FAQs 2018-09-17T10:46:53+00:00
What are the differences between a ready-to-use panel and a custom panel? 2018-09-17T11:12:25+00:00

Ready-to-use panels are initiated and designed by the Paragon Genomics R&D team that aim to provide the best solutions to specific research areas and/or clinical needs. The contents of these panels, as well as the performance, are internally validated to ensure that it passes our high standards.
Custom panels are designed to satisfy the need of customers from all areas of research. It aim to strike the perfect balance of flexibility and quality. There is no limit on what the target species are and what the regions might be. Our design team will work with the customers to come up with the best NGS solution for the problem at hand. Thanks to the robust CleanPlex® chemistry, our custom panels delivers consistent high quality results often similar to our read-to-use panels. However, due to the diverse nature of the requests, it is often difficult to provide performance validations for these panels. Custom panels may require simple optimization in determining the best PCR cycles. If you would rather have our bench scientist perform these optimization steps for your custom panel, please contact [email protected] for a quote.

What types of custom panels can you design? 2018-09-17T11:13:09+00:00

We can design targeted panels for any target with a reference sequence.  Some examples of target species include humans, animals, plants, insects, bacteria, and viruses. Our sample types include, but not limited to, cfDNA, FFPE samples, blood samples, and tissue samples.

For bacterial panels, a minimum 7 amplicons is required. We can build 7 housekeeping genes to ensure all samples have something being amplified if you’re detecting infection.

For viral panels, the sequence information is very important. HIV for example has many variations and subtypes, which will be difficult to cover entirely. However for viruses with fewer sub-types and less variations, we can be more successful with the design. Please submit your design requests to the ParagonDesigner portal or email technical support if you have questions pertaining to your specific applications.

Is there a minimum number of targets for a custom panel design? 2018-09-14T18:11:36+00:00

Short answer is no. We have designed panels that contain as little as 2 amplicons and we can also offer single-amplicon options.
However, it is recommended that a panel contains at least 10 amplicons per pool, to best utilize our multiplex capabilities, and to achieve the best sequencing results from our technology.

What is the upper amplicon count limit for a custom CleanPlex panel and a CleanPlex UMI panel? 2018-09-17T11:12:02+00:00

We are yet to reach our limit of multiplexibility in our current portfolio of a wide range of panels.

For standard CleanPlex technology, we have designed and delivered panels of up to 20,000 amplicons (approx. 400-genes) with great performance. Panels with higher amplicon targets may require more primer pools, which in tern require more DNA. If the total sample quantity is limited, there could be a cap on how many targets can be assayed. 

For CleanPlex UMI technology, sequencing depth is the limiting factor for how many amplicon targets to include in your custom panel. UMI panels require deeper sequencing to fully utilize the resolving power of the technology. The sequencing reads required per sample is calculated based on how low the allele frequencies percentage you wish to detect, how much DNA input you need, and how many amplicons per panel. 

If you have special needs in creating large panels, please reach out to our design and sales team for evaluations, or our technical support team with any questions.

What is the uniformity of a typical CleanPlex target enrichment library? 2018-09-17T16:36:36+00:00

The uniformity ( ≥ 0.2X mean depth) of most CleanPlex libraries are greater than 95%. For example, the observed uniformity (at ≥ 0.2X mean depth) of the CleanPlex OncoZoom Panel is greater than 99% for both NA12878 (a genomic DNA standard) and Horizon Discovery’s HD780 (a cfDNA standard) (see plots below).

What are the AT and GC biases of CleanPlex target enrichment libraries? 2018-09-17T16:36:11+00:00

The observed AT and GC dropouts are below 3% in CleanPlex® ready-to-use libraries. Below is a typical plot of Coverage Depth vs. GC Content of a library made with CleanPlex® OncoZoom Panel.

Can I use CleanPlex technology to detect SNV, CNV, and Fusions? 2018-09-17T16:26:02+00:00

The CleanPlex system can detect SNV, CNV, and known Fusions. Unknown fusion detection on the other hand is not compatible with our chemistry.

How do I submit a custom panel design request? 2018-09-14T18:28:08+00:00

To submit a custom design request, you will need the following:
1. A Paragon Genomics customer account. Register here.
2. If you are targeting genomes other than human and mouse, be ready to provide the reference genome for design upon request;
3. If your targets are not human genes, please provide the target region in the format of a BED file.

With these ready, you can submit your design request by going to our Paragon Designer web portal here.

We would need to know the target regions and the sample type.

What are primer pools? How many pools do I need for custom panels? 2018-09-14T18:29:09+00:00

We design panels based on our ability to multiplex tens of thousands of amplifications in a single tube. Amplicons that cannot be multiplexed in the same tube are put in separate tubes. Each separate tube of mixture of primers is called a primer pool.
In common cases, hotspot targets that are far apart from one another on the genome can be put in the same pool, no matter how many amplicons are involved; exon targets, or any region targets that are longer than the maximum allowed amplicon length, will need 2 pools to cover fully the region(s) of interest. This is because overlapping amplicons are not compatible in the same multiplexed reaction mix.
In very rare cases, a third or fourth pool, or even larger number of pools might be needed. Adding more pools beyond a 2-pool panel usually helps very little in increasing the in silico coverage of the targets, but might help in other aspects of the panel, such as adding the flexibility of splitting a larger panel into smaller panels post-ordering.

How soon will I get my custom designs after submission? 2018-09-14T18:29:27+00:00

It depends on the difficulty of the design. We have a robust assay design algorithm that handles most designs relatively quickly. One can expect between 3 days to 2 weeks turnaround time (TAT) starting from the moment our design team start working on the requests. The following are the most common reasons for potentially longer design times:
1. non-human targets. We will need to verify and format the reference genome before we can start the design. Additional information might be needed on top of the genomic sequence in some cases;
2. modification of original request after the design has started. If the change to the design request involves change of amplicon length, adding or removing new targets or other changes that require a re-design of the panel, it is our rule that it will be viewed as a new panel request and we shall proceed with the modification only after we finished other customers request that we might be working on at the time.
3. difficult regions that require special attention. Our design usually have a >95% in silico design coverage. However, there are often cases where some regions cannot be covered based on the design requirements or due to the nature of the target sequences. If those regions are deemed ‘must-haves’ for a given design, we will need to treat them as special instances and may delay the delivery of the panel.

Does Paragon offer wetlab optimization services for custom CleanPlex and Cleanplex UMI panels? 2018-09-17T11:10:52+00:00

Yes, please contact [email protected] for a quote.

How do I place an order using a Purchase Order? 2018-09-13T16:18:32+00:00
  • Register an account and place an order online (US customers only)

    • You can pay with credit card at the last step of Checkout, or
    • You can use a Purchase Order number at the last step of Checkout (as shown below)
What are the key steps in the CleanPlex and CleanPlex UMI workflow that I should know prior to starting? 2018-09-14T18:30:44+00:00

Support documents

In addition to the User Guide, please download and review the Quick Guide from the Product Document page for details on some often overlooked but critical steps. You can also refer to the Videos page for the series of short videos that demonstrate the Cleanplex protocol step by step.

DNA Quality and Quantification

The quality and quantity of the starting input DNA is critical in determining the appropriate cycles numbers and cycling conditions in the following steps of the workflow. We recommend using a fluorometric method such as the Qubit High Sensitivity dsDNA kit to determine the concentration of your starting materials. Do not use a spectrometric method such as the Nanodrop because it can significantly overestimate your starting concentration. High quality sample and higher quantity input generally yield higher quality libraries.

Correct Cycling Conditions

Generally, a thermal cycler set with the highest ramp speed, such as 5°C/second and higher, is not recommended. For thermal cyclers with adjustable ramp speed, we recommend 3°C/second up and 2°C/second down speed, or use the default setting (no ramp adjustment).

Making Working Solutions & Vortexing Well

Ensure accurate assembly of working solutions by pipetting viscous  reagents carefully. Remove excess liquid from outside of the pipette tip and pipette mix as needed.
Mixing combined solutions is also CRITICAL. Vortex or pipette mix as needed to ensure all PCR working solutions, Digestion working solutions, and all bead+ sample solutions are homogeneous prior to starting the thermocycling or incubations.

Accurate Bead Dispense

The magnetic bead to sample volume ratio is critical to a clean library. Remove excess liquids from outside of tips and pipette mix as needed. Mix combined solution THOROUGHLY prior to incubation.

Minimize Loss

Cleanplex protocol is designed as a single tube workflow. With the exception of combining multi-pool panel reactions after the multiplex PCR step, solutions do not require tube to tube transfer, thus avoiding loss of sample and volume. Ensure the magnetic rack used is intended for the PCR tube or PCR plate being used. Avoid disturbing and removing beads during any of the washing steps.

Thorough Ethanol Washes

Ensure the cleanest libraries by carefully removing all droplets from supernatant and ethanol washes. This minimizes carry over of byproducts that would be amplified in the final PCR step.

How much DNA should I use to detect 1% somatic variant frequency? 2018-09-14T18:31:15+00:00

Lower DNA input will generally increase the possibility of inaccurate calling of variant frequencies. As shown in the following figure on the left, when less and less DNA was amplified with CleanPlex® OncoZoom Panel, increasingly larger variations were observed for calling alternative alleles at 50% frequency, even though the uniformity may not deteriorate significantly (figure below on the right). Therefore, we recommend using higher amounts of DNA for somatic variant detection. This is especially true when DNA quality is uncertain (such as DNA from FFPE tissues and liquid biopsy). Theoretically, 10 ng human DNA (~1500 cells and ~3000 copies of each locus) will allow detection of 30 somatic alternative alleles at 1% frequency. With CleanPlex® technology, lower amount (even less than 1 ng) of high-quality genomic DNA may be used for detection of germline alterations.

 

      

How much DNA should I use to detect 0.1% somatic variant frequency? 2018-09-19T11:58:44+00:00

With our CleanPlex UMI technoloy, one can detect variations down to 0.1% allele frequency. We recommend using 30-50 ng of genomic DNA input for somatic variant detection at 0.1%, or 20-30 ng for 0.25%.  More input (50-80 ng input) may be necessary to improve variant calling when DNA quality is uncertain (such as DNA from FFPE tissues and liquid biopsy). 

What is the CleanPlex or CleanPlex UMI workflow for a multi-pool panel? 2018-09-14T18:32:40+00:00

For a CleanPlex panel with two or more primer pools, follow the procedure shown below.

For CleanPlex UMI panels, the starting mPCR reaction is 40 uL, but is combined in a likewise fashion for total of 80 uL volume for downstream steps.

What should I expect my NGS library to look like on a fragment analyzer trace? 2018-09-14T18:34:02+00:00

Please see user guide for examples of each ready-to-use panels.

When using a fragment analyzer such as Agilent Bioanalyzer 2100, the library peak(s) should fall between 200-400bp, depending on the specific panel. A library can exhibit a single peak (such as OncoZoom) or multiple peaks (such as TP53) depending on the panel’s amplicon length distribution. Most importantly, look for sharp and well-defined peaks that span the base pair length range specific for the panel. Each custom panel design will also contain an “Amplicon Length Distribution Theoretical Plot” for the user to compare with the completed library.

The fragment analyzer most importantly allows the user to visualize any byproducts such as adapter dimers & digested non-specific products (~150-190bp) and 2nd PCR primer dimers (70-90bp). See following FAQs for details on how to reduce these byproduct peaks.

What are the peaks around 70 – 90 bp in addition to the main library peak on a fragment analyzer trace? 2018-09-13T16:39:20+00:00

Peaks from 70 – 90 bp (see the trace below) are primer dimers from the 2nd PCR that result from incomplete removal of small molecular materials during the final magnetic bead purification. These are usually caused by inaccurate pipetting of magnetic beads volume, poor mixing prior after adding magnetic beads, and/or incomplete removal of supernatant and ethanol during washing steps. If these peaks are significant, one can pool all indexed libraries (that will be sequenced in the same lane) and perform one additional round of 1.2X magnetic bead purification, before the quantification step prior to sequencing.

What is peak around 150 -190 bp in addition to the main library peak on the fragment analyzer trace? 2018-09-13T16:41:37+00:00

Peaks from 150 – 190 bp are residues of digested non-specific amplification products (see examples below).

They come from incomplete removal of small-molecular-weight materials during magnetic bead purification after digestion. The digestion reagent cuts non-specific amplification products into small pieces, which are then removed during magnetic bead purification. Peaks from 150 – 190 bp are usually caused by the following:

Insufficient mixing of solutions: 
Mix all working solutions and magnetic bead + sample solutions thoroughly prior to PCR and incubation steps. Vortex or pipette mix as needed to ensure solutions are homogeneous.
Inaccurate magnetic bead volume:
Again because we’re working with small volumes, it’s important to add accurate volume of beads (see 2nd page of quick guide for details). The specific bead ratio is critical for size selection in removing the smaller fragments, therefore it is crucial that the bead+sample solution is thoroughly mixed to ensure the bead to sample ratio is the same throughout.
Forgetting to add 10 uL of TE for single pool panels after mPCR:
This error is also related to bead to sample volume ratio. Add 26 uL of bead solution to 20 uL of sample ( 10ul of mPCR product + 10 uL TE) for the first purification step.
Poor Ethanol Washes:
With extremely poor ethanol washes, there can be significant carry over of byproducts to 2nd PCR step, and are then preferentially amplified. Take care to remove all supernatant after bead incubation (following the second quick spin step 5, 4, &4 on each of the purification sections), and the ethanol from the second ethanol wash of each purification step.
Extremely low or poor quality DNA input: 
First, minimize freeze thaw cycles for low concentration DNA samples (<10ng/uL). Always determine the DNA concentration immediately prior to library preparation instead of days before. Higher quality samples and higher sample input tends to yield higher quality libraries with less by products.
Some custom panels might be more likely to have these products as compared to other due to the unique nature of some custom designs. If this is the case, one can perform an additional 1.3x bead to sample volume purification after the mPCR step. Or one can pool all indexed libraries (that will be sequenced in the same lane) and perform one additional round of 1.2X magnetic bead purification, before the quantification step prior to sequencing.
What is the typical yield of a CleanPlex or CleanPlex UMI library? 2018-09-17T16:38:08+00:00

CleanPlex and CleanPlex UMI workflow generates a single peak of library of approximately 8,000 – 20,000 pM when measured with a fragment analyzer such as Agilent™ 2100 Bioanalyzer instrument and Agilent™ high sensitivity DNA reagents, or approximately 1.5 – 4 ng/µl when measured by Qubit™ dsDNA HS Assay Kit, depending on each specific panel. It is good practice to QC completed library with a fragment analyzer to confirm the quality of the library prior to sequencing. Qubit measurements only give the total yield, and does not indicate if the library was poorly prepared and had resulted in large amount of byproducts.

Concentrations higher or lower than the typical range of yields may lead to lower uniformity due to uneven amplification of target regions. Concentrations much lower in concentration can have difficulty satisfying Illumina’s loading criteria. Please refer to Illumina’s suggestions for minimum loading concentration and volume.

There is little or no library peak when assayed with a fragment analyzer. 2018-09-13T16:23:45+00:00

Possibilities include:

  • Using incompatible index primers.
  • DNA quantification was inaccurate, especially if using spectrophotometric methods such as nanodrop, OR DNA quality is extremely poor.
  • 30% ethanol instead of 70% ethanol OR using TE/H2O instead of 70% ethanol was used in DNA purification.
  • Forgetting to add magnetic beads for any of the purification steps.
  • Forgetting to add one or both index primers in the 2nd PCR step.
  • Over-digestion, forgetting to add Stop Buffer after digestion (for Cleanplex protocol only) , or pausing after digestion step.
  • Weak or incompatible magnetic rack. Do not use racks designed for 1.5 ml tube.
What should I do when the library yield is lower or higher than expected? 2018-09-14T18:35:40+00:00

If the library yield is low, but other wise clean and free from by-product peak(s), the library may require optimization in 2nd PCR cycles.  Increase or decrease the 2nd PCR cycles by 2 to 3.

If significant by-product peak is present, please refer to FAQ’s above for detailed troubleshooting suggestions.

What is the recommended minimum read coverage for Cleanplex NGS panels? 2018-09-14T18:36:25+00:00

For Genotyping applications, minimum read coverage per amplicon is 20X.

Although, we suggest starting with 500 reads/amplicon/sample for the first sequencing run. For following sequencing runs, the reads/amplicons can be significantly reduced based on the panel’s performance and sequencing quality.

For Somatic mutation applications, minimum read coverage is 500X for 1% allele frequency detection, and 200x for 5% allele frequency detection.

We suggest starting with 1000 reads/amplicon/sample for 1% allele frequency detection for the first sequencing run. For following sequencing runs, The reads/amplicons allocated can be significantly reduced based on the panel’s performance and sequencing quality.

What is the recommended minimum read coverage for Cleanplex UMI NGS panels? 2018-09-18T15:27:14+00:00

The suggested minimum read coverage is 4,000 paired-end reads/ amplicon/ nanogram of DNA input for detection of 0.1% allele frequency, when the minimum DNA input amount is also met. 4,000 paired-end reads equates to 2,000 clusters or single-end reads on the sequencing chip. Please see FAQ for DNA input requirement for detecting 0.1% somatic variant frequency.

What read length should I use on Illumina sequencers for CleanPlex and Cleanplex UMI libraries? 2018-09-18T16:53:52+00:00

All ready-to-use libraries are designed for 2 x 150 bp read length, meaning final library molecules with sequencer-specific sequences are < 300 bp.

Custom panels are typically also designed for 2 x 150 bp read length. However we can also design amplicons of 500 bp for 2 x 250bp reads. Please specify your preference when submitting the design.  

Does Paragon offer bioinformatics and data analysis support for ready-to-use panels and custom panels? 2018-09-13T17:34:04+00:00

We’re actively developing a cloud-based data analysis package and customer facing software using our analysis algorithms. In the near term, our bioinformatics team can provide plug-ins specific to Cleanplex libraries for common data analysis pipelines. The team can also offer support, assistance, and guidance to customers who may need additional help in data analysis for our ready-to-use panels and occasionally custom panels.  Please see the user guide for Data Analysis Recommendations for Illumina Sequenced Libraries.  For additional questions, please contact [email protected]

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