We thank E. Arzt for his support through INM, M. Koch for help with the ESEM, and DENS Solutions for providing the microchips. Research in part supported by the Leibniz Competition 2013.

1. Preparation of Labeling and Fixation Reagents
<ol>
	<li>Prepare labeling buffer (BSA-GEL-PBS) using autoclaved phosphate buffered saline, pH 7.4 (PBS). Supplement with 1% BSA (bovine albumin fraction V, biotin-free) and 0.2% gelatin (GEL, from cold water fish skin) and vortex/shake well. 
	Note: Can be stored for ~5 days at 4 &#176;C.</li>
	<li>Prepare washing buffer (BSA-PBS) from autoclaved phosphate buffered saline, pH 7.4. Supplement with 1% BSA and vortex/shake well. Note: Can be stored for ~5 days at 4 &#176;C.</li>
	<li>Prepare 50 mM borate buffer (BB) from 200 mM boric acid stock solution, pH adjusted to 8.3 with 200 mM sodium tetraborate. Note: Can be stored for ~12 months at 4 &#176;C.</li>
	<li>Prepare 0.1 M cacodylate buffer (CB) from 0.2 M sodium cacodylate stock solution, pH adjusted to 7.4, supplemented with 0.1 M sucrose. 
	Note: Opened stock solution can be stored for ~5 days at 4 &#176;C.</li>
	<li>Prepare 0.1 M glycine in PBS (GLY&#8211;PBS). 
	Note: Can be stored for ~2 months at 4 &#176;C.</li>
	<li>Prepare 3% paraformaldehyde in CB (3% PFA) from freshly made or opened vial of 16% PFA, stock solution, EM grade. 
	Note: Opened stock solution can be stored for ~5 days at 4 &#176;C.</li>
	<li>Prepare 2% glutaraldehyde in CB (2% GA) from freshly made (or opened vial of) 25% GA stock solution, EM grade. 
	Note: Opened stock solution can be stored for ~5 days at 4 &#176;C.</li>
	<li>Prepare 300 nM EGF-biotin labeling solution
	<ol>
		<li>Dilute 6 &#181;M EGF-biotin stock solution in BSA-GEL-PBS, use a volume of 100-200 &#181;l per microchip. Use within a few hr.</li>
	</ol>
	</li>
	<li>Prepare 10 nM STR-QD labeling solution
	<ol>
		<li>Centrifuge the streptavidin-QD stock solution for 4 min at 8,000 x g, take supernatant and dilute 1:10 in BB. Different types of QDs can be used, for example, QD655, or QD800. Dilute to final concentration in BSA-GEL-PBS, use 75-100 &#181;l per microchip. Use within a few hr.</li>
	</ol>
	</li>
</ol>
2. Preparation of Silicon Nitride Membrane Microchips
<ol>
	<li>Clean a microscope glass slide, as used for light microscopy, with a clean room tissue and high performance liquid chromatography (HPLC) grade ethanol.</li>
	<li>In a laminar airflow workbench with a low dust level, open a 100 mm diameter Petri dish, and lay the cleaned microscope glass into the dish; leave the dish open.</li>
	<li>With clean tweezers, preferentially coated with carbon fiber or other coating, take 4-10 microchips with thin (50 nm) silicon nitride membranes, one after the other, out of their storage box and onto the glass slide. Grab the microchips gently at their sides and avoid touching the fragile upper side, consisting of the thin silicon nitride membrane. Take great care to always keep the membrane side up and do not touch it. 
	Note: The microchips should be rectangular and have flat edges perpendicular to their silicon nitride covered face, so that they can be gripped easily.</li>
	<li>Close the lid of the petri dish and bring it to a fume hood. Get a new clean room tissue and deposit it beside the petri dish. Prepare two clean 200 ml glass beakers by filling one with 50-70 ml acetone, and the other with a similar volume of ethanol, both of HPLC grade. Transfer the microchips from the glass slide first onto the clean room tissue.</li>
	<li>From the tissue, transfer microchips one by one into the first beaker with acetone for removing the protective resist layer. Wait 2 min, while gently moving the beaker a few times to ensure that the microchips are well rinsed.</li>
	<li>Transfer the microchips directly into the beaker with ethanol, without letting them dry out. Wait 2 min. Then transfer the beaker to the laminar airflow workbench. Get a new clean room tissue.</li>
	<li>Transfer the chips onto the tissue and let them dry for a few min. Transfer the microchips onto the glass slide in the Petri dish, close the dish, and bring it to the plasma cleaner.</li>
	<li>Deposit the glass slide with the microchips into the plasma cleaner. Run a 5 min cleaning program to render the surface of the silicon nitride membrane hydrophilic. 
	Note: Suitable settings applied for our plasma cleaner are: 70 mTorr, gas flow of 11.5 sccm for O2 and 35.0 sccm for Ar, 50 W forward radio frequency (RF) target, 5 W RF range and max. reflected RF.</li>
	<li>Put the glass slide with the microchip back into the petri dish, and close its lid. Take care to keep the microchips sterile from now on.</li>
	<li>Examine the window areas of the microchips under a light microscope for possible membrane ruptures or remaining dirt particles.</li>
	<li>Bring the petri dish to the laminar airflow workbench.</li>
	<li>Place the following items in the laminar airflow workbench: tubes/bottles containing sterilized 0.01% poly-L-lysine solution (PLL, mol. wt 150,000&#8211;300,000, sterile-filtered, and cell culture tested), HPLC grade water, Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum, DMEM without serum, a new clean room tissue, a sterile 24-well plate (a 12-well plate can also be used), a sterile 96-well plate, sterile pipet tips, a pipette, and flat, rounded tip, tweezers.</li>
	<li>Take the 24-well plate, and fill one well with PLL, and two wells with HPLC grade water, use an approx. volume of 1 ml per well. Take the 96-well plate and fill for every microchip two wells with serum supplemented DMEM and one well with serum-free DMEM, use an approx. volume of 200 &#181;l per well.</li>
	<li>From now on use tweezers with sterile tips to handle the microchips. Achieve this, for instance, by briefly flaming and subsequently cooling off the tips of the tweezers.</li>
	<li>First transfer the microchips from the glass slide onto the clean room tissue. 
	From the tissue, transfer up to six microchips into the PLL-filled well, incubate the microchips for 5 min. 
	Note: For an experiment with more microchips, fill additional wells in the 24-well plate.</li>
	<li>Subsequently rinse the microchips by placing them, one-by-one, into the first water-filled well, and from there into the second water-filled well. Don&#39;t leave the microchips in the water longer than about a minute to avoid removal of PLL.</li>
	<li>Finally place them individually into the DMEM-filled wells of the 96-well plate. Store the 96-well plate in the CO2 incubator until the cell suspension is prepared.</li>
</ol>
3. Preparation of Cells on Microchips
<ol>
	<li>Perform a subculture of COS7 cells to prepare a cell suspension with a concentration of approx. 5 &#215; 105 cells/ml. Make sure the cells are mono-dispersed.</li>
	<li>Take the 96-well plate with the microchips and add one droplet of cell suspension to each microchip. Wait 5 min.</li>
	<li>Start to examine the window areas of the microchips with an inverted microscope to choose the right moment for transferring it into a new well with serum-free DMEM. 
	Note: A good density of cells is achieved with approximately one cell per 2,500 &#181;m2 area, corresponding to 24 COS7 cells on a window size of 150 x 400 &#181;m. Higher cell densities have the risk to hinder maximal flattening of the cells. Be aware that after a cell has settled down onto the SIN membrane, it needs 3-5 min to develop enough adhesion to resist floating off during transfer into the new well.</li>
	<li>Transfer microchip to the next well when sufficient cells have adhered onto the window. Keep the microchips always with the cells facing up and avoid any touching of this side.</li>
	<li>After the transfer, examine each microchip window again. If too many cells were washed off and the number of remaining cells is insufficient, re-triturate the cell suspension (to break up newly formed cell clumps) and repeat steps 3.1-3.5.</li>
	<li>When all transferred microchips have enough adhering cells on their window areas, transfer the chips into the last well with serum-free DMEM. Place the 96-well plate back into the CO2 incubator and let the cells recover and flatten out for several hr, best O/N.</li>
</ol>
4. EGFR Labeling, Fixation, and Fluorescence Microscopy
<ol>
	<li>For the following steps, use a 96-well and a 24-well plate. Pre-fill wells with the respective solutions, and use one row or column per microchip. The 24- well plate is used under the fume hood for the fixation steps with CB, PFA and GA. Always transfer microchips with the cells facing up and avoid any touching of this side.</li>
	<li>Rinse microchips by placing the microchips in a well with fresh BSA-GEL-PBS, then place 96 well plate in incubator at 37 &#176;C for 5 min.</li>
	<li>Incubate with 300 nM EGF-biotin solution for 3 min at 37 &#176;C. Because the cells will start endocytosis of EGF-labeled EGFRs, proceed as fast as possible to step 4.4.</li>
	<li>Rinse 3 times with PBS, and 1 time with CB (now the 96 well plate is again at RT).</li>
	<li>Incubate in 3% PFA for 10 min.</li>
	<li>Rinse chips 1 time with CB, and 3 times with PBS.</li>
	<li>Incubate in GLY-PBS for 2 min, rinse 2 times with 1 time PBS.</li>
	<li>Incubate in 10 nM STR-QD for 10 min.</li>
	<li>Rinse 4 times with BSA-PBS.</li>
	<li>Submerge a microchip in a glass-bottom 35 mm dish pre-filled with BSA-PBS at RT.</li>
	<li>Acquire differential interference contrast (DIC) images and fluorescence images of QD-labeled cells.
	<ol>
		<li>Use a 40X air objective for overview images of cells labeled with QD655. Use an oil immersion objective (for example, 63X) to allow the most efficient collection of the emitted fluorescence light; this is especially needed when imaging cells labeled with QD800.</li>
		<li>Use a filter cube that exposes the QDs to fluorescence excitation light between 340 and 380 nm, and allows the collection of emitted light above 420 nm suitable for most QDs (Filter cube A in this case).</li>
		<li>Minimize the light intensity to avoid too intense light that might damage the sample. Set the exposure time to at least 300 msec, so that the signals of all QDs are captured, noting that the fluorescence of QDs has a blinking behavior.</li>
		<li>Adjust the light intensity, and the amplification to provide bright images without visible image noise, while minimizing the light intensity. 
		Note: Alternatively, shorter exposure time can be used if image series are taken from the same region. These series can be processed to yield a maximum projection image, leading to averaging and thus noise reduction of the resulting image.</li>
	</ol>
	</li>
	<li>Place the imaged microchip back (cells facing up) into a well of a 96-well plate filled with BSA-PBS.</li>
	<li>Rinse microchips 1 times in CB.</li>
	<li>Incubate in 2% GA for 10 min.</li>
	<li>Rinse 1 time in CB, and 3 times with BSA-PBS.</li>
	<li>Store microchips until ESEM imaging, in BSA-PBS pre-filled wells of a new 96-well plate. For every microchip fill a row of four wells with HPLC-grade water. 
	Note: The samples can be kept at 4 &#176;C for several weeks, however, the best results are achieved when the complete imaging series is performed within two days. QDs will slowly start to detach from the sample, it is therefore not recommended to wait more than 10 days before the ESEM imaging is done. 
	Note: Fluorescence images are stored for future analysis. The positions of the cells in the images can easily be correlated with electron microscopy images via localizing the corners of the SiN window and measuring the positions relative to the corners. 
	Note: It is advisable to print high quality fluorescence images and use them for orientation purposes when recording the ESEM images.</li>
</ol>
5. Wet ESEM-STEM of Whole Cells
<ol>
	<li>Prepare the ESEM for wet STEM
	<ol>
		<li>Mount the gaseous secondary electron detector (GSED), and the Peltier stage, containing the STEM detector (located under the sample).</li>
		<li>Start the flow of cooling water through the stage, and set the temperature to 3 &#176;C.</li>
		<li>Set the stage x-y coordinates to zero, and adjust the working distance to approximately 6 mm.</li>
	</ol>
	</li>
	<li>During the ESEM imaging, keep the 96-well plate with the microchips cool, for instance on frozen cooling elements, in a Styrofoam box.</li>
	<li>For loading a sample into the precooled ESEM-STEM stage, take the 96-well plate out of the cooling container and place it close to the opened ESEM stage. Put a new clean room tissue beside it.</li>
	<li>Quickly rinse a microchip (with the cells facing up) by dipping it four times, each time for about a second, into a well filled with HPLC-grade water.</li>
	<li>Blot the backside of the microchip briefly on the clean room tissue and place the microchip into the precooled stage. Make sure that the cells remain wet, this can be recognized by a reflective surface (drying results in a dull surface).</li>
	<li>Wet the sample surface with 3 &#181;l of cooled HPLC-grade water (make sure not to touch the surface with the pipet tip), and fix the sample in the stage.</li>
	<li>Place three additional 3 &#181;l water droplets on the stage close to the sample.</li>
	<li>Close the specimen chamber</li>
	<li>Purge the specimen chamber by cycling the pressure five times between 800 and 1,500 Pa. End the cycling with 800 Pa.</li>
	<li>Switch the electron beam (30 kV) on and examine the sample under the lowest magnification with both detectors, i.e., the GSED, and the STEM detector to search for the location of the microchip window. 
	Note: If the water layer is too thick, the window may not be visible. In this case, start lowering the pressure to 760 Pa and wait a few min. The window first becomes visible with the GSED.</li>
	<li>Position the SiN membrane window in the center of the image by moving the specimen stage and reduce the pressure to 750-720 Pa. 
	Note: When the water layer is thin enough, the window also becomes visible with the STEM detector.</li>
	<li>From now on use the dark field segment of the STEM detector to record images. First, acquire one or two images showing all cells on the complete window area.</li>
	<li>Compare these ESEM images with the light microscopy images, locate the corners of the SiN window.</li>
	<li>Correlate the coordinate frames of both microscopy modalities by comparing the magnification, rotation, and possible mirroring. Locate the same cells in both images.</li>
	<li>Select very small dirt particles or a cell region with small features to adjust imaging settings, such as the eucentric height, the fine focus and the correction of lens stigmatism. Work at a magnification of at least 50.000X, use an electron probe current around 0.6 nA, a pixel-dwell time of 30 &#181;s, and an image size of 1024 x 884 pixels. 
	Note: Other settings may also be used as long as an increase of the electron dose is avoided. 
	Note: Even when working very clean, the chip surfaces will in practice always contain some dirt particles, because the protocol is not executed in a clean room environment.</li>
	<li>To record STEM images from the QD-labeled cells, refer to the fluorescence images and choose a cell of interest. 
	Note: It is best to start with a cell showing strong QD signals.</li>
	<li>Record a STEM overview image of this cell.</li>
	<li>Go to a peripheral region where the cell border is visible and zoom in to reach 50.000X magnification.</li>
	<li>Record images showing individual QDs, use pixel dwell times of 20-30 &#181;sec. In order to avoid radiation damage, image every region only once with high magnification. 
	Note: Usually, at this stage both, the focus and the correction of the stigmatism, need some fine adjustment.</li>
	<li>After the recording of a sufficient number of high magnification images, zoom-out, and acquire another STEM overview image, depicting the regions just recorded with high-magnification as dark rectangles. 
	Note: These overview images will later serve to correlate ESEM and fluorescence images and to determine the precise location of the high magnification images within the cellular context.</li>
	<li>Proceed to the next cell of interest, start with an overview image, record a series of high magnification images, and end with an overview image.</li>
	<li>At the end of an ESEM imaging session, increase the pressure to 850-900 Pa before starting to vent the chamber. 
	Note: This increase of pressure improves the chance of recovering the microchip with a still wetted surface; however, this cannot always be achieved. A sample that has never been dried out can be restored in BSA-PBS and, if desired, reinvestigated in the ESEM at a later point in time.</li>
</ol>

<ol>
	<li>Dukes, M. J., Peckys, D. B., de Jonge, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+fluorescence+microscopy+and+scanning+transmission+electron+microscopy+of+quantum-dot-labeled+proteins+in+whole+cells+in+liquid.">Correlative fluorescence microscopy and scanning transmission electron microscopy of quantum-dot-labeled proteins in whole cells in liquid.</a> ACS Nano. 4, 4110-4116 (2010).</li><li>Nishiyama, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atmospheric+scanning+electron+microscope+observes+cells+and+tissues+in+open+medium+through+silicon+nitride+film.">Atmospheric scanning electron microscope observes cells and tissues in open medium through silicon nitride film.</a> J. Struct. Biol. 169, 438-449 (2010).</li><li>Kinoshita, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immuno-electron+microscopy+of+primary+cell+cultures+from+genetically+modified+animals+in+liquid+by+atmospheric+scanning+electron+microscopy.">Immuno-electron microscopy of primary cell cultures from genetically modified animals in liquid by atmospheric scanning electron microscopy.</a> Microsc. Microanal. 20, 469-483 (2014).</li><li>Jonge, N., Peckys, D. B., Kremers, G. J., Piston, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electron+microscopy+of+whole+cells+in+liquid+with+nanometer+resolution.">Electron microscopy of whole cells in liquid with nanometer resolution.</a> Proc. Natl. Acad. Sci. 106, 2159-2164 (2009).</li><li>Peckys, D. B., de Jonge, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Liquid+Scanning+Transmission+Electron+Microscopy:+Imaging+Protein+Complexes+in+their+Native+Environment+in+Whole+Eukaryotic+Cells.">Liquid Scanning Transmission Electron Microscopy: Imaging Protein Complexes in their Native Environment in Whole Eukaryotic Cells.</a> Microsc. Microanal. 20, 189-198 (2014).</li><li>Peckys, D. B., Baudoin, J. P., Eder, M., Werner, U., de Jonge, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epidermal+growth+factor+receptor+subunit+locations+determined+in+hydrated+cells+with+environmental+scanning+electron+microscopy.">Epidermal growth factor receptor subunit locations determined in hydrated cells with environmental scanning electron microscopy.</a> Sci. Rep. 3, 2621-2626 (2013).</li><li>Peckys, D. B., Korf, U., de Jonge, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Local+variations+of+HER2+dimerization+in+breast+cancer+cells+discovered+by+correlative+fluorescence-+and+liquid+electron+microscopy.">Local variations of HER2 dimerization in breast cancer cells discovered by correlative fluorescence- and liquid electron microscopy.</a> Science Advances. under revision, (2015).</li><li>Giepmans, B. N., Deerinck, T. J., Smarr, B. L., Jones, Y. Z., Ellisman, M. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlated+light+and+electron+microscopic+imaging+of+multiple+endogenous+proteins+using+Quantum+dots.">Correlated light and electron microscopic imaging of multiple endogenous proteins using Quantum dots.</a> Nat. Meth. 2, 743-749 (2005).</li><li>Herbst, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Review+of+epidermal+growth+factor+receptor+biology.">Review of epidermal growth factor receptor biology.</a> Int. J.Radiat. Oncol. Biol. Phys. 59, S21-S26 (2004).</li><li>Normanno, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epidermal+growth+factor+receptor+(EGFR)+signaling+in+cancer.">Epidermal growth factor receptor (EGFR) signaling in cancer.</a> Gene. 366, 2-16 (2006).</li><li>Hofman, E. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EGF+induces+coalescence+of+different+lipid+rafts.">EGF induces coalescence of different lipid rafts.</a> J. Cell Sci. 121, 2519-2528 (2008).</li><li>Irwin, M. E., Mueller, K. L., Bohin, N., Ge, Y., Boerner, 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=Lipid+raft+localization+of+EGFR+alters+the+response+of+cancer+cells+to+the+EGFR+tyrosine+kinase+inhibitor+gefitinib.">Lipid raft localization of EGFR alters the response of cancer cells to the EGFR tyrosine kinase inhibitor gefitinib.</a> J. Cell Physiol. 226, 2316-2328 (2011).</li><li>Lillemeier, B. F., Pfeiffer, J. R., Surviladze, Z., Wilson, B. S., Davis, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasma+membrane-associated+proteins+are+clustered+into+islands+attached+to+the+cytoskeleton.">Plasma membrane-associated proteins are clustered into islands attached to the cytoskeleton.</a> Proc. Natl. Acad. Sci. 103, 18992-18997 (2006).</li><li>Mayawala, K., Vlachos, D. G., Edwards, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heterogeneities+in+EGF+receptor+density+at+the+cell+surface+can+lead+to+concave+up+scatchard+plot+of+EGF+binding.">Heterogeneities in EGF receptor density at the cell surface can lead to concave up scatchard plot of EGF binding.</a> FEBS Lett. 579, 3043-3047 (2005).</li><li>Valley, C. C., Lidke, K. A., Lidke, D. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+spatiotemporal+organization+of+ErbB+receptors:+insights+from+microscopy.">The spatiotemporal organization of ErbB receptors: insights from microscopy.</a> Cold Spring Harb. Perspect. Biol. 6, (2014).</li><li>Peckys, D. B., Bandmann, V., de Jonge, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+fluorescence-+and+scanning+transmission+electron+microscopy+of+quantum+dot+labeled+proteins+on+whole+cells+in+liquid.">Correlative fluorescence- and scanning transmission electron microscopy of quantum dot labeled proteins on whole cells in liquid.</a> Meth. Cell Biol. 124, 305-322 (2014).</li><li>Peckys, D. B., Dukes, M. J., de Jonge, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+fluorescence+and+electron+microscopy+of+quantum+dot+labeled+proteins+on+whole+cells+in+liquid.">Correlative fluorescence and electron microscopy of quantum dot labeled proteins on whole cells in liquid.</a> Methods Mol. Biol. 1117, 527-540 (2014).</li><li>Ring, E. A., Peckys, D. B., Dukes, M. J., Baudoin, J. P., de Jonge, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Silicon+nitride+windows+for+electron+microscopy+of+whole+cells.">Silicon nitride windows for electron microscopy of whole cells.</a> J. Microsc. 243, 273-283 (2011).</li><li>Bogner, A., Thollet, G., Basset, D., Jouneau, P. H., Gauthier, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wet+STEM:+A+new+development+in+environmental+SEM+for+imaging+nano-objects+included+in+a+liquid+phase.">Wet STEM: A new development in environmental SEM for imaging nano-objects included in a liquid phase.</a> Ultramicroscopy. 104, 290-301 (2005).</li><li>Chung, I., Akita, R., Vandlen, R., Toomre, D., Schlessinger, J., Mellman, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+control+of+EGF+receptor+activation+by+reversible+dimerization+on+living+cells.">Spatial control of EGF receptor activation by reversible dimerization on living cells.</a> Nature. 464, 783-787 (2010).</li><li>Stoyan, D., Stoyan, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estimating+pair+correlation+functions+of+planar+cluster+processes.">Estimating pair correlation functions of planar cluster processes.</a> Biom. J. 38, 259-271 (1996).</li><li>Gao, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+tumor-targeted+fluorescence+imaging+using+near-infrared+non-cadmium+quantum+dots.">In vivo tumor-targeted fluorescence imaging using near-infrared non-cadmium quantum dots.</a> Bioconjug. Chem. 21, 604-609 (2010).</li><li>Lippincott-Schwartz, J., Manley, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Putting+super-resolution+fluorescence+microscopy+to+work.">Putting super-resolution fluorescence microscopy to work.</a> Nat. Meth. 6, 21-23 (2009).</li><li>Piston, D. W., Kremers, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+protein+FRET:+the+good,+the+bad+and+the+ugly.">Fluorescent protein FRET: the good, the bad and the ugly.</a> Trends Biochem. Sci. 32, 407-414 (2007).</li><li>Warren, C. M., Landgraf, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Signaling+through+ERBB+receptors:+Multiple+layers+of+diversity+and+control.">Signaling through ERBB receptors: Multiple layers of diversity and control.</a> Cell. Signal. 18, 923-933 (2006).</li><li>Leuchowius, K. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Parallel+visualization+of+multiple+protein+complexes+in+individual+cells+in+tumor+tissue.">Parallel visualization of multiple protein complexes in individual cells in tumor tissue.</a> Mol. Cell. Proteomics. 12, 1563-1571 (2013).</li><li>Hoenger, A., McIntosh, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Probing+the+macromolecular+organization+of+cells+by+electron+tomography.">Probing the macromolecular organization of cells by electron tomography.</a> Curr. Opin. Cell Biol. 21, 89-96 (2009).</li><li>Zhang, J., Leiderman, K., Pfeiffer, J. R., Wilson, B. S., Oliver, J. M., Steinberg, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterizing+the+topography+of+membrane+receptors+and+signaling+molecules+from+spatial+patterns+obtained+using+nanometer-scale+electron-dense+probes+and+electron+microscopy.">Characterizing the topography of membrane receptors and signaling molecules from spatial patterns obtained using nanometer-scale electron-dense probes and electron microscopy.</a> Micron. 37, 14-34 (2006).</li><li>Pinto da Silva, P., Kan, F. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-fracture:+a+method+for+high+resolution+labeling+of+cell+surfaces.">Label-fracture: a method for high resolution labeling of cell surfaces.</a> J. Cell Biol. 99, 1156-1161 (1984).</li><li>Bergersen, L. H., Storm-Mathisen, J., Gundersen, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunogold+quantification+of+amino+acids+and+proteins+in+complex+subcellular+compartments.">Immunogold quantification of amino acids and proteins in complex subcellular compartments.</a> Nat. Protoc. 3, 144-152 (2008).</li><li>Orlov, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+cell+immunogold+labelling+of+RNA+polymerase.">Live cell immunogold labelling of RNA polymerase.</a> II. Sci. Rep. 5, 8324 (2015).</li><li>Fiala, G. J., Kaschek, D., Blumenthal, B., Reth, M., Timmer, J., Schamel, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pre-clustering+of+the+B+cell+antigen+receptor+demonstrated+by+mathematically+extended+electron+microscopy.">Pre-clustering of the B cell antigen receptor demonstrated by mathematically extended electron microscopy.</a> Front. Immunol. 4, 427 (2013).</li></ol>

The authors have nothing to disclose.

Studying the spatial distribution and stoichiometry of the membrane proteins, such as the EGFR, in whole cells provides important context information about its function and possible changes thereof11-14. It is challenging to study the distribution of monomers, dimers, and clusters directly on a subcellular level using commonly used biochemical methods, for which information is lost about the localization of the protein in the cell or differences between cells15. The labeling and microscopy methods described here allow the visualization of protein complexes on a length scale of a few nanometers so that its stoichiometry can be studied within the context of individual, intact cells4-7. This is not possible with (super resolution) light microscopy23 because its resolution is insufficient to distinguish molecules engaged in a protein complex. F&#246;rster Resonance Energy Transfer (FRET) is an ensemble averaging technique24, and does not necessarily detect the dimers and higher order clusters as a consequence of the short range of the energy transfer25. Proximity ligation assays26 do not actually measure distances and are thus not capable of distinguishing between a cellular region with a high protein density, inevitably leading to a number of detected proximities by random chance, from a cellular region with a similar protein density but containing protein complexes exhibiting distinct distances between the labels. The required high resolution to visualize the constituents of protein complexes is achieved by transmission electron microscopy (TEM). However, conventional TEM of whole cells is limited by the requirement of thin samples/sections slicing through the plasma membrane27, or plasma membrane ripping or breaking off28,29 resulting in randomly formed small pieces of membrane. The plasma membrane is thus not imaged as a whole, leading to a lack of possibly crucial cellular context information.
Other experimental challenges for the study of protein stoichiometry in cells arise from the applied protein labeling. Commonly used immunolabeling bears the difficulty that one-to-one stoichiometry between receptor and label can only be achieved with a monovalent probe; this is not the case when primary and secondary antibodies are required. To obtain information about the protein stoichiometry, it is required that a probe binds uniquely to one epitope on the receptor and has one binding site for a fluorescent probe or a high atomic number nanoparticle on the other side. Secondly, the precision achievable with antibody-labeling is restricted on account of their large size30 to 30 nm, so that a discrimination between neighboring single proteins, homodimers, or larger clusters is not feasible, at least not for receptors of the EGFR family. Much smaller specific labels are known in literature and can be applied even for intracellular labeling31 but those are not commonly used. The length of the whole label (EGF-biotin-streptavidin-QD) is of a similar dimension as the EGFR, and sufficiently small to be able to detect the dimer. Moreover, the described two-step labeling procedure avoids labeling induced receptor clustering.
Our method is not very difficult, not much more than fluorescence microscopy of labeled proteins. However, an experiment needs to be carried out with great care, since the number of steps adds up and an error in one of the steps may lead to a complete failure in the experiment. Handling the microchips is not more tedious than the handling of TEM grids but some training empty chips is recommended. ESEM-STEM of whole hydrated cells is probably the most difficult aspect, and requires at least a skilled operator and several days of practice in order to reach a resolution around 3 nm as needed to visualize the QDs. Radiation damage of the specimen under investigation presents a risk. The pressure and the temperature should be carefully watched to maintain a water layer. Moreover, the thickness of the water layer may vary between cells. It is advisable to monitor the presence of the water layer using the gaseous electron detector above the specimen, as described elsewhere6. Cellular regions should only be imaged once or twice to avoid damage.
A key limitation of the method is that high-resolution information about the ultrastructure is absent. Other techniques, for example, cryo TEM, are needed to study the protein structure, and the cellular ultrastructure15. Secondly, studying the dynamic interplay of a multiple of proteins in live cell in time-lapse microscope is not feasible on account of radiation damage, and requires state-of-the-art light microscopy23. Currently, our method is not capable of determining the absolute level of dimers since the labeling efficiency is unknown but we expect to add such data in future. It is nevertheless possible to tell if dimers are present or not. From the literature is it known, that even a labeling efficiency around 15% is enough to detect dimers with sufficient statistical significance32.
It is readily possible to study other membrane-bound receptors via their ligand, via biotinylated Fab fragments of specific antibodies, or via other small linkers7 using the described two-step labeling protocol. Since we have already demonstrated that two different colors (sizes) of QDs can be used, and gold nanoparticle labeling was used in prior work6, it seems feasible to label multiple protein species in the same experiment. The key challenge for the study of the stoichiometry of multiple protein species is ensuring a similar labeling efficiency for both species or at least a normalization of the obtained relative stoichiometry on the respective labeling efficiencies. Yet, the two different QDs used in this study do not seem to differ much in labeling efficiency (see Figures 2 and 4). The described method involving two-step labeling with QDs, and correlative fluorescence microscopy and ESEM-STEM presents a viable method to study the complex interplay of membrane proteins in the intact plasma membranes of whole cells.

A new approach was introduced recently, to image whole cells with labeled proteins in liquid state using STEM1-3, or correlative fluorescence microscopy and STEM4-7. This methodology is capable of studying the spatial distribution of membrane proteins and protein complexes including their stoichiometry at the single molecule level in the intact plasma membranes of whole cells. Here, we describe a protocol involving a two-step specific labeling of receptor proteins with fluorescent quantum dots (QDs)8, and correlative light microscopy and ESEM-STEM. As example, we studied the EGFR, a membrane protein that belongs to the protein kinase superfamily. Upon ligand binding, the receptor activates and then forms a dimer with another activated EGFR; the subsequent signal cascade drives cell proliferation9. Mutations leading to abnormal EGFR expression or activity play a central role in many types of cancer10. Studying the spatial arrangement of this membrane receptor in whole cells provides important context information about its function and possible changes thereof11-14. But it remains challenging to probe the distribution of EGFR monomers, dimers, and clusters directly on a (sub-)cellular level with biochemical- or conventional microscopy methods15. Our method is capable of visualizing EGFRs with a spatial resolution of a few nanometers such that the locations of proteins in a complex can be determined. Most importantly, the obtained information about the stoichiometric distribution relates to the native situation of the protein in the intact cell.
In order to achieve a short distance between the label and the receptor, EGFR was directly labeled via its ligand EGF, using a biotin-streptavidin bond to a QD. This label can be detected both with fluorescence- and electron microscopy. We have used this label for correlative imaging of COS7 cells in liquid enclosed in a microfluidic chamber previously1,16,17. However, on account of a multiple of streptavidin molecules per QD, this label may induce receptor clustering, leading to a low labeling efficiency or rather expensive experiments, and it is not possible to conclude on the stoichiometry of the protein under investigation. Label induced receptor clustering can be avoided altogether by employing the two-step labeling procedure presented here, resulting in a probe comprising biotinylated EGF to which only one streptavidin-quantum dot (STR-QD) is coupled. The cells are first incubated with biotinylated EGF, and then fixed. The fixation step determines the time point at which protein processes are stopped (depending on the speed of fixation). STR-QD is applied thereafter as second step of the labeling protocol. Clustering does not take place since dynamic membrane motion of the proteins is hindered once the proteins are fixed. Moreover, the STR-QD solution, which is the most expensive component of the experiment, can be applied in an optimized low concentration, and this second step of labeling is not time-crucial, i.e., the QD can be coupled in several minutes.
To study the stoichiometry of EGFR as distributed in whole cells, cellular samples were prepared on silicon microchips18. After applying the two-step QD labeling, and the recording of fluorescence microscopy images, the cells were rinsed with pure water, and imaged in hydrated state using ESEM-STEM19. The resulting correlative images provide information about the cellular EGFR distribution, and the stoichiometry in their natural context of hydrated cells. A similar method was used to study HER2 proteins in breast cancer cells in a recent study7.

<table><tbody><tr><td>Inverted fluorescence microscope</td><td>Leica</td><td>AF6000 + DMI6000B</td><td></td></tr><tr><td>ESEM</td><td>FEI Company</td><td>Quanta 400 FEG</td><td></td></tr><tr><td>DMEM (1x) + GlutaMax + glucose + pyruvate</td><td>Gibco</td><td>31966-021</td><td></td></tr><tr><td>DPBS (1x) - Calcium chloride - Magnesium chlorid</td><td>Gibco</td><td>14190-144</td><td></td></tr><tr><td>Fetal bovine serum, Performance Plus (FBS)</td><td>Gibco</td><td>16000-036</td><td></td></tr><tr><td>Cellstripper</td><td>Mediatech, Inc.</td><td>25-056 Cl</td><td></td></tr><tr><td>Water, chromasolv Plus for HPLC</td><td>Sigma-Aldrich</td><td>34877-2.5L</td><td></td></tr><tr><td>PBS Roti-Stock 10x PBS</td><td>Carl Roth</td><td>1058.1</td><td></td></tr><tr><td>Ethanol, Rotisolv HPLC grade</td><td>Carl Roth</td><td>P076.2</td><td></td></tr><tr><td>Albumin Fraction V, biotin-free (BSA)</td><td>Carl Roth</td><td>O163.2</td><td></td></tr><tr><td>Gelatin, from cold water fish skin</td><td>Sigma-Aldrich</td><td>G7041-500G</td><td></td></tr><tr><td>Glycine</td><td>Carl Roth</td><td>T873.1</td><td></td></tr><tr><td>Epidermal Growth Factor</td><td>Molecular Probes</td><td>E3477</td><td>Biotin-XX Conjugate (biotin EGF)</td></tr><tr><td>Sodium teraborate decahydrate</td><td>Sigma-Aldrich</td><td>S9640-500G</td><td></td></tr><tr><td>Boric acid</td><td>Sigma-Aldrich</td><td>B6768-500G</td><td></td></tr><tr><td>Qdot 655 streptavidin conjugate</td><td>Molecular Probes</td><td>Q10121MP</td><td></td></tr><tr><td>Qdot 800 streptavidin conjugate</td><td>Molecular Probes</td><td>Q10173MP</td><td></td></tr><tr><td>Silicon microchips with silicon nitride membranes of 50 nm thickness and dimensions of 50 &amp;micro;m x 0.40 mm</td><td>DENS Solutions</td><td>Custom made</td><td></td></tr></tbody></table>

studying the stoichiometry of epidermal growth factor receptor in intact cells using correlative microscopy

This protocol describes the labeling of epidermal growth factor receptor (EGFR) on COS7 fibroblast cells, and subsequent correlative fluorescence microscopy and environmental scanning electron microscopy (ESEM) of whole cells in hydrated state. Fluorescent quantum dots (QDs) were coupled to EGFR via a two-step labeling protocol, providing an efficient and specific protein labeling, while avoiding label-induced clustering of the receptor. Fluorescence microscopy provided overview images of the cellular locations of the EGFR. The scanning transmission electron microscopy (STEM) detector was used to detect the QD labels with nanoscale resolution. The resulting correlative images provide data of the cellular EGFR distribution, and the stoichiometry at the single molecular level in the natural context of the hydrated intact cell. ESEM-STEM images revealed the receptor to be present as monomer, as homodimer, and in small clusters. Labeling with two different QDs, i.e.,&#160;one emitting at 655 nm and at 800 revealed similar characteristic results.

Figure 1 and 2 show representative images of QD655 labeled, membrane-bound EGFR visualized in intact, fully hydrated COS-7 cells. The DIC image in Figure 1 a gives an impression of the membrane topography of the cells, and the corresponding fluorescence image in Figure 1b depicts the distribution of EGFR after 3 min of EGF-Biotin incubation. Figure 1A and B are each stitched together from two images recorded with a 40X air objective. The EGF activated EGFR is distributed over the entire cellular surface. In most cells a slight enhancement of fluorescence (Figure 1B) can be seen at the cell edges, indicating a locally increased occurrence of EGFR20. Control experiments conducted using a similar protocol verified specific EGFR labeling (data not shown). These controls included: 1) a control labeling without previous incubation of the cells with EGF-biotin, i.e., only incubation with STR-QDs, and 2) an incubation with non-biotinylated EGF and STR-QDs.
The three cells marked with rectangles were further investigated with ESEM-STEM. The ESEM-STEM images depicted in Figure 1C-E show low magnification overviews of these cells. These transmission images reflect whole cells in hydrated state with a thin layer of water residing over the cell placed on a silicon microchip with silicon nitride (SiN) viewing window. The vacuum chamber in the microscope contained water vapor, and the balance between sample temperature and pressure was adjusted to maintain a thin layer of liquid over the cells. The individual cells can be easily recognized on account of their shape and location on the SiN membrane window. The low-magnification ESEM-STEM images reveal fine structures, such as filopodia, extending from the cell edge towards neighboring cells, and some structures inside the thinner cell regions. Thick central cellular regions including the nucleus appear white because transmission through the sample is not possible for electrons of the used energy (30 keV).
High-resolution ESEM-STEM images were recorded in the thinner, peripheral regions. The spatial resolution for ESEM-STEM of nanoparticles in the thin regions of cells in a thin liquid layer was determined in a previous study6 to amount to 3 nm. Four high-resolution images shown in Figure 2A-D were recorded at the locations of the small rectangles in Figure 1C-E. The used magnification was sufficient to discern individual QDs, appearing as bright, bullet-shaped rods, each bound to an individual EGFR. The QD655 has typical dimensions of 6 x 14 nm2.
Figure 2A (corresponding to the rectangular area indicated on the cell in Figure 1C) shows QD labels on a membrane fold diagonally crossing the image. This membrane structure has a higher EGFR density than the surrounding membrane regions. At several locations, two labels were at close proximity. Two examples with distances of 20 and 24 nm are indicated with arrowheads. These pairs of labels are interpreted as belonging to EGFR dimers. Figure 2B gives an example from the edge of a cell (Figure 1D, left rectangle), EGFR monomers and dimers can be seen as well. Figure 2C (Figure 1C, right rectangle) was recorded of a region with a lower fluorescence signal, i.e., lower EGFR density. Nevertheless, EGFR was also found here in dimers as well. In addition, two clusters of 10 or 11 EGFRs were present (see ellipses). Figure 2D shows another example of a membrane fold (Figure 1E) with smaller clusters including 5-6 EGFRs, in addition to several monomers and dimers.
As an example of the kind of information that can be obtained from this data, the pair correlation function21 g(x) was determined for all label positions in Figure 2. Note that this analysis is not part of this protocol and the procedure was described elsewhere6,7. g(x) is a measure of the chance of a particle to be found within a certain radial distance x from a reference particle. g(x) = 1 represents a random distribution, and a larger value is evidence for clustering. The positions of a total of 210 labels were automatically detected in the four images of Figure 2A-D, and g(x) was calculated using a bin size of 5 nm and a smoothing filter with a bandwidth of 10 nm. The g(x) curve (Figure 3) shows a preferred center-to-center QD distance of 25 nm. EGFRs are thus not randomly oriented but a significant fraction of them resides at this preferred distance. From an approximate molecular model6 (Figure 3 inset) we estimate that the center-to-center distance between the QD and the EGF binding pocket amounts to ~14 nm, and center-to-center distance between the two QDs attached to a EGFR dimer to be ~27 nm (this value is likely to vary by a few nanometers due to the flexibility of the linker). The preferred label distance is thus consistent with the expected label distance for the EGFR dimer within the precision of the method. The g(x) curve is larger than unity for distance of up to 300 nm, indicating the presence of clusters, consistent with the data. This analysis shows that the stoichiometry of the EGFR can be studied with our method.
Figure 4 shows a similar result as Figure 2, except that the labeling was performed with EGF-QD800, and a 63X oil immersion objective was used for the fluorescence image. This data confirms that these typical results are also found when the EGFR labeling is done with QDs, which emit in the near red spectrum. Figure 4A is the fluorescence image of a representative cell, Figure 4B shows the same cell in ESEM-STEM overview mode and Figure 4C is a 150,000 x magnification image, recorded at the upper membrane border of the cell (see rectangle in Figure 4B). Similar to Figure 2A and D, the images capture QD-labeled EGFR on a membrane fold and shows monomeric, dimeric, and clustered receptors. Note that the electron dense QD core appears somewhat smaller and rounder than the cores of QDs emitting at 655 nm, consistent with their smaller core size22 of ~5 nm.
<img alt="Figure 1" src="/files/ftp_upload/53186/53186fig1.jpg" /> 
Figure 1. Correlative DIC, fluorescence and ESEM-STEM of EGF-QD655 labeled EGFR on fully hydrated COS-7 cells. (A) DIC image of the cells grown on the SiN membrane window area, giving a topographic impression of the plasma membrane. (B) Fluorescence image showing cells on the centrally located SiN membrane window (marked by the dashed rectangle). (C-E) Three ESEM-STEM low magnification images from the cells marked with rectangles in A and B. These cell overview images serve to determine the precise location of subsequently recorded high-resolution images. The location of four high-resolution images (shown in Figure 2A-D) are marked with white rectangles. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53186/53186fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53186/53186fig2.jpg" /> 
Figure 2. High-resolution ESEM-STEM images depicting membrane bound EGFRs. (A) Micrograph at 75,000X magnification of the area marked in Figure 1B. Many monomers are visible. Several dimers are indicated with the arrowheads. (B) and (C) images acquired at 50,000X magnification of the left, respectively right, membrane areas marked in Figure 1C. Clusters of EGFRs are outlined. (D) image recorded with 50,000X magnification at the location marked in Figure 1D. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53186/53186fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53186/53186fig3.jpg" /> 
Figure 3. Pair correlation function g(x) as function of the radial distance x determined for the inter-particle distances of all 210 labels detected in Figure 2A-D. The peak at 25 nm indicates that a center-to-center QD distance has a much higher chance of occurring than random, whereby a g(x) = 1 indicates a random chance. The dashed line is a guide to the eye representing g(x) of a random distribution. The inset shows an approximate molecular model of the EGFR dimer with bound EGF and streptavidin coated QDs attached via biotin. The models of streptavidin, EGF and the EGFR were obtained from CPK models of the 1stp (streptavidin), 1EGF (EGF), 1NQL, 2JWA, 1M17, 1IVO and 2GS6 (EGFR) structures in the RCSB Protein Protein Databank, created by Jmol Version 12.2.15. The biotin model is as drawn in RCSB Ligand Explorer Version 1.0. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53186/53186fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53186/53186fig4.jpg" /> 
Figure 4. Exemplary COS-7 cell labeled with EGF-QD800 and imaged with correlative fluorescence and ESEM-STEM. (A) Fluorescence image, (B) low magnification overview ESEM-STEM image. (C) High-resolution image recorded at the location of the rectangle in b at 150,000X magnification. Monomeric, dimeric, and clustered EGFRs are detected similar to labeling with EGF-QD655. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53186/53186fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Studying the Stoichiometry of Epidermal Growth Factor Receptor in Intact Cells using Correlative Microscopy at JoVE.com

Studying the Stoichiometry of Epidermal Growth Factor Receptor in Intact Cells using Correlative Microscopy

This protocol describes the labeling of epidermal growth factor receptor (EGFR) on COS7 fibroblast cells, and subsequent correlative light- and electron microscopy of whole cells in hydrated state. The label contained fluorescent quantum dots. The protocol can be used to study the stoichiometry of EGFR at the single molecule level.

This protocol describes the labeling of epidermal growth factor receptor (EGFR) on COS7 fibroblast cells, and subsequent correlative fluorescence ...

studying-stoichiometry-epidermal-growth-factor-receptor-intact-cells

Nanyang Technological University

Research

JoVE Journal

Bioengineering

8.3K Views.  INM-Leibniz Institute for New Materials. This protocol describes the labeling of epidermal growth factor receptor (EGFR) on COS7 fibroblast cells, and subsequent correlative light- and electron microscopy of whole cells in hydrated state. The label contained fluorescent quantum dots. The protocol can be used to study the stoichiometry of EGFR at the single molecule level. 

Video: Studying the Stoichiometry of Epidermal Growth Factor Receptor in Intact Cells using Correlative Microscopy

An Optimized Single-Molecule Pull-Down Assay for Quantification of Protein Phosphorylation

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to measure protein phosphorylation cannot preserve the heterogeneity in phosphorylation across individual proteins. The single-molecule pull-down (SiMPull) assay was developed to investigate the composition of macromolecular complexes via immunoprecipitation of proteins on a glass coverslip followed by single-molecule imaging. The current technique is an adaptation of SiMPull that provides robust quantification of the phosphorylation state of full-length membrane receptors at the single-molecule level. Imaging thousands of individual receptors in this way allows for quantifying protein phosphorylation patterns. The present protocol details the optimized SiMPull procedure, from sample preparation to imaging. Optimization of glass preparation and antibody fixation protocols further enhances data quality. The current protocol provides code for the single-molecule data analysis that calculates the fraction of receptors phosphorylated within a sample. While this work focuses on phosphorylation of the epidermal growth factor receptor (EGFR), the protocol can be generalized to other membrane receptors and cytosolic signaling molecules.

The present protocol describes sample preparation and data analysis to quantify protein phosphorylation using an improved single-molecule pull-down (SiMPull) assay.

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to ...

Biochemistry

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial applications as well as increases basic fundamental biological knowledge. F&ouml;rster (Fluorescence) Resonance Energy Transfer (FRET) between a donor molecule in an electronically excited state and a nearby acceptor molecule has been frequently utilized for studies of protein-protein interactions in living cells. The proteins of interest are tagged with two different types of fluorescent probes and expressed in biological cells. The fluorescent probes are then excited, typically using laser light, and the spectral properties of the fluorescence emission emanating from the fluorescent probes is collected and analyzed. Information regarding the degree of the protein interactions is embedded in the spectral emission data. Typically, the cell must be scanned a number of times in order to accumulate enough spectral information to accurately quantify the extent of the protein interactions for each region of interest within the cell. However, the molecular composition of these regions may change during the course of the acquisition process, limiting the spatial determination of the quantitative values of the apparent FRET efficiencies to an average over entire cells. By means of a spectrally resolved two-photon microscope, we are able to obtain a full set of spectrally resolved images after only one complete excitation scan of the sample of interest. From this pixel-level spectral data, a map of FRET efficiencies throughout the cell is calculated. By applying a simple theory of FRET in oligomeric complexes to the experimentally obtained distribution of FRET efficiencies throughout the cell, a single spectrally resolved scan reveals stoichiometric and structural information about the oligomer complex under study. Here we describe the procedure of preparing biological cells (the yeast Saccharomyces cerevisiae) expressing membrane receptors (sterile 2 &alpha;-factor receptors) tagged with two different types of fluorescent probes. Furthermore, we illustrate critical factors involved in collecting fluorescence data using the spectrally resolved two-photon microscopy imaging system. The use of this protocol may be extended to study any type of protein which can be expressed in a living cell with a fluorescent marker attached to it.

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

By employing a spectrally resolved two-photon microscopy imaging system, pixel-level maps of F&ouml;rster Resonance Energy Transfer (FRET) efficiencies are obtained for cells expressing membrane receptors hypothesized to form homo-oligomeric complexes. From the FRET efficiency maps, we are able to estimate stoichiometric information about the oligomer complex under study.

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial ...

Biology

Validated Immunochemical Assay for Comprehensive Determination of the Human Epidermal Growth Factor Receptor 2 Released from and Bound to Cells

Human epidermal growth factor receptor 2 (HER2) is a well-established cancer marker. It became a very successful diagnostic and therapeutic target, especially in breast cancer and other HER2-expressing cancer types. In the clinic, the gold-standard immunohistochemical diagnostic methods employing the specific anti-HER2 antibodies are used to measure the expression level of the membrane-bound receptor. The soluble extracellular domain (ECD) of HER2 that is released from the overexpressing cells circulates in the blood and can reflect the tissue expression of the receptor. There is a need for accurate and validated assays to correlate the concentration of the circulating HER2 protein with disease clinical manifestations.
Our team has developed and validated the novel sandwich enzyme-linked immunosorbent assay (ELISA) for quantification of the membrane-bound and the released from cells ECD domain of HER2. The assay uses two unique monoclonal antibodies specific to HER2 developed previously. The quantitation range includes HER2 concentration from 1.56-100 ng/mL, which is expected for cancer cells cultured in vitro and shows sensitivity at the level of 0.5 ng/mL. The satisfactory intra- and inter-assay precision and accuracy of the method make it applicable for HER2 quantification in various types of biological samples, including cell culture medium, serum, and solid tumor tissue. Here, we focus on the comprehensive determination of the receptor-associated and secreted by the in vitro cultured cancer cells. The paper presents a step-by-step protocol for the quantification of HER2 protein that can be employed for testing a variety of cell lines, blood, and tissues.

We present a validated sandwich ELISA assay using novel anti-HER2 monoclonal antibodies. This assay enables precise quantification of cell-bound and released HER2 protein from in vitro cultured cells and other samples, including blood and tissues.

Human epidermal growth factor receptor 2 (HER2) is a well-established cancer marker. It became a very successful diagnostic and therapeutic target, ...

Cancer Research

An Analytical Tool that Quantifies Cellular Morphology Changes from Three-dimensional Fluorescence Images

The most common software analysis tools available for measuring fluorescence images are for two-dimensional (2D) data that rely on manual settings for inclusion and exclusion of data points, and computer-aided pattern recognition to support the interpretation and findings of the analysis. It has become increasingly important to be able to measure fluorescence images constructed from three-dimensional (3D) datasets in order to be able to capture the complexity of cellular dynamics and understand the basis of cellular plasticity within biological systems. Sophisticated microscopy instruments have permitted the visualization of 3D fluorescence images through the acquisition of multispectral fluorescence images and powerful analytical software that reconstructs the images from confocal stacks that then provide a 3D representation of the collected 2D images. Advanced design-based stereology methods have progressed from the approximation and assumptions of the original model-based stereology1 even in complex tissue sections2. Despite these scientific advances in microscopy, a need remains for an automated analytic method that fully exploits the intrinsic 3D data to allow for the analysis and quantification of the complex changes in cell morphology, protein localization and receptor trafficking. 
Current techniques available to quantify fluorescence images include Meta-Morph (Molecular Devices, Sunnyvale, CA) and Image J (NIH) which provide manual analysis. Imaris (Andor Technology, Belfast, Northern Ireland) software provides the feature MeasurementPro, which allows the manual creation of measurement points that can be placed in a volume image or drawn on a series of 2D slices to create a 3D object. This method is useful for single-click point measurements to measure a line distance between two objects or to create a polygon that encloses a region of interest, but it is difficult to apply to complex cellular network structures. Filament Tracer (Andor) allows automatic detection of the 3D neuronal filament-like however, this module has been developed to measure defined structures such as neurons, which are comprised of dendrites, axons and spines (tree-like structure). This module has been ingeniously utilized to make morphological measurements to non-neuronal cells3, however, the output data provide information of an extended cellular network by using a software that depends on a defined cell shape rather than being an amorphous-shaped cellular model. To overcome the issue of analyzing amorphous-shaped cells and making the software more suitable to a biological application, Imaris developed Imaris Cell. This was a scientific project with the Eidgen&ouml;ssische Technische Hochschule, which has been developed to calculate the relationship between cells and organelles. While the software enables the detection of biological constraints, by forcing one nucleus per cell and using cell membranes to segment cells, it cannot be utilized to analyze fluorescence data that are not continuous because ideally it builds cell surface without void spaces. To our knowledge, at present no user-modifiable automated approach that provides morphometric information from 3D fluorescence images has been developed that achieves cellular spatial information of an undefined shape (Figure 1). 
We have developed an analytical platform using the Imaris core software module and Imaris XT interfaced to MATLAB (Mat Works, Inc.). These tools allow the 3D measurement of cells without a pre-defined shape and with inconsistent fluorescence network components. Furthermore, this method will allow researchers who have extended expertise in biological systems, but not familiarity to computer applications, to perform quantification of morphological changes in cell dynamics.

We developed a software platform that utilizes Imaris Neuroscience, ImarisXT and MATLAB to measure the changes in morphology of an undefined shape taken from three-dimensional confocal fluorescence of single cells. This novel approach can be used to quantify changes in cell shape following receptor activation and therefore represents a possible additional tool for drug discovery.

The most common software analysis tools available for measuring fluorescence images are for two-dimensional (2D) data that rely on manual settings for ...

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 ...

Confocal Microscopy Reveals Cell Surface Receptor Aggregation Through Image Correlation Spectroscopy

Confocal microscopy provides an accessible methodology to capture sub-cellular interactions critical for the characterization and further development of pre-clinical agents labeled with fluorescent probes. With recent advancements in antibody based cytotoxic drug delivery systems, understanding the alterations induced by these agents within the realm of receptor aggregation and internalization is of critical importance. This protocol leverages the well-established methodology of fluorescent immunocytochemistry and the open source FIJI distribution of ImageJ, with its inbuilt autocorrelation and image mathematical functions, to perform spatial image correlation spectroscopy (ICS). This protocol quantitates the fluorescent intensity of labeled receptors as a function of the beam area of the confocal microscope. This provides a quantitative measure of the state of target molecule aggregation on the cell surface. This methodology is focused on the characterization of static cells with potential to expand into temporal investigations of receptor aggregation. This protocol presents an accessible methodology to provide quantification of clustering events occurring at the cell surface, utilizing well established techniques and non-specialized imaging apparatus.

Antibodies that bind to target receptors on the cell surface can confer conformation and clustering alterations. These dynamic changes have implications for characterizing drug development in target cells. This protocol utilizes confocal microscopy and image correlation spectroscopy through ImageJ/FIJI to quantify the extent of receptor clustering on the cell surface.

Confocal microscopy provides an accessible methodology to capture sub-cellular interactions critical for the characterization and further development of ...

Measuring Proliferation of Vascular Smooth Muscle Cells Using Click Chemistry

The ability of a cell to proliferate is integral to the normal function of most cells, and dysregulation of proliferation is at the heart of many disease processes. For these reasons, measuring proliferation is a common tool used to assess cell function. Cell proliferation can be measured simply by counting; however, this is an indirect means of measuring proliferation. One common means of directly detecting cells preparing to divide is by incorporation of labeled nucleoside analogs. These include the radioactive nucleoside analog 3H-thymidine plus non-radioactive nucleoside analogs such as 5-bromo-2&#8217; deoxyuridine (BrdU) and 5-ethynyl-2&#8242;-deoxyuridine (EdU). Incorporation of EdU is detected by click chemistry, which has several advantages when compared to BrdU. In this report, we provide a protocol for measuring proliferation by the incorporation of EdU. This protocol includes options for various readouts, along with the advantages and disadvantages of each. We also discuss places where the protocol can be optimized or altered to meet the specific needs of the experiment planned. Finally, we touch on the ways that this basic protocol can be modified for measuring other cell metabolites.

Proliferation is a critical part of cellular function, and a common readout used to assess potential toxicity of new drugs. Measuring proliferation is, therefore, a frequently used assay in cell biology. Here we present a simple, versatile method of measuring proliferation that can be used in adherent and non-adherent cells.

The ability of a cell to proliferate is integral to the normal function of most cells, and dysregulation of proliferation is at the heart of many disease ...

Oligomerization Dynamics of Cell Surface Receptors in Living Cells by Total Internal Reflection Fluorescence Microscopy Combined with Number and Brightness Analysis

Despite the importance and ubiquity of receptor oligomerization, few methods are applicable for detecting clustering events and measuring the degree of clustering. Here, we describe an imaging approach to determine the average oligomeric state of mEGFP-tagged-receptor homocomplexes in the membrane of living cells. The protocol is based on Total Internal Reflection Fluorescence (TIRF) microscopy combined with Number and Brightness (N&#38;B) analysis. N&#38;B is a method similar to fluorescence-correlation spectroscopy (FCS) and photon counting histogram (PCH), which are based on the statistical analysis of the fluctuations of the fluorescence intensity of fluorophores diffusing in and out of an illumination volume during an observation time. In particular, N&#38;B is a simplification of PCH to obtain information on the average number of proteins in oligomeric mixtures. The intensity fluctuation amplitudes are described by the molecular brightness of the fluorophore and the average number of fluorophores within the illumination volume. Thus, N&#38;B considers only the first and second moments of the amplitude distribution, namely, the mean intensity and the variance. This is, at the same time, the strength and the weakness of the method. Because only two moments are considered, N&#38;B cannot determine the molar fraction of unknown oligomers in a mixture, but it only estimates the average oligomerization state of the mixture. Nevertheless, it can be applied to relatively small time series (compared to other moment methods) of images of live cells on a pixel-by-pixel basis, simply by monitoring the time fluctuations of the fluorescence intensity. It reduces the effective time-per-pixel to a few microseconds, allowing acquisition in the time range of seconds to milliseconds, which is necessary for fast oligomerization kinetics. Finally, large cell areas as well as sub-cellular compartments can be explored.

We describe an imaging approach for the determination of the average oligomeric state of mEGFP-tagged-receptor oligomers induced by ligand binding in the plasma membrane of living cells. The protocol is based on Total Internal Reflection Fluorescence (TIRF) microscopy combined with Number and Brightness (N&#38;B) analysis.

Despite the importance and ubiquity of receptor oligomerization, few methods are applicable for detecting clustering events and measuring the degree of ...

Graphene Enclosure of Chemically Fixed Mammalian Cells for Liquid-Phase Electron Microscopy

A protocol is described for investigating the human epidermal growth factor receptor 2 (HER2) in the intact plasma membrane of breast cancer cells using scanning transmission electron microscopy (STEM). Cells of the mammalian breast cancer cell line SKBR3 were grown on silicon microchips with silicon nitride (SiN) windows. Cells were chemically fixed, and HER2 proteins were labeled with quantum dot nanoparticles (QDs), using a two-step biotin-streptavidin binding protocol. The cells were coated with multilayer graphene to maintain a hydrated state, and to protect them from electron beam damage during STEM. To examine the stability of the samples under electron beam irradiation, a dose series experiment was performed. Graphene-coated and non-coated samples were compared. Beam induced damage, in the form of bright artifacts, appeared for some non-coated samples at increased electron dose D, while no artifacts appeared on coated samples.

Presented here is a protocol for labeling membrane proteins in mammalian cells and coating the sample with graphene for liquid-phase scanning transmission electron microscopy. The stability of the samples against the damage caused by radiation can also studied with this protocol.

A protocol is described for investigating the human epidermal growth factor receptor 2 (HER2) in the intact plasma membrane of breast cancer cells using ...

Tension Gauge Tether Probes for Quantifying Growth Factor Mediated Integrin Mechanics and Adhesion

Multicellular organisms rely on interactions between membrane receptors and cognate ligands in the surrounding extracellular matrix (ECM) to orchestrate multiple functions, including adhesion, proliferation, migration, and differentiation. Mechanical forces can be transmitted from the cell via the adhesion receptor integrin to ligands in the ECM. The amount and spatial organization of these cell-generated forces can be modulated by growth factor receptors, including epidermal growth factor receptor (EGFR). The tools currently available to quantify crosstalk-mediated changes in cell mechanics and relate them to&#160;focal adhesions, cellular morphology, and signaling are limited. DNA-based molecular force sensors known as tension gauge tethers (TGTs) have been employed to quantify these changes. TGT probes are unique in their ability to both modulate the underlying force threshold and report piconewton scale receptor forces across the entire adherent cell surface at diffraction-limited spatial resolution. The TGT probes used here rely on the irreversible dissociation of a DNA duplex by receptor-ligand forces that generate a fluorescent signal. This allows quantification of the cumulative integrin tension (force history) of the cell. This article describes a protocol employing TGTs to study the impact of EGFR on integrin mechanics and adhesion formation. The assembly of&#160;the TGT mechanical sensing platform is systematically detailed and the procedure to image forces, focal adhesions, and cell spreading is outlined. Overall, the ability to modulate the underlying force threshold of the probe, the adhesion ligand, and the type and concentration of growth factor employed for stimulation make this a robust platform for studying the interplay of diverse membrane receptors in regulating integrin-mediated forces.

TGT surface is an innovative platform to study growth factor-integrin crosstalk. The flexible probe design, specificity of the adhesion ligand, and precise modulation of stimulation conditions allow robust quantitative assessments of EGFR-integrin interplay. The results highlight EGFR as a 'mechano-organizer' tuning integrin mechanics, influencing focal adhesion assembly and cell spreading.

Multicellular organisms rely on interactions between membrane receptors and cognate ligands in the surrounding extracellular matrix (ECM) to orchestrate ...

Studying the Stoichiometry of Epidermal Growth Factor Receptor in Intact Cells using Correlative Microscopy

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