Monarch HMW DNA Extraction from Tissue: Protocol Overview
In this video, we walk through the protocol for extraction of high molecular weight DNA (HMW DNA) from tissues.
In this video, we are going to walk you through the protocol for high molecular weight DNA extraction from animal tissues using the Monarch HMW DNA Extraction Kit. This kit enables the isolation of very large DNA fragments, even megabases in size when working with soft organ tissues.
The protocol is broken into two parts: Tissue Homogenization & Lysis and DNA Binding and Elution.
First, you will need to prepare a master mix of the Tissue Lysis Buffer and Proteinase K. For sample homogenization, you can use either the included microtube pestle or a rotor-stator homogenizer. The pestle allows for effective homogenization without compromising DNA integrity.
When working with soft organ tissues like liver or kidney, pestle homogenization allows for isolation of DNA ranging into the megabases. Rotor-stator homogenization is a convenient solution but usually will not produce DNA much larger than 250 kb.
First, we’ll demonstrate homogenization with a pestle. Start with the desired amount of tissue in a pestle tube and make sure keep fresh samples on ice and frozen samples on dry ice.
Frozen samples should be thawed briefly at room temperature first. To begin the homogenization procedure, spin down the sample in a benchtop minicentrifuge to collect all tissue material at the bottom of the tube.
Grind the sample thoroughly using the pestle to get the thinnest possible layer of tissue. This will help speed up lysis and maximize DNA fragment length. Leave the pestle in the tube. Use a wide-bore pipette tip to add 600 µl of the lysis master mix to the sample. Do not dispose of this tip yet, as it will be used to mix the sample later.
Make sure there is no tissue material left on the pestle, and if there is, transfer it carefully into the tube by wiping the pestle tip along the rim of the tube. Use the wide-bore tip to pipette the lysate up and down a few times. This ensures all tissue pieces are lysed efficiently, which is essential for obtaining the highest DNA integrity.
In this example with mouse kidney powder, you can see that after homogenization, the lysis is very close to being complete. Discard the pipette tip and immediately place the sample in the thermal mixer set at 56°C and at the appropriate agitation speed to start lysis.
Now we will demonstrate homogenization using a rotor-stator homogenizer. Make sure your sample is in a compatible 2 ml tube and add 600 µl of the lysis master mix. Flick or vortex to mix, and make sure nothing is stuck to the walls of the tube.
Insert the tip of the probe into the sample and homogenize on the lowest setting until the tissue pieces are no longer visible; this usually takes 5-10 seconds. Stop as soon as the lysate begins to foam. Some optimization may be required to find the best conditions for your sample type.
Immediately transfer the sample to a 1.5 ml Pestle Tube using a wide-bore pipette tip and incubate in the thermal mixer. Regardless of the homogenization method used, samples should be incubated in a thermal mixer at 56°C for 45 minutes with agitation at the desired speed.
When working with brain, muscle or any low input samples, stop agitation after 15 minutes for best yields; finish the incubation without shaking.
When lysis is complete, samples will have changed from turbid to clear or mostly clear, depending on the tissue type.
The speed of the thermal mixer influences fragment length and lysis efficiency; higher agitation speeds reduce DNA size and sample lysis time. For most applications, including ligation-based nanopore sequencing, maximum agitation speed is recommended and will produce DNA fragments predominantly 50 to 250 kb and ranging up to ~500 kb.
For maximum DNA length, agitate at 500–700 rpm, while mixing the samples by manually inverting every 10–15 minutes to optimize the lysis efficiency. With these low agitation speeds, tissue samples are digested very slowly, and the resulting DNA will be difficult to handle and more challenging to dissolve after elution.
Once the lysis incubation is finished, the removal of RNA can be addressed. Add 10 µl RNase A and invert 5-10 times to mix. Incubate for 10 minutes at 56°C at the same agitation speed previously used.
After this incubation, the lysis is complete, but the Proteinase K treatment typically does not remove all of the protein. Therefore, an additional protein removal step is needed before the DNA can be bound to the beads. Add 300 µl of Protein Separation Solution and invert the sample for 1 minute.
Centrifuge the sample for 10 minutes to create a phase separation. The sample will separate into a large, clear phase containing the DNA on top and a smaller, often brown, phase containing protein on the bottom. For some tissues, the protein phase may be green, yellow or even clear.
Sometimes additional centrifugation time may be needed to facilitate the phase separation, especially when low agitation speeds were used for lysis.
In the meantime, change the heat block in the thermal mixer to 2 ml block, and preheat it to 56°C. It is helpful to prepare and label all the plastics that you will need for part 2 during this centrifugation.
The top layer contains the DNA, but a substantial amount of the DNA will be located close to the interface between the clear upper phase and the lower protein phase. Therefore, it is important to transfer as much of the upper phase as possible in the upcoming transfer step. Use a 1000 µl wide bore tip first to get most of the sample and transfer it to a new Monarch 2 ml Tube. Then, use a 200 µl wide bore tip to transfer the rest of the sample.
Here are some additional tips for the phase separation. For low input samples, use a 200 µl wide-bore tip for the entire transfer. If a lower phase is not visible, leave 30 µl behind to prevent protein carry over. For liver samples, the addition of sodium chloride helps with glycogen removal; see our detailed guidance in the product manual.
Avoid transferring any material from the protein layer, although 1–2 µl will not be detrimental. If a small amount of the protein phase enters the pipette tip, gently push it back into the tube. Typically, the transferred volume will be around 800 µl, or 400 µl for low input samples. If you notice that the volume you’ve transferred is significantly lower, be sure to adjust the volume of isopropanol in the next step so that it equals 0.7 volumes.
Using clean forceps, add 2 DNA Capture Beads to each sample. Then add 550 µl of isopropanol, close the cap, and mix on a vertical rotating mixer at 10 rpm for 5 minutes to attach the DNA to the beads.
If you don’t have access to a vertical rotating mixer, you can invert the sample manually 25-30 times, slowly and gently. A manual inversion is complete when the tube returns to the upright position, and each inversion should take around 5 or 6 seconds. If you carry out the inversions slowly enough, no beads will stick to the bottom of the tube.
After a few inversions, the solution becomes more viscous and the DNA will wrap loosely around the beads. During the following inversions, precipitation of DNA may be visible, especially with larger sample inputs. The DNA complex will often contain small air bubbles and with more inversions, the DNA will completely wrap around the beads, often causing them to stick together. Once the DNA is completely wrapped around the beads, the viscosity of the solution will drop back to normal levels.
After the inversions are complete, remove and discard the liquid by pipetting, and there are two ways to do this. The first option is to keep the tube upright and insert pipette tip, gently pushing the beads aside to remove the liquid. The second way is to angle the tube so that the beads remain at the bottom, and liquid reaches toward tube opening. Then pipette from the liquid surface and continue to tilt the tube as you remove the liquid.
Add 500 µl of gDNA Wash Buffer, close the cap, and mix by inverting 2–3 times. Then, remove the wash buffer by pipetting, using one of the methods just described. Repeat this wash step and once again remove the buffer. This time, you can pour the buffer out using the pipette tip to ensure the beads stay in the tube.
Make sure the bead retainer is inserted into a collection tube and pour the beads into the bead retainer. Discard the used 2 ml tube. A quick pulse spin removes residual wash buffer efficiently, and there is no drying step necessary.
Separate the bead retainer from the collection tube and pour the beads into a new, labeled 2 ml Tube. Insert the used bead retainer into a labeled low bind 1.5 ml microfuge tube – this will be used for elution. You can now discard the used collection tube.
Add 100 µl of elution buffer onto the glass beads and incubate for 5 minutes at 56°C in a thermal mixer with agitation at the lowest speed. Halfway through this incubation, take the sample out of the incubator and tilt the tube almost horizontally and shake it gently. This ensures that the beads can move freely, allowing for complete release of the DNA from the beads. Place the sample back into the thermal mixer and finish the incubation.
Make sure the bead retainer is inserted into the 1.5 ml microfuge tube. After the 5-minute incubation, pour the eluate and the glass beads into that bead retainer and close the cap.
Centrifuge for 30 seconds to separate the eluate from the glass beads. When taking the samples out of the centrifuge, remove and discard the bead retainer with the beads and close the microfuge tube containing your eluate.
Before measuring or using your high molecular weight DNA, you’ll need to ensure it is uniformly dispersed. To do this, first pipette up and down 5-10 times with a wide bore pipette tip. Next, you have three options to help the DNA return to its natural conformation in solution. First, you can heat at 37°C for 30 minutes to 1 hour. Alternatively, you can leave at room temperature overnight. The third option is to leave at 4°C for at least 24 hours.
Any time you want to quantify your DNA, it is important to homogenize the sample by pipetting up and down with a wide bore pipette tip in order to get accurate measurements. If your sample is stored at 4°C, spin down the sample before pipetting to collect any droplets that have formed in the tube. We provide more detailed guidance on this online and in the product manual.
If you have any questions or need help with your preps, our technical support scientists are happy to help; contact us at firstname.lastname@example.org