Library Preparation – The First Step in a NGS Setup

Welcome back! Last quarter we discussed why Next Generation, or Massively Parallel, Sequencing is the next big thing in the world of Molecular Diagnostics. The sensitivity, the depth of coverage and the ability to interrogate many different areas of the genome at the same time were just a few of the benefits of these types of assays. Next, I would like to describe a couple different methods of library preparation, which is the first step necessary to run an NGS assay.

First of all, let’s define “Library.” I find this is the most common question technologists new to this technology ask. Essentially, a library is a specimen’s collection of amplicons produced by the assay that have been barcoded, tagged with appropriate platform adapters and purified. These will serve as the input for the next part of the NGS workflow, clonal amplification (the topic of next quarter’s blog!).  How these libraries are prepared differ depending on platform (i.e, Ion Torrent vs. MiSeq), starting material (RNA vs. DNA), and type of assay (targeted amplicon vs. exome).

Before we begin the library prep discussion, a note about the input specimen. The DNA must be quantitated using a method that is more specific than spectrophotometry – it must be specific for double-stranded DNA. It will lead to an overestimation of the amount of DNA in the specimen, which will lead to over-dilution and consequently, lower quantity of final library. Real-time PCR and a double-stranded kit with fluorometry are two examples of assays that will give accurate concentrations of double-stranded DNA.

Our lab has begun using NGS for some of our oncology assays, so I will focus on the two types we perform currently, but keep in mind, there are many other types of assays and platforms.

Image 1: ion torrent amplicon library preparation. Source: Ion AmpliSeqTM Library Preparation User Guide – MAN0006735, Rev. 10 September 2012.

The assay we use for our Ion Torrent platform is a PCR amplicon based assay. The first step is to amplify up the 207 regions over 50 genes that contain hotspots areas for a number of different cancer types. This all occurs in one well for each specimen. Once those areas are amplified, the next step is to partially digest the primer sequences in order to prepare the ends of amplicons for the adapters necessary for the sequencing step. As shown in the figure above, two different combinations of adapters may be used. The top one, listed as the A adapter (red) and the P1 adapter (green), would be used if only one specimen was to be sequenced on the run. The A and P1 adapters provide universal priming sites so that every amplicon of every sample can be primed with the same primers, rather than having to use gene specific primers each time. The second possibility is listed below that, with the same P1 adapter (green) and a Barcode Adapter labeled X (red and blue) – it still contains the A adapter necessary for sequencing (red), but it also contains a short oligonucleotide sequence called a “barcode” (blue) that will be recognized during the analysis step based on the sequence. For example, Barcode 101’s sequence is CTAAGGTAAC – this will be assigned to specimen 1 in the run and all of the amplicons for that specimen will be tagged with this sequence. Specimen 2 will have the barcode 102 (TAAGGAGAAC) tag on all of its amplicons. During analysis, the barcodes will be identified and all of the reads with the 101 sequence will be binned together and all of the reads with the 102 sequence will be binned together. This allows many specimens to be run at the same time, thus increasing the efficiency of NGS even more. Lastly, the tagged amplicons are purified and normalized to the same concentration.

Image 2: MiSeq amplicon library preparation. Image source:

The assay we use for our MiSeq platform is a hybridization followed by PCR amplicon based assay. The first step is to hybridize probes to 568 regions over 54 genes that contain hotspots for a number of different cancer types. This occurs in one well for each specimen. Once the probes have hybridized, the unbound probes are washed away using a size selection filter plate. Next, the area between the probes is extended and ligated so that each of the 568 amplicons are created. These are then amplified in a PCR step using primers that are complimentary to a universal priming site on the probes, but also contain adapters plus the two indices required for paired end sequencing (the Ion Torrent platform utilizes single-end sequencing – this will be discussed in the sequencing portion in an upcoming blog!). As in the previous method, after PCR, these tagged amplicons are purified and normalized to the same concentration in preparation for the next step – clonal amplification.

Stay tuned for next quarter’s post – clonal amplification!



-Sharleen Rapp, BS, MB (ASCP)CM is a Molecular Diagnostics Coordinator in the Molecular Diagnostics Laboratory at Nebraska Medicine. 

Massively Parallel – the Next Generation of Sequencing

Sounds like a good title for a sci-fi novel, right?  What is the big deal about Next Generation Sequencing (NGS)?  Otherwise known as massively parallel sequencing or high throughput sequencing, NGS has become a technique used by many molecular labs to interrogate multiple areas of the genome in a short amount of time with high confidence in the results.  Throughout the next few blogs, we’ll discuss why NGS has become the next big thing in the world of molecular.  We’ll go through the steps of setting up the specimens to prepare them to be sequenced (library preparation), what types of platforms are available and what technologies they use to sequence.  Lastly, we’ll go through some of the challenges with this type of technology.

Let’s start with a review of dideoxy sequencing, otherwise known as Sanger sequencing, which has been the gold standard since its inception in 1977.  A typical setup in our lab for this assay begins with a standard PCR to amplify a region of the genome that we are interested in, say PIK3CA exon 21, specifically amino acid 1047, a histidine (CAT).  The setup would include primers complementary to an area around exon 21, a 10x buffer, MgCl2, a deoxynucleotide mix (dNTP’s), and Taq polymerase.  After amplification, the resulting products would be purified with exonuclease and shrimp alkaline phosphatase (SAP).  Next, another PCR would be set up using the purified products as the sample and using a similar mix as in the original amp, but with the addition of a low concentration of fluorescently labeled dideoxynucleotides.  These bases have no -OH group, so when they are incorporated into the product, amplification ceases on that strand.  Because they are present in a lower concentration, the incorporation of these is random and will occur at each base in the strand eventually.  The resulting products are then run and analyzed on a capillary electrophoresis instrument that will detect the fluorescent label on the dideoxynucleotides at the end of each fragment.  Shown below is an example of the output of the data:


The bases will be shown as peaks as they are read across the laser.  The base in question in the middle of the picture is, in a “normal” sequence, an adenine (A), as seen in green.  In this case, there is also a thymine (T) detected at that same location, as seen in red.  This indicates that some of the DNA in this tumor sample has mutated from an A to a T at this location.  This causes a change from a histidine amino acid to a leucine (p.His1047Leu) and is a common mutation in colorectal cancers.

So all of this looks great, right?  Why do we need to have another method since we have been using this one for so long and it works so well?  There are a few reasons:

  1. The sensitivity of dideoxy sequencing is only about 20%.  This means lower level mutations could be missed.  The sensitivity of NGS can get down to 5% or even lower in some instances.
  2. The above picture shows the sequencing in the forward direction as well as the reverse direction.  This area then has 2x coverage – we can see the mutation in both reads.  If we could get a higher coverage of this area and be able to sequence it multiple times and see that data, we could feel more confident that this mutation is real.  In our lab, we require each area has 500x coverage so that we feel sure that we have not missed anything.  The picture below displays the same sequenced area as in the dideoxy sequencing above.  This a typical readout from an NGS assay and, as you can see, this base has a total of 4192 reads, so it has been sequenced over four thousand times.  In 1195 of those reads, a T was detected, not an A.  We can feel very confident in these results due to how many times the area was covered.
  3. The steps above detailed only amplifying this one area, but with colorectal cancer specimens, we want to know the status of the KRAS, BRAF, NRAS, and HRAS genes as well as other exons in PIK3CA  Using the dideoxy sequencing method is a lot of time and effort.  NGS can cover these areas in these five genes as well as multiple other areas (our assay looks at 207 areas total) all in the same workflow


Join me for the next installment to discover the first steps in NGS workflow!



-Sharleen Rapp, BS, MB (ASCP)CM is a Molecular Diagnostics Coordinator in the Molecular Diagnostics Laboratory at Nebraska Medicine. 

Association for Molecular Pathology – A Bunch of Party Loving Pathologists…

I was privileged to attend this year’s Association for Molecular Pathology (AMP) meeting in Charlotte, North Carolina, in the beginning of November. I really enjoy this meeting – it is relevant to everything our lab does with sessions offered in topics of Hematopathology, Infectious Diseases, Solid Tumors, Inherited Diseases, and just recently added, Bioinformatics.

It is exciting to meet and discuss with others in this field, especially other laboratory technologists. AMP has done a wonderful job of including those of us who perform the bench work, offering discounted memberships, as well as learning opportunities on their website, and even an award especially for technologists’ exemplary posters/abstracts presented at the annual meeting.

This year’s meeting offered the previously mentioned topics, but an emerging trend was evident – testing cell-free DNA (cfDNA) obtained from sources other than tissue biopsies, such as plasma or urine. This quarter’s post will deal with the reason behind this and the technology for testing such specimens, specifically plasma.

Cell-free DNA has become an attractive source for tumor testing recently. This source can be tested when a tissue biopsy is just not possible, such as when a patient has progressed to the point that surgery is not recommended. Here is the biology behind why this can work as a source of tumor DNA:


Figure 1.

The sources of DNA in a sample of whole blood (as shown in Figure 1) are:

  • white blood cells
  • degraded white blood cells (cfDNA)
  • degraded tumor cells (cfDNA)
  • circulating tumor cells (CTCs).

Because of the biology of tumor cells, they have higher turnover than other cells in the body. Due to this, a larger fraction of the cfDNA in the plasma is from tumor cells. We can take advantage of this with a so called “liquid biopsy” – with 10 cc’s of whole blood, we can attempt to capture about 10ng of cfDNA and test this for possible resistance mutations to the therapies the patient may be on.

Many of the posters and several of the sessions at the AMP meeting dealt with cfDNA. Several pre-analytical steps were stressed in order to have success with this type of specimen.

  1. The whole blood needs to be collected, as any other blood specimen should, with care taken to not lyse any of the cells during collection.
  2. The collection tube type varies depending on how much time it will take to centrifuge the specimen to obtain the plasma. If it can be spun within two to four hours, a simple EDTA tube is sufficient. If it cannot be spun within a short time, then another tube with special preservatives is required. A Streck tube has been the tube of choice in these situations, but others are becoming available on the market as the demand increases. These specific tubes offer a greater amount of time to capture the cfDNA without white blood cell lysis becoming an issue. This is important, because as the white blood cells lyse, the plasma is flooded with the patient’s normal cfDNA that will dilute out the tumor cfDNA fraction, making it even more difficult to detect.
  3. Centrifugation procedures must be altered. The brake should not be applied when stopping the centrifuge because braking can cause the white blood cells to be sheared, which will, again, flood the plasma sample with normal cfDNA. An initial spin should be performed to obtain the plasma, then an additional spin should be performed before extraction of the DNA.

There are multiple kits available on the market for extraction of cfDNA. Once the DNA is extracted it is suggested to measure the DNA fraction with a method that will display the size of the fragments, such as with a Bioanalyzer. Cell-free DNA is about 160-170bp in size and, with the readout from an instrument such as the Bioanalyzer, one can see the size of the DNA, quantitate it, as well as observe any contamination from genomic DNA (shown by a peak >>170bp in size).

Many types of testing are being performed on this cfDNA fraction such as real time PCR, digital droplet PCR, and next generation sequencing. Whichever platform is used, a validation must be performed to ensure a fairly low level of detection (as low as 0.1% or 0.01%) because, many times, the positive tumor cfDNA allele fraction will be very low due to the normal cfDNA in the plasma.

This method of testing non-invasive specimens from patients is an amazing way to help save possibly very sick people from having to undergo a risky surgery. This is yet another use of a new technique in the ever changing world of Molecular Diagnostics!



-Sharleen Rapp, BS, MB (ASCP)CM is a Molecular Diagnostics Coordinator in the Molecular Diagnostics Laboratory at Nebraska Medicine. 

The Exciting World of Molecular Diagnostics

Hello everyone! I am Sharleen Rapp and I’m a Molecular Diagnostics Coordinator at Nebraska Medicine. I feel lucky to be able to discuss all about the exciting world of Molecular Diagnostics. For my first post, I’d like to give you a little background about myself and why I feel I am lucky to be in the career that I’m in.

Ever since I was little, science has intrigued me. Perhaps it was the experiments my Dad performed in our kitchen as practice for his labs for his high school chemistry classes (who doesn’t enjoy watching salt crystals “grow” on string in peanut butter jars?) or watching my brother set up his fruit fly experiment for his high school science class, but I’ve always enjoyed learning about how things work.

I went to a small parochial school in the middle of Nebraska, and unfortunately we didn’t have the funds for elaborate science class labs. Interestingly enough, the event that clinched science for me was a project that I did for my government class. We were responsible for writing, essentially, a textbook, complete with chapters, endnotes, quizzes and tests, on a topic of our choosing. I chose to write about the Human Genome Project. I wrote this in the year 2000, when the Project was in full swing. I had read about it in the previous years, and I was completely amazed by what it accomplished. In the middle of the school year, in fact, Time magazine came out with an issue titled “The Future of Medicine – How genetic engineering will change us in the next century.” It contained nineteen different articles, all focused on how the information from the Human Genome Project would impact the future – one of which discussed the way pharmaceutical companies were designing drugs to combat the mutations in different types of cancer. I knew then I would be a part of that future; I just didn’t know how. At this time, I had no idea how I could go about working in this field. I had never heard of the discipline “Molecular Diagnostics” or medical technology.

I went off to college and got a degree in Biological Sciences with the intent to go to graduate school and study in Genetics, but I still had no real idea about how to get into the field of study of DNA. Through some interesting twists and turns, including working in a fruit fly lab in college and an amazing internship at Washington University under Elaine Mardis, I ended up at a small private company where my job was to sequence mitochondrial DNA and mitochondrial-related genes, and in doing this, I knew I had found my career. I am a self-proclaimed science nerd and I love sequencing, the whole process from wet bench to analysis, more than anything that I have ever done. When I moved over to Nebraska Medicine and began working in the Molecular Diagnostics lab, I was amazed at the work that was being done there. I’ve had some amazing opportunities to work with all different types of sequencing – dideoxy sequencing, pyrosequencing, and now, massively parallel (aka, next generation) sequencing. I am so excited to be sharing some of my experiences and case studies from the work that we do in our lab in future posts.

Thanks for reading!!



-Sharleen Rapp, BS, MB (ASCP)CM is a Molecular Diagnostics Coordinator in the Molecular Diagnostics Laboratory at Nebraska Medicine. 

Blotting and Probing Techniques

“Blotting,” in relation to molecular diagnostics, is a term that refers to the process of detecting the presence and quantity of DNA, RNA, or protein in cells. There are three main types of blotting procedures that those in the field should be familiar with: Southern, Northern, and Western. Three additional blotting procedures are termed Southwestern, Eastern, and Far-Eastern. These are also summarized in the table below.

Southern Blot Steps

  1. DNA is isolated and cut with restriction enzymes.
  2. The DNA fragments are then analyzed by gel electrophoresis and separated by size (see previous blog post on Separation and Detection).
  3. Depurination – Gel is soaked in hydrogen chloride (HCl) to remove the purine bases from the sugar-phosphate backbone. This loosens up larger fragments before denaturation.
  4. Denaturation – The DNA is denatured by exposing the gel to sodium hydroxide (NaOH). Denaturation breaks the hydrogen bonds that hold the DNA strands together.
  5. Blotting – The denatured DNA is transferred to a solid substrate (nitrocellulose) that helps to facilitate probe binding and signal detection.
  6. Pre-hybridization – Prevents non-specific binding of the probe to other sites on the membrane surface.
  7. The membrane is exposed to the hybridization probe, usually a single DNA fragment with a specific sequence to the target DNA. The probe DNA is labelled either with radioactivity or fluorescent dyes.

Importance of the Membrane

Nitrocellulose and nylon membranes are best for smaller sized single stranded DNA fragments. It is compatible with many types of buffers and transfer systems. These membranes work well with protein and nucleic acids.


Capillary Transfer Ÿ Utilizes capillary movement of the buffer from a soaked paper to the dry paper

Ÿ Denatured DNA moves from the gel to the membrane

Electrophoretic Transfer Ÿ Electric current moves the DNA from the gel to the membrane
Vacuum Transfer Ÿ The force from suction moves the DNA from the gel to the membrane


Northern Blots

Northern blots are used in the laboratory to look at RNA structure and quantity. It’s a powerful method that can measure levels of gene expression, as well as structural abnormalities in RNA.

  • Needs to take place in an RNase-free environment.
  • The samples are applied directly to an agarose gel.
  • The sample is cut out from the gel, soaked in ammonium acetate to remove the denaturant (denaturant is inhibitory to the binding of RNA to nitrocellulose membranes), and stained with acridine orange or ethidium bromide.

Western Blots

Western blots detect proteins and separates them according to their molecular weight or charge

  • Run using a polyacrylamide gel with molecular weight standards / markers.
  • Utilizes capillary or electrophoretic transfer methods.
  • Membrane must be blocked with a solution to prevent binding of the primary antibody probe to the membrane.


DNA Probes Southern Blots Complementary to the target gene
RNA Probes Northern Blots Complementary to the target sequence
Protein Probes Western Blots Antibodies bind to the target protein


Method Target Probe Purpose
Southern Blot DNA Nucleic Acid ·      Gene structure
Northern Blot RNA Nucleic Acid ·      RNA transcript structure, processing, and gene expression
Western Blot Protein Protein ·      Protein processing and gene expression
Southwestern Blot Protein DNA ·      DNA binding proteins and gene regulation
Eastern Blot Protein Protein ·      Modification to western blot using enzymatic detection

·      Detection of specific agriculturally important proteins

Far-Eastern Blot Lipids None ·      Transfer of HPLC-separated lipids to PVDF membranes for analysis by mass spectrometry


L Noll Image_small

-LeAnne Noll, BS, MB(ASCP)CM is a molecular technologist in Wisconsin and was recognized as one of ASCP’s Top Five from the 40 Under Forty Program in 2015.




New Zika Test on the Horizon?

According to a recent press release, Rheonix is pursuing a rapid Zika Virus diagnostic test. Lablogatory recently discussed this test with the senior vice president for scientific and clinical affairs at Rheonix.

Lablogatory: I understand this test is a so-called “self-confirming” assay; it corroborates serological results with a molecular confirmation. Can you tell readers a bit about the methodology behind this?

Richard Montagna, PhD, FACB: It works much like the “dual assay” for HIV that we recently developed. Using microfluidics, the sample will be split between two sections of the same cartridge. One section will test for antibodies in a methodology similar to ELIZA. The other section will use LAMP technology to lyse, extract, purify, and amplify Zika-specific RNA sequences.

Lab: Sounds efficient! How long do you anticipate the assay will take to run?

RM: We expect it would take less than an hour to perform. Using our existing equipment base, we anticipate the capacity to perform 24 tests in an hour.

Lab: Given the timeline of the impending outbreak, will you seek Emergency Use Authorization (EUA) from the FDA?

RM: Once development is complete, we’ll discuss EUA with the FDA to determine if that approach is feasible.

Molecular Diagnostics Survey

If you’re involved with molecular diagnostics, then L. J. Lee’s team at the Ohio State University would like your input. They’ve developed a new technology using molecular beacons and would love for lab directors, managers, and bench technologists to answer this short survey.