Tips and Tools

Tips for Successful Dual-Reporter Assays

Posted on by Ben Schmidt

Updated 02/12/2021

Previously, we described some of the advantages of using dual-reporter assays (such as the Dual-Luciferase®, Dual-Glo® Luciferase and the Nano-Glo® Dual-Luciferase® Systems). Another post describes how to choose the best dual-reporter assay for your experiments. For an overview of luciferase-based reporter gene assays, see this short video:

These assays are relatively easy to understand in principle. Use a primary and secondary reporter vector transiently transfected into your favorite mammalian cell line. The primary reporter is commonly used as a marker for a gene, promoter, or response element of interest. The secondary reporter drives a steady level of expression of a different marker. We can use that second marker to normalize the changes in expression of the primary under the assumption that the secondary marker is unaffected by what is being experimentally manipulated.

While there are many advantages to dual-reporter assays, they require careful planning to avoid common pitfalls. Here’s what you can do to avoid repeating some of the common mistakes we see with new users:

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T-Vector Cloning: Questions, Answers and Tips

Posted on by Leah Cronan
Blue/White colony screening helps you pick only the colonies that have your insert.

Q: Can PCR products generated with GoTaq DNA Polymerase be used to for T- vector cloning?

A: Yes. GoTaq® DNA Polymerase is a robust formulation of unmodified Taq Polymerase. GoTaq® DNA Polymerase lacks 3’ →5’ exonuclease activity and displays terminal transferase activity that adds a 3′ deoxyadenosine (dA) to product ends. As a result, PCR products amplified using GoTaq® DNA Polymerases (including the GoTaq® Flexi and GoTaq® G2 polymerases) will contain A-overhangs which makes them suitable for T-vector cloning with the pGEM®-T (Cat.# A3600), pGEM®-T Easy (Cat.# A1360) and pTARGET™ (Cat.# A1410) Vectors.

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I Have My Luciferase Vector, Now What?

Posted on by Joliene Lindholm
Choosing and Optimizing Transfection Methods

Here in Technical Services we often talk with researchers at the beginning of their project about how to carefully design and get started with their experiments. It is exciting when you have selected the Luciferase Reporter Vector(s) that will best suit your needs; you are going to make luminescent cells! But, how do you pick the best way to get the vector into your cells to express the reporter? What transfection reagent/method will work best for your cell type and experiment? Do you want to do transient (short-term) transfections, or are you going to establish a stable cell line?

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qPCR: The Very Basics

Posted on by Michele Arduengo
Real-Time (or quantitative, qPCR) monitors PCR amplification as it happens and allows you to measure starting material in your reaction.
qPCR monitors amplification in real and allows you to measure starting material.

For those of us well versed in traditional, end-point PCR, wrapping our minds and methods around real-time or quantitative (qPCR) can be challenging. Here at Promega Connections, we are beginning a series of blogs designed to explain how qPCR works, things to consider when setting up and performing qPCR experiments, and what to look for in your results.

First, to get our bearings, let’s contrast traditional end-point PCR with qPCR.

End-Point PCRqPCR
Visualizes by agarose gel the amplified product AFTER it is produced (the end-point)Visualizes amplification as it happens (in real time) via a detection instrument
Does not precisely measure the starting DNA or RNAMeasures how many copies of DNA or RNA you started with (quantitative = qPCR)
Less expensive; no special instruments requiredMore expensive; requires special instrumentation
Basic molecular biology techniqueRequires slightly more technical prowess

Quantitative PCR (qPCR) can be used to answer the same experimental questions as traditional end-point PCR: Detecting polymorphisms in DNA, amplifying low-abundance sequences for cloning or analysis, pathogen detection and others. However, the ability to observe amplification in real-time and detect the number of copies in the starting material can quantitate gene expression, measure DNA damage, and quantitate viral load in a sample and other applications.

Anytime that you are performing a reaction where something is copied over and over in an exponential fashion, contaminants are just as likely to be copied as the desired input. Quantitative PCR is subject to the same contamination concerns as end-point PCR, but those concerns are magnified because the technique is so sensitive. Avoiding contamination is paramount for generating qPCR results that you can trust.

  1. Use aerosol-resistant pipette tips, and have designated pipettors and tips for pre- and post-amplification steps.
  2. Wear gloves and change them frequently.
  3. Have designated areas for pre- and post-amplification work.
  4. Use reaction “master mixes” to minimize variability. A master mix is a ready-to-use mixture of your reaction components (excluding primers and sample) that you create for multiple reactions. Because you are pipetting larger volumes to make the reaction master mix, and all of your reactions are getting their components from the same master mix, you are reducing variability from reaction to reaction.
  5. Dispense your primers into aliquots to minimize freeze-thaw cycles and the opportunity to introduce contaminants into a primer stock.

These are very basic tips that are common to both end-point and qPCR, but if you get these right, you are off to a good start no matter what your experimental goals are.

If you are looking for more information regarding qPCR, watch this supplementary video below.

We’re committed to supporting scientists who are using molecular biology to make a difference. Learn more about our qPCR Grant program.  

Are you looking for more in-depth information about qPCR? Check out our qPCR and RT-qPCR Guide!

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Optimizing PCR: One Scientist’s Not So Fond Memories

Posted on by Kelly Grooms

primer_tubesThe first time I performed PCR was in 1992. I was finishing my Bachelors in Genetics and had an independent study project in a population genetics laboratory. My task was to try using a new technique, RAPD PCR, to distinguish clonal populations of the sea anemone, Metridium senile. These creatures can reproduce both sexually and asexually, which can make population genetics studies challenging. My professor was looking for a relatively simple method to identify individuals who were genetically identical (i.e., potential clones).

PCR was still in its infancy. No one in my lab had ever tried it before, and the department had one thermal cycler, which was located in a building across the street. We had a paper describing RAPD PCR for population work, so we ordered primers and Taq DNA polymerase and set about grinding up bits of frozen sea anemone to isolate the DNA. [The grinding process had to be done using a mortar and pestle seated in a bath of liquid nitrogen because the tissue had to remain frozen. If it thawed it became a disgusting mass of goo that was useless—but that is a topic for a different blog.] Since I had never done any of the procedures before, my professor and I assembled the first set of reactions together. When we ran our results on a gel, we had all sorts of bands—just what he was hoping to see. Unfortunately, we realized that we had added 10X more Taq DNA polymerase than we should have used. I repeated the amplification with the correct amount of Taq polymerase, and I saw nothing. Continue reading “Optimizing PCR: One Scientist’s Not So Fond Memories”

Think Restriction Enzymes are so last decade? Not so fast!

Posted on by Leah Cronan
Ribbon diagram of EcoRI homodimer bound to doublestranded DNA
Ribbon diagram of EcoRI homodimer bound to doublestranded DNA

Restriction enzymes sometimes get a lot of flak. In the not-so-distant past, they were the workhorses of molecular biology. Restriction enzymes played a huge role in developing early DNA sequencing techniques. They chop DNA in a predictable manner, which makes cutting and pasting genes of interest manageable and relatively easy, enabling the development of  genetic engineering and recombination technologies. These technologies are now moving beyond restriction enzymes toward more modern methods, with the most talked-about method being CRISPR /Cas9. As technology continues to advance at such a rapid pace, restriction analysis  and other “ancient” technologies feel antiquated. But this is not necessarily the case.

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Restriction Enzyme Digestion: Capabilities and Resources

Posted on by Promega
Image of someone pipetting while setting up a restriction enzyme digestion

Restriction enzymes recognize short DNA sequences and cleave double-stranded DNA at specific sites within or adjacent to these sequences.  These enzymes are the workhorse in many molecular biology applications such as cloning, RFLP, methylation-specific restriction enzyme analysis of DNA, etc.  Restriction enzymes with enhanced capabilities can help you streamline and shorten these workflows and improve success of restriction enzyme digestion.

A subset of Promega restriction enzymes offer capabilities that  include rapid digestion of DNA in 15 minutes or less, ability to completely digest DNA directly in the GoTaq® Green Master Mix, and Blue/White Cloning Qualification which allows for rapid, reliable detection of transformants.

To learn more about restriction enzymes and applications, check out Restriction Enzyme Resource on the web. The resource provides everything from information on restriction enzyme biology to practical information on how to set up and design a restriction enzyme digestion. This resource also contains useful online tools, including the Restriction Enzyme Tool, to help you use enzymes more effectively. It helps you choose the best reaction buffer for double digests, find the commercially available enzyme that cuts your sequence of interest, find compatible ends, and search for specific information on cut site, overhang isoschizomers and neoschizomers by enzyme name.

For added convenience, you can download the mobile app available for iOS devices and use the Restriction Enzyme Tool to plan your next digest.

For additional information regarding Restriction Enzyme Digest, reference the supplementary video below.

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A Quick Method for A Tailing PCR Products

Posted on by Isobel Maciver
PCR experiment and products, pipette tip, tube in researcher's hand.
PCR is a common technique used in research labs to amplify DNA.

Some thermostable DNA polymerases, including Taq, add a single nucleotide base extension to the 3′ end of amplified DNA fragments. These polymerases usually add an adenine, leaving an “A” overhang. There are several approaches to overcome the cloning difficulties presented by the presence of A overhangs on PCR products. One method involves treating the product with Klenow to create a blunt-ended fragment for subcloning. Another choice is to add restriction sites to the ends of your PCR fragments. You can do this by incorporating the desired restriction sites into the PCR primers. After amplification, the PCR product is digested and subcloned into the cloning vector. Take care when using this method, as not all restriction enzymes efficiently cleave at the ends of DNA fragments, and you may not be able to use every restriction enzyme you desire. There is some useful information about cutting with restriction sites close to the end of linear fragments in the Restriction Enzyme Resource Guide. Also, some restriction enzymes require extra bases outside the recognition site, adding further expense to the PCR primers as well as risk of priming to unrelated sequences in the genome.

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Selecting the Right Colony: The Answer is There in Blue and White

Posted on by Kelly Grooms
Agar plate containing colonies important for research, blue and white.
Agar plate containing colonies important for research.

Ah, the wonders and frustrations of cloning. We’ve all been there. After careful planning, you have created the cloned plasmid containing your DNA sequence of interest, transformed it into bacterial cells and carefully spread those cells on a plate to grow. Now you stand at your bench gazing down at your master piece: a plate full of tiny bacterial colonies. Somewhere inside those cells is your DNA sequence, happily replicating with its plasmid host. But wait – logic tells you that not ALL of those colonies can contain your plasmid.  There must be hundreds of colonies. Which ones have your plasmid? You begin to panic. Visions of yourself old and grey and still screening colonies flash through your mind. At the next bench, your lab-mate is cheerfully selecting colonies to screen. Although there are hundreds of colonies on her plate as well, some are white and some are blue. She is only picking the white colonies. What does she know that you don’t?

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Lab Sustainability: Easy as 1-2-3

Posted on by Darcia Schweitzer

Sustainability is a bit of buzzword lately—for good reason—but knowing how to be more sustainable and actually putting sustainable practices in action are not the same thing. This may be one reason why scientists have been slow to adopt change in their laboratories. By sponsoring My Green Lab, we’re hoping to help spread the message that there are simple changes researchers can make in their labs to significantly impact sustainability.

Here are some easy ways to reduce energy, water and waste in your lab and start making your research more sustainable.

1. Energy

Compared to office buildings on campus, academic lab buildings consume 5 times more energy. To put that into perspective, labs typically consume 50% of the energy on a university campus despite occupying less than 30% of the space. Fortunately, reducing energy usage can be one of the easiest ways to make your lab more sustainable.

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