Biology
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Analysis of LINE-1 Retrotransposition at the Single Nucleus Level
Chapters
Summary
Here we employ FISH methodology to track LINE-1 retrotransposition at the single nuclei level in chromosome spreads of HepG2 cell lines stably expressing synthetic LINE-1.
Transcript
The overall goal of this procedure is to visualize and analyze L-1 retrotransposition rates and patterns at the single nucleus level using fluorescence in situ hybridization, or FISH. The major advantage of this methodology is the fact that it allows the experimentalist to avoid over-or underestimation of rates and patterns of retrotransposition, and when complemented with other methodologies, allows for the study of L-1 retrotransposition at the single nuclei level. Performing this procedure today is Dr.Pasano Bojang, a senior fellow in the laboratory.
To begin, after preparing a 0.7%to 1%agarose gel in TAE buffer according to the text protocol, use CY3 or FITC to label the SNeo probe and incubate at 37 degrees Celsius for one hour according to the manufacturer's protocol. Mix the labeled SNeo probe solution with 1X DNA loading buffer and load into the wells of the agarose gel. Then run the gel at 75 to 100 volts for 15 to 20 minutes.
Using a UV illuminator, visualize the probe band. And with a clean blade, cut the labeled SNeo probe from the gel. Then use a PCR cleanup kit according to the manufacturer's protocol to solubilize and purify the SNeo probe.
After purification, rerun five microliters of SNeo labeled probe with an unlabeled SNeo probe on a 0.7%agarose gel to confirm the size increase and loss of signal intensity. To quantify the amount of labeled SNeo probe, measure the absorbance at 260 nanometers and dilute the SNeo probe to 10 nanograms per microliter aliquots in nuclease-free water. Store the aliquots at negative 20 degrees Celsius.
After generating stable clones expressing retrotransposition-competent L-1, or vector backbone, grow HepG2 cells stably expressing the control, or L-1, vector. Add 0.4 micrograms per milliliter of colcemid to the culture medium. And incubate for 90 minutes to arrest cells at metaphase.
Use 10 milliliters of 1X Dulbecco's PBS, or DPBS, to wash the cells two times. Then add 0.25%trypsin solution and incubate for five minutes. Media containing 10%FBS was added to inactivate the trypsin.
Next, transfer the cells to a 15 milliliter tube and centrifuge at 1, 000 Gs and four degrees Celsius for two minutes. Then aspirate the medium from the cell pellet and use DPBS to wash the cells. Aspirate all but 200 microliters of DPBS and use it to resuspend the cells by flicking or gentle pipetting to ensure cells are mixed well and contain no clumps.
While rotating the tube horizontally, in a drop-wise manner, add five milliliters of prewarmed hypotonic solution and incubate at 37 degrees Celsius for 20 minutes. Centrifuge the cells at 120 Gs and four degrees Celsius for five minutes. Then remove all but 200 microliters of hypotonic solution.
Wash with fixative solution three times, leaving approximately 200 microliters of fixative solution in the cells between washes. After the three washes, check that the pellet is visibly white and swollen. Then, with 200 microliters of Carnoy fixative solution, resuspend the pellet, and use the fixative to make the following dilutions of the resuspended cells.
Drop 10 microliters of each dilution onto a dry, clean slide from approximately one centimeter above, and immediately expose the spread-free side of the slide to hot steam from boiling water for 30 seconds. After drying the spreads at room temperature, stain with DAPI-containing mounting medium and mount a cover slip. Following the incubation, use DPBS to wash the slides three times.
To view the spreads, use a fluorescence-based contrast microscope at 40X magnification. After optimizing spread preparation, it is not necessary to stain them. To mark good spreads, use a diamond point marker on the opposite side of the slide.
To stabilize and dehydrate metaphase chromosome spreads, apply 200 microliters of RNase A to the samples, and incubate at 37 degrees Celsius for one hour. With 2X SSC, wash the slides two times for five minutes each, and use 10 millimolar HCl to rinse the samples. Then add 1%pepsin to the spreads and incubate at 37 degrees Celsius for 10 minutes.
After using deionized water to rinse the slides, use 2X SSC to wash the samples two times for five minutes each. Then add 4%paraformaldehyde to the spreads and incubate for 10 minutes, before carrying out two additional 2X SSC buffer washes. Following the SSC washes, dehydrate the spreads by incubating the samples for two minutes each in the following ethanol series.
Add 30 nanograms of SNeo to hybridization buffer and heat at 72 degrees Celsius for 10 minutes, before cooling at room temperature for two minutes. Then, pipette 30 microliters of the SNeo probe onto each spread, and use a cover slip to cover the sample. With rubber cement, seal the edges, ensuring no bubble formation occurs.
Keep the slide on a heat block at 72 degrees Celsius for five minutes. Then gradually drop the temperature to 37 degrees Celsius. Incubate overnight in the dark in a humidified chamber at 37 degrees Celsius.
To apply a secondary antibody, begin by immersing the slides in 2X SSC buffer to remove the cover slips. Then immerse the slides in 2X SSC at 45 degrees Celsius for five minutes to wash them, before transferring the samples to wash buffer and washing two times at 45 degrees Celsius for five minutes each. Next, immerse the slides in 0.1X SSC buffer at 45 degrees Celsius for 10 minutes.
Then wash the slides in 2X SSC buffer at 45 degrees Celsius for 10 minutes. After cooling the slides to room temperature, for direct labeled probes, use 2X SSC to wash the slides two times for five minutes each. Equilibrate the slides in detection buffer for 10 minutes at RT.Then add DAPI-containing mounting medium and counterstain the samples for 10 minutes.
Mount a cover slip and use nail polish to seal the edges. Finally, analyze L-1 retrotransposition using a fluorescence microscope at 40X magnification. As shown here, the density of metaphase chromosome spreads was determined by preparing dilutions and evaluating each spread for density, distribution, and distance of spreads from the nucleus.
Low-density spreads with even distribution were chosen for subsequent analysis. These images represent chromosome spreads that were stained with Hoechst dye DAPI, and the spread qualities were evaluated based on the length of chromosomes, the roundness of the spreads, and inter-chromosome distance. If cells were incubated for a short period in hypotonic solution, chromosome spreads became tightly knotted and individual chromosomes were difficult to visualize.
However, longer incubation in hypotonic solution resulted in rupture of the nuclei, scattering of chromosomes, and/or loss of chromosomes. Longer incubations in colcemid increased the number of cells in metaphase, but led to condensation of chromosomes. As illustrated here, a 90-minute incubation in colcemid, 20 minutes in hypotonic solution at 37 degrees Celsius, and the use of a hot steam to burst the cells were found to be optimal conditions for the generation of high-quality spreads.
In this experiment, chromosome spreads were stained for L-1 retrotransposition using probes that target SNeo. The probe was labeled with both FITC and CY3 to show that in both cases, L-1 retrotransposition is seen only in cells expressing wild-type L-1. Once mastered, this experiment can actually be completed in six to eight hours, especially if hybridization times are caught from two to four hours and the technology is performed properly.
While performing the technology, it is important to wash thoroughly, to optimize hybridization times and to chemically label the dNTP probes to avoid contamination. In order to answer additional questions like chromosomal location of the insertion, additional technologies like L-1 sequencing and G-banding can be completed. After watching this video, you will have a good understanding of how to label the L-1 probes, carry out the chromosomal spreads, and measure patterns and rates of L-1 retrotransposition utilizing fluorescence in situ hybridization technology.
This technology has paved the way for the study of patterns of L-1 retrotransposition as well as sites of L-1 insertion. It is important to remember that when working with toxic agents, biological reagents, as well as ultraviolet light, precautions such as wearing gloves and protective gear is essential.
Tags
LINE-1 Retrotransposition Single Nucleus Level Fluorescence In Situ Hybridization FISH Retrotransposition Rates Retrotransposition Patterns Agarose Gel TAE Buffer CY3 FITC SNeo Probe DNA Loading Buffer PCR Cleanup Kit UV Illuminator Size Increase Signal Intensity AbsorbanceRelated Videos
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