Name: mammalian cell division in 3d matrices via quantitative confocal reflection microscopy
Uploaded: 2017-11-29T14:00:00.000+00:00
Duration: 10 min 22 s
Description: Watch this Scientific Journal Video about Mammalian Cell Division in 3D Matrices via Quantitative Confocal Reflection Microscopy at JoVE.com

This work was supported by NIH grants R01CA174388 and U54CA143868. The authors would like to acknowledge the PURA award from the Johns Hopkins University for support of Wei-tong Chen. This material is based upon work supported by the National Science Foundation Graduate Research Fellowship under Grant No. 1232825.

The protocol provided follows the guidelines of The Homewood Institutional Review Board (HIRB).
1. Stable Expression of H2B-mCherry as a Marker for Cell Mitosis
<ol>
	<li>Generation of lentiviral particles from human embryonic kidney 293T (HEK 293T) cells
	<ol>
		<li>Plate the HEK 293T cells on a 10 cm cell culture dish at the density of 5 x 106 cells/dish in cell culture medium (Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM) containing high glucose (4.5 g/L), sodium pyruvate, 10% Fetal Bovine Serum (FBS), and 1% penicillin-streptomycin (Pen/Strep)). Incubate for 24 h at 37 &#176;C and 5% carbon dioxide (CO2). The desired confluency of the cells on the day of transfection is about 70 - 80%. 
		NOTE: HEK 293T cells are used here to produce viral particles, which will be used later to transduce MDA-MB-231 cells in order to generate a cell line stably expressing H2B-mCherry. Make sure that cells are evenly distributed throughout the plate. Addition of the cells drop wise to the plate, and gently moving the plate back and forth can aid in even distribution. Cells can also be seeded in plates or flasks of other sizes, which will result in the collection of different volumes of viral particles.</li>
		<li>Transfect the HEK 293T cells using a transfection reagent with three plasmids (lentiviral vector, CMV &#916;R 8.91, and pMDG-VSVG). The plasmid encoding H2B-mCherry is cloned in a lentiviral vector with phosphoglycerate kinase promoter (PGK). CMV &#916;R 8.91 contains three required HIV genes, gag, pol, and rev. pMDG-VSVG contains the VSV-G envelope gene. 
		NOTE: There are multiple transfection agents to choose from.
		<ol>
			<li>Allow the vial of transfection reagent to reach room temperature (RT) before use. Invert or vortex the vial briefly.</li>
			<li>Mix 16 &#181;g of DNA, which consists of 6 &#181;g of H2B-mCherry plasmid, 8 &#181;g of CMV &#916;R 8.91, and 2 &#181;g of pMDG-VSVG, in 1 mL of reduced serum media. Incubate at RT for 5 min. 
			NOTE: The amount and ratio of the three vectors is varied and optimized for different lentiviral transfer vectors.</li>
			<li>Add 48 &#181;L (use 1:3 ratio of DNA: transfection reagent) of the transfection reagent into the above DNA solution, and then incubate at RT for at least 15 min. 
			NOTE: The ratio of the DNA vs. transfection reagent is varied and optimized for different lentiviral transfer vectors. Make sure that the transfection reagent does not contact the side of the 1.5 mL centrifuge tube.</li>
			<li>Add the DNA-lipid complex solution made in step 1.1.2.3 drop-wise to the cells. Gently swirl the plate to ensure the even distribution of the complex in the plate.</li>
		</ol>
		</li>
		<li>Approximately 6 h after transfection, aspirate the medium and add fresh cell culture medium (DMEM with high glucose (4.5 g/L), sodium pyruvate, 10% FBS, and 1% Pen/Strep).</li>
		<li>Harvest the supernatant 24, 48, and 72 h post-transfection. Replace the medium after each harvest (DMEM, 4.5 g/L glucose, sodium pyruvate, with 10% FBS, and 1% Pen/Strep).</li>
		<li>Filter the lentiviral particles through a 0.45 &#181;m filter to remove the cellular debris. Alternatively, spin down the supernatant to separate out the cell debris. The supernatant can be used for transduction right away, or can be stored at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Generation of cell lines stably expressing H2B-mCherry 
	NOTE: This protocol is described in detail for MDA-MB-231 cells. Before use in the experiment, human breast carcinoma cells MDA-MB-231 (Physical Sciences Oncology Center, NIH) are cultured in DMEM containing high glucose (4.5 g/L), sodium pyruvate, 10% FBS, and 1% Pen/Strep.
	<ol>
		<li>Plate the MDA-MB-231 cells at the density of 1 x 105 cells in a 35 mm culture dish. 
		NOTE: The density of plating for different cell lines should be optimized, and therefore may be different from the optimal density for the MDA-MB-231 cells.</li>
		<li>24 h after plating the cells, add 1 mL of virus and 1 mL of fresh medium (DMEM, high glucose (4.5 g/L), sodium pyruvate, 10% FBS, and 1% Pen/Strep).</li>
		<li>Incubate the cells for a time period of 6 h to overnight. 
		NOTE: Both the volume of the virus and the incubation time need to be optimized for different cell lines. For example, if the cells do not look healthy a few hours after the addition of the virus, use 1 mL of virus and 2 mL of fresh medium (DMEM, high glucose (4.5 g/L), sodium pyruvate, 10% FBS, and 1% Pen/Strep) for step 1.2.2. The length of the incubation time in step 1.2.3 can also be reduced.</li>
		<li>Aspirate the medium and add 10 mL of fresh medium (DMEM, high glucose (4.5 g/L), sodium pyruvate, 10% FBS and 1% Pen/Strep). Incubate for between 24 to 72 h at 37 &#176;C and 5% CO2.</li>
		<li>Check the expression of H2B-mCherry in cells using an epifluorescence microscope. 
		NOTE: If the transduction efficiency is low in the cells, the volume of virus in step 2.2 and the incubation time in step 2.3 could be increased.</li>
	</ol>
	</li>
</ol>
2. Synchronization of the Cells Stably Expressing H2B-mCherry
<ol>
	<li>Plate the cells at 50 to 60% confluency, i.e. plate 2 x 104 MDA-MB-231 cells in each well of a 24-well plate.</li>
	<li>Incubate the culture at 37 &#176;C and 5% CO2 for 24 h.</li>
	<li>Replace the growth medium with 0.5 mL of medium (DMEM, high glucose (4.5 g/L), sodium pyruvate, 10% FBS, and 1% Pen/Strep) containing 2 mM thymidine and leave in the incubator for 24 h. 
	NOTE: Cells exposed to thymidine are arrested at the phase of cell growth (G1)/DNA synthesis (S) transition and throughout S-phase due to the inhibition of DNA synthesis. The length of the incubation time should be varied and optimized for different cell lines.</li>
	<li>Release the cells from thymidine exposure by washing them with phosphate buffered saline (PBS) three times. Then, incubate cells in normal cell culture medium (DMEM, high glucose (4.5 g/L), sodium pyruvate, 10% FBS and 1% Pen/Strep) for 5 h. 
	NOTE: The release of the cells from the thymidine exposure allows the cells to progress to the cell growth (G2)/mitotic (M) phase for cells previously arrested at the G1/S phase, and to the G1 phase for cells previously arrested at the S phase. The length of release time should be varied and optimized for different cell lines.</li>
	<li>Block the cells with 250 ng/mL of nocodazole for 12 h. 
	NOTE: All of the cells exposed to nocodazole are arrested at the G2/M phase. Nocodazole is cytotoxic. Prolonged exposure to nocodazole can cause apoptosis. Adjust the period or concentration of exposure for different cell lines if cell deaths are observed. Cells that are successfully synchronized will exhibit a spherical morphology.</li>
	<li>Shake the cells for 45 s to 1 min using an orbital shaker at 150 to 200 rpm. 
	NOTE: Mitotic cells, which have little adherence to the substrate, will be shaken off during the process.</li>
	<li>Remove the medium to extract cells, by pipetting the medium into a centrifuge tube, and then add 0.5 mL of fresh medium (DMEM, high glucose (4.5 g/L), sodium pyruvate, 10% FBS, and 1% Pen/Strep) to each well of the plate.</li>
	<li>Repeat steps 2.6 and 2.7 three times.</li>
	<li>Centrifuge the collected medium containing the mitotic cells at 800 x g for 3 min. 
	NOTE: This step is used to remove the nocodazole from the cell medium.</li>
</ol>
3. Incorporation of the Synchronized Cells into the Collagen I Matrices
NOTE: Type I collagen is the most abundant protein in the human body and in the ECM of connective tissues, and thus is widely used to investigate how eukaryotic cell functions are modulated by a 3D environment17,23,24. Collagen is soluble in acetic acid. After neutralizing and warming the collagen solution to 20 - 37 &#176;C, collagen monomers polymerize into a meshwork of collagen fibrils.
<ol>
	<li>Prepare the 10x&#160;DMEM solution by dissolving a packet of DMEM powder, 3.7 g of sodium bicarbonate (NaHCO3) and 1 g of 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) in 50 mL of distilled water. Filter the solution, and then prepare 1 M of sodium hydroxide (NaOH) by dissolving 2 g of NaOH pellets in 50 mL of distilled water. Filter and aliquot the solution into 1.5 mL centrifuge tubes. 
	NOTE: Normal DMEM solution should not be used in this step. The addition of significant volume of the collagen solution will dilute the medium. Therefore the concentrated DMEM solution is prepared to ensure that the final concentration of DMEM in the collagen matrix will be the same as the normal DMEM.</li>
	<li>Continue to work with the cells collected from step 2.9. Aspirate the medium, and re-suspend cells in about 0.25 - 0.5 mL of fresh cell culture medium (DMEM, high glucose (4.5 g/L), sodium pyruvate, 10% FBS, and 1% Pen/Strep). 
	NOTE: To reach a specific cell density in the collagen matrix, the initial density of the cells in the suspension cannot be too low. Thus, the volume of the medium used to re-suspend the cells will depend on the total number of available cells.</li>
	<li>Place 10 &#181;L of the re-suspended cell solution from step 3.2 on a hemocytometer and count the density of the cells in the solution.</li>
	<li>Determine the volumes needed for all components to make the 3D collagen matrix. 500 &#181;L of 2 &#181;g/&#181;L collagen gel is used here as an example.
	<ol>
		<li>Calculate the volume of the cell solution needed to obtain 40,000 cells/mL (or 20,000 cells for the 500 &#181;L of collagen gel). This number in &#181;L is X. 50 &#181;L of 10x&#160;DMEM will be needed. 50 &#181;L of FBS will be needed.</li>
		<li>Calculate the volume of collagen I stock needed. The concentration of collagen I is around 4 &#181;g/&#181;L. The amount of collagen needed in &#181;L is Y.</li>
		<li>Calculate the volume of sodium hydroxide need to neutralize the acetic acid in the collagen solution: Y &#181;L collagen * 0.023 * 1000 = Z &#181;L NaOH needed.</li>
		<li>Determine the volume of the filler solution: (500 &#181;L total gel - X &#181;L cells - 50 &#181;L 10X DMEM - 50 &#181;L of FBS - Y &#181;L collagen - Z &#181;L NaOH) = R &#181;L 
		NOTE: The filler solution is distilled water. If the calculated volume of the filler solution is a negative number, the original cell density in the cell suspension is too low. The cells need to be spun down again and re-suspended in a smaller volume of medium (DMEM, high glucose (4.5 g/L), sodium pyruvate, 10% FBS, and 1% Pen/Strep).</li>
	</ol>
	</li>
	<li>Add the components in the following order to a clean 1.5 mL centrifuge tube: cells in medium, 10x&#160;DMEM, FBS, deionized (DI) water, collagen, NaOH. Place all of the components on ice to slow down the gelation of the collagen solution. Use proper pipetting techniques to prevent the formation of bubbles.</li>
	<li>After adding the NaOH, mix the solution carefully with a 1 mL pipet. 
	NOTE: Gelation of the collagen will start right immediately after the addition of NaOH. Mixing should be done carefully and quickly.</li>
	<li>Once the solution is well mixed, add 500 &#181;L of the solution to each well of the 24-well plate. Place the 24-well plate in the 37 &#176;C incubator. Place fresh cell culture medium in the 37 &#176;C water bath. 
	NOTE: Plastic bottom plates are used for low-magnification live-cell microcopy in which the working distance of the lens is above 1 mm. Glass bottom plates are used for high-magnification live-cell microscopy due to the short working distance of the high magnification lens.</li>
	<li>Add 500 &#181;L of pre-warmed medium (DMEM, high glucose (4.5 g/L), sodium pyruvate, 10% FBS and 1% Pen/Strep) to the top of the gel 30 min after completing step 3.7.</li>
</ol>
4. Live Cell Imaging of the Cells Dividing in the 3D Collagen Matrices (Low Magnification)
NOTE: Images of cells are collected at 2 min intervals using a charge coupled device (CCD) camera mounted on a phase contrast microscope that is equipped with a 10X objective and controlled by imaging software.
<ol>
	<li>Turn on the live cell unit mounted on top of the objective lens. Wait until the temperature stabilizes at 37 &#176;C, the concentration of CO2 is at 5% and the humidity is at 75%.</li>
	<li>Put the 24-well plate of gels into the live cell unit on the microscope. Find the bottom of the plate and then move the objective lens up until the microscope focuses at about 500 &#181;m from the bottom of the plate. 
	NOTE: Cells in the collagen matrix that are too close to the bottom of the plate are not completely embedded in the 3D matrix. Imaging cells that are 500 &#181;m away from the bottom of the plate will ensure that the cells are not affected by edge effects16,17,25.</li>
	<li>Move the stage around to find the cells, and select multiple positions for imaging.</li>
	<li>Set up the time-lapse experiment at 2-min intervals. 
	NOTE: The maximum number of positions that can be taken during the 2 min interval is limited by the moving speed of the motorized stage, and the length of the exposure time for capturing images.</li>
</ol>
5. Collagen Network Deformation During Cell Division (High Magnification Microscopy)
<ol>
	<li>Turn on the live cell unit mounted on top of the objective lens. Wait until the temperature stabilizes at 37 &#176;C, the concentration of CO2 is at 5% and the humidity is at 75%.</li>
	<li>To visualize collagen fibers in unstained 3D collagen matrices, configure a confocal microscope to capture only reflected light (488-nm) from the 488-nm laser used to illuminate the sample. The laser uses a 60X&#160;water-immersion objective, NA = 1.2, WD = 200 &#181;m, and is controlled by imaging software.</li>
	<li>Put the sample (cells in collagen matrices) into the live cell unit. Find the bottom of the plate and then move the objective lens up until the microscope focuses at about 100 &#181;m from the bottom of the plate. Imaging cells that are 100 &#181;m away from the bottom of the plate will ensure that the cells are not affected by edge effects. 
	NOTE: Here a water-immersion objective is used instead of an oil-immersion lens because the working distance of the oil-immersion lens is only about 100 &#181;m. The use of the oil-immersion lens poses a problem as the thickness of the glass on the bottom of the plate is normally about 100 &#181;m.</li>
	<li>Move the stage around to find the cells, and select multiple positions for imaging. 
	NOTE: Confocal microscopy optically sections the sample and only captures signals from the focal plane. Live cells might be moving in and out of focus during the time-lapse experiment. To enable the capturing of cell images over long periods of time, Z-stack is used to collect images from successive slices at 5 &#181;m intervals. A total of 5 slices spanning 25 &#181;m are imaged.</li>
	<li>Set up the time-lapse experiment at 5 min intervals. 
	NOTE: 5 minutes is used here as the interval instead of 2 min used in section 4 because multiple scanning modes, including fluorescence scanning for H2B-mCherry, reflection scanning for collagen, and Z-stack scanning, are applied to each position. The use of multiple scanning modes reduces the speed of scanning a position when compared to the low magnification imaging.</li>
	<li>Quantify the deformation of the collagen matrix in the vicinity of a cell due to the forces exerted by that cell using particle imaging velocimetry (PIV) with sub-pixel resolution20. Performing this analysis on 2D images allows for the clear visualization of both the signals of the H2B-mCherry and the collagen fibers. Apply the anisotropic low pass filtering to enhance the signal of the collagen network20.</li>
	<li>In order to measure the local displacement of a sub-image region of interest located at (x,y) from frame k to frame k+1, extract a regional window of 15&#215;15 pixels centered at (x,y) at frame k. Then identify the best matching locations (x,y)* through the locations that feature a maximum normalized cross-correlation coefficient in the image obtained at frame k+1. The deformation vector is calculated as (x,y)* - (x,y).</li>
</ol>

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Visualisation of dynamic interactions with the substratum and the distribution of surface structures via a novel confocal reflection imaging technique.</a> Adv Exp Med Biol. 417, 97-103 (1997).</li><li>Geraldo, S., Simon, A., Vignjevic, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revealing+the+cytoskeletal+organization+of+invasive+cancer+cells+in+3D.">Revealing the cytoskeletal organization of invasive cancer cells in 3D.</a> J Vis Exp. (80), e50763 (2013).</li><li>Lleres, D., James, J., Swift, S., Norman, D. G., Lamond, A. 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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biophysical+control+of+invasive+tumor+cell+behavior+by+extracellular+matrix+microarchitecture.">Biophysical control of invasive tumor cell behavior by extracellular matrix microarchitecture.</a> Biomaterials. 33 (16), 4157-4165 (2012).</li><li>Langan, T. J., Chou, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synchronization+of+mammalian+cell+cultures+by+serum+deprivation.">Synchronization of mammalian cell cultures by serum deprivation.</a> Methods Mol Biol. 761, 75-83 (2011).</li><li>Jackman, J., O'Connor, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+synchronizing+cells+at+specific+stages+of+the+cell+cycle.">Methods for synchronizing cells at specific stages of the cell cycle.</a> Curr Protoc Cell Biol. 8, (2001).</li><li>Zwanenburg, T. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Standardized+shake-off+to+synchronize+cultured+CHO+cells.">Standardized shake-off to synchronize cultured CHO cells.</a> Mutat Res. 120 (2-3), 151-159 (1983).</li><li>Harper, J. 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S., Zaman, 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=Quantitative+analysis+of+the+effect+of+cancer+invasiveness+and+collagen+concentration+on+3D+matrix+remodeling.">Quantitative analysis of the effect of cancer invasiveness and collagen concentration on 3D matrix remodeling.</a> PLoS One. 6 (9), e24891 (2011).</li><li>Wolf, K., 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=Collagen-based+cell+migration+models+in+vitro+and+in+vivo.">Collagen-based cell migration models in vitro and in vivo.</a> Semin Cell Dev Biol. 20 (8), 931-941 (2009).</li><li>Petroll, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+interference+contrast+and+confocal+reflectance+imaging+of+collagen+organization+in+three-dimensional+matrices.">Differential interference contrast and confocal reflectance imaging of collagen organization in three-dimensional matrices.</a> Scanning. 28 (6), 305-310 (2006).</li><li>Fraley, S. 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=Three-dimensional+matrix+fiber+alignment+modulates+cell+migration+and+MT1-MMP+utility+by+spatially+and+temporally+directing+protrusions.">Three-dimensional matrix fiber alignment modulates cell migration and MT1-MMP utility by spatially and temporally directing protrusions.</a> Sci Rep. 5, 14580 (2015).</li><li>Ikeda, Y., Collins, M. K., Radcliffe, P. A., Mitrophanous, K. A., Takeuchi, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+transduction+efficiency+in+cells+of+different+species+by+HIV+and+EIAV+vectors.">Gene transduction efficiency in cells of different species by HIV and EIAV vectors.</a> Gene Ther. 9 (14), 932-938 (2002).</li><li>Ma, H. T., Poon, R. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synchronization+of+HeLa+cells.">Synchronization of HeLa cells.</a> Methods Mol Biol. 761, 151-161 (2011).</li></ol>

The authors declare that they have no competing financial interests.

The previous study of cell division in 3D was not efficient due to experimental limitations and technical challenges18,19. The critical steps for efficient study of mammalian cell division in 3D collagen matrices are: (1) the incorporation of fluorescence-labeled mitotic markers to the cells; (2) the synchronization of cell division; and (3) the monitoring of division events in 3D matrices using live-cell imaging technique, time-resolved confocal reflection microscopy, and quantitative imaging analysis.
Mitotic cells on 2D substrates can be distinguished from the interphase cells based on their morphology, i.e. mitotic cells are round and barely attach to the substrate whereas interphase cells spread out and attach firmly to the substrate. In 3D matrices, however, the cell morphology is not a reliable marker for mitotic cells since some cells barely spread out and remain round in the matrix27,28. Thus, it is essential to introduce a mitotic marker to the cells for the study of cell division in 3D matrices. We stably expressed H2B-mCherry in MDA-MB-231 cells, which served as a reliable marker for different mitotic phases including prophase, prometaphase, metaphase, anaphase, and telophase/cytokinesis. We previously used this approach to distinguish mitotic cells from interphase cells in 3D matrices. With the help of this marker, we were also able to measure the length of the mitotic phase for another cell line, HT1080 cells, dividing on 2D substrates and in 3D matrices19.
There are multiple ways to synchronize cells, including serum starvation29, mitotic shake-off30,31, double thymidine blocking32, and nocodazole32. We combined the thymidine treatment and the nocodazole treatment to efficiently synchronize MDA-MB-231 cells to the G2/M phase. Cells exposed to thymidine are arrested at the G1/S transition and throughout S phase due to the inhibition of DNA synthesis by thymidine. The release of the cells from the thymidine exposure let the cells progress to G2/M phase for cells arrested at G1/S phase, and to G1 for cells arrested at S phase. All the cells exposed to nocodazole are arrested at the G2/M phase. The rounded-up cells entering mitotic phase were then shaken-off from the plate and directly encapsulated into the collagen matrices. We showed that about 70% of the cells divide within 2 h after they are incorporated into the collagen matrices (Figure 2). Alternatively, cells could be embedded in collagen matrices before synchronization, however, the cross-linked network of the collagen matrix presents a physical barrier and reduces the rate of biomolecular diffusion and convection33,34. Indeed, we attempted to synchronize cells in collagen matrices using thymidine and nocodazole, but failed to obtain efficient synchronization. This result might be due to the inefficient diffusion and convection of the drugs through the collagen matrix.
Reflection is an intrinsic optical property of many biopolymers, including collagen. The confocal reflection microscopy technique visualizes and quantitates the microtopography of porous biomaterials prepared from synthetic polymers and 3D collagen matrices23,24,35,36,37. In our lab, we have established techniques to monitor changes in collagen fiber polarity as the concentration of collagen varies38. Here, we describe the method to monitor the deformation of collagen fibers based on the time-lapse video of the reflective confocal images to denote the cell-matrix interaction. The representative results presented here show that the matrix deformation for the mitotic MDA-MB-231 cells is significantly smaller than those cells in interphase, which suggests that mammalian cells have minimal attachment and interactions with the surrounding matrix when they enter the mitotic phase19.
Previously, we used confocal reflection microscopy to monitor and quantify cell-matrix interaction before, during and after the mitosis of HT1080 cells. We also monitored the matrix deformation by &#946;1-integrin knock-down HT1080 cells during both interphase and mitotic phase. Depleting &#946;1-integrin significantly reduces the matrix deformation by the cell during interphase. However, there is no difference in matrix deformation during the mitotic phase of the round &#946;1-integrin knockdown (KD) cells and the HT1080 wild type cells19.
An alternative approach to visualize collagen fibers is to employ fluorescence-conjugated type I collagen. We previously used this approach to image tracks in the collagen matrices generated by cells19. This approach, however, requires the labeling of collagen with fluorescent dye such as fluorescein isothiocyanate (FITC), which is both time consuming and less efficient. On the other hand, confocal reflection microscopy can be directly applied to unmodified collagen to save time and resources, and exclude the issues associated with fluorescence photobleaching. Moreover, this method does not require an individual fluorescent channel, and is therefore compatible with all fluorescent dyes.
The method presented in this paper can be potentially applied to any type of mammalian cells that divide in a 3D collagen matrix. The incorporation of the H2B-mCherry marker into other mammalian cells by lentiviral transduction will follow exactly the same procedures as described in the paper, although different types of cells may have varied efficiencies for transduction by lentivirus39. Both the density of the cells upon transduction and the titer of the virus could be optimized for efficiency. If high transduction efficiency cannot be achieved, cells could be selected by fluorescence assisted cell sorting (FACS). Thymidine blocking and nocodazole are applied to successfully synchronize several other types of mammalian cells, such as HeLa40. Mechanical shake-off could be applied to any cell types that round up and barely attach to the substrate during the mitotic phase30,31. Moreover, the imaging of collagen fibers using confocal reflection microscopy, and the quantification of the matrix deformation can be directly applied to all other types of dividing mammalian cells.
The method presented here is an efficient and general approach to study mammalian cell division and cell-matrix interactions in a 3D environment. The approach facilitates our probe into the molecular basis of the development of normal tissue and diseases, and potentiates the design of novel diagnostic and therapeutic approaches in the future.

Cell mitosis is a critical event in cellular life, the regulation of which plays crucial roles in tissue and organ development. Abnormal mitosis is implicated in natural genetic variations, human aging processes, and the progression of cancer1,2,3,4,5. The increased rate of proliferation of tumor cells compared with normal cells is one of the hallmarks of cancer, despite the fact that cell behaviors are quite heterogeneous among different types of tumors and even among patients. In spite of promising preclinical results, some newly-developed antimitotic drugs have not shown to be effective in clinical trials6,7,8,9,10,11.The relevance of experimental and preclinical models has to be considered. Many types of normal mammalian and cancer cells divide in three-dimensional (3D) matrices, such as fibroblasts and fibrosarcoma cells in collagen I-rich 3D connective tissues, and metastatic cancer cells in the 3D stromal extracellular matrix (ECM). However, the vast majority of mammalian cell division experiments and assays have been performed on cells cultured on two-dimensional (2D) substrates. An engineered 3D matrix could better recapitulate the microstructure, mechanical properties, and biochemical signals of the 3D ECM of both normal and pathologic tissues12,13,14,15,16,17.
The study of how mammalian cell division is regulated in 3D environments remains largely unexplored despite both the physiological relevance and the therapeutic significance18,19. Possible reasons include the technical difficulties and experimental challenges associated with studying cell division in 3D matrices. Cell mitosis constitutes a small temporal fraction in the whole cell cycle20. Previous work has shown that the proliferation rate of many mammalian cells, such as human breast adenocarcinoma MCF-7, human osteosarcoma U2OS, and human liver HepG2, is much lower in 3D matrices compared with their counterparts on 2D substrates21,22. Furthermore, cells embedded in 3D matrices move in and out of focus during live-cell imaging. All of these factors contribute to the extremely low efficiency of capturing cell-division events in 3D culture using imaging techniques.
Interactions between the ECM and cells play critical roles in regulating cell divisions. Here, we describe an approach to efficiently study mammalian cell division in 3D collagen matrices. The method includes the incorporation of mitotic markers to the cells, synchronization of cell division, as well as the monitoring of division events in 3D matrices using the live-cell imaging technique, time-resolved confocal reflection microscopy, and quantitative imaging analysis. Fluorescence-labeled histone protein H2B is first introduced into the cells as a marker to differentiate mitotic and interphase cells. Then the cells are synchronized using the combination of thymidine blocking and nocodazole treatment, followed by a mechanical shake-off technique. Synchronized cells are then directly encapsulated into 3D collagen matrices. Cell division events of multiple cells are monitored efficiently using low-magnification time-lapse live-cell imaging. The deformation of collagen fibers, which is an indicator of cell-matrix interaction, is monitored using confocal reflection microscopy at high-magnification.
We have previously used this technique to monitor and quantify cell-matrix interaction before, during and after the mitosis of two metastatic cancer cell lines, human invasive ductal carcinoma MDA-MB-231 and human fibrosarcoma HT1080 cells, in 3D collagen matrices19. The methods presented here provide an efficient and general approach to study both mammalian cell division in a 3D environment and cell-matrix interactions. The MDA-MB-231 cell line is used as an example throughout the paper. This protocol provides novel insights into the molecular basis of the development of normal tissue and diseases, and could also allow for the design of novel diagnostic and therapeutic approaches.

<table><tbody><tr><td>Human embryonic kidney 293T</td><td>ATCC</td><td></td><td></td></tr><tr><td>MDA-MB-231</td><td>Physical Sciences Oncology Center, NIH</td><td></td><td></td></tr><tr><td>DMEM</td><td>Corning</td><td>10-013-CV</td><td></td></tr><tr><td>DMEM powder</td><td>ThermoFisher Scientific</td><td>12100-046</td><td></td></tr><tr><td>Fetal bovine serum</td><td>Hyclone</td><td>SH30910.03</td><td></td></tr><tr><td>Penicillin-Streptomycin 100X</td><td>Sigma-Aldrich</td><td>P0781</td><td></td></tr><tr><td>Fugene HD</td><td>Promega</td><td>E2311</td><td></td></tr><tr><td>Lipofectamine 2000</td><td>Life technologies</td><td>11668-07</td><td></td></tr><tr><td>Plasmid encoding H2B-mCherry in a lentiviral vector</td><td>Addgene</td><td>plasmid 21217</td><td></td></tr><tr><td>Thymidine</td><td>Sigma-Aldrich</td><td>T1895</td><td></td></tr><tr><td>Nocodazole</td><td>Sigma-Aldrich</td><td>M1404</td><td></td></tr><tr><td>Opti-MEM</td><td>Life Technologies</td><td>31985-070</td><td></td></tr><tr><td>Sodium bicarbonate</td><td>GibcoBRL</td><td>11810-025</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>113375-100</td><td></td></tr><tr><td>Collagen</td><td>Corning</td><td>354236</td><td></td></tr><tr><td>NaOH</td><td>J.T. Bake</td><td>3722-01</td><td></td></tr><tr><td>Millex-HV syringe filter unit, 0.45-&amp;mu;m, PVDF, 33 mm</td><td>Millipore</td><td>SLHVM33RS</td><td></td></tr><tr><td>Nikon TE2000E epifluorescence microscope</td><td>Nikon</td><td>TE2000E</td><td></td></tr><tr><td>Cascade 1K CCD camera</td><td>Roper Scientific</td><td></td><td></td></tr><tr><td>NIS-Elements AR imaging software</td><td>Nikon</td><td></td><td></td></tr><tr><td>Nikon A1 confocal microscope</td><td>Nikon</td><td>A1</td><td></td></tr></tbody></table>

mammalian cell division in 3d matrices via quantitative confocal reflection microscopy

The study of how mammalian cell division is regulated in a 3D environment remains largely unexplored despite its physiological relevance and therapeutic significance. Possible reasons for the lack of exploration are the experimental limitations and technical challenges that render the study of cell division in 3D culture inefficient. Here, we describe an imaging-based method to efficiently study mammalian cell division and cell-matrix interactions in 3D collagen matrices. Cells labeled with fluorescent H2B are synchronized using the combination of thymidine blocking and nocodazole treatment, followed by a mechanical shake-off technique. Synchronized cells are then embedded into a 3D collagen matrix. Cell division is monitored using live-cell microscopy. The deformation of collagen fibers during and after cell division, which is an indicator of cell-matrix interaction, can be monitored and quantified using quantitative confocal reflection microscopy. The method provides an efficient and general approach to study mammalian cell division and cell-matrix interactions in a physiologically relevant 3D environment. This approach not only provides novel insights into the molecular basis of the development of normal tissue and diseases, but also allows for the design of novel diagnostic and therapeutic approaches.

The goal of this article is to present an imaging-based method to study mammalian cell division processes in 3D matrices, and to quantify the interactions between the cell and the 3D extracellular matrix during and after cell division. To facilitate the imaging of cell mitosis, we incorporated H2B-mCherry into MDA-MB-231 cells using lentiviral transduction. H2B conjugated with fluorescent proteins is used as a mitotic marker to distinguish mitotic cells from interphase cells, and to define different stages during cell mitosis19,20,26. Using this method, we were able to monitor the entire division process of MDA-MB-231 cells stably expressing H2B-mCherry in 3D collagen matrices (Figure 1A). Mitotic phases started with the dissolution of the nuclear membrane (prophase; frame 1, Figure 1A); the re-organization of the chromosomes (prometaphase: frame 2); the alignment of the chromosomes in the middle of the cell body (metaphase; frame 3); the separation of the chromosomes (anaphase; frame 4); the reorganization of the chromosomes and nuclear membrane, and the separation of the bodies of the two daughter cells (telophase/cytokinesis, frame 5).
The proliferation rate of cells in a 3D matrix is usually much lower than their counterparts on a 2D substrate, which renders the monitoring of cell division in 3D less efficient19. To increase the efficiency of studying 3D cell division, we employed a method combining thymidine, nocodazole, and a shake-off technique to synchronize and select MDA-MB-231 cells that are at the mitotic phase. Synchronized cells were then embedded into collagen matrices. The division process of multiple cells was monitored in real-time using live-cell imaging at 10X magnification. About 70% of the cells divided within the first 2 h after the formation of collagen matrix (Figure 2), thus allowing for efficient monitoring of mitosis in 3D (Figure 2).
To monitor the interaction between cells and their surrounding collagen matrices, we combined confocal reflection microscopy to image collagen fibers, and fluorescence microscopy to image the cells. A 60X lens allows for the capture of high-quality images of collagen fibers. Imaging using a high-magnification lens, which captures fewer cells in each field of view, is much lower-throughput compared to the use of low-magnification at 10X or 20X. Successful synchronization of the cells greatly enhances the efficiency and throughput of such an experiment, since most of the synchronized cells divide within the first 2 h after the formation of the collagen matrix. The matrix deformation during and after cell division is visualized (representative snapshots are shown in Figure 1B) and quantified using a custom PIV software. We quantified and compared matrix deformation during interphase and mitotic phase for cells synchronized into the mitotic phase. We observed that matrix deformation changed very little during the mitotic phase (Figure 3A), and is less than the deformation observed in post-mitotic phases (Figure 3B). This result shows that mammalian cells have minimal attachment and interactions with the surrounding matrix while they are in the mitotic phase.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56364/56364fig1.jpg" /> 
Figure 1: Representative micrographs obtained from a high-magnification live-cell imaging video of a MDA-MB-231 cell embedded in a collagen matrix that stably expresses H2B-mCherry. (A) Different phases of the mitotic progression of the MDA-MB-231 cell are defined by H2B-mCherry as labeled in red. (B) The collagen fibers (white) during the mitotic process are visualized by time-dependent confocal reflection microscopy. Scale bar = 20 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56364/56364fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56364/56364fig2.jpg" /> 
Figure 2: Division of the synchronized cells in 3D collagen matrices. Cells were synchronized to G2/M phase and embedded in a collagen matrix. About 70% of synchronized cells divide within the first 2 h, whereas control cells without synchronization divide randomly. Error bar = SEM (standard error of the mean). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56364/56364fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56364/56364fig3.jpg" /> 
Figure 3: Quantification of the matrix deformation for MDA-MB-231 cells during interphase and mitosis. (A) The change in the magnitude of matrix deformation for matrix-embedded MDA-MB-231 cells. The green arrow indicates prophase and red arrow indicates telophase. (B) Quantification of matrix deformation for MDA-MB-231 cells during interphase and mitotic phase, indicating that matrix deformation is minimal during cell division. Error bar = SEM (standard error of the mean). * p &#60;0.05. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56364/56364fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Mammalian Cell Division in 3D Matrices via Quantitative Confocal Reflection Microscopy at JoVE.com

Mammalian Cell Division in 3D Matrices via Quantitative Confocal Reflection Microscopy

This protocol efficiently studies mammalian cell division in 3D collagen matrices by integrating synchronization of cell division, monitoring of division events in 3D matrices using live-cell imaging technique, time-resolved confocal reflection microscopy and quantitative imaging analysis.

The study of how mammalian cell division is regulated in a 3D environment remains largely unexplored despite its physiological relevance and therapeutic ...

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9.1K Views.  Johns Hopkins University. Dit protocol bestudeert efficiënt zoogdieren celdeling in 3D collageen matrices door de integratie van de synchronisatie van de celdeling, toezicht op divisie gebeurtenissen in 3D-matrices met behulp van live-cel beeldvormende techniek, time-resolved confocal reflectie microscopie en kwantitatieve analyse van beeldvorming.

Video: Mammalian Cell Division in 3D Matrices via Quantitative Confocal Reflection Microscopy

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound healing, tissue regeneration, and cancer metastasis, cells must navigate through complex, three-dimensional extracellular tissue. To better understand the mechanisms behind these biological processes, it is important to examine the roles of the proteins responsible for driving cell migration. Here, we outline a protocol to study the mechanisms of cell migration using the epithelial cell line (MDCK), and a three-dimensional, fibrous, self-polymerizing matrix as a model system. This optically clear extracellular matrix is easily amenable to live-cell imaging studies and better mimics the physiological, soft tissue environment. This report demonstrates a technique for directly visualizing protein localization and dynamics, and deformation of the surrounding three-dimensional matrix. 
Examination of protein localization and dynamics during cellular processes provides key insight into protein functions. Genetically encoded fluorescent tags provide a unique method for observing protein localization and dynamics. Using this technique, we can analyze the subcellular accumulation of key, force-generating cytoskeletal components in real-time as the cell maneuvers through the matrix. In addition, using multiple fluorescent tags with different wavelengths, we can examine the localization of multiple proteins simultaneously, thus allowing us to test, for example, whether different proteins have similar or divergent roles. Furthermore, the dynamics of fluorescently tagged proteins can be quantified using Fluorescent Recovery After Photobleaching (FRAP) analysis. This measurement assays the protein mobility and how stably bound the proteins are to the cytoskeletal network.
By combining live-cell imaging with the treatment of protein function inhibitors, we can examine in real-time the changes in the distribution of proteins and morphology of migrating cells. Furthermore, we also combine live-cell imaging with the use of fluorescent tracer particles embedded within the matrix to visualize the matrix deformation during cell migration. Thus, we can visualize how a migrating cell distributes force-generating proteins, and where the traction forces are exerted to the surrounding matrix. Through these techniques, we can gain valuable insight into the roles of specific proteins and their contributions to the mechanisms of cell migration.

Cellular processes such as cell migration have traditionally been studied on two-dimensional, stiff plastic surfaces. This report describes a technique for directly visualizing protein localization and analyzing protein dynamics in cells migrating in a more physiologically relevant, three-dimensional matrix.

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound ...

MAME Models for 4D Live-cell Imaging of Tumor: Microenvironment Interactions that Impact Malignant Progression

We have developed 3D coculture models, which we term MAME (mammary architecture and microenvironment engineering), and used them for live-cell imaging in real-time of cell:cell interactions. Our overall goal was to develop models that recapitulate the architecture of preinvasive breast lesions to study their progression to an invasive phenotype. Specifically, we developed models to analyze interactions among pre-malignant breast epithelial cell variants and other cell types of the tumor microenvironment that have been implicated in enhancing or reducing the progression of preinvasive breast epithelial cells to invasive ductal carcinomas. Other cell types studied to date are myoepithelial cells, fibroblasts, macrophages and blood and lymphatic microvascular endothelial cells. In addition to the MAME models, which are designed to recapitulate the cellular interactions within the breast during cancer progression, we have developed comparable models for the progression of prostate cancers. 
Here we illustrate the procedures for establishing the 3D cocultures along with the use of live-cell imaging and a functional proteolysis assay to follow the transition of cocultures of breast ductal carcinoma in situ (DCIS) cells and fibroblasts to an invasive phenotype over time, in this case over twenty-three days in culture. The MAME cocultures consist of multiple layers. Fibroblasts are embedded in the bottom layer of type I collagen. On that is placed a layer of reconstituted basement membrane (rBM) on which DCIS cells are seeded. A final top layer of 2% rBM is included and replenished with every change of media. To image proteolysis associated with the progression to an invasive phenotype, we use dye-quenched (DQ) fluorescent matrix proteins (DQ-collagen I mixed with the layer of collagen I and DQ-collagen IV mixed with the middle layer of rBM) and observe live cultures using confocal microscopy. Optical sections are captured, processed and reconstructed in 3D with Volocity visualization software. Over the course of 23 days in MAME cocultures, the DCIS cells proliferate and coalesce into large invasive structures. Fibroblasts migrate and become incorporated into these invasive structures. Fluorescent proteolytic fragments of the collagens are found in association with the surface of DCIS structures, intracellularly, and also dispersed throughout the surrounding matrix. Drugs that target proteolytic, chemokine/cytokine and kinase pathways or modifications in the cellular composition of the cocultures can reduce the invasiveness, suggesting that MAME models can be used as preclinical screens for novel therapeutic approaches.

We have developed 3D coculture models for live-cell imaging in real-time of interactions among breast tumor cells and other cells in their microenvironment that impact progression to an invasive phenotype. These models can serve as preclinical screens for drugs to target paracrine-induced proteolytic, chemokine/cytokine and kinase pathways implicated in invasiveness.

We have developed 3D coculture models, which we term MAME (mammary architecture and microenvironment engineering), and used them for live-cell imaging in ...

Medicine

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a rapid and high-throughput manner. S2 cells have proven amenable to both fixed- and live-cell imaging applications. Notably, live-cell imaging can yield valuable information about how loss or knockdown of a gene can affect the kinetics and dynamics of key events during cell division, including mitotic spindle assembly, chromosome congression, and segregation, as well as overall cell cycle timing. Here we utilize S2 cells stably transfected with fluorescently tagged mCherry:&#945;-tubulin to mark the mitotic spindle and GFP:CENP-A (referred to as &#8216;CID&#8217; gene in Drosophila) to mark the centromere to analyze the effects of key mitotic genes on the timing of cell divisions, from prophase (specifically at Nuclear Envelope Breakdown; NEBD) to the onset of anaphase. This imaging protocol also allows for the visualization of the spindle microtubule and chromosome dynamics throughout mitosis. Herein, we aim to provide a simple yet comprehensive protocol that will allow readers to easily adapt S2 cells for live imaging experiments. Results obtained from such experiments should expand our understanding of genes involved in the cell division by defining their role in several simultaneous and dynamic events. Observations made in this cell culture system can be validated and further investigated in vivo using the impressive toolkit of genetic approaches in flies.

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Cell divisions can be visualized in real time using fluorescently tagged proteins and time-lapse microscopy. Using the protocol presented here, users can analyze cell division timing dynamics, mitotic spindle assembly, and chromosome congression and segregation. Defects in these events following RNA interference (RNAi)-mediated gene knockdown can be assessed and quantified.

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a ...

Genetics

Discovery of Metastatic Regulators using a Rapid and Quantitative Intravital Chick Chorioallantoic Membrane Model

Recent advances in cancer research has illustrated the highly complex nature of cancer metastasis. Multiple genes or genes networks have been found to be involved in differentially regulating cancer metastatic cascade genes and gene products dependent on the cancer type, tissue, and individual patient characteristics. These represent potentially important targets for genetic therapeutics and personalized medicine approaches. The development of rapid screening platforms is essential for the identification of these genetic targets.
The chick chorioallantoic membrane (CAM) is a highly vascularized, collagen rich membrane located under the eggshell that allows for gas exchange in the developing embryo. Due to the location and vascularization of the CAM, we developed it as an intravital human cancer metastasis model that allows for robust human cancer cell xenografting and real-time imaging of cancer cell interactions with the collagen rich matrix and vasculature. 
Using this model, a quantitative screening platform was designed for the identification of novel drivers or suppressors of cancer metastasis. We transduced a pool of head and neck HEp3 cancer cells with a complete human genome shRNA gene library, then injected the cells, at low density, into the CAM vasculature. The cells proliferated and formed single-tumor cell colonies. Individual colonies that were unable to invade into the CAM tissue were visible as a compact colony phenotype and excised for identification of the transduced shRNA present in the cells. Images of individual colonies were evaluated for their invasiveness. Multiple rounds of selections were performed to decreases the rate of false positives. Individual, isolated cancer cell clones or newly engineered clones that express genes of interest were subjected to primary tumor formation assay or cancer cell vasculature co-option analysis. In summary we present a rapid screening platform that allows for anti-metastatic target identification and intravital analysis of a dynamic and complex cascade of events.

This is an effective method to screen for suppressors or drivers of cancer metastasis. Cells, transduced with an expression library, are injected into the chicken chorioallantoic membrane vasculature to form metastatic colonies. Colonies having decreased or increased invasiveness are excised, expanded, reinjected to confirm their phenotype, and finally, analyzed using high throughput sequencing.

Recent advances in cancer research has illustrated the highly complex nature of cancer metastasis. Multiple genes or genes networks have been found to be ...

Cancer Research

Concentric Gel System to Study the Biophysical Role of Matrix Microenvironment on 3D Cell Migration

The ability of cells to migrate is crucial in a wide variety of cell functions throughout life&#160;from embryonic development and wound healing&#160;to tumor and cancer metastasis. Despite intense research efforts, the basic biochemical and biophysical principles of cell migration are still not fully understood, especially in the physiologically relevant three-dimensional (3D) microenvironments. Here, we describe an in vitro assay designed to allow quantitative examination of 3D cell migration behaviors. The method exploits the cell&#8217;s mechanosensing ability and propensity to migrate into previously unoccupied extracellular matrix (ECM). We use the invasion of highly invasive breast cancer cells, MDA-MB-231, in collagen gels as a model system. The spread of cell population and the migration dynamics of individual cells over weeks of culture can be monitored using live-cell imaging and analyzed to extract spatiotemporally-resolved data. Furthermore, the method is easily adaptable for diverse extracellular matrices, thus offering a simple yet powerful way to investigate the role of biophysical factors in the microenvironment on cell migration.

The mechanical properties and microstructure of the extracellular matrix strongly affect 3D migration of cells. An in vitro method to study the spatiotemporal cell migration behavior in biophysically variable environments, at both population and individual cell levels, is described.

The ability of cells to migrate is crucial in a wide variety of cell functions throughout life&#160;from embryonic development and wound healing&#160;to ...

Quantification of Breast Cancer Cell Invasiveness Using a Three-dimensional (3D) Model

It is now well known that the cellular and tissue microenvironment are critical regulators influencing tumor initiation and progression. Moreover, the extracellular matrix (ECM) has been demonstrated to be a critical regulator of cell behavior in culture and homeostasis in vivo. The current approach of culturing cells on two-dimensional (2D), plastic surfaces results in the disturbance and loss of complex interactions between cells and their microenvironment. Through the use of three-dimensional (3D) culture assays, the conditions for cell-microenvironment interaction are established resembling the in vivo microenvironment. This article provides a detailed methodology to grow breast cancer cells in a 3D basement membrane protein matrix, exemplifying the potential of 3D culture in the assessment of cell invasion into the surrounding environment. In addition, we discuss how these 3D assays have the potential to examine the loss of signaling molecules that regulate epithelial morphology by immunostaining procedures. These studies aid to identify important mechanistic details into the processes regulating invasion, required for the spread of breast cancer.

Quantification of Breast Cancer Cell Invasiveness Using a Three-dimensional 3D Model

This article provides detailed methodologies for the use of three-dimensional (3D) assays to quantify breast cancer cell invasion. Specifically, we discuss the procedures required to set up such assays, quantification, and data analysis, as well as methods to examine the loss of membrane integrity that occurs when cells invade.

It is now well known that the cellular and tissue microenvironment are critical regulators influencing tumor initiation and progression. Moreover, the ...

Mapping the Emergent Spatial Organization of Mammalian Cells using Micropatterns and Quantitative Imaging

A fundamental goal in biology is to understand how patterns emerge during development. Several groups have shown that patterning can be achieved in vitro when stem cells are spatially confined onto micropatterns, thus setting up experimental models which offer unique opportunities to identify, in vitro, the fundamental principles of biological organisation.
Here we describe our own implementation of the methodology. We adapted a photo-patterning technique to reduce the need for specialized equipment to make it easier to establish the method in a standard cell biology laboratory. We also developed a free, open-source and easy to install image analysis framework in order to precisely measure the preferential positioning of sub-populations of cells within colonies of standard shapes and sizes. This method makes it possible to reveal the existence of patterning events even in seemingly disorganized populations of cells. The technique provides quantitative insights and can be used to decouple influences of the environment (e.g., physical cues or endogenous signaling), on a given patterning process.

The method presented here uses micropatterning together with quantitative imaging to reveal spatial organization within mammalian cultures. The technique is easy to establish in a standard cell biology laboratory and offers a tractable system to study patterning in vitro.

A fundamental goal in biology is to understand how patterns emerge during development. Several groups have shown that patterning can be achieved in vitro ...

Developmental Biology

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Neuroscience

Three-dimensional Imaging of Bacterial Cells for Accurate Cellular Representations and Precise Protein Localization

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be affected by cell shape. Bacterial cells are three-dimensional (3D) objects, although they are rarely treated as such. Most microscopy techniques result in two-dimensional (2D) images leading to the loss of data pertaining to the actual 3D cell shape and localization of proteins. Certain shape parameters, such as Gaussian curvature (the product of the two principal curvatures), can only be measured in 3D because 2D images do not measure both principal curvatures. Additionally, not all cells lie flat when mounting and 2D imaging of curved cells may not accurately represent the shapes of these cells. Accurately measuring protein localization in 3D can help determine the spatial regulation and function of proteins. A forward convolution technique has been developed that uses the blurring function of the microscope to reconstruct 3D cell shapes and to accurately localize proteins. Here, a protocol for preparing and mounting samples for live cell imaging of bacteria in 3D both to reconstruct an accurate cell shape and to localize proteins is described. The method is based on simple sample preparation, fluorescent image acquisition, and MATLAB-based image processing. Many high-quality fluorescent microscopes can be simply modified to take these measurements. These cell reconstructions are computationally intensive and access to high-throughput computational resources is recommended, although not necessary. This method has been successfully applied to multiple bacterial species and mutants, fluorescent imaging modalities, and microscope manufacturers.

This protocol explains how to prepare and mount bacterial samples for live three-dimensional imaging and how to reconstruct the three-dimensional shape of E. coli from those images.

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be ...

Immunology and Infection

Super-resolution Imaging of the Bacterial Division Machinery

Bacterial cell division requires the coordinated assembly of more than ten essential proteins at midcell1,2. Central to this process is the formation of a ring-like suprastructure (Z-ring) by the FtsZ protein at the division plan3,4. The Z-ring consists of multiple single-stranded FtsZ protofilaments, and understanding the arrangement of the protofilaments inside the Z-ring will provide insight into the mechanism of Z-ring assembly and its function as a force generator5,6. This information has remained elusive due to current limitations in conventional fluorescence microscopy and electron microscopy. Conventional fluorescence microscopy is unable to provide a high-resolution image of the Z-ring due to the diffraction limit of light (~200 nm). Electron cryotomographic imaging has detected scattered FtsZ protofilaments in small C. crescentus cells7, but is difficult to apply to larger cells such as E. coli or B. subtilis. Here we describe the application of a super-resolution fluorescence microscopy method, Photoactivated Localization Microscopy (PALM), to quantitatively characterize the structural organization of the E. coli Z-ring8.
	PALM imaging offers both high spatial resolution (~35 nm) and specific labeling to enable unambiguous identification of target proteins. We labeled FtsZ with the photoactivatable fluorescent protein mEos2, which switches from green fluorescence (excitation = 488 nm) to red fluorescence (excitation = 561 nm) upon activation at 405 nm9. During a PALM experiment, single FtsZ-mEos2 molecules are stochastically activated and the corresponding centroid positions of the single molecules are determined with &lt;20 nm precision. A super-resolution image of the Z-ring is then reconstructed by superimposing the centroid positions of all detected FtsZ-mEos2 molecules.
	Using this method, we found that the Z-ring has a fixed width of ~100 nm and is composed of a loose bundle of FtsZ protofilaments that overlap with each other in three dimensions. These data provide a springboard for further investigations of the cell cycle dependent changes of the Z-ring10 and can be applied to other proteins of interest.

We describe a super-resolution imaging method to probe the structural organization of the bacterial FtsZ-ring, an essential apparatus for cell division. This method is based on quantitative analyses of photoactivated localization microscopy (PALM) images and can be applied to other bacterial cytoskeletal proteins.

Bacterial cell division requires the coordinated  assembly of more than ten essential proteins at midcell1,2. Central  to this process is the formation of ...

Mammalian Cell Division in 3D Matrices via Quantitative Confocal Reflection Microscopy

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