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How do you dilute forward and reverse primers for a master mix?

How do you dilute forward and reverse primers for a master mix?


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I apologize for the very naive question, but I'm just starting out in a high school biology lab and I am very confused. If I have stocks of 100μM for forward and reverse primers separately, I can dilute them to 10μM very easily. But what if I want to make a master mix? I have 1000μL of 10μM for each primer, forward and reverse (not mixed). Would I just add 500μL of my forward primer and 500μL of my reverse primer for a mix of 1000μL in which both concentrations are still 10μM?


Would I just add 500μL of my forward primer and 500μL of my reverse primer for a mix of 1000μL in which both concentrations are still 10μM?

No. Diluting your 10μM solution in half will half the concentration. Mixing equal parts of 10μM primer will make a master mix where each primer is 5μM

But in general, primers are added to these reactions in great excess, so 5μM of primer might be okay.


Preparing a PCR Mastermix

Today, I had to figure out the right concentrations for a PCR master mix.

We will be using two primer sets, thus making 2 master mixes, which will go into 10 tubes.

We have three different DNA samples: BW (control), 9916Bxx, and 9916Bx. On the third one, we will perform three tests of 1x DNA, 1/10x DNA, and 1/100x. For this, the DNA must be diluted.

The first step in making a master mix is diluting the primers. They come in 100μM stock concentrate, and we must dilute it to 10μM, so to do that you put 10μL of the forward and 10μL of the reverse into 80μL of H2O. Then, you take 1μL of the newly diluted primer and put it into your master mix. You will have 1μL / 50μL, so it will be 0.2μM of primer in the end. You also must put 25μL of the Taq mix, 1μL of the template DNA, and 23μL of H2O, to make a 50μL total master mix. We will do this for the other primer too. HOWEVER, when you make the master mix, you cannot put the DNA in yet.

We will make the two master mixes up until they are 49μL in volume and have no template DNA. Then, we will put 49μL into each tube, 10 tubes total. In the first 3 of each set of 5 tubes, the DNA will be added at 1x. This means you simply add 1μL of the DNA. For 1/10x, you take 10μL of the 1x DNA, add it to 90μL of water to make 1/10x. You take 10μL of that and put it in 90μL of water for 1/100x.

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Our Top 10 Tips for Consistent qPCR

qPCR requires a certain amount of technical finesse to ensure consistent data across experiments. The main challenges encountered when starting out with this technique are contamination issues or inconsistency between replicates. The cost of running qPCR is much higher than end-point PCR, so getting every experiment right may also be critical for your lab budget.

Below are our top 10 tips to help you to get consistent qPCR data every time!

1. Always Mix the Reagents Well Before Use

qPCR reagents include dyes, nucleotides and enzymes that may settle while sitting in the freezer or refrigerator. Make sure to mix your individual reagents thoroughly before preparing your master mix. Similarly, pipette your master mix thoroughly before aliquoting into your plate or tubes to avoid uneven distribution of reagents between reactions.

2. Store Primers in a Buffer to Protect Their Stability

When your primers arrive, avoid resuspending the master stock in water. The pH of water can be low (especially if it is DEPC-treated), leading to primer degradation over time. Instead, use a buffered solution at neutral pH to protect your primers from acid hydrolysis. Tris-EDTA (TE) is a common choice. The EDTA (1 mM) will inhibit potential DNAse activity, and when you dilute the primers for working stocks, the EDTA should be sufficiently diluted so as not to interfere with Taq polymerase activity.

3. Aliquot the Primers

Once you have a master stock (usually 100-200 mM), you should make working stocks to avoid multiple freeze/thaw cycles of your original primer solution. Prepare these working stocks (10-20 mM) in TE buffer in volumes to suit your needs. Limit yourself to three freeze/thaw cycles of these working stocks. Repeated freezing/thawing can lead to primer degradation, which may negatively impact your qPCR results e.g. reduced qPCR efficiency and reduced sensitivity.

Preparing aliquots will also help you out in the event of primer contamination. If you accidentally contaminate one vial of primer, you can throw it away and take a fresh one without worrying about contaminating the master vial.

4. Use Pipettes Calibrated for Low Volumes

If you require absolute accuracy in quantification, use pipettes calibrated for low volume pipetting (such P2 or P10) to prepare your standard curves and qPCR reactions.

Using the right pipettes ensures reproducibility between replicates. This is important when you are measuring qPCR efficiency based on standard curves, as you need to be sure that you are truly measuring qPCR efficiency and not your pipetting skills. Also, make sure that your pipettes are accurate before you start out!

5. Perform a Standard Curve for Every New Primer Pair

Don’t assume that every set of primers you order is going to work as well as the last. qPCR efficiency can be influenced by a number of factors. The best practice is to run a 5-point standard curve with 10-fold dilutions for every new primer pair and make sure you can get at least 90% qPCR efficiency with control DNA.

6. Follow the Three Room Rule

One of the biggest sources of contamination is using the same pipettes for all parts of the qPCR workflow i.e. DNA extraction, PCR and PCR product handling post-run. This is not advisable even if you use aerosol resistant tips at all times. For qPCR, always use a set of pipettes that are solely dedicated to qPCR reaction set up.

In addition to using these dedicated pipettes, you should keep them away from the room used for DNA/RNA extraction. The ideal set up is to have three rooms one for nucleic acid extraction, one for reaction set up (with a hood containing a UV lamp to pre-treat pipettes and plastics between users), and one for the qPCR cycler.

This is the safest way to minimize the risk of contamination in your negative controls.

7. Double Check the Cycling Conditions

This is important if you are using a shared instrument. Even if you have your own template file set up, double check your run cycle before hitting start. Someone may have made small changes to your cycling template (e.g. annealing temperature, hot start activation time) without your knowledge.

8. Dilute the Template (Less May Be More)

Depending on the gene(s) of interest, you might actually be starting with too much template. qPCR is so sensitive that less template often gives a more accurate measurement.

Ideally, you want your samples to cross the cycle threshold between cycles 20-30. Samples that cross the threshold before cycle 15 will fall into the default baseline setting on most instruments, and this will lead to a subtraction of fluorescence signal from other samples in the run. You can remedy this by adjusting the baseline setting, but if you are unfamiliar with your instrument, you may need to call technical service for help.

Also, if there were any inhibitors in the sample from the purification step (e.g. guanidine salts or ethanol), diluting the sample will minimize their impact on the results, boosting your chances of accurately quantifying your target.

The best approach for a new sample is to perform a standard curve – even just a 3-point dilution series – to determine the template concentration that results in a Cq within range of your qPCR efficiency standard curve.

9. Make Dilutions Fresh

Nucleic acids stick to plastic so if you want to store a dilution series for future runs, you will need to prevent the nucleic acids from absorbing to the tube walls, thus becoming diluted over time.

You can achieve this by using a carrier nucleic acid, such as tRNA, or by using specially treated plasticware that does not bind nucleic acids. Several manufacturers offer low retention tubes or silicon-treated tubes to circumvent this issue.

If you do store dilutions in non-treated tubes, you may want to recheck the most concentrated dilutions on a Nanodrop before use to make sure they match the expected concentration.

10. Make Sure Your Data Is Publication Worthy – Know the MIQE Guidelines!

There isn’t much point in spending time, money and energy setting up qPCR if you can’t publish your data. In 2009, a group of UK-based researchers compiled a checklist of the minimum information required to publish qPCR data. Their goal was to streamline qPCR approaches to achieve reliability of results, integrity of scientific literature, consistency between labs, and to increase experimental transparency (1).

This list is called the MIQE guidelines, and it should supplement your initial manuscript submission to a journal. Full disclosure of all reagents, sequences, and analysis methods used is necessary to enable other investigators to reproduce results. It is also stipulated that MIQE details are published either in abbreviated form or as online supplementary material.


Ask TaqMan Episode 13 — How TaqMan Works

TaqMan genotyping assays (TaqMan® SNP Genotyping Assays and TaqMan® Drug Metabolism Genotyping Assays) consist of pre-optimized PCR primer pairs and two probes for allelic discrimination .

  • A pair of unlabeled primers
  • Two TaqMan probes (one with a FAM dye label and one with a VIC dye label) on the 5’ end and minor groove binders (MGB) and nonfluorescent quenchers (NFQ) on the 3´ end.

TaqMan genotyping assays are used to amplify and detect specific alleles in genomic DNA (gDNA). The figure below depicts the TaqMan SNP Genotyping Assay process.

  1. Genomic DNA is introduced into a reaction mixture consisting of TaqMan® Genotyping Master Mix, forward and reverse primers and two TaqMan® MGB Probes.
  2. Each TaqMan MGB Probe anneals specifically to a complementary sequence, if present, between the forward and reverse primer sites. When the probe is intact, the proximity of the quencher dye to the reporter dye suppresses the reporter fluorescence.
  3. The exonuclease activity of AmpliTaq Gold® DNA Polymerase cleaves only probes hybridized to the target. Cleavage separates the reporter dye from the quencher dye, increasing fluorescence by the reporter. The increase in fluorescence occurs only if the amplified target sequence is complementary to the probe. Thus, the fluorescence signal generated by PCR amplification indicates which alleles are in the sample.

TaqMan® Copy Number Assays are run together with a TaqMan® Copy Number Reference Assay in a duplex real-time Polymerase Chain Reaction (PCR). The Copy Number Assay detects the target gene or genomic sequence of interest and the Reference Assay detects a sequence that is known to be present in two copies in a diploid genome.

a. A TaqMan® Copy Number Assay, a TaqMan® Copy Number Reference Assay, TaqMan® Genotyping Master Mix, and a gDNA sample are mixed together in a single well or tube.

b. The gDNA template is denatured and each set of assay primers anneals to its specific target sequences. Each TaqMan® probe anneals specifically to its complementary sequence between forward and reverse primer binding sites. When each oligonucleotide probe is intact, the proximity of the quencher dye to the reporter dye causes the reporter dye signal to be quenched.

c. During each round of PCR, the target and reference sequences are simultaneously amplified by AmpliTaq® Gold DNA Polymerase. This enzyme has a 5′ nuclease activity that cleaves probes that are hybridized to each amplicon sequence. When an oligonucleotide probe is cleaved by the AmpliTaq Gold DNA Polymerase 5′ nuclease activity, the quencher is separated from the reporter dye increasing the fluorescence of the reporter. Accumulation of PCR products can be detected in real time by monitoring the increase in fluorescence of each reporter dye at each PCR cycle.

This method of relative quantitation is used to determine the relative copy number of the target of interest in a gDNA sample, normalized to the known copy number of the reference sequence.


Site Directed Mutagenesis

Site-directed mutagenesis (SDM) is a method to create specific, targeted changes in double stranded plasmid DNA. There are many reasons to make specific DNA alterations (insertions, deletions and substitutions), including:

  • To study changes in protein activity that occur as a result of the DNA manipulation.
  • To select or screen for mutations (at the DNA, RNA or protein level) that have a desired property
  • To introduce or remove restriction endonuclease sites or tags


Method Overview:

SDM is an in vitro procedure that uses custom designed oligonucleotide primers to confer a desired mutation in a double-stranded DNA plasmid. Formerly, a method pioneered by Kunkel (Kunkel, 1985) that takes advantage of a strain deficient in dUTPase and uracil deglycosylase so that the recipient E. coli degrades the uracil-containing wild-type DNA was widely used. Currently, there are a number of commercially available kits that also require specific modification and/or unique E. coli strains (for example, the Phusion ® Site-Directed Mutagenesis from Thermo and the GeneArt ® system from Life). The most widely-used methods do not require any modifications or unique strains and incorporate mutations into the plasmid by inverse PCR with standard primers. For these methods, primers can be designed in either an overlapping (QuikChange ® , Agilent) or a back-to-back orientation (Q5 ® Site-Directed Mutagenesis Kit) (Figure 1). Overlapping primer design results in a product that will re-circularize to form a doubly-nicked plasmid. Despite the presence of these nicks, this circular product can be directly transformed into E. coli, albeit at a lower efficiency than non-nicked plasmids. Back-to-back primer design methods not only have the advantage of transforming non-nicked plasmids, but also allow exponential amplification to generate significantly more of the desired product (Figure 2). In addition, because the primers do not overlap each other, deletions sizes are only limited by the plasmid and insertions are only limited by the constraints of modern primer synthesis. Currently, by splitting the insertion between the two primers, insertions up to 100 bp can routinely be created in one step using this method.

Before primers are designed, it is important to determine which mutagenesis workflow is to be used. Here we present a comparison of three commercially available kits (Figure 3) and a brief description of important features.

Before you plan your next SDM experiment, be sure to read through our list of important experimental considerations.

Kunkel, T.A. (1985) Proc Natl Acad Sci U.S.A. 82(2):488-492. PMID: 3881765

Figure 1: Site-specific mutagenesis proceeds in less than 2 hours.

The use of a master mix, a unique multi-enzyme KLD enzyme mix, and a fast polymerase ensures that, for most plasmids, the mutagenesis reaction is complete in less than two hours.

Figure 2: Q5 Site-Directed Mutagenesis Kit Overview.

This kit is designed for rapid and efficient incorporation of insertions, deletions and substitutions into doublestranded plasmid DNA. The first step is an exponential amplification using standard primers and a master mix fomulation of Q5 Hot Start High-Fidelity DNA Polymerase. The second step involves incubation with a unique enzyme mix containing a kinase, a ligase and DpnI. Together, these enzymes allow for rapid circularization of the PCR product and removal of the template DNA. The last step is a high-efficiency transformation into chemicallycompetent cells (provided).

Figure 3: Primer Design for the Q5 Site-Directed Mutagenesis Kit

Substitutions, deletions and insertions are incorporated into plasmid DNA through the use of specifically designed forward (black) and reverse (red) primers. Unlike kits that rely on linear amplification, primers designed for the Q5 Site-Directed Mutagenesis Kit should not overlap to ensure that the benefits of exponential amplification are realized. A) Substitutions are created by incorporating the desired nucleotide change(s) (denoted by *) in the center of the forward primer, including at least 10 complementary nucleotides on the 3´side of the mutation(s). The reverse primer is designed so that the 5´ ends of the two primers anneal back-to- back. B) Deletions are engineered by designing standard, non-mutagenic forward and reverse primers that flank the region to be deleted. C) Insertions less than or equal to 6 nucleotides are incorporated into the 5´ end of the forward primer while the reverse primer anneals back-to-back with the 5´ end of the complementary region of the forward primer. D) Larger insertions can be created by incorporating half of the desired insertion into the 5´ ends of both primers. The maximum size of the insertion is largely dictated by oligonucleotide synthesis limitations.

Choose Type:

This product is covered by one or more patents, trademarks and/or copyrights owned or controlled by New England Biolabs, Inc (NEB).

While NEB develops and validates its products for various applications, the use of this product may require the buyer to obtain additional third party intellectual property rights for certain applications.

For more information about commercial rights, please contact NEB's Global Business Development team at [email protected]

This product is intended for research purposes only. This product is not intended to be used for therapeutic or diagnostic purposes in humans or animals.


  • Sterile filter pipette tips
  • Sterile 1.5 mL screw-top microcentrifuge tubes (CLS430909)
    • PCR tubes and plates, select one to match desired format:
    • Individual thin-walled 200 μL PCR tubes (Z374873 or P3114)
    • Plates
      - 96-well plates (Z374903)
      - 384-well plates (Z374911)
    • Plate seals
      - ThermalSeal RTS™ Sealing Films (Z734438)
      - ThermalSeal RT2RR™ Film (Z722553)

    In the example given below, the primer concentrations can be adjusted according to the results of optimization procedures (Primer Concentration Optimization, Primer Optimization Using Temperature Gradient and Assay Optimization and Validation).


    Protocol for Enrichment of mRNAs, excluding Globin mRNA, from Whole Blood Total RNA

    This protocol describes the enrichment of poly(A) mRNA followed by globin mRNA & rRNA depletion (Section 1 and 2). The enriched RNA contains only mRNA (excluding globin) and not non-coding RNA. The enriched RNA is then used as input for directional RNA library preparation for sequencing on an Illumina instrument (Section 3).

    This protocol requires the following NEBNext products:

    • NEBNext Poly(A) mRNA Magnetic Isolation Module (NEB #E7490)
    • NEBNext Globin and rRNA Depletion Kit (Human/Mouse/Rat) with RNA Sample Purification Beads (NEB #E7755)
    • NEBNext Ultra II Directional RNA Library Prep for Illumina with Sample Purification Beads (NEB #E7765)

    RNA Sample Requirements

    RNA Integrity:
    Assess the size and quality of the input RNA by running the RNA sample on an Agilent Bioanalyzer RNA 6000 Nano/Pico Chip to determine the RNA Integrity Number (RIN). For Poly(A) mRNA enrichment, high quality RNA with RIN Score >7 is required.

    RNA Sample:
    The RNA sample should be free of salts (e.g., Mg 2+ , or guanidinium salts) or organics (e.g., phenol and ethanol). RNA must be free of DNA. gDNA is a common contaminant in RNA preps. It may be carried over from the interphase of organic extractions or when the silica matrix of solid phase RNA purification methods is overloaded. If the total RNA sample may contain gDNA contamination, treat the sample with DNase I (not provided in this kit) to remove all traces of DNA. After treatment, the DNase I should be removed from the sample. Any residual DNase I may degrade the oligos necessary for the enrichment.

    Input Amount:

    This protocol has been tested with 100 ng human whole blood total RNA (DNA-free) in a maximum of 50 µl of nuclease-free water, quantified by an RNA-specific dye-assisted fluorometric method (Qubit ® ) and quality checked by Bioanalyzer.

    Keep all buffers on ice, unless otherwise indicated.

    1.0. Poly(A) mRNA Enrichment using the NEBNext Poly(A) mRNA Magnetic Isolation Module (NEB #E7490)

    1.1. Dilute the total RNA with nuclease-free water to a final volume of 50 &mul in a nuclease-free 0.2 ml PCR tube and keep on ice.

    1.2. To wash the Oligo (dT) beads, add the components from the table below to a 1.5 ml nuclease-free tube. If preparing multiple libraries, beads for up to 10 samples can be added to a single 1.5 ml tube for subsequent washes (use magnet NEB #S1506 for 1.5 ml tubes). The purpose of this step is to bring the beads from the storage buffer into the binding buffer. The 2X Binding Buffer does not have to be diluted for this step.

    NEBNext RNA Binding Buffer (2X)

    Total volume

    1.3. Wash the beads by pipetting up and down six times.

    1.4. Place the tube on the magnet and incubate at room temperature until the solution is clear (

    1.5. Remove and discard all of the supernatant from the tube. Take care not to disturb the beads.

    1.6. Remove the tube from the magnetic rack.

    1.7. Add 100 &mul RNA Binding Buffer (2X) to the beads and wash by pipetting up and down six times. If preparing multiple libraries, add 100 µl RNA Binding Buffer (2X) per sample. The Binding Buffer does not have to be diluted.

    1.8. Place the tubes on the magnet and incubate at room temperature until the solution is clear (

    1.9. Remove and discard the supernatant from the tube. Take care not to disturb the beads.

    1.10. Add 50 &mul RNA Binding Buffer (2X) to the beads and mix by pipetting up and down until beads are homogenous. If preparing multiple libraries, add 50 &mul RNA Binding Buffer (2X) per sample.

    1.11. Add 50 &mul beads to each RNA sample from Step 1.1 Mix thoroughly by pipetting up and down six times. This binding step removes most of the non-target RNA.

    1.12. Place the tube in a thermocycler and close the lid. Heat the sample at 65°C for 5 minutes and cool to 4°C with the heated lid set at &ge 75°C. This step will denature the RNA and facilitate binding of the mRNA to the beads.

    1.13. Remove the tube from the thermocycler when the temperature reaches 4°C.

    1.14. Mix thoroughly by pipetting up and down six times. Place the tube on the bench and incubate at room temperature for 5 minutes to allow the mRNA to bind to the beads.

    1.15. Place the tube on the magnetic rack at room temperature until the solution is clear (

    1.16. Remove and discard all of the supernatant. Take care not to disturb the beads.

    1.17. Remove the tube from the magnetic rack.

    1.18. To remove unbound RNA add 200 &mul of Wash Buffer to the tube. Gently pipette the entire volume up and down 6 times to mix thoroughly.

    1.19 Spin down the tube briefly to collect the liquid from the wall and lid of the tube.

    Note: It is important to spin down the tube to prevent carryover of the Wash Buffer in subsequent steps.

    1.20. Place the tube on the magnetic rack at room temperature until the solution is clear (

    1.21. Remove and discard all of the supernatant from the tube. Take care not to disturb the beads containing the mRNA.

    1.22. Remove the tube from the magnetic rack.

    1.24. Add 11 &mul of nuclease-free water to each tube. Gently pipette up and down 6 times to mix thoroughly.

    1.25. Place the tube in the thermocycler. Close the lid and heat the samples at 80°C for 2 minutes, then cool to 25°C with the heated lid set at &ge 90°C to elute the mRNA from the beads.

    1.26. Remove the tube from the thermocycler when the temperature reaches 25°C.

    1.27 Immediately place the tube on the magnet at room temperature until the solution is clear (

    1.28. Collect the purified mRNA by transferring 10 &mul of the supernatant to a clean nuclease-free PCR tube.

    1.29. Place the RNA on ice and proceed to the Globin and rRNA Depletion in Section 2.

    2.0. Globin and rRNA Depletion using the NEBNext Globin and rRNA Depletion Kit (NEB #E7750/E7755)

    2.1 Probe Hybridization to RNA

    2.1.2. Assemble the following RNA/Probe hybridization reaction on ice:

    RNA/PROBE HYBRIDIZATION REACTION

    (white) mRNA in nuclease-free water (Step 1.29)

    (white) NEBNext Globin and rRNA Depletion Solution

    (white) NEBNext Probe Hybridization Buffer

    Total Volume

    2.1.3. Mix thoroughly by gently pipetting up and down at least 10 times. Note: It&rsquos crucial to mix well at this step.

    2.1.4. Briefly spin down the tube in a microcentrifuge to collect the liquid from the side of the tube.

    2.1.5. Place the tube in a pre-heated thermocycler and run the following program with the heated lid set at 105°C. This program will take approximately 15&ndash20 minutes to complete:

    2.1.6. Briefly spin down the tube in a microcentrifuge, and place on ice. Proceed immediately to the RNase H digestion.

    2.2. RNase H Digestion

    2.2.1. Assemble the following RNase H digestion reaction on ice:

    (white) NEBNext Thermostable RNase H

    (white) RNase H Reaction Buffer

    Total Volume

    2.2.2. Mix thoroughly by gently pipetting up and down at least 10 times.

    2.2.3. Briefly spin down the tube in a microcentrifuge.

    2.2.4. Incubate the tube in a pre-heated thermocycler for 30 minutes at 50°C with the lid set at 55°C.

    2.2.5. Briefly spin down the tube in a microcentrifuge, and place on ice. Proceed immediately to the DNase I digestion.

    2.3. DNase I Digestion

    2.3.1. Assemble the following DNase I digestion reaction on ice:

    DNASE I DIGESTION REACTION

    RNase H treated RNA (Step 1.2.5)

    (white) DNase I Reaction Buffer

    (white) NEBNext DNase I

    Total Volume

    2.3.2. Mix thoroughly by pipetting up and down at least 10 times.

    2.3.3. Briefly spin down the tube in a microcentrifuge.

    2.3.4. Incubate the tube in a pre-heated thermocycler for 30 minutes at 37°C with the lid set at 40°C or off.

    2.3.5. Briefly spin down the tube in a microcentrifuge, and place on ice. Proceed immediately to the RNA Purification step.

    2.4. RNA Purification Using Agencourt RNAClean XP Beads or NEBNext RNA Sample Purification Beads

    2.4.1. Vortex the Agencourt RNAClean XP Beads or NEBNext RNA Sample Purification Beads to resuspend.

    2.4.2. Add 90 &mul (1.8X) beads to the RNA Sample from Step 2.3.5 and mix thoroughly by pipetting up and down at least 10 times.

    2.4.3. Incubate the tube for 15 minutes on ice to bind the RNA to the beads.

    2.4.4. Place the tube on a magnetic rack to separate the beads from the supernatant.

    2.4.5. After the solution is clear, carefully remove and discard the supernatant. Be careful not to disturb the beads which contain the RNA.

    2.4.6. Add 200 µl of freshly prepared 80% ethanol to the tube while in the magnetic rack. Incubate at room temperature for 30 seconds, and then carefully remove and discard the supernatant. Be careful not to disturb the beads, which contain the RNA.

    2.4.7. Repeat Step 2.4.6 once for a total of two washes.

    2.4.8. Completely remove residual ethanol, and air dry the beads for up to 5 minutes while the tube is on the magnetic rack with the lid open.

    Caution: Do not over-dry the beads. This may result in lower recovery of RNA target. Elute the samples when the beads are still dark brown and glossy looking, but when all visible liquid has evaporated. When the beads turn lighter brown and start to crack they are too dry.

    2.4.9. Remove the tube from the magnetic rack. Elute the RNA from the beads by adding 7 &mul of nuclease-free water. Mix thoroughly by pipetting up and down at least 10 times and briefly spin the tube.

    2.4.10. Incubate the tube for 2 minutes at room temperature.

    2.4.11. Place the tube on the magnetic rack until the solution is clear (

    2.4.12. Remove 5 µl of the supernatant containing RNA and transfer to a nuclease-free tube.

    2.4.13. Place the tube on ice and proceed with RNA-Seq library construction (protocol below) or other downstream application. Alternatively, the sample can be stored at -80°C.

    Note: The next step provides a fragmentation incubation time resulting in an RNA insert of

    200nt. Refer to Appendix (Section 4 of the NEBNext Ultra II Directional RNA Library Prep for Illumina Manual) for fragmentation conditions if you are preparing libraries with large inserts (>200 bp).

    3.1.3. Incubate the sample for 15 minutes at 94°C in a thermocycler with the heated lid set at 105°C.

    3.1.4. Immediately transfer the tube to ice for 1 minute.

    3.1.5 Perform a quick spin to collect all liquid from the sides of the tube and proceed to First Strand cDNA Synthesis.

    3.2. First Strand cDNA Synthesis

    3.2.1. Assemble the first strand synthesis reaction on ice by adding the following components to the fragmented
    and primed RNA from Step 3.1.5:

    FIRST STRAND SYNTHESIS REACTION

    Fragmented and primed RNA (Step 3.1.5)

    (brown) NEBNext Strand Specificity Reagent

    (lilac) NEBNext First Strand Synthesis Enzyme Mix

    Total Volume

    3.2.2. Mix thoroughly by pipetting up and down 10 times.

    3.2.3. Incubate the sample in a preheated thermocycler with the heated lid set at &ge 80°C as follows:

    Note: If you are following recommendations in Section 4 of the NEBNext Ultra II Directional RNA Library Prep for Illumina Manual for libraries with longer inserts (>200 bases), increase the incubation at 42°C from 15 minutes to 50 minutes at Step 2 below.

    3.2.4. Proceed directly to Second Strand cDNA Synthesis.

    3.3. Second Strand cDNA Synthesis

    3.3.1. Assemble the second strand cDNA synthesis reaction on ice by adding the following components into the first strand synthesis product from Step 3.2.4.

    SECOND STRAND SYNTHESIS REACTION

    First-Strand Synthesis Product (Step 3.2.4)

    (orange) NEBNext Second Strand Synthesis Reaction Buffer
    with dUTP Mix (10X)

    (orange) NEBNext Second Strand Synthesis Enzyme Mix

    Total Volume

    3.3.2. Keeping the tube on ice, mix thoroughly by pipetting up and down at least 10 times.

    3.3.3. Incubate in a thermocycler for 1 hour at 16°C with the heated lid set at &le 40°C (or off).

    3.4. Purification of Double-stranded cDNA Using SPRIselect Beads or NEBNext Sample Purification Beads

    3.4.1. Vortex SPRIselect Beads or NEBNext Sample Purification Beads to resuspend.

    3.4.2. Add 144 &mul (1.8X) of resuspended beads to the second strand synthesis reaction (

    80 &mul). Mix well on a vortex mixer or by pipetting up and down at least 10 times.

    3.4.3. Incubate for 5 minutes at room temperature.

    3.4.4. Briefly spin the tube in a microcentrifuge to collect any sample on the sides of the tube. Place the tube on a magnetic rack to separate beads from the supernatant. After the solution is clear, carefully remove and discard the supernatant. Be careful not to disturb the beads, which contain DNA. Caution: do not discard beads.

    3.4.5. Add 200 &mul of freshly prepared 80% ethanol to the tube while in the magnetic rack. Incubate at room temperature for 30 seconds, and then carefully remove and discard the supernatant.

    3.4.6. Repeat Step 3.4.5 once for a total of 2 washing steps.

    3.4.7. Air dry the beads for up to 5 minutes while the tube is on the magnetic rack with lid open.

    Caution: Do not over-dry the beads. This may result in lower recovery of DNA target. Elute the samples when the beads are still dark brown and glossy looking, but when all visible liquid has evaporated. When the beads turn lighter brown and start to crack they are too dry.

    3.4.8. Remove the tube from the magnetic rack. Elute the DNA from the beads by adding 53 &mul 0.1X TE Buffer (provided) to the beads. Mix well on a vortex mixer or by pipetting up and down at least 10 times. Quickly spin the tube and incubate for 2 minutes at room temperature. Place the tube on the magnetic rack until the solution is clear.

    3.4.9. Remove 50 µl of the supernatant and transfer to a clean nuclease-free PCR tube.

    Note: If you need to stop at this point in the protocol samples can be stored at &ndash20°C.

    3.5. End Prep of cDNA Library

    3.5.1. Assemble the End Prep reaction on ice by adding the following components to the second strand synthesis product from
    Step 3.4.9.

    Second Strand cDNA Synthesis Product (Step 3.4.9)

    (green) NEBNext Ultra II End Prep Reaction Buffer

    (green) NEBNext Ultra II End Prep Enzyme Mix

    Total Volume

    If a master mix is made, add 10 µl of master mix to 50 µl of cDNA for the End Prep reaction.

    3.5.2. Set a 100 &mul or 200 &mul pipette to 50 &mul and then pipette the entire volume up and down at least 10 times to mix thoroughly. Perform a quick spin to collect all liquid from the sides of the tube.

    Note: It is important to mix well. The presence of a small amount of bubbles will not interfere with performance.

    3.5.3. Incubate the sample in a thermocycler with the heated lid set at &ge 75°C as follows.

    3.5.4. Proceed immediately to Adaptor Ligation.

    3.6. Adaptor Ligation

    Note: If you are selecting for libraries with larger insert size (>200 nt) follow the size selection recommendations in Appendix, Section 4 of the NEBNext Ultra II Directional RNA Library Prep for Illumina Manual.

    3.7.1. Add 87 &mul (0.9X) resuspended SPRIselect Beads or NEBNext Sample Purification Beads and mix well on a vortex mixer or by pipetting up and down at least 10 times.

    3.7.2. Incubate for 10 minutes at room temperature.

    3.7.3. Quickly spin the tube in a microcentrifuge and place the tube on a magnetic rack to separate beads from the supernatant. After the solution is clear (

    5 minutes), discard the supernatant that contains unwanted fragments. Caution: do not discard beads.

    3.7.4. Add 200 &mul of freshly prepared 80% ethanol to the tube while in the magnetic rack. Incubate at room temperature for 30 seconds, and then carefully remove and discard the supernatant.

    3.7.5. Repeat Step 3.7.4 once for a total of 2 washing steps.

    3.7.6. Briefly spin the tube and put the tube back in the magnetic rack.

    3.7.7. Completely remove the residual ethanol, and air-dry beads until the beads are dry for up to 5 minutes while the tube is on the magnetic rack with the lid open.

    Caution: Do not over-dry the beads. This may result in lower recovery of DNA target. Elute the samples when the beads are still dark brown and glossy looking, but when all visible liquid has evaporated. When the beads turn lighter brown and start to crack they are too dry.

    3.7.8. Remove the tube from the magnetic rack. Elute DNA target from the beads by adding 17 &mul 0.1X TE (provided) to the beads. Mix well on a vortex or by pipetting up and down. Quickly spin the tube and incubate for 2 minutes at room temperature. Place the tube in the magnet until the solution is clear.

    3.7.9. Without disturbing the bead pellet, transfer 15 &mul of the supernatant to a clean PCR tube and proceed to PCR enrichment.

    Note: If you need to stop at this point in the protocol, samples can be stored at &ndash20°C.

    3.8. PCR Enrichment of Adaptor Ligated DNA

    Check and verify that the concentration of your oligos is 10 &muM on the label.

    Use Option A for any NEBNext Oligos kit where index primers are supplied in tubes. These kits have the forward and reverse primers supplied in separate tubes.

    Use Option B for any NEBNext Oligos kit where index primers are supplied in a 96-well plate format. These kits have the forward and reverse primers (i7 and i5) combined.

    3.8.1. Set up the PCR reaction as described below based on the type of oligos (PCR primers) used.

    3.8.1A. Forward and Reverse Primers Separate

    Adaptor Ligated DNA (Step 2.11.9)

    (blue) NEBNext Ultra II Q5 ® Master Mix

    Universal PCR Primer/i5 Primer*,**

    Total Volume

    * NEBNext Oligos must be purchased separately from the library prep kit. Refer to the corresponding NEBNext Oligo kit manual
    for determining valid barcode combinations.

    ** Use only one i7 primer/ index primer per sample. Use only one i5 primer (or the universal primer for single index kits) per sample.

    3.8.1B. Forward and Reverse Primers Combined

    Adaptor Ligated DNA (Step 2.11.9)

    (blue) NEBNext Ultra II Q5 Master Mix

    Index (X) Primer/i7 Primer Mix*

    Total Volume

    * NEBNext Oligos must be purchased separately from the library prep kit. Refer to the corresponding NEBNext Oligo kit manual
    for determining valid barcode combinations.

    ** Use only one i7 primer/ index primer per sample. Use only one i5 primer (or the universal primer for single index kits) per sample

    3.8.2. Mix well by gently pipetting up and down 10 times. Quickly spin the tube in a microcentrifuge.

    3.8.3. Place the tube on a thermocycler with the heated lid set to 105°C and perform PCR amplification using the following PCR cycling conditions (refer to Table 3.8.3A and Table 3.8.3B):

    * The number of PCR cycles should be adjusted based on RNA input.

    ** It is important to limit the number of PCR cycles to avoid overamplification.
    If overamplification occurs, a second peak

    1,000 bp will appear on the Bioanalyzer trace (See Figure 5.2 of the NEBNext Ultra II Directional RNA Library Prep for Illumina Manual).

    Table 3.8.3B: Recommended PCR cycles based on total RNA input amount:

    * The PCR cycles are recommended based on high quality human whole blood total RNA. To prevent over-amplification, the number of cycles may require optimization based on the sample quality and the fraction of globin mRNA. For RNA where globin mRNA is > than 50% of the transcripts (once rRNA is removed), follow the higher cycle recommendation for that input.

    3.9. Purification of the PCR Reaction using SPRIselect Beads or NEBNext Sample Purification Beads

    3.9.1. Vortex SPRIselect Beads or NEBNext Sample Purification Beads to resuspend.

    3.9.2. Add 45 &mul (0.9X) of resuspended beads to the PCR reaction (

    50 &mul). Mix well on a vortex mixer or by pipetting up and down at least 10 times.

    3.9.3. Incubate for 5 minutes at room temperature.

    3.9.4. Quickly spin the tube in a microcentrifuge and place the tube on a magnetic rack to separate beads from the supernatant. After the solution is clear (about 5 minutes), carefully remove and discard the supernatant. Be careful not to disturb the beads that contain DNA targets. Caution: do not discard beads.

    3.9.5. Add 200 &mul of freshly prepared 80% ethanol to the tube while in the magnetic rack. Incubate at room temperature for 30 seconds, and then carefully remove and discard the supernatant.

    3.9.6. Repeat Step 3.9.5 once for a total of 2 washing steps.

    3.9.7. Air dry the beads for up to 5 minutes while the tube is on the magnetic rack with the lid open.

    Caution: Do not over-dry the beads. This may result in lower recovery of DNA target. Elute the samples when the beads are still dark brown and glossy looking, but when all visible liquid has evaporated. When the beads turn lighter brown and start to crack they are too dry.

    3.9.8. Remove the tube from the magnetic rack. Elute the DNA target from the beads by adding 23 &mul 0.1X TE (provided) to the beads. Mix well on a vortex mixer or by pipetting up and down ten times. Quickly spin the tube in a microcentrifuge and incubate for 2 minutes at room temperature. Place the tube in the magnetic rack until the solution is clear.

    3.9.9. Transfer 20 &mul of the supernatant to a clean PCR tube and store at &ndash20°C.

    3.10. Library Quantification

    3.10.1. Use a Bioanalyzer or TapeStation to determine the size distribution and concentration of the libraries.

    3.10.2. Check that the electropherogram shows a narrow distribution with a peak size approximately 300 bp.

    80 bp (primers) or 128 bp (adaptor-dimer) is visible in the bioanalyzer traces, bring up the sample volume (from Step 3.9.9) to 50 &mul with 0.1X TE buffer and repeat the SPRIselect Bead or NEBNext Sample Purification Bead Cleanup Step (Section 3.9).

    Figure 3.9.1 Example of RNA library size distribution on a Bioanalyzer.


    Colony PCR or PCR with Growing culture in Broth - Trying to detect the cloned fragment in living cells (Jan/18/2007 )

    Hey Guys
    Another Question for you.
    What is the best technique to do PCR on living cells, cells from colony or can I directly take cell from overnight cultures which I use for plasmid extraction. I tried it once while taking 1 microliter from the broth culture in a total volume of 25 microliter. It worked and I got the amplification but I am not sure, will it work in every case.
    Any Suggestions are highly appreciated.
    And if any body has a nice protocol for this, I will request to post it here too.
    Thanks in advance.

    if the cells are oldish (growing for >16hrs) I would avoid using them for PCR. This is true for both colonies and cultures.. cells growing more then 16hrs greatly increase their production of liposaccarides (slime!) which intefers with the PCR reaction.

    Otherwise (provide sufficient dilution) and the cells are 'fresh-young' there is no difference between the two methods. Personally I prefer doing my PCR on colonies.. i use colony PCR for screening purposes and doing the screen on colonies saves me a day.

    Use a 96 well plate to hold my colony dilutions.

    Fill 96 well plate with 50ul sterile distilled water
    Pick 1 isolated colony with pipette tip and place said colony into a well.
    Repeat as many times as desired.

    PCR mix per reaction (but this is usually made a single master mix)
    4.22 ul sterile distilled water
    1ul 2mM dNTP
    1ul 10X taq buffer
    0.5ul primer forward
    0.5ul primer reverse
    0.08ul Taq
    0.2ul 50mM MgCl2

    Add PCR mix into 96 Well PCR plate. Then add

    1 = 95 Celsius (4min)
    2 = 94 Celsius (30 sec) melting
    3 = Tm (30 sec) anealing
    4 = 72 Celsius (1min) extention

    Of course the above conditions can be changed. the time for each step is a little too long, but I don't bother to optimise it for each individual screen.

    EDIT: The primer concentration is 10mM, made by diluting a master stock (100mM) with sterile distilled water.

    if the cells are oldish (growing for >16hrs) I would avoid using them for PCR. This is true for both colonies and cultures.. cells growing more then 16hrs greatly increase their production of liposaccarides (slime!) which intefers with the PCR reaction.

    Otherwise (provide sufficient dilution) and the cells are 'fresh-young' there is no difference between the two methods. Personally I prefer doing my PCR on colonies.. i use colony PCR for screening purposes and doing the screen on colonies saves me a day.

    Use a 96 well plate to hold my colony dilutions.

    Fill 96 well plate with 50ul sterile distilled water
    Pick 1 isolated colony with pipette tip and place said colony into a well.
    Repeat as many times as desired.

    PCR mix per reaction (but this is usually made a single master mix)
    4.22 ul sterile distilled water
    1ul 2mM dNTP
    1ul 10X taq buffer
    0.5ul primer forward
    0.5ul primer reverse
    0.08ul Taq
    0.2ul 50mM MgCl2

    Add PCR mix into 96 Well PCR plate. Then add

    1 = 95 Celsius (4min)
    2 = 94 Celsius (30 sec) melting
    3 = Tm (30 sec) anealing
    4 = 72 Celsius (1min) extention

    Of course the above conditions can be changed. the time for each step is a little too long, but I don't bother to optimise it for each individual screen.

    if the cells are oldish (growing for >16hrs) I would avoid using them for PCR. This is true for both colonies and cultures.. cells growing more then 16hrs greatly increase their production of liposaccarides (slime!) which intefers with the PCR reaction.

    Otherwise (provide sufficient dilution) and the cells are 'fresh-young' there is no difference between the two methods. Personally I prefer doing my PCR on colonies.. i use colony PCR for screening purposes and doing the screen on colonies saves me a day.

    Hi, I've been getting negative results all this time for my colony pcr screen! White colonies with negative results really depressing
    My colonies have been on the plate for 2 days (unfortunately) cause they were really too small to be picked.
    Would it also matter if I had resuspended the colonies in LB (5ul) instead of sterile water? Had realised you used 50ul!!
    Maybe i should try out your dilution method as well.

    yes, I certainly agree. Do dilute your colony solution. If the colony solution is too concentrated, the PCR reaction will either fail or become very smeary.

    If only a small amount of LB entered the PCR mix, 2ul or so, there should not be any effect. But personally I use sterile distilled water. Should I decide to keep the colonies in the 96 well plate for an extended time (>2 days), I later add LB to the wells.

    Hi perneseblue,
    I just added another 50ul of distilled water as u suggested.
    But unfortunately, still nothing turned up
    Have tried on 36 samples !

    Did u do any serial dilutions, or change the volumes of Colony dilutons for your PCR screen?
    Anyway, I'll be repeating the TA cloning step once more since my plate is about 4days old now

    Is there any other factors other than colony dilutions? I've browsed through the forum and it seems that there are also other members facing similar problems: picking up white colonies but not finding the inserts

    I'd like to foresee any upcoming problems and be mentally prepared

    Hi perneseblue,
    I just added another 50ul of distilled water as u suggested.
    But unfortunately, still nothing turned up
    Have tried on 36 samples !

    Did u do any serial dilutions, or change the volumes of Colony dilutons for your PCR screen?
    Anyway, I'll be repeating the TA cloning step once more since my plate is about 4days old now

    Is there any other factors other than colony dilutions? I've browsed through the forum and it seems that there are also other members facing similar problems: picking up white colonies but not finding the inserts

    I'd like to foresee any upcoming problems and be mentally prepared

    1. Add 100 uL water to desired number of wells in a PCR plate or tubes.
    2. Toothpick colony into each well or add 10 uL of overnight culture
    3. Heat in thermal cycler to 95C for 5 or 10 minutes. Heat blocks are not recommended, as the protocol does not work as well if the temperature is 93C rather than 95C.
    4. Spin down the plate or tubes so that any cell debris is pelleted at the bottom of the tube/well. This removes the inhibitory components from the supernatant.
    5. Carefully transfer 25 uL of supernatant to the PCR reaction, being careful not to disturb the bottom of the tube/well, even if you can't see any pellet.
    6. PCR with forward and reverse to check for single/double inserts. Use vector only for a control.

    hi makosad05 and tfitzwater,

    Thanks for your replies!
    I use the same primers as my inserts, 40cycles
    my PCR reaction is 10ul

    Hi perneseblue,
    I just added another 50ul of distilled water as u suggested.
    But unfortunately, still nothing turned up
    Have tried on 36 samples !

    Did u do any serial dilutions, or change the volumes of Colony dilutons for your PCR screen?
    Anyway, I'll be repeating the TA cloning step once more since my plate is about 4days old now

    Is there any other factors other than colony dilutions? I've browsed through the forum and it seems that there are also other members facing similar problems: picking up white colonies but not finding the inserts

    I'd like to foresee any upcoming problems and be mentally prepared

    The PCR product size. I find that under colony PCR conditions, Taq polymerase is reliably only for amplification of 900bp products and below. Anything larger, and the reaction's success becomes uncertain. If your product is on the large side, I would try reducing the extention temperature (to around 70 Celsius - 68 Celsius) compensating by increased extention time. (50% longer)

    tm- what is the tm of your primers? Like any PCR reaction, all rules concerning primers design apply. There shouldn't be too large a difference between primers melting temperature.

    The rule of the tumb for annealing tempearture is (tm - 5 Celsius). You can reduce the annealing temperature, a little further.

    I would also consider testing more colonies, up to 72.

    Lastly, there is there is the possibility that there the colony PCR isn't working because there aren't any colonies which are positive.


    Principles of Molecular Techniques

    Faramarz Naeim , . Wayne W. Grody , in Atlas of Hematopathology , 2013

    Related Amplification Techniques

    A large number of innovations to the basic PCR technique have been developed over the years to address particular applications or to circumvent certain pitfalls. Included are such techniques as nested PCR, whole-genome amplification, inverse PCR, hot-start PCR, allele-specific PCR, cold PCR, and many others. For the most part they are beyond the scope of this chapter, but will be mentioned in the context of particular disease applications where relevant elsewhere in the book. In addition, a number of non-PCR amplification techniques have been developed over the years, such as Q-β replicase, ligation chain reaction, etc., but for the most part they have fallen by the wayside in favor of PCR, at least for applications relevant to hematopathology (some are used in molecular microbiology and genetics testing).


    Watch the video: Dilution Problems - Chemistry Tutorial (May 2022).


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