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Methods

Posted by Karen on May 7th, 2015  ⟩  0 comments

One day your PCR works; then when you repeat it, you get no results, and when you try yet again, you get nonspecific binding. It’s these situations that drive you to superstitious rituals and prayers to the PCR gods for mercy. Unfortunately, divine forms of troubleshooting yield few results.

#PCRLuckyBuddies

For a veteran in life science research, PCR has become second nature; however, I have seen undergraduate and graduate students highly stressed out about PCR. I have heard them utter to one another, “I’m about to see if this works. Wish me luck.” I knew one undergraduate student who struggled for an entire semester trying to make PCR work only to end up switching majors. After investing years into scientific coursework and research, we don’t want it to come to that. Instead, we’re here to help.

Rather than having a huge troubleshooting article with every single PCR tip known to scientists, I’ll break this into a series: nonspecific binding, no results, smearing, weak results and contamination.

4 PCR Tips When Encountering Nonspecific Binding:


1.  Aliquot Aliquot Aliquot: If you remember the article about fridge & freezer organization, certain areas of your upright freezer have a greater risk of unintentionally exposing your reagents to freeze-thaws. Outside of the freezer, you also run the risk of contamination to your reagents that might degrade them. Simply put, protect your supplies. Aliquot DNTPs, primers, etc. Only move your working vials into easily accessible freezer boxes. Store the rest of your stock in a more protected area at -80 °C.

2.  Negative Controls: This is a must in every PCR setup. In fact, my mentor made it a practice to set up the first two tubes and last two tubes to be positive and negative controls. This let both he and I see if contamination was ever a reason for a bad PCR. And it let us see whether or not it occurred throughout the whole process.

3.  Increase Annealing Temp: By increasing the annealing temperature, you’re driving specificity. In general, you want to use an annealing temperature that is 5 °C lower than the Tm of your primers.

4.  Touch-down PCR: In this process, the first stages of PCR should have a high annealing temperature, even higher than the estimated Tm of your primers. Following cycles have incrementally lower annealing temperatures. This gradual adjustment stops when you have reached the calculated annealing temperature of your PCR primers. The tricky part will be deciding on the incremental decreases you want to use. With the higher annealing temperature being used at the beginning, the resulting sequence will be the most specific and able to out-compete nonspecific results.

Stay tuned for more PCR tips in the following article. We hope that some of these tips begin to help you attain the results you’re hoping for.


    
              Karen Martin
GoldBio Marketing Coordinator


"To understand the universe is to understand math." My 8th grade
math teacher's quote meant nothing to me at the time. Then came
college, and the revelation that the adults in my past were right all
along. But since math feels less tangible, I fell for biology and have
found pure happiness behind my desk at GoldBio, learning, writing
and loving everything science. 



Category Code: 79103 79108

Posted by Karen on February 25th, 2015  ⟩  0 comments

There are a lot of products in the lab that do similar things but were developed for a specialized purpose. And sometimes you’re in a position where only the alternative is available, but you’re unsure whether you can trust it.

Let’s take a look at a few of these products and see exactly what the difference is in order to determine which one is more appropriate, given a certain experiment.

1. Tris vs. Tris HCl

     vs.  

Both can be used for electrophoresis, but why would you choose one or the other? Tris is much less expensive compared to tris HCl; however, tris HCl is meant to simplify the buffer-making process.

On its own, tris is the basic component of the buffer, while the acidic component of the buffer would come from adding HCl. Rather than working with HCl or NaOH (to adjust the pH of tris HCl), tris and tris HCl can be blended together to reach the desired pH.

Making a tris buffer solution with tris-HCl also prevents overshoot, which occurs when too much acid is accidentally added, meaning NaOH will need to be added to correct the situation.

Verdict:
In the end, it’s handy to have both in the lab. But either or can be used, so long as you have the acid or base to properly adjust it.

2. XTT vs. MTT

          vs.      

Once again, the most obvious difference is price. XTT is priced higher than MTT, yet they are both very similar in application. So how do they differ? And what justifies the difference in price point?

XTT holds some significant advantages over MTT. For example, XTT is very sensitive and has a higher dynamic range. Reactions with XTT result in a soluble formazan dye. This means the final solubilizing step is eliminated, which is not the case when using MTT. Removing this step also reduces the risk of error, such as air bubbles from Triton X-100 or SDS. Lastly, OD reading can be immediately taken after incubation when using XTT, while MTT usually requires a longer incubation for the solubilization of precipitate.

Verdict:
Whether you choose XTT or MTT, the job will get done. Choosing between the two depends on your preference. If you want to eliminate steps and reduce the potential for error, then XTT is your guy. But if those advantages are not worth the additional cost, which is very significant, then MTT is a great alternative. Both are very reliable for evaluating cell populations.

3. DTT vs. β-mercaptoethanol vs. TCEP HCl

      vs.          vs.     

If you have worked with β-mercaptoethanol (βME) in the past, then you are too familiar with the stench. Even after you dispose of your gloves, the smell lingers in your nose for some time. Despite its unforgettable scent and toxicity, it is much less expensive than DTT or TCEP. Sometimes saving money is worth the obstacles; however, there are some preferred applications for each of the three.

TCEP is known for its stability and lack of odor. According to some posts on Research Gate, many researchers prefer to use it for storage. By only storing protein stock in TCEP, you use less and incur a lower expense. TCEP is also useful if you’re doing UV detection of protein in buffer. This is because TCEP absorbs less UV than the other two reagents.

During protein purification, βME or DTT are the popular choices. DTT is a strong reducer, 7-fold stronger than βME, and it doesn’t have the odor that comes with βME. On the other hand, βME is far less stable. It evaporates from solution which means its concentration in solution will decrease with time. And to maintain equilibrium, more βME is required; otherwise, proteins won’t be sufficiently reduced, causing electrophoretic bands to be fuzzy.

Ultimately, it’s a matter of preference between the two, however some researchers suggest there being a benefit to using DTT for protein purification and for using βME when purifying smaller molecules.

Verdict:
In the end, this is yet another series where all candidates should have a place in your lab. Though if you must boil it down to two, choose DTT and TCEP. Neither of the two products smell quite as bad, and GoldBio’s prices justify the subtle advantages.

4. DTT vs. DTE

     vs.   

With all of this talk about reducing agents in the previous example, you might now be wondering about DTT vs. DTE (Cleland’s Reagents). This is one case where it’s simply a matter of preference and price. In Cleland’s original article, he stated that there appears to be no significant difference. They are epimers: the hydroxyl groups of DTE are in the cis form, while in DTT, they’re in the trans form.

Verdict:
Either or is fine. Go with the more cost effective product or the product you have the most experience with. At least now you know that if you’re ever in a situation where you must “borrow a cup of sugar” from another lab, either product you’re given will work just fine.

5. Ampicillin (Sodium) vs. Carbenicillin (Disodium)

    vs.     

Ampicillin is widely used for selection during cloning experiments; however it has its drawbacks. Carbenicillin is a more stable substitute, but like the previous examples, advantages always come with a cost. So do the benefits of carbenicillin outweigh that cost? And, are there times where one antibiotic is more advantageous than the other?

During selection, cells containing the bla gene from transformation will show resistance to ampicillin by expressing beta-lactamase, which inactivates ampicillin. The problem is that beta-lactamase is secreted by the bacterial cells, and when enough extracellular accumulation occurs, ampicillin in culture can be inactivated. What this ultimately means is that you can have a lower yield of desired cells in liquid culture, and satellite colonies may appear on agar plates (“These aren’t the cells you’re looking for”). As your plates age, the risk for satellite colonies increases.

Carbenicillin is more stable than ampicillin. It has a higher resistance to heat, it won’t degrade as easily in a lower pH, and it has an increased shelf life. But the other benefit is that satellite colonies are less likely to form since carbenicillin lasts longer and is less susceptible to hydrolysis by beta-lactamase. While all that sounds good, the risk when using carbenicillin is that its potency may kill cells before they have time to manufacture the resistance.

Verdict:
Keep both in the lab, and know when to use one over the other. When you’re dealing with quantification and longer incubation times, it’s safer to use carbenicillin. But when you’re doing a ligation, your cells are slow growing or your DNA is fragile, it’s safer to use ampicillin.

So there you have it, a starter guide for some common reagents that do very similar things. We know this is a list of only five categories. So if you think we need to have a part two, please chime in with your suggestions, questions or even your answers about other products!


    
              Karen Martin
GoldBio Marketing Coordinator


"To understand the universe is to understand math." My 8th grade
math teacher's quote meant nothing to me at the time. Then came
college, and the revelation that the adults in my past were right all
along. But since math feels less tangible, I fell for biology and have
found pure happiness behind my desk at GoldBio, learning, writing
and loving everything science. 



Category Code: 88253 79108 79107

Posted by Chris on June 27th, 2013  ⟩  0 comments

“Before, beside us, and above
        The firefly lights his lamp of love.”
                        by Bishop Reginald Heber

Bioluminescence is one of the premier tools that scientists have in research, whether studying in vitro or in vivo. Few devices allow for the range, versatility, and ease of use as our adaption of the firefly’s twinkling star. But the firefly luminescence was only the beginning, and biologists have found many other species (mostly in shallow, coastal waters) which have developed the ability to light their own way and which we can copy for our own use and benefit.

Firefly luciferase, with its substrate luciferin, is still by far the most popular system for use in bioluminescent imaging (BLI), with Gaussia luciferase, and its substrate coelenterazine, a close second. Over the years, these two systems have been combined in various methods or kits in order to provide a more expansive research device. Most often, one or the other is used as a system control while the other pulls the heavy load. And there have been many attempts to expand the system even further, such as altering the luciferase cDNA and changing its emission spectrum in order to add a third BLI wavelength. But most of this work has been done in vitro, where the “trouble” of dealing with more than one substrate in a system can be a burden. But in vivo researchers are less bothered by such minor complications.

In animal studies, there are other things to worry about. For instance, how well does a new system handle the body temperature of the model? Can the substrate get to the test site, how quickly/slowly, and which route of injection works best? Will the substrate be broken down in the system? Can we visualize the BLI through the thousand-fold layers of cells in the animal model? There have also been many combinations of BLI and fluorescence systems in order to expand the in vivo systems as well, but with many models displaying autofluorescence, the advantages of doing so is somewhat muted by comparison. Into that line of discovery, enter Dr. Casey Maguire and his group from Harvard Medical School/Massachusetts General Hospital.

Maguire et al. wanted to develop a system in which three different luciferase signals could help report cancer cells and their cellular interactions. Firefly and Gaussia luciferases were a given, but they needed another, and decided on Vargula (or Cyprindina) luciferase. This relatively new luciferase was found in Vargula hilgendorfii (previously called Cypridina hilgendorfii), a crustacean sometimes called a sea shrimp or sea-firefly. V-Luc (or sometimes C-Luc) utilizes a substrate called Vargulin to produce a blue colored light around the 450nm wavelength. Using a mouse model, Maguire injected cancer cells intracranially which had been modified with either VLuc, FLuc or GLuc cDNA. Ultimately, they wanted to test their ability to “monitor the effect of an adeno-associated virus (AAV)-mediated soluble tumor necrosis factor-related apoptosis-inducing ligand (sTRAIL) therapy against intracranial glioma tumors.”

The results were outstanding, barring a few caveats which you can read for yourself in the discussion section of their article. There was little to no overlap in the BLI signals between the three substrates and all three were clearly visible, even in deep tissue, like the brain. The use of luciferin, coelenterazine, and now vargulin, as triple BLI reporters makes for the best of a cost-effective, sensitive and easy-to-use reporting system. And at GoldBio, that’s just the way we like it!

Triple Bioluminescence

 
 

Maguire, Casey A., et al. "Triple Bioluminescence Imaging for In Vivo Monitoring of Cellular Processes." Molecular Therapy—Nucleic Acids 2.6 (2013): e99.

Category Code: 88241 88231

Posted by Chris on June 20th, 2013  ⟩  0 comments

If you’re anything like me (geek that I am), every new technological device tends to get your blood pumping and invokes an involuntary reflex to reach for your wallet. That’s even truer for the ever-popular “i”-products which tend to grab our collective-geek attention even faster with every new device. Now there are some clever scientists from the University of Bonn in Germany who have developed a new reporter system utilizing Gaussia luciferase, the “iGLuc”!

While researching the inflammasome process, and specifically IL-1β, a primary target of caspase-1, Bartok et al. hit a frustrating road block. Inflammasomes are large, multiprotein oligomers that are intregal parts of the immune response system. They are a platform which supports an inflammatory cascade after sensing damage-associated molecular patterns. Caspase-1 is an enzyme that’s utilized by the inflammasome cascade in order to proteolytically cleave specific proteins (such as IL-1β precursor) into active, mature peptides. Once cleaved, IL-1β can finally bind to its receptor in order to induce a variety of cellular responses, such as pyroptosis; a form of programmed cell death that is in response to inflammation.

Bartok was looking for a better way to analyze IL-1β. ELISA techniques were not sensitive enough to distinguish between the IL-1β precursors and mature IL-1β, and Western blotting was too time consuming and useless for high throughput analysis. So, instead they devised a fusion protein of pro-IL-1β and GLuc (Gaussia Luciferase) and called it iGLuc! Unexpectedly, they first saw virtually no luciferase signal from the fusion, even though they were seeing high expression levels of luciferase in the system. But they discovered that pro-IL-1β tends to form a protein aggregate which acts to restrict the release of the signaling C-terminal portion of GLuc. But with the simple addition of caspase-1, pro-IL-1β was cleaved and a corresponding bioluminescent signal could be measured.

The resulting process seems to make for an excellent reporter assay for inflammasome activity! Bartok tested the system both in vitro and in vivo and the system showed good sensitivity and specificity as well as a great signal to noise ratio. The system also shows a lot of promise that it can be further applied to other proteases as well! So, if you’re in the field of inflammasomes (or if you have to own every new device), be sure to keep an “i” out for the iGLuc system. It may become the next, best geeky thing on the technological front! You can find their complete article here.

iGLuc in vivo images

Category Code: 88221 88241

Posted by Patrick on January 21st, 2013  ⟩  0 comments

For today’s blog post, we’re going to shine a spotlight on a widely used technique in the field of molecular biology, Blue-White screening.  While this technique has been around for some time, it is still very important to researchers doing genetic engineering because it is a quick and easy way to determine if your gene of interest has been successfully ligated into a host plasmid. In this article, we’ll go over the technology behind Blue-White screening and what’s necessary to make this technique works.

To begin with, we will first need to talk about the β-galactosidase enzyme (or β-gal for short) and some if its properties. β-gal is a naturally occurring enzyme in E. coli cells that’s main function is to cut lactose molecules into glucose and galactose, so the cells can utilize lactose as food source. Typically this enzyme is encoded by a single gene in the bacterial chromosome, the LacZ gene, and in its active form it’s composed of 4 identical sub-units. Later, researchers discovered a mutant form of the LacZ gene in the M15 E. Coli strain that was missing a short string of amino acids from its N-Terminus, and it produced a non-functional β-gal (known as the ω-peptide). However, researchers then found that if they produced the missing fragment (the α-peptide) in another part of the cell, the two fragments would join together and the enzyme functionality would be restored. This technique is known as α-complementation and was published in 1967 by Ullman, Jacob, and Monod in the Journal of Molecular Biology. This may seem like ancient history in the field of molecular biology, but it was a revolutionary new discovery in the way that proteins are made and interact, and we are still utilizing this today.

In order to utilize the α-complementation technique for the purpose of Blue-White screening, you need a few supplies. First you will need an E. coli cell line that contains the mutant LacZ gene for the ω-peptide, typically denoted in the genotype as Δ(lacZ)M15. Next we will need a plasmid that contains the gene for the α-peptide with a Multiple Cloning Site (MCS), and of course your gene of interest. These items are commercially available, and typically both of the genes will be under the control of the lac operon, which means they are inducible with IPTG (Isopropyl-β-D-thiogalactoside, a lactose analog). Next we will need a chromogenic substrate, such as X-Gal (5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside). In a cell line that has a full LacZ gene and produces a functional β-gal, when they are plated on media containing X-Gal the β-gal enzyme will cleave the colorless X-Gal and form an insoluble bright blue precipitate, which will turn the bacterial colony blue. If the β-gal enzyme is not functional the colony will remain white on the plate.

By inserting your gene of interest into the MCS of the host plasmid, you disrupt the coding for the α-peptide. Then once the plasmid is transformed into the bacterial cell, it will not bind to the ω-peptide, and will not form a functional β-gal enzyme. This allows you to quickly screen a transformation plate to determine if your gene of interest was successfully ligated into the host plasmid, because the correct colonies will be white due to the knocked out α-peptide, while the background colonies will be blue due to the functional α-peptide.  While this technique isn’t completely foolproof, it can be an incredibly useful tool for genetic engineering, especially when cloning into a blunt-end cut site, or other techniques that show high levels of background recombination.

For more information on the preparation of Blue-White screening media and the necessary chemical reagents, a protocol can be found on our website, or you can contact us any time at techsupport@goldbio.com.  Thanks for reading!

Be sure to watch our YouTube video on Blue-White Screening here:

1. Ullmann, A.; Jacob, F.; Monod, J. (1967). "Characterization by in vitro complementation of a peptide corresponding to an operator-proximal segment of the beta-galactosidase structural gene of Escherichia coli". Journal of Molecular Biology 24 (2): 339–343.

Category Code: 88221 79105 88231 79108