FAQ's & Technical Support

Nonacus is committed to providing high quality products and support. If you have a question about any of our products, please check our FAQ’s below or complete the support form at the bottom of the page and one of our team will be back to you as soon as possible.

Frequently asked questions

1. Cell3™ Target Library preparation: methods, equipment & reagents

I extracted and stored my cfDNA at -20˚C prior to starting the library prep. What should I be aware of before I start?

For best results, we recommend that you extract cfDNA and quantify just before you start your library prep. However, sometimes there is a need to extract and store cfDNA at an earlier time. In this case we advise you to store extracted cfDNA in a low-bind tube at -20˚C and quantify your samples after thawing when you are ready to start the library preparation.

Can I combine Cell3 Target enrichment probes with libraries prepared from library prep kits from other manufacturers? If, so, what issues do I need to be aware of?

We don’t recommend mixing Cell3™ Target enrichment probes with libraries prepared from other library prep kits because, we cannot guarantee the results. However, if you choose to do this you should be aware that some compatibility issues may arise. For example, the blockers in our kits are designed for our library prep kits and are for hybridisation-based kits with 8nt or lower indexing. Therefore, depending on the kit you used, you may see a high percentage of off target reads.

I am using Cell3™ Xtract for cfDNA extraction from plasma or serum. I notice that your elution buffer contains EDTA, which is incompatible with my downstream protocols. Can I elute in water instead of the supplied buffer?

Our elution buffer consists of 10 mM Tris-HCl, pH 8.5, 0.1 mM EDTA; therefore, this low level of EDTA should not affect most downstream applications, including NGS library prep. It  is perfectly fine to use water in place of the supplied buffer to elute your cfDNA. If doing so, please ensure that the pH is >6.0.  However, if you are storing your samples the buffer should be used. In addition, it is advisable to only extract cfDNA at the point where you plan to proceed with the library preparation.

I don't have a cycler where I can adjust the volume. Is this important?

We have successfully run the protocol using cyclers that have a set max volume of 50 or 100ul.

The maximum temperature for the lids of our cyclers is 99˚C. Is it possible to reduce temperature of the denaturation steps in step 1D (library amplification) to 95˚C with a lid heated to 99˚C?

A heated lid that has a max of 99˚C should work fine. However, we would not recommend adjusting the denaturation temperature to 95 ˚C in the library amplification step and recommend keeping this at 98 ˚C.

Is there a maximum time for the Hybridization reaction (Chapter 2)?

We recommend the hybridisation to run between 4 and 16 hours. However, we regularly run the hybridization overnight. For example, start the reaction at 4/5pm and then perform probe capture at 9 or 10am the following day. We have tested longer hybridisation times only up to 24 hours. If you want to break in between, you can store the dried down library pool with COT-1 DNA and universal blockers overnight at 4˚C if you want to delay starting the hybridisation reaction.

How should the beads look after drying? Are they always "fine" after 5 minutes? Or should I wait until they have cracks?

A. Often, we find that the beads dry quicker than 5 minutes, so it is advisable to check at 3 mins. Ensure that they are not shiny and try not to wait until they are cracking and overdried as beads are more difficult to resuspend and you may recover less of your sample (see Fig.1 below for an example)

beads

Figure 1. example of dried clean up beads showing matt appearance. Well E shows a slight sheen on the beads just before completely drying.

The Cell3 Target protocol suggests using 16 cycles for targets <0.04Mb. My targeted region is very small <0.01Mb. Should I still only conduct 16 cycles of post capture PCR?

Section 2C Captured library amplification, of the Cell3 Target protocol gives general guidelines on the suggested number of post capture PCR cycles. For very small targets the number of cycles would need to be increased. For example, 0.004Mb may need an additional 2 cycles (total 18) to ensure there is enough library for sequencing. Therefore, please note that for very small targets you may need to optimise the number of post capture PCR cycles.

The protocol states using M-270 Streptavidin beads. Can other types of beads, such as DynaOne beads be used instead?

The correct type of Dyna Beads is critical to the success of the capture. Only M-270 Streptavidin beads are suitable and will give the correct post capture yield. Any other beads are likely to be a different diameter and concentration and have shown to give a significant decrease in final yield.

I am preparing to do whole genome-seq library prep on FFPE DNA with low DIN values but high concentration. How much starting material would you advise I use for the library preps? Are there any parts of the protocol I would need to change for a whole genome seq library prep (vs exome-seq), such as fragmentation time?

As you have plenty of DNA for your WGS, and therefore not using the UMIs, we recommend performing a PCR free method to eliminate PCR bias. To do this you would need a minimum of 100ng of DNA to input and if your FFPE DNA has very low DIN scores (e.g. less than 3) you would need to increase your DNA input by 5-10 fold (see pg. 11 in Cell3 Target protocol).

I would like to run PCR free method of library prep. How is this done and what are the important points?

A.    The following are the steps for running a PCR free method for Cell3 Target library prep:

  1. Fragmentation/End repair/A-tailing (use 1 ul for tapestation to verify size)
  2. Ligation of adapters
  3. Bead clean-up
  4. Quantitation by qPCR (not Qubit and TapeStation)

When conducting a PCR free library prep, the QC at the end (following adapter ligation) will not give accurate Qubit readings (they will appear lower than what they should be) and the Tapestation will not show fragments in the right range. This is due to the ligation of Y-shaped adapters. The best way to QC is using qPCR. If you want to confirm the average fragment size, you could use 1 ul of the fragmentation reaction (post incubation) and run that on the tapestation. The average fragment length + adapters (144bp) will give the final library average fragment length.

What volume of probe set is included in the enrichment and capture kit for the Cancer 50 Panel?

Quantity of probe set is 18 ul for a 4x reaction kit for the cancer 50 panel. This allows you to prepare 4 x hybridisation pools with one probe set.

How many libraries can I pool together in one hybridisation reaction for the Cancer 50 Panel or the exome kit?

Library pooling guidelines for all custom and off the shelf products are listed on pg. 27 of the protocol. For the Cancer 50 panel up to 16 libraries can be pooled into 1 hybridisation reaction and you use 4 ul of probe set per reaction. For the exome panel, up to 8 libraries can be pooled. By all means, you can pool less than this number. However, you will need to ensure that the concentration of the combined pool reaches a total of 1000ng.

2. Cell3™ Target Library preparation: Quality controls (QC) & troubleshooting

I performed a QC step, using a TapeStation High Sensitivity D1000 screen tape, on my extracted plasma cell free DNA (cfDNA) prior to starting my library prep. What should I look for in the TapeStation analysis traces?

A. The TapeStation profile and yield of extracted cfDNA will vary depending on the input sample. The following descriptions and images give examples of these:

a) cfDNA extracted from cancer patients will be a mixture of cfDNA from the normal process of apoptosis and circulating tumour DNA (ctDNA) shed from the patient’s tumour. The amount of ctDNA shed into the blood varies with tumour type, stage and whether the patient has received treatment. There is a size difference observed in fragment length between cfDNA derived from the tumour and the patient’s normal background cfDNA. The normal cfDNA is generally around 166 bp (160-180bp peak), which corresponds to the length of DNA wrapped around the nucleosome and is likely to be the result of normal apoptosis. Peaks in multiples of 160-180 bp are often observed. The ctDNA portion in the sample is generally a smaller peak and reveal a shorter fragment length of approx. 145bp.

b) cfDNA extracted from patients with inflammatory conditions, infections, late stage cancer or transplant rejection may reveal larger peaks of cfDNA on the TapeStation analysis. You may see several peaks of differing sizes as multiples of 166bp (e.g. 332bp, 498bp, 664bp…).

profile-DNA

Fig.2. Example of cell free DNA profile for patient with an inflammatory condition

What QC steps do you recommend following extraction of DNA and prior to library preparation?

A. In all cases it is important to accurately determine the DNA concentration in your sample, especially when using <100 ng of DNA as input. To do this we recommend using a fluorometric method (such as the Qubit assay, Invitrogen).  Additionally, you may want to perform different QC steps on your input material according to your material type.

a) FFPE derived DNA may be compromised due to formalin fixation so we recommend performing a quality check by running your samples on a genomic screen tape (TapeStation) to determine the DIN (DNA integrity) score. This allows you to optimise the DNA input adding more material with lower DIN scores.

b) DNA derived from plasma may be contaminated with gDNA and the extent of this can be determined using a High Sensitivity D1000 Screen Tape on the TapeStation. Contamination with gDNA can arise during the processing of plasma due to haemolysis of white blood cells. A peak at >1500 bp is evidence of sample contamination with gDNA. You may also want to run this check to determine that the TapeStation profile reveals fragments of the expected size range (see Fig. 2 above).

The enzymatic shearing doesn’t look like it worked properly as the fragments are much longer than expected. (See image Fig.3 below for example).

enzymatic

Figure.3. Fragment size distribution of unsuccessful library prepared with 10 ng of input high molecular weight genomic DNA. The presence of a tail in the long fragment size range suggests that the sample was not entirely sheared during enzymatic fragmentation. The small 160 bp peak (indicated by the arrow) represents the presence of a small amount of adapter-dimers.

There are several possible causes for this. Accurate quantification of material and dilution upfront is a primary concern if you have an under fragmented library. Additionally, please ensure that the correct time and temperature have been used for fragmentation. If you are not using 10-100ng and this is your first use of Cell3™Target with this sample type, we recommend first trying 3 different times to find the best option for your sample type and input quantity.

Have you got EDTA in your DNA extraction elution buffer? EDTA affects the enzymatic shearing and causes poor/under fragmentation. DNA stored in EDTA buffers should be cleaned up with bead or column clean up prior to use. Ideally non EDTA elution buffers such as EB buffer (10mM Tris HCL pH 7.5-8) should be used.

Did you prepare the Enzymatic shearing reaction on ice and mix adequately before incubation? Mixing should be quite vigorous by pipetting or vortexing.

 

My TapeStation trace shows a peak at ~160bp representing adapter-dimers, what should I do?

A. A small amount of adapter-dimers seen at the pre-capture library preparation QC (as seen in fig.3 Q16 above) should not be carried over to the final library pool. If your libraries show a significant adapter-dimer peak (see fig. 4 below) this may be because you have input less than 50ng of DNA into the initial reaction without diluting the adapters to 1.5 µM (1:10 dilution). Always ensure you have accurately determined the concentration in your samples immediately prior to library preparation (see Q.1). You can attempt to reduce the adapter-dimer peak by performing a 0.9X (bead to sample ratio) clean-up step. However, this will result in sample loss and you will need to perform additional PCR cycles to ensure you have sufficient material to continue.

adapter-dimers

Figure 4. Library preparation from cfDNA showing significant adapter-dimer peak at 152bp

3. Sequencing Cell3™ Target libraries

Can I sequence a mix of Cell3™ Target prepared libraries when some have been Enzymatically sheared and others Covaris sheared?

A. Yes. Provided all libraries are Cell3™ Target libraries and they all have different sample indexes then a mix of Enzymatically sheared and Covaris sheared is fine.

How many samples can I load on a NextSeq or NovaSeq to achieve 2X coverage for a whole human genome?

A. Sequencing coverage can be estimated by using the Illumina Sequencing Coverage Calculator, which can be found at

https://support.illumina.com/downloads/sequencing_coverage_calculator.html

Below is an example for 2x coverage for WGS:

genome-table

Can I sequence libraries prepared from cfDNA together with libraries prepared from FFPE DNA?

A. This is possible; however, it is relevant to note that the read length you set may alter from your usual parameters. cfDNA is on average 166bp in length therefore, we usually recommend 2x75bp read length. For FFPE DNA fragments are often longer, and we would recommend a 2x150 bp read length. If you run at the longer read length you will sequence into adapter sequences and need to remove these during analysis. Alternatively, you can sequence all your libraries at the shorter read length of 2x75bp, however, this will impact the achievable depth of coverage for your FFPE DNA libraries.

4. Cell3™ Xtract & Cell3™ Preserver

My question relates to the plasma isolation step before using Cell3™ Xtract kit for cfDNA extraction.

Our own lab validated protocol uses a different sample "spin-down" protocol than the Nonacus recommendation. In our lab the second spin is 3000 g for 20 minutes, however the Nonacus protocol suggests a minimum of 10000 g for 10 minutes. We would like to run all samples together using the same protocol. Do you have any experience using different plasma isolation parameters and is the one we use still suitable for use with Cell3™ Xtract?

The protocol you are using would still work fine to remove excess intact cells in the 2nd spin however, it may not remove cell debris which usually floats around in plasma and requires higher speeds to pellet. In order to get a cleaner plasma sample, we recommend you perform the second spin at 10,000g or higher. Nevertheless, the Cell3 Xtract kit will still work with either isolation protocol.

I am noticing a pink colouration to my plasma when using Cell3™ Preserver tubes. I am concerned about contamination. Is this normal?

We regularly see a slight pink colouration to plasma that is extracted from our Cell3 Preserver tubes. This is nothing to worry about. Our tubes protect and prevent white blood cells (containing DNA) from breaking down, however they do not prevent completely the red cells with thinner walls (not containing DNA) from breaking down. The pink colour is due to some of the red blood cells breaking open and this does not affect or create any gDNA contamination.

5. Cell3™ Target: ordering & purchasing

Can I buy a combination of fragmentation and non-fragmentation kits or do I need to order a full kit for each?

A mixture of fragmentation and non-fragmentation kits can be purchased. For example, 48 fragmentation, 48 non-frag or 16 sample of one and 32 of the other so that both FFPE and cfDNA can be prepared.

6. Cell3™ Direct

What qPCR machines are compatible with Cell3™ Direct?

A full compatibility matrix is available here or alternatively can be found in the protocol for a specified product, all protocols can be downloaded in the product specific resource section.

My Non Template Controls (NTCs) or negative controls are amplifying?

Cell3™ Direct qPCR technology is extremely sensitive, as few as 1-3 copies can be amplified. Amplification in the NTCs or negative controls that is random in nature is likely due to low level contamination. We recommend the following actions:

  • Use clean working practises to minimise the potential of template contamination.
  • Where possible use separate areas for PCR mix preparation, template addition and qPCR reaction running.
  • Set up qPCR reaction in a PCR or biological safety cabinet using sterile / PCR-clean equipment and consumables only.

How should I analyse my results?

Each qPCR machine is different and therefore we recommend validation of our kit in each laboratory. However, we have provided some guidelines that in most instances provide a robust analysis approach.

Cell3™ FS Direct Analysis Guidelines v1.2

Cell3™ RHD Direct Analysis Guidelines v1.3

What is an inconclusive result and what should I do if I see one?

An inconclusive result is where there are not enough markers to be certain that the sample is positive, but some replicates have been amplified and therefore it is not clearly a negative result. This could be for a number of reasons including low level contamination, poor PCR efficiency or there is simply not enough cell free fetal DNA to amplify reliably. If an inconclusive result is obtained we recommend re-running the assay direct from plasma or alternatively extracting cell free DNA from the remaining plasma and running using the cell free DNA extracted protocol option.

Technical Support

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