanning speed can increase image resolution, this can also have the unfortunate side effect of damaging the resin block and distorting subsequent images in the imaging series. Here we present a routine protocol for the preparation of biological tissue samples that preserves cellular ultrastructure and diminishes tissue charging. We also provide imaging considerations for the rapid acquisition of high-quality serial-images with minimal damage to the tissue block.

Serial Block-Face Scanning Electron Microscopy SBF-SEM of Biological Tissue Samples

This protocol outlines a routine method for using serial block-face scanning electron microscopy (SBF-SEM), a powerful 3D imaging technique. Successful application of SBF-SEM hinges on proper fixation and tissue staining techniques, as well as careful consideration of imaging settings. This protocol contains practical considerations for the entirety of this process.

Serial block-face scanning electron microscopy (SBF-SEM) allows for the collection of hundreds to thousands of serially-registered ultrastructural images, ...

A 3D Cartographic Description of the Cell by Cryo Soft X-ray Tomography

Imaging techniques are fundamental in order to understand cell organization and machinery in biological research and the related fields. Among these techniques, cryo soft X-ray tomography (SXT) allows imaging whole cryo-preserved cells in the water window X-ray energy range (284-543 eV), in which carbon structures have intrinsically higher absorption than water, allowing the 3D reconstruction of the linear absorption coefficient of the material contained in each voxel. Quantitative structural information at the level of whole cells up to 10 &#181;m thick is then achievable this way, with high throughput and spatial resolution down to 25-30 nm half-pitch. Cryo-SXT has proven itself relevant to current biomedical research, providing 3D information on cellular infection processes (virus, bacteria, or parasites), morphological changes due to diseases (such as recessive genetic diseases) and helping us understand drug action at the cellular level, or locating specific structures in the 3D cellular environment. In addition, by taking advantage of the tunable wavelength at synchrotron facilities, spectro-microscopy or its 3D counterpart, spectro-tomography, can also be used to image and quantify specific elements in the cell, such as calcium in biomineralization processes. Cryo-SXT provides complementary information to other biological imaging techniques such as electron microscopy, X-ray fluorescence or visible light fluorescence, and is generally used as a partner method for 2D or 3D correlative imaging at cryogenic conditions in order to link function, location, and morphology.

Here, a protocol describing the sample preparation and data collection steps required in cryo soft X-ray tomography (SXT) to image the ultrastructure of whole cryo-preserved cells at a resolution of 25 nm half pitch, is presented.

Imaging techniques are fundamental in order to understand cell organization and machinery in biological research and the related fields. Among these ...

Sample Preparation by 3D-Correlative Focused Ion Beam Milling for High-Resolution Cryo-Electron Tomography

Cryo-electron tomography (cryo-ET) has become the method of choice for investigating cellular ultrastructure and molecular complexes in their native, frozen-hydrated state. However, cryo-ET requires that samples are thin enough to not scatter or block the incident electron beam. For thick cellular samples, this can be achieved by cryo-focused ion beam (FIB) milling. This protocol describes how to target specific cellular sites during FIB milling using a 3D-correlative approach, which combines three-dimensional fluorescence microscopy data with information from the FIB-scanning electron microscope. Using this technique, rare cellular events and structures can be targeted with high accuracy and visualized at molecular resolution using cryo-transmission electron microscopy (cryo-TEM).

Here, we present a pipeline for 3D-correlative focused ion beam milling on guiding the preparation of cellular samples for cryo-electron tomography. The 3D position of fluorescently tagged proteins of interest is first determined by cryo-fluorescence microscopy, and then targeted for milling. The protocol is suitable for mammalian, yeast, and bacterial cells.

Cryo-electron tomography (cryo-ET) has become the method of choice for investigating cellular ultrastructure and molecular complexes in their native, ...

Preparing Adherent Cells for X-ray Fluorescence Imaging by Chemical Fixation

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