We would like to thank members of Temple Bioengineering for valuable discussions. We thank David Ambrose at the flow cytometry core (Lewis Katz School of Medicine) for his assistance with cell sorting and Tony Boehm from the IDEAS Hub (College of Engineering, Temple University) for help with the 3D printing. We also thank our funding resources: American Cancer Society Research Scholar Grant 134415-RSG-20-034-01-CSM, Conquer Cancer Now / Young Investigator Award, National Institutes of Health, R00 CA172360 and R01 CA230777, all to BG.

1. Fabrication of a Spheroid Imaging Device (SID) to optimize spheroid embedding (Duration 1 day)
<ol>
	<li>Create the spacer using a 3D printer (Figure 1A, B&#160;and Supplementary File 1).</li>
	<li>Weigh out a 10:1 (wt/wt) ratio of base polymer:crosslinker in a plastic cup [e.g., 20 g of ethylbenzene base polymer and 2 g of silicone resin crosslinker to create polydimethylsiloxane (PDMS)].</li>
	<li>Thoroughly mix the PDMS solution in the plastic cup using a disposable pipette.</li>
	<li>Place the plastic cup into a vacuum chamber to remove the air bubbles from the mixture. Quickly release the vacuum pressure to remove the small amount of air trapped at the surface of the mixture and dissipate the remaining air bubbles.</li>
	<li>Incubate the 3D printed spacer at 100 &#176;C for 5 min to increase its flexibility.</li>
	<li>Clean the two glass plates that will be used to construct the PDMS mold thoroughly by wiping them with 100% isopropanol. If the glass plates were previously used to cast PDMS, be sure to remove any remaining old PDMS by gently scraping the glass plates with a razor blade and wiping them with 100% isopropanol.</li>
	<li>Construct the mold by placing the 3D printed spacer flush in-between the two clean glass plates.</li>
	<li>Seal the mold using large binder clips on the outside edges of the glass plates. Place two binder clips on the bottom edge and one on the top corner.</li>
	<li>Inspect the top part of the mold to ensure that the spacer is flush with the glass plates. This guarantees that no deformations will be present and that an even sheet of PDMS will be created. 
	NOTE: If the resulting sheet is not of uniform thickness, the clips need to be adjusted slightly.</li>
	<li>Cut the tip of a disposable pipette (~2 cm from the tip) and add the PDMS mixture at a slow and constant rate to the top left corner of the mold. Pour the mixture slowly to prevent the creation of large air pockets.</li>
	<li>Place the mold into a vacuum chamber to remove air bubbles that formed during the pour.</li>
	<li>Cure the PDMS by incubating the mold at 100 &#176;C for 1 h.</li>
	<li>Retrieve the mold from the incubator and allow it to cool to touch.</li>
	<li>Remove the binder clips and glass plates from the spacer containing the cured PDMS.</li>
	<li>Use a razor blade to cut through the seal that was created on all four sides of the mold between the spacer and the glass plate. With all four sides cut into, begin pulling apart the mold to reveal the PDMS sheet in the spacer.</li>
	<li>Carefully peel the new PDMS sheet off of the spacer using tweezers.</li>
	<li>On a cutting mat, punch out 17.5 mm diameter PDMS disks from the sheet, and then punch three evenly distributed 5.5 mm diameter holes, using different size biopsy punches. Here these 3-hole PDMS disks are referred as &#8220;inserts&#8221; (Figure 1C). 
	NOTE: Nonuniform cuts made into the PDMS, or a faulty bind via plasma treatment, may result in the future leaks.</li>
	<li>Clean each insert by gently removing any dust particles using tape.</li>
	<li>Stick the inserts onto a piece of double-sided tape and wrap the tape around the lid of a 10 cm Petri dish.</li>
	<li>Place the lid of the Petri dish in the plasma machine, along with the open 35 mm glass bottom dishes.</li>
	<li>Activate the surface of the inserts and of the glass via plasma treatment for 1 min at 300 mTorr. A hand-held plasma wand can be used.</li>
	<li>Using tweezers, quickly attach the upside, i.e., treated side, of one insert (Figure 1D) to the glass part of a glass bottom dish (Figure 1E). Repeat for all dishes.</li>
	<li>Use pointer finger and thumb to apply an even pressure while rotating the glass bottom dish. This will ensure stable securement of the insert to the glass bottom dish.</li>
	<li>Incubate the SIDs (Figure 1F) at 60 &#176;C for 20 min to strengthen the adhesion between glass and PDMS.</li>
	<li>Perform a second round of plasma treatment on the SIDs, using the same settings as in step 1.21 or with the hand-held wand. This will render the free PDMS surface adhere to the poly-L-lysine in the coating solution (see 1.26).</li>
	<li>Freshly prepare the following solutions.
	<ol>
		<li>Coating solution: 1x PBS containing 0.01% (vol/vol) poly-L-Lysine [e.g., add 10 &#181;L of 0.1% (vol/vol) poly-L-Lysine to 90 &#181;L of 1x PBS].</li>
		<li>Crosslinking solution: Distilled water containing 1x (vol/vol) glutaraldehyde [e.g., add 10 &#181;L of 10x glutaraldehyde to 90 &#181;L of distilled water].</li>
		<li>Storage solution: 1x PBS containing 10x (vol/vol) Penicillin-Streptomycin [e.g., add 1 mL of 100x Penicillin-Streptomycin to 9 mL of 1x PBS].</li>
	</ol>
	</li>
	<li>Add 35 &#181;L of coating solution per each hole and incubate for 1 h at room temperature.</li>
	<li>Aspirate the poly-L-Lysine and rinse the entire SID 3 times with distilled water.</li>
	<li>Add 35 &#181;L of crosslinking solution per each hole incubate for 30 min at room temperature.</li>
	<li>Aspirate the crosslinking solution and rinse the entire SID 3 times with distilled water.</li>
	<li>Add 70% ethanol into each SID and place under ultraviolet (UV) light for 30 min.</li>
	<li>Under the hood, aspirate ethanol and rinse the SID 3 times with distilled water.</li>
	<li>Add 2.5 mL of storage solution per each SID. 
	NOTE: At this stage, the SIDs can be stored at 4 &#176;C for a week. It was observed that the binding strength between PDMS and glass decreases over time. Beyond a week, users are recommended to test the SIDs for potential leakage before use. To do so, aspirate the storage solution and add 35 &#181;L of 1x PBS into each hole. After use, PDMS inserts can be peeled off the glass bottom dishes. To achieve optimal cleaning of the glass bottom dishes, several washes with isopropanol and hydrochloric acid should be used to remove PDMS residues.</li>
</ol>
2. Spheroid formation and embedding into collagen (Duration 4 days)
NOTE: For live imaging of spheroids, longitudinally or in time-lapse videos, use a cell line expressing a cytoplasmic and/or nuclear fluorescent protein. If such a cell line is available, follow the steps described in this section. Alternatively, in the section 3, a protocol is proposed to label cancer cells in spheroids using a cytoplasmic dye.
<ol>
	<li>Form spheroids of 4T1 and/or 67NR cells using the hanging drop technique, as previously described4,5,6,7, with 3,000 cells/40 &#181;L droplet and a 3-day incubation time. Add the bovine atelocollagen I solution last and maintain all solutions on ice, at all time.</li>
	<li>Identify the correctly formed spheroids using a bright-field microscope.</li>
	<li>Fill a 15 mL conical tube with 8 mL of pre-warmed complete medium.</li>
	<li>Collect spheroids with a P1000 pipette and transfer to the 15 mL conical tube. Wet the pipette tip by pipetting some complete medium in and out to prevent spheroids from sticking to the inside walls of the pipette tip and limit spheroid loss.</li>
	<li>Allow spheroids to sink to the bottom of the tube and carefully wash the spheroids by exchanging the medium. Repeat twice.</li>
	<li>Prepare a 5 mg/mL collagen I solution according to the manufacturer&#8217;s recommendations (alternate gelation procedure, see exemplary calculation below). Replace the distilled water with complete medium. When calculating the volume of medium to be used, take into account that spheroids will be added in 20 &#181;L of complete medium [e.g., for one SID, 3 x 30 + (3 x 30) x 20 % = 108 &#181;L of 5 mg/mL collagen I is needed (prepare 20 % extra to account for pipetting loss when handling viscous fluids); 22 &#181;L of complete medium + 10.8 &#181;L of 10x PBS + 1.2 &#181;L of 1 M NaOH + 54 &#181;L of 10 mg/mL collagen I stock].</li>
	<li>Collect all spheroids in 20 &#181;L of medium, using a P200 pipette, and add them to the collagen I solution prepared in step 2.6.</li>
	<li>Slowly mix the solution by pipetting up and down to avoid heterogeneity in the collagen I concentration, limiting bubble formation. Keep the solution on ice.</li>
	<li>Remove the storage solution from the SID(s) and wash 3 times with 3 mL of 1x PBS. After the final wash, leave the SID(s) dry so not to dilute the collagen I solution.</li>
	<li>Start a timer and dispense 30 &#181;L of the collagen I solution containing one spheroid into one of the three holes of the SID. Visually ensure that a single spheroid is contained in the 30 &#181;L.</li>
	<li>Repeat step 2.10 twice more to fill all the 3 holes of a SID.</li>
	<li>Use a 10 &#181;L pipette tip to re-center the spheroid if it is located close to the PDMS border. If two or three spheroids end up dispensed in one of the holes, same pipette tip can be used to separate the spheroids from each other. Stop the timer. 
	NOTE: Frequently inverting of the SID upside-down and upside-up, throughout the period of the collagen I polymerization and solidification, ensures that the spheroid is positioned at the vertical center of the ECM layer, and prevents invasion of cells in 2D. The frequency of inverting (flipping) should be maximized. As the flipping frequency is controlled by the spheroid &#8220;dispensing time&#8221; measured in 2.10-2.12, the dispensing time should be minimized. In our lab, dispensing time is 2 min on average.</li>
	<li>To vertically center the spheroid in the collagen layer, flip the SID upside-down and incubate at 37 &#176;C for dispensing time.</li>
	<li>Flip the SID upside-up and incubate at 37 &#176;C for dispensing time. 
	NOTE: The length of time that the spheroids spend in the upside-down orientation should be equal to the time spent in the upside-up orientation.</li>
	<li>Repeat steps 2.13 and 2.14 for 30 min, until the collagen I polymerizes.</li>
	<li>Add 2.5 mL of complete medium/SID and if needed, acquire an image of the spheroids for the initial timepoint.</li>
	<li>Repeat steps 2.10-2.16 if multiple SIDs are used.</li>
</ol>
3. Fluorescence labeling of spheroids
<ol>
	<li>Live imaging (Duration 6-7 days) 
	NOTE: If a cell line expressing a cytoplasmic and/or nuclear fluorescent protein is available, follow the steps described in the section 2. Alternatively, for cytoplasmic labeling, the following protocol is proposed.
	<ol>
		<li>Follow steps 2.1-2.5 of the protocol described in the section 2.</li>
		<li>Dilute the cytoplasmic dye to 25 &#181;M in 200 &#181;L of serum-free medium. 
		NOTE: While a red cytoplasmic dye is used in this experiment, any other available color should be suitable. Cancer cells were labeled inside spheroids using nuclear dyes diluted to 20 &#181;M in 200 &#181;L of serum-free medium (see Table of Materials).</li>
		<li>After the final wash, resuspend spheroids in the cytoplasmic or nuclear dye solution and incubate at room temperature for 20 min, protected from light. 
		NOTE: With this approach, labeling of the cells in the spheroid center will not be efficient. To achieve labeling of all cells, incubation can be extended overnight, by placing the dish on a rocker. Alternatives are described in Discussion.</li>
		<li>Wash spheroids 3 times in complete medium.</li>
		<li>Proceed to steps 2.6-2.17 of the protocol described in the section 2.</li>
		<li>Image spheroids via time-lapse imaging every 10 min for 24-72 h (Figure 2), or via longitudinal imaging, daily, for up to 7 days (Figure 3). Use a laser scanning confocal microscope, 10x air objective (0.4 numerical aperture and 3.1 mm working distance), 1024 x 1024 pixels, exposure time 8 &#181;s/pixel, pinhole 90 &#181;m, 6 x 15 &#181;m z-steps and 3 fields of view-each containing one spheroid. 
		NOTE: For time-lapse imaging, use minimal laser power and equip the microscope with an environmental chamber with temperature, humidity and gas control. Culture the embedded spheroids at 37 &#176;C for &#62; 8 h or overnight prior to imaging to minimize the time on the microscope.</li>
	</ol>
	</li>
	<li>Immunofluorescence staining of spheroids (Duration 2 days) 
	NOTE: The procedure described here is adapted and optimized from previously published protocols8,9. This method can be used after the protocol outlined in the sections 2 and 3.1.
	<ol>
		<li>Freshly prepare the following solutions.
		<ol>
			<li>Fixing solution: 1x PBS containing 4% PFA (vol/vol) [e.g., add 5 mL of 16% (vol/vol) PFA to 15 mL of 1x PBS].</li>
			<li>Fixing and permeabilizing solution: 1x PBS containing 4% PFA (vol/vol) and 0.5% Triton X-100 (vol/vol) [e.g., add 50 &#181;L of Triton X-100 to 10 mL of fixing solution].</li>
			<li>Blocking solution: 1x PBS containing 1% FBS (vol/vol) and 1% BSA (wt/vol) [e.g., dissolve 100 mg of BSA in 10 mL of 1x PBS, then add 100 &#181;L of FBS].</li>
			<li>Washing solution: 1x PBS containing 0.05% Tween 20 (vol/vol) [e.g., add 25 &#181;L of Tween 20 to 50 mL of 1x PBS].</li>
		</ol>
		</li>
		<li>Remove the culture medium from the SID(s) and wash once with warm 1x PBS.</li>
		<li>Add 2 mL of fixing and permeabilizing solution per SID and incubate at room temperature for 5 min.</li>
		<li>Remove the fixing and permeabilizing solution and add 2 mL of fixing solution per SID and incubate at room temperature for 20 min.</li>
		<li>Wash 3 times with the washing solution.</li>
		<li>Add 2 mL of blocking solution per SID and incubate at 4 &#176;C for 24 h with mild shaking. 
		NOTE: At this step, samples can be incubated at 4 &#176;C, over the weekend. Please note that a longer blocking time might interfere with the immunofluorescence labeling procedure.</li>
		<li>Dilute primary antibodies in blocking solution.</li>
		<li>Add 150 &#181;L of blocking solution with primary antibodies/well in a 48-well plate.</li>
		<li>Using fine tweezers, carefully detach the plug of collagen I containing a spheroid and transfer into a well. Repeat if multiple spheroids are labeled. Wipe any liquid that might remain on the tweezers when working with different antibodies, to prevent contamination.</li>
		<li>Incubate overnight at 4 &#176;C with mild shaking.</li>
		<li>Add 300 &#181;L of washing solution to empty and full wells.</li>
		<li>Carefully transfer the plug of collagen I containing a spheroid into a &#8220;wash&#8221; well.</li>
		<li>Wash 3 times with washing solution for 2 h, at room temperature and with mild shaking. 
		NOTE: Transferring the plugs of collagen I containing a spheroid, instead of aspirating the solutions, can help reducing sample loss and sample damage.</li>
		<li>Dilute secondary antibodies, 4&#8217;,6-diamidino-2-phenylindole (DAPI) and/or phalloidin in blocking solution.</li>
		<li>Add 150 &#181;L of blocking solution with secondary antibodies, DAPI and/or phalloidin per well.</li>
		<li>Using tweezers, carefully transfer the collagen plug containing the spheroids into the well. Repeat if multiple spheroids are labeled.</li>
		<li>Incubate at room temperature for 1 h with mild shaking.</li>
		<li>Repeat steps 3.2.11 and 3.2.12.</li>
		<li>Wash 3 times with washing solution for 30 min, at room temperature with mild shaking.</li>
		<li>Using a razor blade, cut a 3-hole PDMS insert into three parts so that each part contains a hole.</li>
		<li>Place two pieces of PDMS onto a microscope slide.</li>
		<li>Add a drop of mounting solution using a P200 tip with the end cut off. Prevent bubble formation.</li>
		<li>Carefully transfer a collagen I plug into each hole. 
		NOTE: If the spheroid was positioned at the top of the collagen plug, invert the collagen plug so that the spheroid ends up closer to the glass coverslip.</li>
		<li>Place a coverslip on top and seal using tape.</li>
		<li>Let the sample dry at room temperature for 10 min, protected from light.</li>
		<li>Store samples at 4 &#176;C, protected from light until imaging.</li>
	</ol>
	</li>
</ol>
4. Image processing to analyze cancer invasion over time
NOTE: The format required for this macro is a single-channel x,y,t image saved as a .tiff file.
<ol>
	<li>If multiple z-slices were acquired (i.e. x,y,z,t image), open the image in Fiji and select Image | Stacks | Z Project. It is recommended to use the Max Intensity option. Alternatively, a single z-slice can be used to run the macro. Save the image as a .tiff file.</li>
	<li>Create a separate &#8220;Processing&#8221; folder on the Desktop.</li>
	<li>Select File | Save As | Image Sequence. Use the TIFF format, update the digits number according to the number of frames, check the box to use slice label as file name and select Ok. Select the &#8220;Processing&#8221; folder created in step 2.</li>
	<li>Open one image from the &#8220;Processing&#8221; folder. It should be an x,y image, corresponding to a single timepoint.</li>
	<li>Select Image | Adjust | Auto Threshold. In the dropdown menu select the method Try all and check the box for white objects on black background.</li>
	<li>A montage image appears showing the result for each automated thresholding method.</li>
	<li>Identify the best automated thresholding method, for example RenyIEntropy.</li>
	<li>To confirm the choice for the automated thresholding method, open any other image from the &#8220;Processing&#8221; folder and test the chosen thresholding method using Image | Adjust | Auto Threshold.</li>
	<li>Close all images.</li>
	<li>Download the SpheroidAreaTime macro (Supplementary File 2).</li>
	<li>Open the Fiji macro by drag-and-drop.</li>
	<li>In line 58, update the automated thresholding method as needed.</li>
	<li>In line 62, update the size range according to the cell dimensions.</li>
	<li>Select Run.</li>
	<li>Select the &#8220;Processing&#8221; folder and type in &#8220;Processing&#8221; as the Parent folder, then select Ok.</li>
	<li>Once the run is over, save the table &#8220;Summary&#8221;. The first column indicates the name of the image; the third column indicates the spheroid area; the sixth, seventh and eight columns specify the parameters for an ellipse fitted onto the spheroid.</li>
	<li>The &#8220;Processing&#8221; folder now contains a processed image for each time point, with the extension _SpheroidArea. 
	NOTE: If the spheroid center is dim, fill it with white using the selection tools, for all timepoints, and then proceed to step 3. Similarly, the space around the spheroid can be filled with black to &#8220;clean&#8221; the image. If the image is clean, comment the line 61 to speed up the run.</li>
</ol>

<ol>
	<li>Hanahan, D., Weinberg, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hallmarks+of+cancer:+The+next+generation.">Hallmarks of cancer: The next generation.</a> Cell. 144, 646-674 (2011).</li><li>Noone, A., 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=.">.</a> SEER Cancer statistics review 1975-2015, based on November 2017 SEER data submission. , (2019).</li><li>Yamada, K. M., Cukierman, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+tissue+morphogenesis+and+cancer+in+3D.">Modeling tissue morphogenesis and cancer in 3D.</a> Cell. 130, 601-610 (2007).</li><li>Foty, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Simple+hanging+drop+cell+culture+protocol+for+generation+of+3D+spheroids.">A Simple hanging drop cell culture protocol for generation of 3D spheroids.</a> Journal of Visualized Experiment. , e2720 (2011).</li><li>Berens, E. B., Holy, J. M., Riegel, A. T., Wellstein, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+cancer+cell+spheroid+assay+to+assess+invasion+in+a+3D+setting.">A cancer cell spheroid assay to assess invasion in a 3D setting.</a> Journal of Visualized Experiments. 2015, (2015).</li><li>T&#246;nisen, F., 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=EP4+receptor+promotes+invadopodia+and+invasion+in+human+breast+cancer.">EP4 receptor promotes invadopodia and invasion in human breast cancer.</a> European Journal of Cell Biology. 96, 218-226 (2017).</li><li>Bayarmagnai, B., 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=Invadopodia-mediated+ECM+degradation+is+enhanced+in+the+G1+phase+of+the+cell+cycle.">Invadopodia-mediated ECM degradation is enhanced in the G1 phase of the cell cycle.</a> Journal of Cell Sciences. 132, 227116 (2019).</li><li>Dekkers, J. F., 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=High-resolution+3D+imaging+of+fixed+and+cleared+organoids.">High-resolution 3D imaging of fixed and cleared organoids.</a> Nature Protocols. 14, 1756-1771 (2019).</li><li>Cukierman, E., Pankov, R., Stevens, D. R., Yamada, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Taking+cell-matrix+adhesions+to+the+third+dimension.">Taking cell-matrix adhesions to the third dimension.</a> Science. 294, 1708-1712 (2001).</li><li>Boutin, M. E., 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=A+high-throughput+imaging+and+nuclear+segmentation+analysis+protocol+for+cleared+3D+culture+models.">A high-throughput imaging and nuclear segmentation analysis protocol for cleared 3D culture models.</a> Science Reports. 8, 11135 (2018).</li><li>Pampaloni, F., Richa, R., Ansari, N., Stelzer, E. H. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+spheroid+formation+recorded+with+light+sheet-based+fluorescence+microscopy.">Live spheroid formation recorded with light sheet-based fluorescence microscopy.</a> Methods in Molecular Biology. 1251, 43-57 (2015).</li><li>Marcello, M., Richards, R., Mason, D., S&#233;e, V., Dmitriev, R. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+cell+invasion+using+a+multicellular+spheroid+model+and+light-sheet+microscopy.">Live imaging of cell invasion using a multicellular spheroid model and light-sheet microscopy.</a> Advances in Experimental Medicine and. , 155-161 (2017).</li><li>Tinevez, J. Y., 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=TrackMate:+An+open+and+extensible+platform+for+single-particle+tracking.">TrackMate: An open and extensible platform for single-particle tracking.</a> Methods. 115, 80-90 (2017).</li></ol>

The authors have nothing to disclose.

The 3D printed spacer was designed to create 1-mm thick sheets of PDMS that can then be used to easily create various shapes of PDMS, as required by the experimental applications. Due to the simplicity of its fabrication and the freedom to alter the design, this method of PDMS casting was chosen for the initial design of the SID. If high volume of SIDs is required, production can be made more efficient by creating a 3D-printed mold, which already contains PDMS disks with three equally spaced holes, and reducing the proce...

During cancer progression, cancer cells can acquire a motile and invasive phenotype, enabling them to escape the tumor mass and invade into the surrounding tissues1. Eventually, these invasive cancer cells can reach and grow inside secondary organs, a process called cancer metastasis1. Metastasis causes more than 90% of cancer-related deaths2. One reason for this is that, while localized tumors are clinically manageable, no efficient methods exist for the elimination of invasive cancer cells once metastatic spreading has occurred. Therefore, the emergence of invasive cancer cells and the transition from a localized to an invasive disease is posing a major clinical challenge. Determining how cancer cells initiate and sustain an invasive behavior may lead to the development of novel potent therapies.
The 3D spheroid model is an ideal platform to investigate the motile behavior of cancer cells under controlled, yet physiologically relevant conditions3. Indeed, in this assay, spheroids of cancer cells are embedded inside extracellular matrix (ECM), for example collagen I, which mimics a simplified tumor. Then, imaging is used to visualize the invasion of cancer cells from the spheroid into the collagen matrix. However, multiple challenges limit this procedure.
The first challenge occurs at the embedding step, where the liquid collagen matrix can spread across the dish surface, causing the spheroid to touch the bottom of the dish. Consequently, cells from the spheroid spread on the two-dimensional (2D) surface, breaking the three-dimensional (3D) spheroid morphology. Increasing the volume of collagen is an efficient, but costly solution. To prevent cells from spreading on the 2D surface, while maintaining a minimal volume of collagen, we developed a spheroid imaging device (SID) by bounding a 1 mm-thick, 3-hole polydimethylsiloxane (PDMS) insert onto a glass bottom dish.
The second challenge of the spheroid assay is the labeling of cancer cells in spheroids, which is limited by the poor penetration of antibodies and fluorescent dyes, an effect that increases with the spheroid size. While the ideal solution for labeling cells is the establishment of cell lines stably expressing fluorescent protein(s), this option is mostly restricted to immortalized cell lines and is limited by the availability of fluorescent protein chimeras. Here, we describe an optimized protocol for immunofluorescence staining of fixed spheroids, as well as the efficient use of a cytoplasmic dye to label cells immediately before embedding the spheroid.
The third challenge of the spheroid assay is the lack of simple Fiji macros for semi-automated quantification of cell invasion over time. To address this challenge, we describe a simple methodology to analyze the spheroid area over time. We illustrate the advantages of this protocol using the 4T1 and 67NR cell lines as examples.

<table>
	<tbody>
		<tr>
			<td>1 N NaOH</td>
			<td>Honeywell Fluka</td>
			<td>60-014-44</td>
			<td></td>
		</tr>
		<tr>
			<td>10X Dulbecco&#8217;s phosphate-buffered saline (PBS)</td>
			<td>Gibco</td>
			<td>SH30028.LS</td>
			<td></td>
		</tr>
		<tr>
			<td>16% paraformaldehyde (PFA)</td>
			<td>Alfa Aesar</td>
			<td>43368-9M</td>
			<td></td>
		</tr>
		<tr>
			<td>1X Dulbecco&#8217;s phosphate-buffered saline (PBS)</td>
			<td>Gibco</td>
			<td>20012027</td>
			<td></td>
		</tr>
		<tr>
			<td>4&#8217;,6-diamidino-2-phenylindole (DAPI)</td>
			<td>Invitrogen</td>
			<td>D1306</td>
			<td></td>
		</tr>
		<tr>
			<td>48-well plate</td>
			<td>Falcon</td>
			<td>T1048</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 647 phalloidin</td>
			<td>Life Technologies</td>
			<td>A20006</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti cortactin antibody</td>
			<td>Abcam</td>
			<td>ab33333</td>
			<td>1 to 200 dilution</td>
		</tr>
		<tr>
			<td>Anti E-cadherin antibody</td>
			<td>Invitrogen</td>
			<td>13-1900</td>
			<td>1 to 100 dilution</td>
		</tr>
		<tr>
			<td>Bovine atelocollagen I solution (Nutragen)</td>
			<td>Advanced Biomatrix</td>
			<td>501050ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum albumin (BSA)</td>
			<td>Sigma Aldrich</td>
			<td>A4503-50G</td>
			<td></td>
		</tr>
		<tr>
			<td>CellTracker Red CMTPX Dye</td>
			<td>Invitrogen</td>
			<td>C34552</td>
			<td></td>
		</tr>
		<tr>
			<td>Conical tubes</td>
			<td>Falcon</td>
			<td>352095</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>FisherBrand</td>
			<td>12-548-5E</td>
			<td></td>
		</tr>
		<tr>
			<td>Disposable container</td>
			<td>Staples</td>
			<td></td>
			<td>Plastic cups</td>
		</tr>
		<tr>
			<td>Disposable transfer pipette</td>
			<td>Thermo Scientific</td>
			<td>202</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Fisher Scientific</td>
			<td>11965118</td>
			<td></td>
		</tr>
		<tr>
			<td>Double-faced tape</td>
			<td>Scotch</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Sigma Aldrich</td>
			<td>E7023-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>Bio-Techne</td>
			<td>S11550</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoromount-G</td>
			<td>eBioscience</td>
			<td>00-4958-02</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde</td>
			<td>Sigma Aldrich</td>
			<td>G5882-100mL</td>
			<td></td>
		</tr>
		<tr>
			<td>Hoescht nuclear stain</td>
			<td>Thermo Fischer</td>
			<td>62249</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropanol</td>
			<td>Thermo Fischer</td>
			<td>S25371A</td>
			<td></td>
		</tr>
		<tr>
			<td>MatTek dish (glass bottom dish)</td>
			<td>MatTek Corporation</td>
			<td>P35G-1.5-14-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Methyl cellulose</td>
			<td>Sigma Aldrich</td>
			<td>M6385-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>MilliQ water</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Penicilin/streptomycin solution</td>
			<td>Thermo Fischer</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Corning</td>
			<td>353003</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipet tips</td>
			<td>Fisherbrand</td>
			<td>02-707</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipets</td>
			<td>Gilson</td>
			<td>F167300</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-L-Lysine</td>
			<td>Sigma</td>
			<td>P8920</td>
			<td></td>
		</tr>
		<tr>
			<td>Primary antibodies, user specific</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Rat Tail Collagen I</td>
			<td>Corning</td>
			<td>47747-218</td>
			<td></td>
		</tr>
		<tr>
			<td>Razor Blade</td>
			<td>Personna</td>
			<td>74-0001</td>
			<td></td>
		</tr>
		<tr>
			<td>Secondary antibodies, user specific</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Slides</td>
			<td>Globe Scientific</td>
			<td>1354W-72</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard 184 Silicone</td>
			<td>Dow Corning</td>
			<td>4019862</td>
			<td></td>
		</tr>
		<tr>
			<td>Tape</td>
			<td>Scotch</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X100</td>
			<td>Sigma Aldrich</td>
			<td>10789704001</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween 20</td>
			<td>Sigma Aldrich</td>
			<td>655204-100ML</td>
			<td></td>
		</tr>
	</tbody>
</table>

time-resolved fluorescence imaging and analysis of cancer cell invasion in the 3d spheroid model

The invasion of cancer cells from the primary tumor into the adjacent healthy tissues is an early step in metastasis. Invasive cancer cells pose a major clinical challenge because no efficient method exist for their elimination once their dissemination is underway. A better understanding of the mechanisms regulating cancer cell invasion may lead to the development of novel potent therapies. Due to their physiological resemblance to tumors, spheroids embedded in collagen I have been extensively utilized by researchers to study the mechanisms governing cancer cell invasion into the extracellular matrix (ECM). However, this assay is limited by (1) a lack of control over the embedding of spheroids into the ECM; (2) high cost of collagen I and glass bottom dishes, (3) unreliable immunofluorescent labeling, due to the inefficient penetration of antibodies and fluorescent dyes and (4) time-consuming image processing and quantification of the data. To address these challenges, we optimized the three-dimensional (3D) spheroid protocol to image fluorescently labeled cancer cells embedded in collagen I, either using time-lapse videos or longitudinal imaging, and analyze cancer cell invasion. First, we describe the fabrication of a spheroid imaging device (SID) to embed spheroids reliably and in a minimal collagen I volume, reducing the assay cost. Next, we delineate the steps for robust fluorescence labeling of live and fixed spheroids. Finally, we offer an easy-to-use Fiji macro for image processing and data quantification. Altogether, this simple methodology provides a reliable and affordable platform to monitor cancer cell invasion in collagen I. Furthermore, this protocol can be easily modified to fit the users&#8217; needs.

Due to its biocompatibility, PDMS is widely used for microfabrication of confining wells, stamps and molds, which revolutionized micropatterning and microfluidic devices. In the method described here, it is used to create SIDs, customizable wells that optimize spheroid embedding and imaging procedure. Figure 1 illustrates the major components used in the fabrication of the SIDs. To cast the PDMS mold, a 1-mm thick spacer is 3D printed (Figure 1A,B</stron...

Watch this Scientific Journal Video about Time-Resolved Fluorescence Imaging and Analysis of Cancer Cell Invasion in the 3D Spheroid Model at JoVE.com

Time-Resolved Fluorescence Imaging and Analysis of Cancer Cell Invasion in the 3D Spheroid Model

Presented here is a protocol for the fabrication of a spheroid imaging device. This device enables dynamic or longitudinal fluorescence imaging of cancer cell spheroids. The protocol also offers a simple image processing procedure for the analysis of cancer cell invasion.

The invasion of cancer cells from the primary tumor into the adjacent healthy tissues is an early step in metastasis. Invasive cancer cells pose a major ...

time-resolved-fluorescence-imaging-analysis-cancer-cell-invasion-3d

Research

JoVE Journal

Biology

6.3K Views.  Temple University. Presented here is a protocol for the fabrication of a spheroid imaging device. This device enables dynamic or longitudinal fluorescence imaging of cancer cell spheroids. The protocol also offers a simple image processing procedure for the analysis of cancer cell invasion.

Video: Time-Resolved Fluorescence Imaging and Analysis of Cancer Cell Invasion in the 3D Spheroid Model

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Quantitative Analysis of Autophagy using Advanced 3D Fluorescence Microscopy

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, traditional chemotherapeutic approaches have been largely unsuccessful. Hormone therapy is effective at early stage, but often fails with the eventual development of hormone-refractory tumors. We have been interested in developing therapeutics targeting specific metabolic deficiency of tumor cells. We recently showed that prostate tumor cells specifically lack an enzyme (argininosuccinate synthase, or ASS) involved in the synthesis of the amino acid arginine1. This condition causes the tumor cells to become dependent on exogenous arginine, and they undergo metabolic stress when free arginine is depleted by arginine deiminase (ADI)1,10. Indeed, we have shown that human prostate cancer cells CWR22Rv1 are effectively killed by ADI with caspase-independent apoptosis and aggressive autophagy (or macroautophagy)1,2,3. Autophagy is an evolutionarily-conserved process that allows cells to metabolize unwanted proteins by lysosomal breakdown during nutritional starvation4,5. Although the essential components of this pathway are well-characterized6,7,8,9, many aspects of the molecular mechanism are still unclear - in particular, what is the role of autophagy in the death-response of prostate cancer cells after ADI treatment? In order to address this question, we required an experimental method to measure the level and extent of autophagic response in cells - and since there are no known molecular markers that can accurately track this process, we chose to develop an imaging-based approach, using quantitative 3D fluorescence microscopy11,12.
Using CWR22Rv1 cells specifically-labeled with fluorescent probes for autophagosomes and lysosomes, we show that 3D image stacks acquired with either widefield deconvolution microscopy (and later, with super-resolution, structured-illumination microscopy) can clearly capture the early stages of autophagy induction. With commercially available digital image analysis applications, we can readily obtain statistical information about autophagosome and lysosome number, size, distribution, and degree of colocalization from any imaged cell. This information allows us to precisely track the progress of autophagy in living cells and enables our continued investigation into the role of autophagy in cancer chemotherapy.

Autophagy is a ubiquitous process that enables cells to degrade and recycle proteins and organelles. We apply advanced fluorescence microscopy to visualize and quantify the small, but essential, physical changes associated with the induction of autophagy, including the formation and distribution of autophagosomes and lysosomes, and their fusion into autolysosomes.

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Rapid Analysis and Exploration of Fluorescence Microscopy Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized image analysis tools are available, most traditional image-analysis-based workflows have steep learning curves (for fine tuning of analysis parameters) and result in long turnaround times between imaging and analysis. In particular, cell segmentation, the process of identifying individual cells in an image, is a major bottleneck in this regard.
Here we present an alternate, cell-segmentation-free workflow based on PhenoRipper, an open-source software platform designed for the rapid analysis and exploration of microscopy images. The pipeline presented here is optimized for immunofluorescence microscopy images of cell cultures and requires minimal user intervention. Within half an hour, PhenoRipper can analyze data from a typical 96-well experiment and generate image profiles. Users can then visually explore their data, perform quality control on their experiment, ensure response to perturbations and check reproducibility of replicates. This facilitates a rapid feedback cycle between analysis and experiment, which is crucial during assay optimization. This protocol is useful not just as a first pass analysis for quality control, but also may be used as an end-to-end solution, especially for screening. The workflow described here scales to large data sets such as those generated by high-throughput screens, and has been shown to group experimental conditions by phenotype accurately over a wide range of biological systems. The PhenoBrowser interface provides an intuitive framework to explore the phenotypic space and relate image properties to biological annotations. Taken together, the protocol described here will lower the barriers to adopting quantitative analysis of image based screens.

Here we describe a workflow for rapidly analyzing and exploring collections of fluorescence microscopy images using PhenoRipper, a recently developed image-analysis platform.

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized ...

An Efficient and Flexible Cell Aggregation Method for 3D Spheroid Production

Monolayer cell culture does not adequately model the in vivo behavior of tissues, which involves complex cell-cell and cell-matrix interactions. Three-dimensional (3D) cell culture techniques are a recent innovation developed to address the shortcomings of adherent cell culture. While several techniques for generating tissue analogues in vitro have been developed, these methods are frequently complex, expensive to establish, require specialized equipment, and are generally limited by compatibility with only certain cell types. Here, we describe a rapid and flexible protocol for aggregating cells into multicellular 3D spheroids of consistent size that is compatible with growth of a variety of tumor and normal cell lines. We utilize varying concentrations of serum and methyl cellulose (MC) to promote anchorage-independent spheroid generation and prevent the formation of cell monolayers in a highly reproducible manner. Optimal conditions for individual cell lines can be achieved by adjusting MC or serum concentrations in the spheroid formation medium. The 3D spheroids generated can be collected for use in a wide range of applications, including cell signaling or gene expression studies, candidate drug screening, or in the study of cellular processes such as tumor cell invasion and migration. The protocol is also readily adapted to generate clonal spheroids from single cells, and can be adapted to assess anchorage-independent growth and anoikis-resistance. Overall, our protocol provides an easily modifiable method for generating and utilizing 3D cell spheroids in order to recapitulate the 3D microenvironment of tissues and model the in vivo growth of normal and tumor cells.

Here, we describe a rapid and flexible protocol for the formation of 3D cell spheroids through cell aggregation. This is easily adapted to multiple cell types and is suitable for use in a variety of applications including cell migration, invasion, or anoikis assays, and for imaging and quantifying cell-matrix interactions.

Monolayer cell culture does not adequately model the in vivo behavior of tissues, which involves complex cell-cell and cell-matrix interactions. ...

Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to hyperglycemia and severe, life-threatening consequences. Understanding how the beta-cells operate under physiological conditions and what genetic and environmental factors might cause their dysfunction could lead to better treatment options for diabetic patients. The ability to measure calcium levels in beta-cells serves as an important indicator of beta-cell function, as the influx of calcium ions triggers insulin release. Here we describe a protocol for monitoring the glucose-stimulated calcium influx in zebrafish beta-cells by using GCaMP6s, a genetically encoded sensor of calcium. The method allows monitoring the intracellular calcium dynamics with single-cell resolution in ex vivo mounted islets. The glucose-responsiveness of beta-cells within the same islet can be captured simultaneously under different glucose concentrations, which suggests the presence of functional heterogeneity among zebrafish beta-cells. Furthermore, the technique provides high temporal and spatial resolution, which reveals the oscillatory nature of the calcium influx upon glucose stimulation. Our approach opens the doors to use the zebrafish as a model to investigate the contribution of genetic and environmental factors to beta-cell function and dysfunction.

Beta-cell functionality is important for blood-glucose homeostasis, which is evaluated at single-cell resolution using a genetically encoded reporter for calcium influx.

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to ...

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 ...

Mass Spectrometry Analysis to Identify Ubiquitylation of EYFP-tagged CENP-A (EYFP-CENP-A)

Studying the structure and the dynamics of kinetochores and centromeres is important in understanding chromosomal instability (CIN) and cancer progression. How the chromosomal location and function of a centromere (i.e., centromere identity) are determined and participate in accurate chromosome segregation is a fundamental question. CENP-A is proposed to be the non-DNA indicator (epigenetic mark) of centromere identity, and CENP-A ubiquitylation is required for CENP-A deposition at the centromere, inherited through dimerization between cell division, and indispensable to cell viability.
Here we describe mass spectrometry analysis to identify ubiquitylation of EYFP-CENP-A K124R mutant suggesting that ubiquitylation at a different lysine is induced because of the EYFP tagging in the CENP-A K124R mutant protein. Lysine 306 (K306) ubiquitylation in EYFP-CENP-A K124R was successfully identified, which corresponds to lysine 56 (K56) in CENP-A through mass spectrometry analysis. A caveat is discussed in the use of GFP/EYFP or the tagging of high molecular weight protein as a tool to analyze the function of a protein. Current technical limit is also discussed for the detection of ubiquitylated bands, identification of site-specific ubiquitylation(s), and visualization of ubiquitylation in living cells or a specific single cell during the whole cell cycle.
The method of mass spectrometry analysis presented here can be applied to human CENP-A protein with different tags and other centromere-kinetochore proteins. These combinatory methods consisting of several assays/analyses could be recommended for researchers who are interested in identifying functional roles of ubiquitylation.

Mass Spectrometry Analysis to Identify Ubiquitylation of EYFP-tagged CENP-A EYFP-CENP-A

CENP-A ubiquitylation is an important requirement for CENP-A deposition at the centromere, inherited through dimerization between cell division, and indispensable to cell viability. Here we describe mass spectrometry analysis to identify ubiquitylation of EYFP-tagged CENP-A (EYFP-CENP-A) protein.

Studying the structure and the dynamics of kinetochores and centromeres is important in understanding chromosomal instability (CIN) and cancer ...

Staining and High-Resolution Imaging of Three-Dimensional Organoid and Spheroid Models

In vitro three-dimensional (3D) cell culture models, such as organoids and spheroids, are valuable tools for many applications including development and disease modeling, drug discovery, and regenerative medicine. To fully exploit these models, it is crucial to study them at cellular and subcellular levels. However, characterizing such in vitro 3D cell culture models can be technically challenging and requires specific expertise to perform effective analyses. Here, this paper provides detailed, robust, and complementary protocols to perform staining and subcellular resolution imaging of fixed in vitro 3D cell culture models ranging from 100 &#181;m to several millimeters. These protocols are applicable to a wide variety of organoids and spheroids that differ in their cell-of-origin, morphology, and culture conditions. From 3D structure harvesting to image analysis, these protocols can be completed within 4-5 days. Briefly, 3D structures are collected, fixed, and can then be processed either through paraffin-embedding and histological/immunohistochemical staining, or directly immunolabeled and prepared for optical clearing and 3D reconstruction (200 &#181;m depth) by confocal microscopy.

Here, we provide detailed, robust, and complementary protocols to perform staining and subcellular resolution imaging of fixed three-dimensional cell culture models ranging from 100 &#181;m to several millimeters, thus enabling the visualization of their morphology, cell-type composition, and interactions.

In vitro three-dimensional (3D) cell culture models, such as organoids and spheroids, are valuable tools for many applications including development and ...