Bioengineering
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Mammalian Cell Division in 3D Matrices via Quantitative Confocal Reflection Microscopy
Chapters
Summary November 29th, 2017
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.
Transcript
The overall goal of this procedure is to study mammalian cell division and cell matrix interactions in a physiologically relevant 3D environment. The main advantage of this technique is that it provides an efficient and general approach to study mammalian cell division and cell matrix interactions. This method can help answer key questions in cell biology such as the molecular basis of the development of normal and disease tissue.
A day prior to transfection, plate five million HEK-293 T-cells in a 100 milliliter cell culture dish in 10 milliliters of normal cell culture medium. Incubate these cells at 37 degrees Celsius for 24 hours in a humidified incubator with 5%carbon dioxide. The next day just before starting transfections, remove a vial of the transfection reagent from four degrees Celsius and leave it out to reach room temperature.
Then to start, add one milliliter of reduced serum medium to a sterile microfuge tube. To this, add specified amounts of lentiviral vector plasmids used for transfection and mix by pipetting. Incubate the tube at room temperature for five minutes.
After five minutes, add 48 microliters of the transfection reagent into this tube and mix gently by pipetting. Subsequently, incubate this mixture at room temperature for at least 15 minutes. Once the incubation is complete, remove the plate with HEK-293 T-cells from the incubator.
Add the prepared DNA transfection reagent mixture drop wise onto the cells and gently swirl the plate to ensure even distribution in the plate. Place the plate back into the incubator. Approximately six hours after transfection, remove the plate and aspirate the medium.
Add 10 milliliters of fresh culture medium and transfer the plate back into the incubator. At multiple time points, 24, 48, and 72 hours post-transfection, harvest the culture medium containing the lentiviral particles in separate tubes. Next, filter the harvested culture medium through a sterile 0.45 micron filter to remove cellular debris.
Replace with fresh culture medium following each harvest. The filtrate containing the lentiviral particles can be used for transduction right away or can be stored at minus 80 degrees Celsius. Plate human breast carcinoma cells MDA-MB-231 in a 35 millimeter culture dish in normal culture medium.
Incubate these cells at 37 degrees Celsius for 24 hours in a humidified incubator with 5%carbon dioxide. The next day, aspirate the culture medium from the plate and add one milliliter of the lentiviral particles together with one milliliter of fresh culture medium. Incubate the cells in the incubator for about 16 hours.
After the incubation with viral particles is complete, remove the plate to aspirate the medium. Then add two milliliters of fresh medium and transfer back to the incubator for 24 to 72 hours. Check the expression of H2B-mCherry in these transduced cells using an epifluorescence microscope.
To start the synchronization experiments, first plate MDA-MB-231 cells expressing H2B-mCherry at 50 to 60%confluency in a 24-well plate. Incubate the culture at 37 degrees Celsius and 5%carbon dioxide for 24 hours. The next day, replace the culture medium with 0.5 milliliters of medium containing two millimolar Thymidine.
Leave the plate in the incubator for 24 hours. Once the incubation is complete, wash the cells three times with PBS to release them from the Thymidine treatment. Then leave the cells in 0.5 milliliters of normal culture medium for five hours in the incubator.
After this, to block these cells in mitosis, add Nocodazole to the culture medium. Leave the cells in the incubator for 12 hours. After the Nocodazole treatment is complete, take out the plate and using an orbital shaker, shake the cells for 45 seconds to one minute to release mitotic cells.
Subsequently, transfer the medium containing the extracted mitotic cells into a new centrifuge tube. Then add 0.5 milliliters of fresh medium to each well. After, repeat the shaking and extraction of cells two more times.
Pool the total collected medium and then centrifuge it to pellet the mitotic cells. After the spin, resuspend the cell pellet in 0.25 milliliters of fresh cell culture medium. Using a hemocytometer, count the number of cells.
At the start of this step, prepare 50 milliliters of 10x DMEM solution and filter sterilize it. Next, determine the volumes of all the components that will be used to make the 3D collagen matrix as specified in the text protocol. Pre-chill all these components on ice to slow down the gelation of collagen.
Now, to a pre-chilled 1.5 milliliter centrifuge tube, add the required numbers of cells in medium then 10x DMEM, FBS, deionized water, collagen, and finally sodium hydroxide. Mix carefully using a one milliliter pipette. Once the solution is well mixed, add 500 microliters of the mixture into each well of the 24-well plate.
Place the 24-well plate in a 37 degree Celsius incubator. After 30 minutes, take out the plate and add 500 microliters of pre-warmed culture medium on top of the gel. Prior to imaging, turn on the live cell units of the phase contrast and confocal microscopes.
To capture dividing cells, place the 24-well plate containing the cells embedded in collagen matrices into the live cell unit of the fluorescence microscope. Using the 10X objective, find the bottom of the plate and then move it up until the microscope focuses at about 500 micrometers from the bottom of the plate. Move the stage around to find cells in different stages of mitosis and select multiple positions for imaging.
Set up the timelapse experiment with two-minute intervals. Next, to image the collagen network, configure the confocal microscope to capture only reflected light from the 488 nanometer laser. Then place the plate containing cells in collagen matrices into the live cell unit of this microscope.
Find the bottom of the plate and then move the objective lens up until it focuses at about 100 micrometers from the bottom of the plate. Move the stage around to find cells and select multiple positions for imaging. Set up the timelapse experiment with five-minute intervals.
Fluorescent images of a dividing H2B-mCherry expressing MDA-MB-231 cell embedded in collagen matrix are shown here. Different phases of mitosis are readily observed as expected. The distribution of collagen fibers in white visualized simultaneously using confocal reflection microscopy changes very little during the course of mitosis.
Cell synchronized G2-M phase and embedded in a collagen matrix complete cell division in the first two hours while control asynchronous cells divide randomly. Quantification of the distribution of collagen fibers during interphase and mitotic phases suggest that matrix deformation is minimal during cell division and higher in interphase cells. Once mastered, this technique can be completed in seven days starting from generating the stable cell line to analyzing the video cell division in a 3D matrix.
After watching this video, you hopefully have a good understanding of how to synchronize and monitor mammalian cell division in 3D matrices using live cell imaging and how to determine matrix deformation using confocal reflection microscopy and quantitative imaging analysis techniques.
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