qPCR: quicker and easier but don't be sloppy

(Image credit)

Quick post just to direct your attention to a new article in Nature Methods, under the "Technology Feature" section, discussing qPCR. This is a very powerful technique, but care must be taken when setting it up in order to arrive to the right conclusions.

Go check it out!

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Two interesting papers on ChIP-Seq

Quick post just to direct your attention to two recent interesting papers using ChIP-Seq [See also [Techniques] Analyzing the genome-wide chromatin landscape: ChIP-Seq].

Comparing genome-wide chromatin profiles using ChIP-chip or ChIP-seq. Bioinformatics. 2010 Mar 5. [Epub ahead of print]

Annotation of gene promoters by integrative data-mining of ChIP-seq Pol-II enrichment data. BMC Bioinformatics. 2010 Jan 18;11 Suppl 1:S65.

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Fourth time is the charm: the quest for the final plasmid

In a previous post, I highlighted the wonders of using yeast recombinational cloning (YRC) as an alternative to “classic” cloning, particularly when under a high-throughput approach [See An alternative cloning strategy: yeast recombinational cloning].

Just to refresh your memory, and in a nutshell, the idea is to co-transform the DNA segment to be cloned into yeast along with the linearized target plasmid, provided that this DNA segment bears homology to defined plasmid sequences. By homologous recombination, yeast machinery will directly “ligate” your DNA segment into the linearized vector (See fig.1 in the preceding post).

As I stated in the previous post, my interest was to use YRC to clone promoter sequences upstream of a reporter gene (and I have MANY promoter sequences to clone, so this is a suitable approach), and after discussing the methodology and how it applied to my own research, I ended that post by saying:

(…) we have generated a yeast strain which contains a plasmid with our promoter of interest controlling the reporter gene. Now, it’s time to obtain that plasmid in working concentrations, sequence it and then transform it into our model organism.

How do we do this? How do we get the resulting plasmid out of yeast?

The purpose of this post is to discuss how we do this: how we get our plasmid out of yeast so that we can use it to transform our model organism to study the promoter’s biology in vivo.

The answer? A “Smash & Grab” approach (reflecting that you’d smash yeast cells and get what’s rightfully yours: your plasmid).

When I first got to the lab, my PI gave me a protocol that consisted on picking one of the yeast transformants (a yeast colony that grew on URA- media plate after transformation with the DNA segment of interest and the linearized plasmid), streaking it onto a new selective plate and growing it O/N. The next day, you’d scrap the clone and transfer it to a Eppi containing 300 ul of S&G buffer (which has Triton X-100, SDS, NaCl, Tris pH 8 and EDTA), resuspend, add glass beads and phenol/chloroform and vortex for a minute.
You’d then centrifuge and transfer part of the aqueous solution to a new Eppi containing sodium acetate 3M pH 5.2. Then, after an ethanol precipitation, you would have yeast DNA, both plasmid and chromosomal of origin.
After resuspending the DNA in 50 ul of water or TE, you’d take 3 ul and transform E. coli (remember that the plasmid we are using is a shuttle vector).

Even though this seems pretty easy, the efficiency of such method was disastrous in our hands (we use chemocompetent E. coli). My PI insisted this was routinely used in his previous lab (where he did his post-doc and used overpriced commercial electrocompetents cells) with great results (lots of E. coli colonies), but we just couldn’t get it to work with our cells nor we had those commercial electrocompetent cells available. This drove me to literature in order to find a protocol that would give good results in our lab.

I dug up a paper from 1995 (see ref below¹) describing a protocol which was very similar to the one we were using, although it claimed to yield plasmid DNA without chromosomal contamination, which is perfect.

Briefly, two/three loopfuls of yeast cells (taken directly from patches on minimal media, grown O/N), were resuspended on a solution containing NaCl, Tris-HCl pH 8, EDTA and SDS. Then, glass beads were added, and the mixture was vortexed for 2 mins. An ice cold solution containing NaOH and Triton X-100 was then added, and after mixing by inversion, an aliquot of sodium acetate 3M pH 4.8 was added, and everything was then mixed thoroughly. After a short incubation in ice, you’d add an equal volume of phenol chloroform-isoamyl alcohol (25:24:1, v/v), vortex the mixture and then centrifuge. Another phenol-chloroform extraction of the aqueous solution was next. Then, the aqueous solution was ethanol-precipitated.

This actually worked like a charm the first time in my hands, but my colleagues couldn’t get it to work. Afterwards, and due to the “magic” of life in the lab, I couldn’t get it to work either, despite several tries!
I didn’t put much thought into it and instead, looked for another protocol, considering that this thing only worked once, and only for one person. I went back online and found another somewhat related protocol², which “takes advantage of the finding that phenol/chloroform extraction in the presence of LiCl and Triton X-100 solubilises plasmid DNA, while precipitating cellular proteins and denatured chromosomal DNA”.

No good. Transformation efficiency was still dismal in our hands.

Decided to tackle this problem as soon as possible, and with a little help from Google & Pubmed, I found what is now the standard Smash & Grab protocol in our lab: “A simple and highly efficient procedure for rescuing autonomous plasmids from yeast” (my emphasis), an article published in NAR in 1992³. How did I miss this in my first search? This is exactly what I was trying to achieve!

So, how is this protocol different?

[from the article]

In order to take advantage of the extremely powerful techniques of molecular genetics which are available in the yeast Saccharomyces cerevisiae, it is often necessary to recover shuttle vectors replicating in yeast back into E. coli. However, many workers have found this step troublesome due to what has been described as a 'persistent inhibitor of E. coli transformation' which accompanies yeast DNA preparations. None of the previously published protocols completely eliminates this problem.

(my emphasis)

Notably, one of the referenced articles which “did not eliminate the problem”, is the one I used in my third try! So after just reading the first paragraph of this one-page article, I became excited.

Besides its claimed higher efficiency (which again cited the article I used in my third try, stating that it was 1-2 orders of magnitude more efficient), this method doesn’t require organic solvents (phenol and chloroform), which is always appreciated :)

The idea behind this protocol is simple: let’s simply boil yeast!

In the words of Montell Jordan, “this is how we do it”: We grow an O/N 5 ml yeast culture under selective conditions (i.e in URA- minimal media), harvest the cells by centrifugation, resuspend the pellet in a buffer containing sucrose, Tris pH8, EDTA and Triton X-100, add glass beads and vortex vigorously for 5 mins. We then place the tubes in a boiling water bath for 3 mins. After a brief cool-down on ice, we spin the cells and transfer the supernatant to a fresh tube with 7.5 M ammonium acetate and incubate for an hour at -20C. Interestingly, the authors claim that:

At this stage, residual chromosomal DNA, large RNA species, and the putative impurities which inhibit E.coli transformation are precipitated

Then, we centrifuge again and add an aliquot of the supernatant to ice cold ethanol and incubate again for an hour at -20C. Later, we centrifuge, wash with 70% ethanol and resuspend in water. We then use 10ul to transform E. coli.

This has reproducibly given great results in our lab (not only in my hands this time ;-)), so we are happy to have finally nailed down this part of the protocol.

But let’s remember that the idea is to get the plasmid out of yeast in working concentrations, sequence it and later use it to transform our model organism.

Now that we have the correctly assembled plasmid in bacteria (which we check by PCR), the aforementioned steps are simple: we just grow a colony O/N in selective media, do a miniprep and send an aliquot out for sequencing.

And that’s it!

In just a week, you can have your segment of interest cloned in a particular plasmid and send it out for sequencing, thanks to the power of yeast genetics (and its high levels of homologous recombination) and to a simple, cheap and efficient protocol for getting plasmid DNA out of yeast and into E. coli(*).

We are now ready to transform our model organism to study the promoter in vivo.
What reporter do we use? How do we monitor it? Be sure to check the next post on this series of articles. Stay tuned!

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(*) There are some “yeast miniprep kits” available, but as expected, they are pricey and, in my opinion, not a justifiable expense considering the protocol I just detailed, which is not only cheap, but also easy and amenable for upscaling.


¹Sobanski, M., & Dickinson, J. (1995). A simple method for the direct extraction of plasmid DNA from yeast Biotechnology Techniques, 9 (3), 225-230 DOI: 10.1007/BF00157083

²Ward AC (1990). Single-step purification of shuttle vectors from yeast for high frequency back-transformation into E. coli. Nucleic acids research, 18 (17) PMID: 2205843

³Robzyk K, & Kassir Y (1992). A simple and highly efficient procedure for rescuing autonomous plasmids from yeast. Nucleic acids research, 20 (14) PMID: 1641351

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Method of the year 2009

Nature Methods has selected induced pluripotent stem (iPS) cells as its Method of the Year 2009, a contest we discussed a few months ago [See The Method of the Year].

The fantastic discovery that somatic cells can be reprogrammed to a pluripotent state by the expression of a transcription factor cocktail has had a great impact for basic research, diagnosis and cell therapy. Indeed, these cells have potential uses in in vitro disease modeling, toxicology and pharmacology, and regenerative medicine.

Be sure to check Nature's Special Feature, in which "a series of articles—and the related video —showcase how induced pluripotency is coming into its own in 2009 as a tool for discovery in both basic and disease biology and explore the incredible impact this area promises to have in biological research".

Here's the aforementioned video:

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An alternative cloning strategy: yeast recombinational cloning

As part of my Ph.D thesis, I have to generate a lot of transcriptional fusions (constructs in which a promoter of choice is cloned in front of a reporter gene in order to evaluate transcriptional regulation. Such plasmids can then be transformed into your model organism to study this regulation in vivo).

Traditionally, this involves amplifying the region of interest (in my case a promoter region) by PCR using primers that harbor the recognition site of particular restriction enzymes. Sometimes you can use the PCR product directly in a digestion reaction with the proper enzymes (although it’s recommended that you column purify your PCR product first) in order to clone it into the vector of interest, but many prefer to clone it first into pGEMT. In order to do the latter, you’ll have to do a TA tailing, as high fidelity enzymes (used for cloning) typically produce blunt ends. Once again, it’s recommended that you column purify your PCR product before tailing.

Once tailed, you can clone the PCR product into pGEMT and transform E. coli in order to propagate the plasmid, which can now be sent out for sequencing. After everything checks out, you would have to set up a digestion reaction in order to get the segment out of pGEMT and clone it into the final plasmid. After that digestion, and in order to get this segment into the final plasmid, you would digest the target plasmid with the same enzymes, set up a ligation reaction and then use an aliquot of the reaction to transform E. coli.

Ideally you’ll use two different enzymes for target plasmid linearization, which would avoid plasmid re-circularization, but sometimes things are not that easy. If you can only use one enzyme for linearization (due to plasmid design, enzyme availability in your lab, etc.) then you are going to have to dephosphorylate the linearized plasmid (typically using Calf intestinal phosphatase) after digestion to avoid re-circularization. Some phosphatases can be heat inactivated, but others can’t. In any case, after digesting and dephosphorylating your plasmid, you’ll have to do a phenol:chlorophorm extraction followed by ethanol precipitation to get your dephosphorylated plasmid, which can now be used for ligation with the segment of interest. After setting up this ligation, an aliquot is used for transformation into E. coli.
You can then check your plasmid by colony PCR of the antibiotic-resistant colonies resulting from the transformation.

At this point you’ll realize that generating several constructs following this protocol can take some time. This strategy, then, does not fit my cloning needs.

We’ve taken a different approach to satisfy most of our cloning requirements, which makes use of my favorite organism: Saccharomyces cerevisiae.

The cloning strategy we use routinely in our lab is called Yeast Recombinational Cloning (YRC), a strategy which has been around for some time (Ma et al., 1987) and has been refined over the last ~13 years (Oldenburg et al., 1997; Gibson, 2009).

The concept behind this strategy is simple: if the segment you’d like to clone into a particular plasmid bears homology to defined plasmid sequences, you can directly “ligate” it into the linearized vector by in vivo recombination: yeast machinery will take care of it. This alleviates the need for an in vitro ligation reaction.

Initially, it was shown that a DNA restriction fragment containing appropriate sequence homology could serve as a substrate for such “recombinational repair” (it’s called repair, as you’ll repair the gap in the linearized plasmid). However, an important advance in the use of such recombination-based methods for gene cloning in yeast, involved the use of PCR (rather than restriction assays) to generate the DNA fragment to be used.

As it was shown that the length of sequence homology needed to promote efficient recombination between the segment of interest and the plasmid was small (~20-40 bp), it was quickly realized that these sequences could be included as part of PCR primers used to amplify a segment of interest. This “recombination-mediated PCR targeting” is very efficient and I consider it now one of my favorite techniques.

So, how do you generate your plasmid through YRC? The linearized target plasmid containing a selectable marker (e.g. URA3 ) is co-transformed into yeast with the PCR fragment of interest: this fragment has 20-40 nt of homology at each end to the region of the plasmid at which recombination is to occur. These nucleotides were added to the fragment as part of the primers. You don’t even have to purify the PCR product before transformation; an aliquot taken directly from the PCR reaction tube works fine. By homologous recombination, some of the cut plasmids are recircularized (due to the integration of the segment of interest into the plasmid) and the plasmid can now be propagated in yeast. Recombinants are then selected as Ura+ transformants (See figure 1).

FIGURE 1. Basic outline of yeast recombinational cloning. Figure based on the one at the “Yeast Model Systems Genomics Group” website.

You can then do yeast colony PCR on the Ura+ transformants to check for your plasmid. Note that the linearized plasmid will not lead to Ura+ transformants, as such a plasmid cannot be propagated.

This protocol may take longer (in days) than the traditional approach outlined at the beginning of this post (as you'll have to wait for the transformed yeast to grow in selective media), but it has significantly less steps, uses less restriction enzymes (as you'll only use them to linearize the target plasmid), uses no ligase, does not require any sort of purification and the only methods involved are digestion, PCR and yeast transformation (the latter is very simple, efficient and requires reagents typically found in a molbio lab. We use the Liac/SS Carrier DNA/PEG method with excellent results).

Further, this approach is more amenable to high-throughput cloning than the traditional strategy (See Colot et al., 2006).

In addition, homologous recombination in yeast can be used to build complex constructs from multiple overlapping constituent parts (see figure 2). Generating such constructs through traditional approaches would take considerably longer. Note that the target vector doesn’t necessarily have to be the final vector. You can use YRC to assembly this complex construct and then the DNA insert can be cut out of the yeast vector and placed into any other vector.

FIGURE 2. Multiple overlapping segments can be cloned into a target plasmid (in this case a shuttle plasmid) through homologous recombination in yeast.

OK, enough about the methodology. How is this strategy used for the generation of my transcriptional fusion constructs? The target plasmid we use is linearized just upstream of the coding region of a reporter gene. Due to proper primer design, the PCR amplicon of the promoter of interest bears homology to particular plasmid sequences that allow its integration upstream of this reporter gene through homologous recombination. The particular target plasmid that we use can be propagated both in yeast and bacteria (i.e it’s a shuttle plasmid) and can also be transformed into our model organism, where by homologous recombination, will integrate in a particular locus in the genome so that we can study the promoter’s biology in vivo.

So, in summary, we have generated a yeast strain which contains a plasmid with our promoter of interest controlling the reporter gene. Now, it’s time to obtain that plasmid in working concentrations, sequence it and then transform it into our model organism.

How do we do this? How do we get the resulting plasmid out of yeast? That will be the matter of a follow-up post, so stay tuned!

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Some of the articles discussed in this post:

MA, H., KUNES, S., SCHATZ, P., & BOTSTEIN, D. (1987). Plasmid construction by homologous recombination in yeast Gene, 58 (2-3), 201-216 DOI: 10.1016/0378-1119(87)90376-3

Oldenburg KR, Vo KT, Michaelis S, & Paddon C (1997). Recombination-mediated PCR-directed plasmid construction in vivo in yeast. Nucleic acids research, 25 (2), 451-2 PMID: 9016579

Gibson, D. (2009). Synthesis of DNA fragments in yeast by one-step assembly of overlapping oligonucleotides Nucleic Acids Research, 37 (20), 6984-6990 DOI: 10.1093/nar/gkp687

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The Method of the year

Is that time of the year again. The time to vote for the "Method of the year", a recognition given by Nature Methods since '07. Every once in a while a method is developed with the potential to greatly advance our knowledge in a particular field, allowing us to ask new questions or to refine our answers to long-standing problems in the life sciences. The idea is to "select a methodology with a demonstrated potential to strongly influence the pace and direction of scientific inquiry".

In 07, Nature Methods nominated "next-generation" sequencing for the award (although the name is far from useful, see Post-apres-next generation sequencing), and "super-resolution fluorescence microscopy" was their choice in 08.

You can now participate by nominating and voting for methodological developments to be considered for the recognition at http://www.nature.com/nmeth/votemoy2009 ¹

You may nominate any recent method or class of methods, published anywhere in the scientific literature, that you believe is likely to have a profound impact on future biological research. You are welcome to nominate a method that you yourself developed, but please acknowledge your connection to it. ²

After reading the editorial in Nature Methods I was left with the impression that even though they are hosting an open poll for choosing the Method of the Year, ultimately, they'll have the final saying in deciding, even if it doesn't match the most voted one:

We will take the results of the popular vote into consideration when choosing the Method of the Year 2009, and the votes will also serve as inspiration for picking the Methods to Watch that accompany it.²

Anyway, it's a good chance to log in and nominate your favorite technique!

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¹ You'll need a Nature account to participate.
² Nature Methods 6, 547 (2009)

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New MS protocol at Nature Protocols

I've been away from blogging for a while, because tomorrow I turn in my PhD Thesis Project, and a few weeks after that, I'll be taking my qualification exam: this means that for the last few weeks I've been reading my eyes out and studying, giving me little time to review articles and other items to highlight here at MolBio Research Highlights.

As part of my project (actually as an alternative approach) I've been thinking about using mass spectrometry (MS) to identify if the particular protein I'm studying is regulated through phosphorylation (sorry, can't give more details at the time). This is great as you can sequence phosphopeptides even if they are present at very low levels.
By some coincidence, while thinking about this, Nature Protocols alerted me of their latest FREE featured paper, a new MS protocol for the detection of peptide post-translational modifications.

Here's the link and reference:

Unwin RD, et al. (2009) A sensitive mass spectrometric method for hypothesis-driven detection of peptide post-translational modifications: multiple reaction monitoring-initiated detection and sequencing (MIDAS). Nature Protocols 4,870-877.

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The Plant Cell's struggle with qPCR

Earlier this year, I posted an article from The Plant Cell listing a series of recommendations for qPCR analysis [Eleven golden rules of quantitative RT-PCR].

The Plant Cell (and I can only guess because they are tired of receiving crappy, unreproducible or badly analyzed qPCR results or, even worse, "semiquantitative" RT-PCRs -see below- ) has recently published yet another article along the same line which "provides guidelines for the experimental design and statistical analysis of qRT-PCR data from the statistician's perspective"¹.

From The Plant Cell's Editor in Chief ²:

these are guidelines; any attempt to impose such analysis as standard while we are still struggling to persuade authors of the deficiencies of "semiquantitative" RT-PCR would be a difficult, if not impossible, task.

C'mon people.... nonquantitative semiquantitative RT-PCR? Really? And trying to get it into The Plant Cell (that by the way, has the highest impact factor of primary research journals in plant biology)?

She also commented (although on a previous Editorial)³:

Over the past 2 years, The Plant Cell has taken steps to remove "semiquantitative" RT-PCR from the pages of the journal

Good for The Plant Cell.

Please make the Editors's physical pain (which I assume they get when they see authors drawing quantitative conclusions in PCR analysis, from gels stained with EtBr) stop and follow their not-yet-imposed guidelines.


¹ Rieu I, Powers SJ. (2009) Real-Time Quantitative RT-PCR: Design, Calculations, and Statistics. The Plant Cell 21:1031-1033 (2009)
² Martin C (2009) Guidelines for Quantitative RT-PCR. The Plant Cell 21:1023
³ Martin C (2008) Refining Our Standards. The Plant Cell 20:1727

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[Techniques] Analyzing the genome-wide chromatin landscape: ChIP-Seq

By AMM

New technologies arise every few years that help us address important biological questions from a new angle. An example of this is the DNA microarray. First developed in the 1990s, this important tool has been essential for example, for profiling gene expression in specific cell types.
Nowadays, high-throughput DNA sequencing serves as an example of such important tools. In 2007, Johnson et al.¹ described a technique they called “ChIP-Seq” (from CHromatin ImmunoPrecipitation and SEQuencing) to study the binding sites of a specific transcription factor across the entire human genome using the advance DNA sequencing technology from Solexa/Illumina.

Before explaining what “ChIP-Seq” is, let me tell you a little about the high-throughput technologies used before that to study the binding sites of proteins to DNA.
The most common method of locating these sites in vivo is known as chromatin immunoprecipitation (ChIP). In this technique, cells are treated with a reagent (typically formaldehyde) that crosslinks proteins to DNA. Every protein that is bound to DNA in the moment the reagent is applied will remain bound to DNA. Afterwards, the chromatin is isolated, sheared and incubated with an antibody directed to the protein of interest. This will precipitate the protein and the DNA bound by it (as they are crosslinked). After reverse-crosslinking, the precipitated DNA is analyzed. At first, this technique was restricted to studying if a particular protein was bound to a determined gene (to its promoter, protein coding region, etc), so the analysis of the precipitated DNA fragments was restricted to PCR (with the gene’s-specific primers) to see if the protein indeed allowed the precipitation of that sequence (which would result in an enrichment of the gene in the precipitated DNA vs the control, where no specific antibody is used).
About 6 years ago, a more ‘genomic’ approach was derived from this technique, resulting in the “ChIP-chip” method. The difference with the previously described technique is that after the incubation with the antibody, all the precipitated DNA fragments are used as probes on a DNA microarray (chip). This allows for a high-throughput analysis of the DNA binding sites of a specific protein (limited of course, by the sequences represented on the chip, See "Advantages" below) instead of studying single genes.

In a way to improve this technique, Johnson et al.¹ replaced the “chip” by direct DNA sequencing.
What does this mean? After the precipitation and reverse-crosslinking steps, DNA fragments are sequenced (in that specific paper, using Solexa/ Illumina technology). After sequencing, the reads are mapped to the genome to determine their locations. In this way, the genome-wide DNA binding sites of a particular protein can be assessed [See figure]. The control simply omits the antibody (again, you will be looking for enrichment in your treated sample when compared to the control).
What are the advantages of ChIP-Seq vs ChIP-chip? First, you avoid all complications arising from array hybridization (probes with different optimal temperatures for binding to their complementary strands, probes that hybridize to more than one DNA sequence, interference of hybridization by DNA secondary structure). Second, you are no longer limited to what’s represented on a chip. Tiling arrays may limit such an advantage, though. Nevertheless, ChIP-Seq is cheaper (compared, for example, to whole-genome human tiling arrays)².
Finally, you can apply ChIP-Seq regardless of whether a microarray chip has been developed for a particular species.
This technique, along with the fact that prices of high-throughput DNA sequencing are continuously becoming more accessible, will allow for the identification of the binding sites of not only transcription factors, but chromatin remodeling complexes, structural components, etc, across an entire genome.

[Image is from ref 2.]

¹Johnson DS, Mortazavi A, Myers RM, Wold B (2007) Genome-wide mapping of in vivo protein-DNA interactions. Science 316, 1497.
²Fields S (2007) Molecular biology. Site-seeing by sequencing. Science. 316(5830):1441-2.

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