核小体阵列从重组核心组蛋白和核小体定位的DNA组装

This experiment aims to assemble nucleosomal arrays that contain a series of tandemly repeated nucleosome positioning DNA sequences with an equal number of nucleosomes first refold hisone complexes by dialyzing the core histones out of denaturing buffer. Then separate the histone emer using a size exclusion column. Next, combine the histone ER with template DNA at a range of molar ratios and deposit on the DNA as nucleosomes by step dialysis from high to low salt.

Next, test the samples by sedimentation velocity to determine the proper molar ratio for the desired level of saturation. Results can show the level of saturation of the nucleosomal arrays based on their migration through solution under centrifugal force, or based on an atomic force microscopy generated image. The main advantage of nucleosomal arrays rather than mono nucleosomes is that the nucleosomes are connected by linker, DNA, imposing an additional structural constraint.

Though this method can provide insights into chromatin structure, it can also be applied to other systems such as in vitro transcription study on chromatin templates. Ryan and Anna will now demonstrate the assembly of nucleosomal arrays from recombinant core histones in nucleosome positioning DNA. Begin with the purified lyophilized recombinant core hisone proteins H two A, H two BH three, and H four dissolve approximately five milligrams of each in three milliliters of unfolding buffer and incubate for at least one hour at room temperature with agitation.

Now measure the absorbance of each histone solution at 276 nanometers using unfolding buffer as a reference. Then calculate the molarity of each histone using their extinction coefficients. Compare the absorbent determined concentration of histone to the lyophilized dry weight and estimate if the majority of the protein is dissolved.

Identify the histone sample that contains the fewest moles as this will be the limiting number of moles for all histones. Then combine the histones in equal molar amounts. Dilute the histone mix with unfolding buffer to a final concentration of one milligram per milliliter.

Now transfer the sample into dialysis tubing. Seal the tubing and place into two liters of cold refolding buffer. Dialyze the sample for 18 hours at four degrees Celsius with stirring.

Ensure the dialysis tubing can rotate freely and vigorously to avoid his stone precipitation. Change the dialysis buffer every six hours, even if a precipitate forms do not discard the sample, some octi may be recovered, although the yield will be decreased. Connect the S 200 column to an FPLC system ensuring that air bubbles do not enter the system.

Clean the S 200 column overnight with 0.2 micrometer filtered water using a flow rate of 0.3 milliliters per minute and the back pressure limit of 0.5 megapascals next equilibrate the system with one liter of 0.2 micrometer filtered refolding buffer for about two hours. Also equilibrate a centrifugal concentrator with one milliliter of refolding buffer. Now remove the sample from the dialysis tubing.

Centrifuge the sample at four degree Celsius to remove any precipitates. Pipette the sample supernatant into the concentrator. Concentrate the sample to a volume of approximately 500 microliters.

Transfer the concentrated Optimus sample to a new container. Rinse the concentrator with one milliliter of Refolding buffer. Concentrate the rinse down to 500 microliters and add it to the ER sample.

Spin the sample in a 1.5 milliliter micro fuge tube for five minutes at 10, 000 RPM at four degrees Celsius. Transfer the supernatant to a new tube. Now load the sample onto the S 200 column.

Elute the sample with 400 milliliters of freshly made refolding buffer and collect two milliliter fractions. Monitor the absorbance at 280 nanometers during elution. His stone aggregates usually elute at approximately 45 milliliters EMR at about 65 milliliters and timer.

At around 84 milliliters. Analyze the elucian fractions from peaks of interest by running samples on an 18 to 20%SDS page. Fractions containing purified emus should have equal molar amounts of the four core histone proteins.

Now pool the fractions that contain purified OCRs and concentrate with a centrifugal filter to determine the final octr concentration. Measure the absorbance at 280. Nan calculate concentration using the extinction coefficient.

Reconstitute nucleosomal arrays from DNA and the purified ERs as detailed in the protocol text. Then dilute the sample to an absorbance of 0.5 at 260 nanometers. Load 400 microliters of the sample into a cell with a two sector centerpiece with the sample on one side and the reference 10 buffer on the other.

Although the level of liquid in both sectors should be close to the same. Keep the reference meniscus higher than the sample meniscus. Load the cells into the rotor and align the cells properly.

Using the hash marks on the bottom of the cells on rotor, gently dust the lenses of the cells using compressed air. Turn on the X-L-A-X-L-I centrifuge. Insert the rotor and attach and secure the optics as described in the manual.

Open the proteome lab software and select file new. Set up a single scan run at 3000 RPM at a temperature of 25 degrees Celsius. For each cell, name the samples, select a wavelength, select absorbance and choose a save file location on your computer.

Under options, select stop. After last scan and run radial calibration, begin the single scan run. Use the single scan to determine the appropriate radial scan length.

Using the X-L-A-X-L-I control panel. Enter a speed of zero and press star to equate the chamber. Using the software, set the desired speed and number of scans.

Higher centrifuge speeds increase the resolution of the sedimentation coefficient. However, one should collect at least 20 scans before the entire sample has sedimented to the bottom of the cell. Monitor the first couple of scans to ensure proper operation.

Overlay the last several scans in order to monitor the progress of the run and check for potential problems such as leaking cells, Analysis of sedimentation velocity results, as well as a protocol for visualizing arrays using atomic force. Microscopy are detailed in the protocol text. In this experiment.

Lyophilized xenopus core histones were reconstituted in the octr purified by FPLC on an S 200 column. The non-specific histone aggregate salute first followed by histone octr, H three, H four tetramers, and H two A.H two B dimers analysis of the purified octr fractions by SDS page indicates equimolar amounts of the four histone proteins. Note that xip H two A and H two B have molecular weights that differ by only about 200 daltons, and hence, they often appear as a single band on SDS gels.

Fraction 64 to 67 were pooled and saved for nucleosomal array reconstitution. A proteome lab software screen capture of a single scan collected at 3000 RPM was used to set the range of measurements for the a UC cell scan. Then a series of sedimentation velocity scans is edited to generate a data set for the red and green lines are two different methods for viewing the distribution of sedimentation coefficients.

In UltraScan. The red line is the integral distribution of sedimentation coefficients while the green is the derivative in study of chromatin organization. In vitro atomic force microscopy compliments analytical ultracentrifugation to establish the extent of template saturation after reconstitution.

Here, the majority of the 27 SRAs used in imaging contained 10 to 11 nucleosomes. This 500 by 500 nanometer scan clearly shows nucleosomal arrays with the linker.DNA. Also shown are the amplitude trace, the phase trace, and the same height trace with a free hand line drawn through the nucleosomes to determine the height profile of the nucleosomes Once mastered, this technique can be done in three to four days if performed properly.

While attending this procedure, it is very important to remember to be very precise measuring concentrations of your reagents Following this procedure, you can also test the saturation levels of nucleosomal arrays using techniques like atomic force microscopy After its development. This technique paved the way for researchers in the field of chromatin to study chromatin structure and chromatin compaction.