See how this groundbreaking technology works. http://www.lifetechnologies.com/iontorrent Leveraging consumer technology for scientific breakthroughs Ion Torrent™ technology takes an entirely new approach to sequencing, making it faster, simpler, and more affordable than ever before. Unlike other sequencing technologies, Ion Torrent™ systems sequence DNA using a semiconductor chip, similar to the chip found in your digital camera. While the chip in your camera has a sensing layer covered with millions of pixels that translate light into digital information, an Ion chip has millions of wells covering those pixels. Whenever a nucleotide is incorporated into a single strand of DNA, a hydrogen ion is released. This is how the Ion Torrent system sequences DNA, by reading this chemical change directly in the well on the chip. In essence, each well works as the world's smallest pH meter.
Submit your real-time PCR questions at http://www.lifetechnologies.com/asktaqman Just how does TaqMan work? Sr. Field Applications Specialist Doug Rains explores the specific mechanism by which TaqMan® achieves its unparalleled specificity and sensitivity. Around the world, researchers rely on TaqMan® for gene expression, SNP gentoyping , protein expression, pathogen detection and quantification, and more. For many years, TaqMan has been the gold-standard chemistry for real-time PCR. It's famed for its unparalleled specificity, sensitivity, and ease of use. So it's not surprising that users want to know what Shrikant at ICL College in India asked recently: namely, "How does TaqMan work?" I'm glad you asked. Just like any PCR, TaqMan-based reactions require a double-stranded template, as well as two fairly standard, target-specific primers. But unlike those used in regular PCR, TaqMan Assays require a third, sequence-specific oligo called a probe. TaqMan probes are quite different from the primers in two ways. First, they can't be extended by our friendly enzyme, Taq Polymerase, since they lack a free hydroxyl group. What's more, TaqMan probes are covalently joined to two other molecules. On the 5'-end, there's a fluorescent molecule known as the reporter -- called that because it reports signal to us as we generate more and more product. On the 3'-end is a molecule known as the quencher, which quenches the fluorescent signal from the reporter under certain circumstances. Let's see what those circumstances are. Here we're looking at an intact probe, with the reporter in green, the quencher in red. Normally, when we zap the probe with light, we expect the reporter to get excited and fluoresce. But because the quencher is in close proximity to the reporter, instead what happens is, the energy gets transferred from reporter to quencher. The transfer of energy is known as FRET, or Fluorescent Resonance Energy Transfer. The important thing to note here is that, as long as the probe remains intact, there is no permanent increase in fluorescent signal from the reporter. However, if the reporter and quencher are permanently separated during the reaction, and then light strikes the reaction, the Reporter does in fact fluoresce, producing signal the instrument can detect. The basic idea, then, is this: each time we create a new PCR amplicon, we want to permanently split the reporter and quencher. By doing so, florescence will always increase proportionally with product, allowing us to effectively monitor what's happening to our reactions throughout the run. Here it is in action. We begin our reactions (CLICK) by denaturing our template at a high temperature. As we lower the temperature, our probe and primers bind. Taq now comes in, finds the primers, and begins the extension phase of PCR by creating new complementary strands of DNA. But wait a second: there's a probe sitting in the way. It's a showdown in the making! What will the polymerase do? Stop in its tracks? Turn back in fear? Nay, friends, not Taq Polymerase. You see, our enzyme has what's referred to as "exonuclease activity." Meaning? It pretty much eats DNA for lunch. So when Taq reaches the probe, it simply chews it to bits on its way to creating new amplicon. As a result, the reporter and quencher are physically separated, creating a permanent increase in fluorescence that, not coincidentally, perfectly accords with our doubling of product. And, of course, that our real-time instrument can monitor and record this increase in fluorescence after each cycle, generating an amplification plot that's more than a little useful for interpreting our data.
Learn more at http://www.lifetechnologies.com/transfection Optimized protocol for Lipofectamine LTX & Plus reagent: http://tools.invitrogen.com/content/sfs/manuals/LipofectamineLTX_PLUS_Reag_protocol.pdf For technical questions, please reach out to Technical Support at: '[email protected]' or check http://www.invitrogen.com/site/us/en/home/support/Contact-Us.html for contact details. ---------- Audio transcript: How to perform Plasmid DNA transfection with Lipofectamine® LTX and Plus™ Reagent protocol. Superior plasmid delivery and protein expression. In this video, we will perform a plasmid DNA transfection experiment using Lipofectamine® LTX & Plus™ reagent. As always, use good cell culture practices and wear your personal protective equipment. Be sure to clean your cell culture hood and work surface by spraying and wiping them down with 70% ethanol. The day prior to your transfection, seed your cells so that they will be 70% to 90% confluent at the time of your experiment. For this transfection experiment you will need: - Lipofectamine® LTX and Plus™ Reagent - Opti-MEM® Reduced-Serum Medium - Plasmid DNA at 1 microgram per microliter. We will be using a Green Fluorescent Protein plasmid to serve as a positive control for transfection. - Five, 1.5 mL microcentrifuge tubes in a rack - A P200 and P10 pipette and appropriate tips - A marker and a timer - And a 24-well plate with 70% to 90% confluent cells. We will be following the 24-well plate format of the Lipofectamine® LTX & Plus™ Reagent protocol. Prepare 4 tubes each with 50 microliter of Opti-MEM® Medium, and label them 1 to 4. Add 2 microliters of Lipofectamine® LTX Reagent to tube 1, 3 microliters to tube 2, 4 microliters to tube 3 and 5 microliters to tube 4. Mix each tube well by vortexing or flicking the tube. Prepare a tube with 250 microliters of Opti-MEM® medium and add 5 micrograms of plasmid DNA. Since, our DNA concentration is at 1 microgram per mircoliter we are adding 5 microliters. Next add 5 microliters of Plus™ Reagent and mix well. Add 50 microliters of the diluted DNA to each of the Lipofectamine® LTX dilutions in tubes 1 to 4. Incubate the complex for 5 minutes at room temperature. After the 5-minute incubation, remove your 24-well plate containing your cells from the incubator and bring it to the workspace in the hood. Add 50 microliters of the DNA-reagent complex from tubes 1 to 4 to wells 1 to 4 of the 24-well plate, respectively. You should have enough volume to run duplicates on the same plate if desired. Place your 24-well plate back into the incubator and grow cells for 1 to 3 days at 37 Celsius. After incubating your cells, assess the transfection efficiency in each well by viewing GFP fluorescence. Examine each well using a FLoid cell imaging station or microscope to determine which concentration of reagent provided the highest transfection efficiency. In this experiment dilution 3 provided the highest transfection efficiency. For transfection protocols, FAQ's, troubleshooting and tips & tricks visit http://www.lifetechnologies.com/transfection
Download the free Sanger sequencing handbook at http://www.thermofisher.com/sangerhandbook If you have more questions on sequencing, submit your question at https://www.thermofisher.com/ask Let’s go back to the basics and explore the technology platform that has been regarded as the gold standard for many years. Yea, you guessed it! I am talking about Sanger Sequencing by capillary electrophoresis. Many might ask, “why is it called Sanger Sequencing?” Sanger Sequencing is named after the inventor of this ground breaking technology, Dr. Frederick Sanger, who developed this method over 40 years ago in the mid-70s. So, what are the basics of Sanger Sequencing? It all starts by having a short primer binding next to the region of interest. In the presence of the 4 nucleotides, the polymerase will extend the primer by adding on the complementary nucleotide from the template DNA strand. To find the exact composition of the DNA sequence, we need to bring this reaction to a defined stop that allows us to identify the base of the very end of this particular DNA fragment. Sanger did this by removing an oxygen atom from the ribonucleotide. Such a nucleotide is called a dideoxynucleotide. This is analogous to throwing a wrench into a gear. The polymerase enzyme can no longer add normal nucleotides onto this DNA chain. The extension has stopped and we now need to identify what it is. We identify the chain terminating nucleotide by a specific fluorescent dye, 4 specific colors to be exact. Sanger sequencing results in the formation of extension products of various lengths terminated with dideoxynucleotides at the 3′ end. The extension products are then separated by Capillary Electrophoresis or CE. The molecules are injected by an electrical current into a long glass capillary filled with a gel polymer. During CE, an electrical field is applied so that the negatively charged DNA fragments move toward the positive electrode. The speed at which a DNA fragment migrates through the medium is inversely proportional to its molecular weight. This process can separate the extension products by size at a resolution of one base. A laser excites the dye labeled DNA fragments as they pass through a tiny window at the end of the capillary. The excited dye emits a light at a characteristic wavelength that is detected by a light sensor. Software can then interpret the detected signal and translate it into a base call. When the sequencing reaction is performed in the presence of all four terminated nucleotides, you eventually get a pool of DNA fragments that are measured and separated base by base. What you will get in the end is a data file showing the sequence of the DNA in a colorful electropherogram and a text file which you can use to answer the questions you may be asking. And that in a nutshell is Sanger Sequencing.
https://www.thermofisher.com/global/en/home/references/gibco-cell-culture-basics.html?cid=BID_R01_PJT3313_BID88888_VI_YUT_OD_KT_365 The handbook and videos provide an introduction to cell culture, with a focus on maintaining cell health throughout the processes of culturing, freezing, thawing and passaging cells. In this video, we focus on how to passage cells.
Short educational video explaining the principle of Ion Torrent Next Generation Sequencing (NGS) Technology – how it works, and its applications use in science and medicine such as targeted sequencing in oncology.
See all Ph.Diddy videos here: - "Ph.Diva and the Mystery Band" - http://youtu.be/ryhSGUzzz8U - "Ph.Diddy at the Conference" - http://youtu.be/ajdbuj8P784 - "Behind the scened at Halloween with Ph.Diddy": http://youtu.be/r_NdyX70MF8 PCR Arnold Young (aka the Ph.Diddy) is a biotech Ph.D candidate. Arnold was an A+ student, and is now ready to take over the world of science! He soon discovers how life in the lab is filled with drive and devotion, frustrations and fulfillment, hard work, late hours, repeated experiments, peer review and a strive for respect and recognition. How will the ups and downs of life in the lab shape our Ph.Diddy on his journey to have his first scientific paper published? Acknowledgements: This video is inspired by all the dedicated everyday-heroes-of-science. Invitrogen recognizes your passion and admires your perseverance! BY POPULAR DEMAND - the song is now available on iTunes! http://itunes.apple.com/us/album/the-ph.diddy-is-on-the-scene/id478987896?i=478987900&ign-mpt=uo%3D4 For more information on PCR, please visit www.lifetechnologies.com/elevatepcr -------------------- Lyrics: I was the black belt of my class, marked out as an achiever. First day on the job, enter the gene-weaver. My future colleagues look up from the PCR machine, checking me out man. The Ph.Diddy's on the scene. So I claim my pipette and pimp it up to halt the theivin'. Get the thermocycler going, my samples cool chilling. Yeah, I shoot tips like I've been here all my life. Waiting for instrument time with my free pizza slice. I run my first Western blot. Man, this lab scene is easy. But wait up, hold a minute. I'm feeling kinda queasy. Is that Trizol I can smell? Oh, my head's in a spin! 've gone and dropped my freakin' sample in the biohaz bin. He's a super fly grad. Life plans to shape the future, that's right! Sees his name in lights and a centre spread in Nature. All right! I'm always splitting cells, always working late. I've cleared the biohazard bin so much I herniate. Got samples botched, samples lost, samples dropped on the floor. Messy benches, empty buffers, lab politics galore. My PI's buggin'. Man he sounds like my father. Western blot once again, but development shows nada. I'm doin' my time in the cell culture hood - at the weekend bro. It's the only time I could! Then suddenly my data is starting to make sense. Even Western blot bands start to make an appearance. So I repeat once again all the steps I need to follow. But every culture well... turned yellow. He's a super fly grad. Life plans to shape the future, that's right! Sees his name in lights and a centre spread in Nature. All right! I'm talkin' 'bout the ooh, ooh... Serum and media get swiped with no apology. And all my cells are showing abnormal morphology. So now my PI's freakin' "Find something you can prove!" But contamination aggravation knocks me off my groove. So I stay late once again as the lab detainee. Yeah, its 9 pm but the thermocycler's free I don't understand! What's happened now? That's it. Yeah, time to give up. I'm throwing in the towel. Ooh, ooh, supergrad. I'm talkin' 'bout the ooh, ooh... Super bad lab grad, yeah! Ooh, ooh, supergrad. I'm talkin' 'bout the ooh, ooh... Super bad lab grad, yeah! My transfection worked - confirmed with Pol II chIP - mutagenesis determined the TF targeted. I'm not going down! I can still do BioChem! My genes will express, I'm gonna beat it out of them. And yeah, I must have results that no one can undress. And quickly move my paper from in prep to in press. I'm finally finished writing the discussion, whilst my PI once again, "tweaks" my introduction. So, here we go, yo. Here's the scenario: Peer review's due to hit. When exactly? I don't know. My paper will be published. My research is legit. OK it's not quite Nature but, I think I nailed it! He's a super fly grad. Life plans to shape the future, that's right! Sees his name in lights and a centre spread in Nature. All right! He's a super fly grad. Life plans to shape the future, that's right! Sees his name in lights and a centre spread in Nature. All right! Ooh, ooh, supergrad. I'm talkin' 'bout the ooh, ooh... Super bad lab grad, yeah!
Learn more at http://www.lifetechnologies.com/transfection How to perform siRNA transfection with Lipofectamine® RNAiMAX protocol. Superior siRNA/miRNA delivery and gene knockdown. In this video, we will perform an siRNA transfection experiment using Lipofectamine® RNAiMAX reagent. As always, use good cell culture practices and wear your personal protective equipment. Clean the cell culture hood and work surface by spraying and wiping them down with 70% ethanol. The day prior to your transfection, seed your cells so that they will be 60% to 80% confluent at the time of your experiment. The day prior to your transfection, seed your cells so that they will be 60% to 80% confluent at the time of your experiment. For this transfection experiment you will need: - Lipofectamine® RNAiMAX reagent - Opti-MEM® Reduced-serum Medium - siRNAs diluted to a working concentration of 10 micromolar. We will be using two Ambion® Silencer® Select siRNAs, a BLOCK-iT™ Alexa Fluor® red fluorescent siRNA, and a negative control siRNA - Five, 1.5 milliliter microcentrifuge tubes in a rack - A P200 and P10 pipette and appropriate tips - A marker and a timer - And a 24-well plate with 60% to 80% confluent cells We will be following the 24-well plate format of the Lipofectamine® RNAiMAX protocol. Because we have 4 siRNAs, we will prepare a master mix of RNAiMAX. Add 200 microliters of Opti-MEM® medium and 12 micoliters of RNAiMAX in a tube labelled "Master Mix" Mix well by vortexing or flicking the tube. Add 50 microliters of Opti-MEM® Medium into each of 4 tubes and label them 1, 2, Positive, and Negative. Add 3 microliters of each 10 micromolar siRNA stock to its corresponding tube. Mix well. Add 50 microliters of the RNAiMAX master mix to each of the siRNA dilutions in tubes 1, 2, Positive, and Negative. Incubate the complexes for 5 minutes at room temperature. After the 5-minute incubation, remove your 24-well plate containing your cells from the incubator and bring it to the workspace in the hood. Add 50 microliters of the siRNA-reagent complex from tubes 1, 2, Positive, and Negative to wells 1 to 4 of the 24-well plate, respectively. You should have enough volume to run duplicates if desired. Place your 24-well plate back into the incubator and grow cells for 1 to 3 days at 37 Celsius. After incubating the cells 24 hours at 37 degrees Celsius, assess the transfection efficiency of the BLOCK-iT™ Alexa Fluor® red fluorescent siRNA using the FLoid cell imaging station or microscope. To assess gene knockdown use a quantitative method such as Ambion® Cells-to-Ct™ Kit and Real-Time PCR. For transfection protocols, FAQs, troubleshooting, and tips & tricks visit http://www.lifetechnologies.com/transfection
Episode 1 in a 6-part mini documentary series, see the full series here: http://www.lifetechnologies.com/exosomesdocumentary We asked 10 prominent scientists to share their thoughts on science and in particular the field of exosomes research. The video-series tell the story and history of this exciting new area of research, and its impact on other research fields such as cancer and immunology. The video-series also discusses the potential future therapeutic and diagnostic applications that may come from exosome research. Contributors: - Xandra Breakefield, Ph.D........... Professor, Massachusetts General Hospital - Jan Lötvall, MD., Ph.D .............. Professor, University of Gothenburg, President of ISEV - Suresh Mohla, Ph.D.................. Chief (TBMB) and Division Associate Director, NIH - Esther Nolte-'t Hoen, Ph.D......... Senior Scientist, Utrecht University - Michiel Pegtel, Ph.D................. Assistant Professor, VUmc, Amsterdam - Graça Raposo-Benedetti, Ph.D.. Director of Research, CNRS, Institut Curie - Phillip A. Sharp, Ph.D............... Nobel Laureate, Professor, MIT - Johan Skog, Ph.D.................... CSO, Exosomes Diagnostics - Dima Ter-Ovanesyan.................Ph.D Student, Harvard University - Clotilde Thery, Ph.D................. Director of Research, INSERM, Institut Curie, Secretary General of ISEV Acknowledgements: Members of the research groups of Michiel Pegtel (Cancer Center Amsterdam, VUmc), Graça Raposo-Benedetti (Structure and Membrane Compartments, CNRS, Institut Curie) and Clotilde Théry (Immunity and Cancer, INSERM, Institut Curie) Special thanks to the ISEV society for their strong contribution in leading the field of exosome research forward, and to the participants at the ISEV RNA Workshop in New York City October 1st-2nd 2012 Kudos to all scientists that have the courage to follow their curiosity! If you want to learn about the exosomes-specific products available from Life Technologies: http://www.lifetechnologies.com/us/en/home/life-science/cell-analysis/exosomes.html?cid=exoep1
Submit your questions at http://www.thermofisher.com/forensicfocus Everyone wants to know how much DNA is in their extract, but then they ask: how can I tell if my estimate is accurate? The standard curve holds the answers. A standard curve is a tool that allows us to estimate the DNA concentration of unknown samples by comparing them to standards with known DNA concentrations. In this example, the standards consist of a 10-fold dilution series ranging from 50 ng/ul down to 5 pg/ul. During each PCR cycle, the amount of fluorescent signal for each standard in the dilution seies is measured. When the fluorescent signal crosses the detection threshold the cycle number is recorded as a Ct value, or threshold cycle value. The Ct value is what ultimately is used to create the standard curve. The Ct values are inversely proportional to the concentration of DNA in the standards. The high-concentration, 50 ng/ul standard will cross the detection threshold first, generating a “low” Ct. The low-concentration, 5 pg/ul standard will take many more cycles to cross the same threshold - and therefore the Ct will be higher. The Ct values for each dilution of the standard curve are plotted on a graph, and the software generates a regression line that fits the data. Because the standards are 10-fold dilutions, we expect the change in Ct from one standard to the next to be uniform. An uneven distribution of Ct values might indicate that the dilution series was not accurately pipetted. Let’s take a look at the standard curve for a specific DNA target, the small autosomal target. The X axis is the log of the known standard concentrations. The Y axis is the Ct value of each standard. Now, Do you see the quality metrics at the bottom of the screen? Let’s review Slope, Y intercept, and R2.? The slope measures the efficiency of the PCR reaction. In a perfect world, a slope of -3.3 indicates that the PCR reaction is 100% efficient; the target DNA is doubled each cycle. Two copies become four; four become eight; and so on. The Y intercept is the expected Ct value for a 1ng/ul sample. The R2 value measures how well the regression line fits the data points. A line that fits the data points perfectly has an R2 of 1. If your data points are scattered, the R2 value for the line will be lower. The Ct values of your standards affect the slope, the Y intercept, and the R2 value. It is very important to prepare the standard dilution series carefully to ensure consistent and accurate results! Running the standards in duplicate can help ensure you have a high quality standard curve. Once your standard curve passes the metrics test, it can be used to evaluate an unknown sample! The Ct value of the unknown sample is measured, and compared to the standard curve to estimate the DNA concentration of the unknown sample. Couldn’t be simpler! That’s it for today. If you have other questions, just click on the link below. And don’t forget- when in doubt, refer Back to the Bases!
Submit your Real-Time PCR questions and watch the rest of our videos at http://ow.ly/bQh0l. Life Technologies Sr. Field Application Specialist Doug Rains helps with the understanding of baselines in Real-Time PCR. We're looking at a fairly standard real-time amplification plot. We have some nice curves, each of which has the familiar geometric phase, linear phase, and plateau phase. So far, so good. But what's all this . . . junk in the early cycles? Well, friends, if you said "junk," you were right. That's right, I said it -- junk, trash, waste, detritus, garbage, otherwise known as noise. It's the stuff we see before our actual signal from amplification gets high enough to overcome that noise. And, as the rather impolite adjectives I used a second ago would suggest, it's completely useless to us. This noise does have an effect on our curves. Our job is to minimize that effect by effectively subtracting out the noise. We do that by establishing what's known as a baseline -- a cycle-to-cycle range over which only noise can be seen, prior to the appearance of curves. Once established, the software will effectively subtract out the noise on a well-by-well basis, greatly improving the quality of our data. Let's switch the Y-axis to linear scale for a moment to illustrate the effect of baseline subtraction. Here's our data prior to baselining. Note how every sample begins from a slightly different spot on the Y-axis, causing our geometric phase data— this curvy part over here when we're in a linear scale— to look horrible. But once we subtract noise, every sample begins from the same point 0. And as a result, the data clean up nicely. The value we get after normalizing for background noise is something called delta-Rn. If you ever look closely at a log-scale amplification curve— the one we're used to seeing— you'll notice that delta-Rn is what's graphed on the Y-axis. But before you go, just note that there are two ways to set baselines in Applied Biosystems® real-time PCR software: manually, and automatically. If you do it the manual way, you set the baseline range under Analysis Settings. You either set it for a single assay, in which case all wells for that assay get the same subtraction . . . or you can go under Advanced Settings and set wells individually. Better yet, just use the default setting of Auto Baselining. With this selected, the software figures out how much noise needs to be subtracted from each well individually, and, as such, generally produces the best results. So why have a manual feature? Well, Auto does fail on occasion, especially with some SYBR® Assays and non-standard chemistries. You'll know auto has malfunctioned by the shapes of your curves. If they look more S-shaped than they should, it could be that auto has misapplied the baseline and set the End cycle too low. As a result, not enough noise is being subtracted, and the curves take on a strange shape. To fix the problem, switch over to manual mode for that assay and raise the End cycle until the curves take on a regular shape.
https://www.thermofisher.com/global/en/home/references/gibco-cell-culture-basics.html?cid=BID_R01_PJT3313_BID88888_VI_YUT_OD_KT_366 The handbook and videos provide an introduction to cell culture, with a focus on maintaining cell health throughout the processes of culturing, freezing, thawing and passaging cells. In this video, we focus on how to thaw cells.
Submit your Real-Time PCR questions and watch the rest of our videos at http://ow.ly/bQh0l. Life Technologies Sr. Field Application Specialist Doug Rains helps with the understanding of thresholds in Real-Time PCR. The threshold is a horizontal line in our amplification plot that can be moved up or down on the Y-axis. Its purpose? As we'll see in a minute, it tells the software where to take data. Of course, not all places on the Y-axis are equal. Some places we want to avoid. Specifically, we don't want to be too low, otherwise we get down into the noise. Conversely, if we go too high, we're in the linear or plateau phase of amplification, where data are less predictable. A happy spot? Some place where all of our curves are straight and parallel to one another. What we really want is to put the threshold wherever the precision of our replicates is highest. That's generally somewhere toward the middle of the geometric phase, or maybe slightly higher. In any case, with a really robust assay, hitting a bad spot is quite difficult. The default on all Applied Biosystems® real-time PCR software is Auto Threshold, meaning, the software sets thresholds for us the second we click Analyze. Notice that it sets a different threshold for each assay separately, which is good since not all assays have the same sweet spot. I could go switch any one or all of my thresholds to Manual mode, then move the line up or down with my mouse. Once the threshold is set and we click analyze, all the samples get their respective Ct values. Now, the attentive viewer might be tempted to ask: if the threshold can be moved up or down, doesn't that change the Cts? The answer is, "Yes." But here's the thing: as long as we keep the threshold firmly within the geometric phase, the relative, or delta Ct between any two samples stays constant. This fact allows us to do things like calculate fold changes in expression from sample to sample, and to get quantity information from a standard curve.
Submit your question at http://www.thermofisher.com/ask What do you call a collection of millions of DNA fragments sharing the same short sequences on the 5’ and 3’ ends? The answer is a next generation sequencing, or NGS, library. Today, we are going to focus on the four basic steps of NGS library preparation that can be broadly applied across different preparation methods. A key step in the NGS workflow is preparing the input for sequencing, known as creating a library. An NGS library is a collection of similarly sized DNA fragments with known adapter sequences added to the 5’ and 3’ ends. A library corresponds to a single sample and multiple libraries, each with their own unique adapter sequences, can be pooled and sequenced in the same sequencing run. NGS library preparation has four general steps: 1. DNA Fragmentation or Target Selection, 2. Addition of adapter sequences, 3. Size selection, and 4. Final library quantification and QC. Let’s take a look at our lab book The first step, DNA fragmentation or Target Selection. In order to get the starting DNA into smaller pieces, isolated DNA may be fragmented using physical or enzymatic methods. These libraries are referred to as fragment libraries. Alternatively, if the sequence of specific DNA targets is known, PCR amplification of those targets may be used to produce DNA amplicons within the desired size range. These libraries are referred to as amplicon libraries. Next, specific DNA adapter sequences are annealed to the 5’ and 3’ ends of the fragmented or amplicon DNA. The double-stranded DNA adapters are approximately 20 to 40bp fragments that contain known sequences. Generally, there are two different adapter sequences that can anneal to the DNA fragments in either the 5’ or 3’ orientation. One adapter sequence contains the primer annealing site for the sequencing primer, while the second adapter sequence is generally used to anchor the DNA fragment to a surface for sequencing; for example beads or a solid surface containing a complimentary DNA sequence. Now we have our DNA fragments with known adapter sequences on either end. The next step is to select the library fragment sizes we need for our sequencing run. There are two common size selection methods, the first is a gel electrophoresis based-method, while the second is bead-based size selection method. For the gel-based method, the adapted library fragments are run on a gel to separate the fragments by size and the band corresponding to the size of interest is collected. Using the bead-based method, magnetic beads are used with varying concentrations of buffers to isolate the DNA fragment sizes of interest. Final library fragment size is important for efficient, high quality DNA sequencing downstream. Bonus, when preparing amplicon libraries, size selection is usually not necessary, as long as the PCR products were already designed to be within the desired size range. We are almost done! The last and very important step is library quantification and QC. Accurate library quantification is important for successful template preparation and sequencing. There are a few library quantification methods commonly used. The first is analysis by the BioanalyzerTM system. This method gives you both library concentration and fragment size information. The second is qPCR. This method provides the most accurate library quantification information, as it only measures amplifiable library fragments, but lacks library size information. Which method you prefer is entirely up to you (and probably what’s available in your lab). And that’s NGS library preparation in a nutshell. Of course there are variations on this theme depending on your application, for example, gene expression or DNA methylation analysis, but the fundamentals stay the same. Since your library quality dictates the success of all downstream processes from template preparation to sequencing, understanding the library preparation is important to help ensure you get the highest quality sequencing data. I hope this video was helpful on NGS library, and I am sure you’ll have more questions. Submit your question at thermofisher.com/ask and subscribe to our channel to see more videos like this. And remember, when in doubt, just Seq It Out
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Sharing Eachclip Videos
If you find a video you particularly enjoy and want to share with others, you have multiple options for sharing it. Email sharing, along with sharing options for every major social networking service.
Simply click the Share button for a video and you're presented with several options for sharing it with friends and family.
If you want to simply copy and paste the video page link to share it somewhere, you can do this using the shortened link provided beneath the social share buttons after clicking Share.
Download Videos You Want to Watch Later
Since there's such a wealth of content on Eachclip, the platform makes it easy to save videos you want to watch another time to your Watch Later list or a playlist you created. To download a video to your Watch Later list, just click the Download button and then choose the format you want to add the video.
Is Eachclip Safe?
Eachclip is rated as 93% safe with 80% child safety parameter. Still, to improve child safety, one needs to use additional filters over the browser. As this site usually follow a different kind of ads that are not rated as good for kids so to ensure 100% safety it is better to add other filters on the channel. The other idea to improve ad safety is just to add some ad blockers. If we talk about viruses and malware, then Eachclip is found to be 100% safe for your system.
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