Simultaneous Mapping and Quantitation of Ribonucleotides in Human Mitochondrial DNA

Here we describe a method amenable to simultaneously quantitate and genome-wide map ribonucleotides in highly intact DNA at single-nucleotide resolution, combining enzymatic cleavage of genomic DNA with its alkaline hydrolysis and subsequent 5´-end sequencing.

Transcript

The overall goal of this experiment is to map the location of incorporated ribonucleotides at single nucleotide resolution level, and to quantitate the number of ribonucleotides present in human mitochondrial DNA. This method can help answer key questions regarding DNA integrity, DNA replication mechanisms, DNA repair mechanisms, and how defects in any of these mechanisms make cause disease. The main advantage of this method is that both the location and the number of incorporated ribonucleotides in mitochondrial DNA can be determined in a single experiment.

Though this method can provide insight into ribonucleotide content in human mitochondrial DNA, it can also be applied to study nuclear DNA and ribonucleotide content in sequence genomes of other model organisms. Genomic DNA previously isolated from HeLa cells and described in sections two and three of the text protocol is used in this experiment. To digest the DNA, carefully mix the following in a microfuge tube.

One microgram of DNA, five microliters of 10X buffer 3.1, one microliter of HincII, and nuclease-free water to a final volume of 50 microliters. Incubate at 37 degrees celsius for 30 minutes. When the digest is completed, add 1.8 volumes of paramagnetic beads to each sample, carefully mixed by pipetting, and incubate at room temperature for 10 minutes.

Place the tubes on a magnetic rack for five minutes to pellet the beads, and then remove and discard the supernatant. Wash the pellet with 150 microliters of 70%ethanol for about 30 seconds, then remove and discard the supernatant. Repeat the wash with 200 microliters of 70%ethanol for another 30 seconds.

Dry the samples at room temperature for about 15 to 20 minutes. Next, remove the tubes from the magnetic rack. Elute each pellet in 45 microliters of elution buffer, mixed by pipetting carefully, and incubate for five minutes.

Pellet the beads on the magnetic rack, and transfer 45 microliters of each DNA sample to a new tube. To each 45 microliters of isolated or purified DNA, add five microliters of three molar potassium hydroxide or five microliters of three molar potassium chloride, creating a total volume of 50 microliters. The eight reactions are summarized in this table.

Incubate in a hybridization oven at 55 degrees celsius for two hours, then incubate the samples on ice for five minutes. Precipitate the DNA by adding 10 microliters of three molar sodium acetate at pH 5.2, and 125 microliters of cold, 100%ethanol and incubate on ice for five minutes. Pellet the DNA by centrifuging at 21, 000 times G at four degrees celsius for five minutes.

Discard the supernatant and wash each DNA pellet with 250 microliters of cold, 70%ethanol, centrifuge again, and discard the supernatant. Let the pellet dry in the open tube for about five to 10 minutes, until any visible fluid has evaporated. Lastly, add 20 microliters of elution buffer, and let the DNA pellet dissolve at room temperature for 30 minutes.

Begin this procedure by preparing the reaction mix for each sample as a master mix. For each sample, add 2.5 microliters of 10x T4 polynucleotide kinase reaction buffer, 2.5 microliters of ATP, and one microliter of three prime phosphatase-minus T4 polynucleotide kinase. Transfer 19 microliters of each DNA sample into a new 200 microliter tube, and denature at 85 degrees celsius in a thermocycler for three minutes.

Then, cool the DNA samples on ice, and add six microliters of the prepared reaction mix to each sample. Incubate the reaction mixes at 37 degrees celsius for 30 minutes, followed by 65 degrees celsius for 20 minutes to stop the reaction. Purify the DNA using paramagnetic beads, as demonstrated previously, but elute with 14 microliters of elution buffer.

Prepare the reaction mix as a master mix in advance, containing the following for each sample. Five microliters of 10x T4 RNA ligase reaction buffer, five microliters of hexamminecobalt(III)chloride, 0.5 microliters of ARC140 oligonucleotide, 0.5 microliters of ATP, and 25 microliters of 50%PEG 8000. Mix well by pipetting.

Transfer 13 microliters of each purified DNA to a new 200 microliter tube, and denature at 85 degrees celsius in a thermocycler for three minutes. Cool the DNA on ice, and then add 36 microliters of reaction mix to each sample, mix by pipetting, and spin down briefly. Add one microliter of T4 RNA ligase to each reaction, mix by pipetting, and spin down briefly.

Incubate the samples at room temperature in the dark overnight. When the single strand DNA ligation is complete, purify the DNA as demonstrated earlier, but use 0.8 volumes of paramagnetic beads and pellet the beads for 10 minutes. Elute in 20 microliters of elution buffer, and transfer 20 microliters of each DNA sample to a new 200 microliter tube.

Repeat the purification with 0.8 volumes of paramagnetic beads, and elute in 14 microliters of elution buffer. For each sample, prepare the reaction mix as a master mix consisting of two microliters of 10x T7 DNA polymerase reaction buffer, two microliters of ARC76/77, two microliters of dNTPs, and 0.8 microliters of BSA. Transfer 12.8 microliters of purified DNA to a new 200 microliter tube, and denature at 85 degrees celsius in a thermocycler for three minutes.

Cool the DNA on ice, and then add 6.8 microliters of reaction mix to each sample, mix by pipetting, spin down briefly, and incubate at room temperature for five minutes. Add 0.4 microliters of T7 DNA polymerase to each reaction, and incubate at room temperature for five minutes. After purifying the DNA using paramagnetic beads, elute in 11 microliters of elution buffer.

To begin the PCR amplification, prepare the reaction mix separately for each sample in a new 200 microliter tube consisting of 7.5 microliters of ARC49, 7.5 microliters of unique index primer, and 25 microliters of 2X hot start ready mix. Add 10 microliters of DNA sample to each reaction. Amplify the library using the following conditions.

Denature at 95 degrees celsius for 45 seconds, followed by 18 cycles of 98 degrees celsius for 15 seconds, 65 degrees celsius for 30 seconds, 72 degrees celsius for 30 seconds, and ending with a final elongation at 72 degrees celsius for two minutes. Hold the samples at four degrees celsius after amplification. Purify the libraries as described in the text protocol and elute in 20 microliters of TE buffer.

Determine the quality and average fragment size of each library using a digital electrophoresis system. Representative results of suitable library profiles after potassium hydroxide or potassium chloride treatment are shown. After calculating the concentration of each library, pull equal molar amounts of up to 24 libraries amplified with different index primers for sequencing as described in sections nine and 10 in the text protocol.

Add TE buffer to a final volume of 25 microliters and concentration of 10 nanomolar. After sequencing, perform bioinformatic data analysis as specified in section 11 of the text protocol. These graphs show the summarized reads at all HincII sites in light strand and heavy strand human mitochondrial DNA after potassium chloride treatment.

Around 70%of all detected five prime ends localized to the cut sites, demonstrating the high efficiency of the HincII digestion. Treating libraries with potassium hydroxide to hydrolyze the DNA at embedded ribonucleotides decreases the number of reads at HincII to about 40%The relative number of reads in mitochondrial and nuclear DNA reveals differences in coverage and illustrates the importance of appropriate normalization to remove strand bias. Normalizing read counts to HincII gives a quantitative measure of the number of ribonucleotides per mitochondrial genome on the heavy or light strand.

The reads after potassium hydroxide treatment for each ribonucleotide, normalized to the sequence composition of each strand, show a ratio different than one, indicating a non-random distribution of reads and suggesting a distinct ribonucleotide pattern and a high library quality. Normalizing the reads at the sites of embedded ribonucleotides to those at HincII sites, as well as to the genome nucleotide content, generates a quantitative measure of how many of each ribonucleotide is incorporated per 1, 000 complimentary bases. Once mastered, this protocol can be done in two days if it is performed properly.

While attempting this procedure, it is important to remember to keep the DNA highly in tact and avoid introducing any double strand breaks during library preparation. Following this procedure, other methods like can be performed in order to confirm the ribonucleotide quantification data. After watching this video, you should have a good understanding of how to prepare libraries for the mapping and quantification of ribonucleotides and DNA using next generation sequencing.

Don't forget that working with chemicals like potassium hydroxide can be hazardous, and precautions such as wearing eye protection and gloves should always be taken while performing this procedure.

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Journal (Biology)

Journal (Biology)

Simultaneous Mapping and Quantitation of Ribonucleotides in Human Mitochondrial DNA

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