Tracking membranes by imaging – mCLING and surface glycans

Living cells exhibit many types of membranes which participate in most biological precesses, one way or another. Imaging membranes is usually acheived by two types of reagents: chemical dyes or fluorescent proteins that are targeted to the membrane itself or inside an organelle.

The chemical dyes are usually targeted to an organelle based on a specific chemical property of that organelle.

For example:

Rhodamine 123, tetramethylrosamine, and Mitotracker  are dyes that preferentially target mitochondria, due to its membrane potential. Mitotracker has thiol groups that allow it to bind to matrix proteins, thus making it more resistant to disruption of the membrane potential (e.g. by fixation).

Lysotracker are lypophilic, mildly basic dyes, which accumulate in the acidic lysosomes.

ER-tracker is a BODIPY (boron-dipyrromethene; a group of relatively pH insensitive dyes that are almost all water insoluble) based dyes which are linked to glibenclamide – a sulfonylurease – which binds to ATP sensitive Potassium channels exclusively resident in the ER membrane.

Long chain carbocyanines like DiL, DiO and DiD are lipophylic fluorescent molecules, which are weakly fluorescent in water, but highly fluorescent when incorporetaed into membranes, particularly the plasma membrane.

FM lipophylic styryl dyes bind the plasma membranes in a reversible manner and are also incorporated into internal vesicles.

On the other hand, fluorescent proteins (FP) are targeted to membranes or organelles by fusing them to either whole proteins that localize to a specific organelle, or to short peptides that carry a localization signal. Thus, a nuclear localization signal (NLS) targets the to the nucleus, mitochondrial targeting signal (MTS) to the mitochondria and a palmitoylation signal to the plasma membrane and endocytic vesicle.

There are advantages and disadvantages to each system, relating to ease of use, specificity, photostability etc… I do not want to go into that.

Here, I would like to mention two new methods to image the plasma membrane.

Tracking surface glycans in live cells

Many (most? all?) proteins are modified by linking to glycans – short or long sugar chains. Tracking surface proteins can enable studying of membrane structure and dynamics. Ben Ovryn’s lab developed a combined metabolic and chemical methods to fluorescently label specific surface glycans (i.e. specific glycans linked to surface proteins) using click chemistry.  They then used TIRF microscopy (to allow imaging only the membrane surface without backgound from the cell body) with very short laser pulses (just a few milliseconds). This allowed imaging of single molecule surface glycans without blurring. Bleaching the majority of the fluorophores also reduced the signal to allow single-molecule resolution.

Image of fluorescently lableld N-linked sialic acid - Alexa647 on the surface of Met-1 cells with excitation of 2ms A) or 50ms B).  Source:  Jiang et al. 2015) Angewandte Chemie vol 54:1765.

TIRF image of fluorescently lableld N-linked sialic acid – Alexa647 on the surface of Met-1 cells, with excitation of 2ms (A) or 50ms (B). Source: Jiang et al. (2015) Angewandte Chemie vol 54:1765.

Imaging of these surface glycans, and tracking their trajectories, they were able to calculate diffusion of global surface proteins, discovering areas of sub-diffusion (transient immobilization) as well as highly mobile glycans. This method also allowed studying a particulalry fine membrane structure called membrane nanotube (NT, a.k.a. tunneling nanotubes). These structurs are difficult to image (i know that from personal experience). The surface glycans reveal the outline of the NT and the dynamyc movements of proteins on the surface (as has been suggested for Sonic Hedghog protein).  Here’s a video.

mCLING for imaging plasma membrane and endocytic vesicles

As I mentioned above, there are many dyes to label the plasma membrane. Yet, most probes used for live cell imaging are either lost during fixation of the cells (e.g. for immunostaining) or even transfer to another organelle. Unlike organic dyes, membrane targeted fluorescent proteins remain on the membrane also after fixation (though many may have diminished brightness). However, it requires also requires cloning and transfection of the cells with the FP. This is not always that straight forward, and is more difficult if you wish to study primary cells or even whole organs.

The Group of Silvio Rizzoli from University Medical Center Gottingen developed a new type of membrane probe that is much more stable both in live and fixed cells, so it can also be used for super-resolution. The probe is an octapeptide, composed of one cystein (couples to a fluorophore) and 7 lysines, one of which is bound to a palmitoyl tail. This tail anchors the peptide to the membrane.

Their paper shows very nice images of both live and fixed cells with their plasma membrane and endocytic vesicles labeled with mCLING, much better than FM dyes mentioned above. They could show vesicles involved in ligand-receptor recycling or trafficking, study the composition of synaptic vesicles or membrane recycling.

Comparing mcLING to FM dyes. Revelo NH et al. 2014) JCB 205:591-606.

Comparing mcLING to FM dyes. Source: Revelo NH et al. (2014) JCB 205:591-606.

Looking at vesicles in synapses.

Looking at vesicles in synapses. Red: mCLING; Green: immunofluorescence of synaptic markers. Source: Revelo NH et al. (2014) JCB 205:591-606.

This seemed like a very usefull probe and I wanted to try it myself.

Unfortunately, it did not go well.  I compared mCLING-Atto647N to a membrane-targeted TagRFP-T.

Here’s the TagRFP-T image:

U2OS cell expressing membrane targeted TagRFP-T

U2OS cell expressing membrane targeted TagRFP-T

You can see clearly the membrane, its protrusions, and some vesicles or organelles inside.

Now the mCLING:

U2OS cells labeled with mCLING-Atto647N

Same U2OS cell labeled with mCLING-Atto647N

It looks like a negative image, right? What I think happened here is that it also attached to the glass bottom (which is coated with fibronectin – essential for my purposes) and so it is very bright compared to the cell. The “black” area is where the cell “protected” the glass from mCLING. Taking movies, I did see instances where the dark “shadow” remained were a cell used to be.

The vesicles inside are clearly visible. Most do not overlap with the TagRFP-T, indicating these are differnt vesicles.

Composite image of TagRFP-T red) and mCLING-Atto647N Cyan). The bottom cell does not express the TagRFP-T

Composite image of TagRFP-T (red) and mCLING-Atto647N (cyan). The bottom cell does not express the TagRFP-T

mCLING was clearly much brighter and more photostable than the TagRFP-T: it barely bleached for the entire imaging session, whereas TagRFP-T bleached fast. (Imaging: 14 z sections, 10 sec intervals, 50ms and 25ms exposure TagRFP-T & mCLING).

I did try to label the cells with mCLING  prior to plating them. It was better in a sense that I didn’t get any “shadows”. But by the time I got to image the cells (it takes at least 30min for the cells to attach), all of the dye was already in endocytic vesicles.

Vidoe of U2OS cell expressing membrane tagged TagRFP-T. Note the fast bleaching

Vidoe of U2OS cell expressing membrane tagged TagRFP-T. Note the fast bleaching

Same cell as above, labeled with mcLING-Atto647N. You can see the Airy discs around some of the vesicles. Thats because of out of focus Z-slices in the max projection.

Same cell as above, labeled with mCLING-Atto647N. Note the brightness and stability of the dye. You can see the Airy discs around some of the vesicles. Thats because of out of focus Z-slices in the max projection.

Actually, in this case, the TagRFP-T and mCLING labeled vesicles co-localize. Maybe it’s just a matter of time that the mCLING labels all endocytic vesicles here but not when I label the cells after they attach to the glass.

I asked Prof. Rizzoli concerning these issues. Here’s the reply:

We are aware of the speed of endocytosis – it unfortunately depends on the biological system you are trying to look for. Labeing cells in solution, followed by plating, is difficult, and takes indeed too long. The problem with mCLING on the coverslip comes from having a negatively charged glass surface, to which mCLING binds. I suggest using some blocking medium before applying the mCLING, such as tryptone/peptone. Alternatively, one could use another coverslip coating, such as BSA.

Prof. Rizzoli  also suggested going to confocal instead of widefield imaging. However, as much as I wanted to, all of these suggestions are unsuited for my needs. Too bad.

ResearchBlogging.orgJiang H, English BP, Hazan RB, Wu P, & Ovryn B (2015). Tracking surface glycans on live cancer cells with single-molecule sensitivity. Angewandte Chemie (International ed. in English), 54 (6), 1765-9 PMID: 25515330
Revelo NH, Kamin D, Truckenbrodt S, Wong AB, Reuter-Jessen K, Reisinger E, Moser T, & Rizzoli SO (2014). A new probe for super-resolution imaging of membranes elucidates trafficking pathways. The Journal of cell biology, 205 (4), 591-606 PMID: 24862576

Visualizing translation: insert TRICK pun here

Unlike transcription, it is much harder to image translation at the single molecule level. The reasons are numerous. For starters, transcription sites (TS) are fairly immobile, whereas mRNAs, ribosomes and proteins move freely in the cytoplasm, often very fast. Then there are only a few TS per nucleus, but multiple mRNAs are translating in the cytoplasm. Next, there’s the issue of signal to noise – at the transcription site, the cell often produces multiple RNAs, thus any tagging on the RNA is amplified at the transcription site.  Last, it is fairly easy to detect the transcription product – RNA – at a single-molecule resolution due to multiple tagging on a single molecule (either by FISH or MS2-like systems). However, it is much more difficult to detect a single protein, be it by fluorescent protein tagging, or other ways (e.g. FabLEMs).

The rate of translation is ~5 amino acids  per second, less than 4 minutes to a protein 1000 amino-acids long. This is faster than the folding and maturation rate of most of even the fastest-folding fluorescent proteins. This means that by the time the protein fluoresce, it already left the ribosome. However, attempts were made in the past with some success.

The Singer group has used the FlAsH-ReAsH system, where a tetracysteine motif binds to a biarsenical dye, to detect newly synthesized peptides in fibroblasts, but have abandoned this approach because of the chemical toxicity to cells and the background of the dye. Moreover, This system still does not allow for single molecule resolution.

Fluorescent non-canonical amino acid tagging (FUNCAT) is system developed in Erin Schuman’s lab to visualize protein production in neurons.  Though it is a fun system, it provides information on total protein production, not of a specific protein.  Furthermore, the shortest incubation time with the tag was 10 minutes – fairly long for single molecule resolution. However, there are reports of even a 5 second pulse. I wasn’t convinced.

 I’ve heard about TRICK when I joined the Singer lab three years ago.  Translating RNA imaging by coat protein knock-off (TRICK) is a system developed by Jeff Chao (then a postdoc at the lab, now has his own lab in Basel, Switzerland). The idea is fairly simple – instead of trying to capture the protein being produced, let us examine the “translational state” of the mRNA.

TRICK takes advantage of both the MS2 and the PP7 systems. The MBS (MS2 cap protein binding sequence) tags the 3’UTR, and is therefore a stable tag on the mRNA. However, the PBS (PP7 cap protein binding sequence) was distributed in the coding sequence (usually just upstream to the stop codon). Expressing MCP-RFP and PCP-GFP yields a dualy-labeled mRNA. Once the mRNA is translated, the ribosome traverses the coding sequence and, upon translating the PBS, displaces the PCP-GFP bound to the stem-loops, yielding a red-only mRNA. Thus the loss of the GFP signal indicates that the mRNA was translated.

 The TRICK system. Halstead et al., 2015) Science 347 (6228):  1367-1671

The TRICK system. Halstead et al., (2015) Science 347 (6228): 1367-1671

Indeed, most of the cytoplasmic mRNAs are tagged only by RFP, indicating that they have been translated at least once. Here also lies a disadvantage of this system – it only allows detection of the first round of translation, because once displaced, the PCP-GFP (which has a nuclear localization signal, NLS) quickly diffuses away from the mRNA and, presumably is imported back to the nucleus.  [The NLS is required, in order to reduce cytoplasmic background].

To prove the point, Jeff shows that addition of translation inhibitors results in mostly dual-colored mRNAs.

Translation in the nucleus has been the subject of much debate (see also the paper I linked to above, re: FUNCAT in 5 seconds). The TRICK system allowed examining this issue, at least for a specific mRNA. Indeed, TRICK analysis did not suggest that any significant translation occurs in the nucleus, at least not of that specific mRNA and at least not the entire length of the coding sequence.

TRICK was further used to examine the distance by which mRNAs are being translated for the first time after nuclear export (most are translated fairly soon after export, though some mRNA molecules are being translated minutes after their export) as well as the translational state of the mRNAs upon stress and assembly into P-bodies.

Many mRNAs are being translated only at a specific location or at a specific time-point. For example, upon budding, the ASH1 mRNA in yeast is translationally repressed during its transport to the bud tip. OSKAR mRNA in translated only when it reaches the pole plasm area of a stage 10 oocyte. TRICK is a great system to detect where and when such mRNAs are first translated. So, in collaboration with Anne Ephrussi, they showed indeed that OSKAR-TRICK mRNA is not translated until the exact time in development.

 OSKAR-TRICK is being tranlated only at a certain time and location during fly oogenesis. Halstead et al.,  2015) Science 347 (6228):  1367-1671

OSKAR-TRICK is being tranlated only at a certain time and location during fly oogenesis. Halstead et al., (2015) Science 347 (6228): 1367-1671

The first round of translation is also the cell’s quality control mechanism, in particular for nonsense-mediated decay (NMD). Since OSKAR-TRICK did not go through translation until a late stage in oogenesis, it implies that only at that developmental stage OSKAR mRNA goes through NMD.  The greater implication is that though for some mRNAs NMD may occur immediately upon export, translationally-repressed/ transported mRNAs may experience quality control only upon arrival at their destination or temporal activation.

TRICK can be used to ask many other questions, such as questions related to NMD and mRNA decay in general; questions about “non-coding RNAs”; Questions about IRES and other forms of alternative translation and many more.

I’m happy for Jeff that its finally published.
ResearchBlogging.orgHalstead JM, Lionnet T, Wilbertz JH, Wippich F, Ephrussi A, Singer RH, & Chao JA (2015). Translation. An RNA biosensor for imaging the first round of translation from single cells to living animals. Science (New York, N.Y.), 347 (6228), 1367-671 PMID: 25792328
Yu, J. (2006). Probing Gene Expression in Live Cells, One Protein Molecule at a Time Science, 311 (5767), 1600-1603 DOI: 10.1126/science.1119623
Dieterich DC, Hodas JJ, Gouzer G, Shadrin IY, Ngo JT, Triller A, Tirrell DA, & Schuman EM (2010). In situ visualization and dynamics of newly synthesized proteins in rat hippocampal neurons. Nature neuroscience, 13 (7), 897-905 PMID: 20543841
Rodriguez, A., Shenoy, S., Singer, R., & Condeelis, J. (2006). Visualization of mRNA translation in living cells The Journal of Cell Biology, 175 (1), 67-76 DOI: 10.1083/jcb.200512137
Baboo S, Bhushan B, Jiang H, Grovenor CR, Pierre P, Davis BG, & Cook PR (2014). Most human proteins made in both nucleus and cytoplasm turn over within minutes. PloS one, 9 (6) PMID: 24911415

Transcription caught on camera part 2: Fab-ulous Histones

In eukaryotes, the DNA is packages tightly in nucleosomes, which are composed primarily out of histone proteins. There are four major types of histones (1,2,3 & 4). Extensive work has been done on how histones facilitate and regulate transcription. It turns out that there are multiple post-translational modifications on histones, such as methylation and acetylation that are linked to transcription regulation. The majority of the studies use a method called chromatin immunoprecipitation (ChIP) to study these modifications. In essence, an antibody specific for a certain modification is used to affinity-purify only modified histones, along with any DNA region they are associated with. Thus, one could get a map of the specific modified histone along the chromosomes and correlate this locations with transcription activity, ChIP maps of other transcription related proteins etc…

There are two problems with this approach. The first, since the cells are fixed, the time resolution is limited to several minutes, at best. Second, the results are an average of the entire cell population, and therefore factors considered linked may not actually be present in the same cell, same genomic location at the same time.

So, Timothy Stasevich et al. tried a different approach by using a novel method to image histone modifications in live cells. This method is called Fab-based live endogenous modification labelling (FabLEM). Basically, these are the antigen binding fragments (Fab) of a specific monoclonal antibody, conjugated to a fluorescent dye. The Fabs are small enough to enter the cell and the nucleus.

By using this system, they reached a temporal resolution of about 10 seconds!

They used a cell line that harbours a 200-copy gene array (i.e. an array of 200 copies of the same gene) whose promoter is controlled by a GFP-labeled glucocorticoid receptor (GR).  Why this cell line? This gene array manufactures a very concentrated fluorescent signal, composed of 200 molecules of whatever dye associates with it, in a very small volume. This allows for a good signal to noise ratio compared to the background of the free or chromatin associated fluorescent-labeled molecules.

For the first experiment, they looked at acetylation on lysine position 27 of histone H3 (H3K27ac), in regard to GR-GFP accumulation (transcription initiation) and the state of RNA polymerase II (RNAP2). The C-terminal domain (CTD) of RNAP2 can be unphosphorylated, or phosphorylated on serine 5 (marking transcription initiation) or serine 2 (marking transcription elongation) of the ~50 heptad repeats that it contains.

a, Illustration of the FabLEM strategy. b, Examples from a FabLEM experiment showing that the gene array (yellow arrow) is acetylated at H3K27  before activation by GFP–GR and accumulation of elongating RNAP2 (Ser 2ph). Scale bar, 5 μm. Source: Stasevich et al (2014) Nature 516:272.

a, Illustration of the FabLEM strategy. b, Examples from a FabLEM experiment showing that the gene array (yellow arrow) is acetylated at H3K27 before activation by GFP–GR and accumulation of elongating RNAP2 (Ser 2ph). Scale bar, 5 μm. Source: Stasevich et al (2014) Nature 516:272.

They initially found that the array is hyper-acetylated before induction, and the level of H3K27ac drops after induction. Here, also, comes the advantage of single-cell experiments. They found variability between cells before induction by the hormone – some cells with high H3K27ac and some with low level. So they checked whether these two populations differ in their transcription kinetics. They found no effect on Ser5 phosphorylation, but significant higher levels of GR-GFP recruitment and Ser2 phosphorylation in the high H3K27ac group. This indicated that H3K27ac levels can affect transcription factor recruitment as well as transcription elongation.

To examine if there is a causal connection between K3K27 acetylation and transcription elongation, the researchers perturbed the acetylation (2 experiments by 2 different approaches) and again found that though initiation (Ser5-P) was unaffected, Ser2-P in cells with reduced H3K27ac was also diminished. Consistent with this notion, they found that the protein AFF4, a key component of the elongation complex, which was shown to interact with the H3K27 acetylase p300, is recruited less efficiently to arrays with low H3K27ac.

To conclude: specific protein modifications has been difficult to image in live cells due to the technical issue of “how to specifically fluorescently tag them”. The use of FabLEMs can solve this problem. From discussing FabLEMs with Tim in the past, I understand that the Fabs are constantly binding on and off every few seconds, so that there is a (small) time window for the endogenous regulating protein which is supposed to bind the antigen to actually bind. But, we should take into account that the FabLEMs probably do perturb the biological function of any specific single protein they bind to. But, in these experiments, with an array of 200 genes, and hundreds or more of H3 proteins on each gene, this effect is limited. Similarly, the CTD of each pol II contains ~50 heptad repeats. though it is not clear how many repeats are phosphorylated when phosphorylation occurs, it is assumed that most, if not all, are phosphorylated. So in this case too its a game of numbers of the FabLEMS vs the regulatory proteins which bind these sites.

Still, these are issues that we need to consider when designing such experiments.



ResearchBlogging.orgStasevich TJ, Hayashi-Takanaka Y, Sato Y, Maehara K, Ohkawa Y, Sakata-Sogawa K, Tokunaga M, Nagase T, Nozaki N, McNally JG, & Kimura H (2014). Regulation of RNA polymerase II activation by histone acetylation in single living cells. Nature, 516 (7530), 272-5 PMID: 25252976

In the right place at the right time: visualizing and understanding mRNA localization

The title of this post is also the title of a review paper that I co-authored  with Adina Buxbaum, a recently graduated PhD student from Rob Singer’s lab. The review was published last week in Nature Reviews Molecular Cell biology.

In this paper we review some of the old and new methods to visualize mRNA. These include mostly FISH and MS2-like systems, which I’ve discussed extensively in this blog. There is also a short section (“box”) on quantitative analysis tools for mRNA localization imaging.

We then discuss the current knowledge on the mechanisms of mRNA localization and how it relates to the biology in two very distinct model systems – unicellular organisms (budding yeast) and the extremely polarized neuronal cell.  We also discuss examples in other organisms from bacteria through fly to frog and mammals.

I’m biased, of course, but I think this turned out to be a balanced, comprehensive, yet not too detailed review paper that will benefit both beginners which are unfamiliar with the RNA localization field, as well as experts which are used to a single method or a single model organism.
ResearchBlogging.orgBuxbaum, A., Haimovich, G., & Singer, R. (2014). In the right place at the right time: visualizing and understanding mRNA localization Nature Reviews Molecular Cell Biology DOI: 10.1038/nrm3918

Transcription caught on camera part 1: Halo transcription factors

Transcription factors (TFs) have a fundamental role is regulating gene expression. The basic model, based on numerous biochemical analyses, has determined where TFs bind (usually at specific sites at or near promoters), when they bind the DNA (at a resolution of minutes/hours) and what do they do there (induce/repress transcription. Duh!).  However, much is yet unknown. One aspect that is fairly unknown is the dynamics of how TFs search for their binding sites, bind them and later dissociate, particularly at the single molecule level. To explore this, the Transcription Imaging Consortium (TIC) at Janelia Research Center (JRC) (it used to be Janelia Farm, but the  “farm” part was removed from the name. oh well) applied sophisticated imaging techniques to measure the dynamics of two TFs, SOX2 and OCT4 in the nuclei of live embryonic stem (ES)cells. Their results were published in Cell almost a year ago.

First, how do you detect a single molecule? I’ve discussed many times in this blog how we can visualize single mRNA molecules by using MS2-like systems. These systems allow multiple fluorescent proteins (FPs) on a single mRNA molecule, thus enhancing signal to noise ratio (SNR). However, it’s not advisable to attach multiple FPs to a single protein – it will probably make this protein inactive, if not worse.

So, they had no choice but to fuse the Sox2 and Oct4 to a single fluorescent tag. Out of the endless choices available these days, the TFs were fused to a HaloTag ( developed by Promega). HaloTag is a system which consists of HaloTag protein (a mutated form of a bacterial haloalkane dehalogenase, DhaA) and a modified ligand molecule. This ligand consists of a reactive linker and a functional group (such as biotin or a fluorescent dye). Once the ligand binds the protein, it forms a covalent bond. HaloTag can be fused to the protein of choice, just like any FP, except it is “dark” unless the ligand is applied. Furthermore, the ligand binds only existing proteins. This means that after the ligand is washed away, any nascent protein that is translated will be “dark”.   The fluorescent dyes are, supposedly, brighter than the common FPs.

Still, there should be hundreds to thousands of Sox2-Halo proteins in the nucleus; how can we distinguish single molecules? -With a nice trick. Since these are TFs, they are supposed to bind DNA and, therefore, stay immobile for at least a short amount of time. A 10 millisecond (ms) exposure time was too short – too many molecules, even just Halo protein alone, were visible as diffraction limited spots. However, with a 500 ms exposure time, fast moving molecules only look like a blur, which greatly reduced the background. Thus, only immobile proteins did not appear as a blur. Furthermore,  only a 2D cross-section of the nucleus was visualized, thus any molecules above or below the plane of imaging also blended into the background. Last, they used a low excitation power, limiting the number of emitting molecules. Using this approach, they found a two-component residence time (i.e. “immobile” molecules) of TFs on the DNA. Most molecules had a residence time of ~0.8 seconds, but some had an average residence time of ~12 seconds. The interpretation is that Sox2 “samples” the DNA in search of the specific binding sequence. This sampling takes, on average, 0.8s. Once bound, the protein stays on the DNA for 12 seconds. A control experiment with Sox2 without the DNA binding domain showed only the short residence time.

Imaging transcription factor dynamics . A) approch taken for imaging. B) images taken of Sox2-Halo, Sox2-TAD-Halo no DNA binding domain) of Halo-NLS. C,D) distribution function of residence time. Adaped from Chen J et al Cell 2014.

Imaging transcription factor dynamics . A) approch taken for imaging. B) images taken of Sox2-Halo, Sox2-TAD-Halo no DNA binding domain) of Halo-NLS. C,D) distribution function of residence time. Adapted from Chen J et al Cell 2014.

This data, from in vivo experiments, was then verified  in a “clean” in vitro experimental set-up of single molecule binding. In this set-up, fluorescently-labeled DNA molecules are attached to a glass slide. The protein in question is then applied on the DNA and the residence time of the Halo-tagged protein on the DNA is measured by total internal reflection fluorescence (TIRF)  microscopy. [TIRF allows imaging only to ~100nm hight above the glass surface, thus imaging only bound molecules and greatly reducing background. TIRF is also very useful in imaging the under surface of adherent cells]. This assay allowed examining different DNA lengths or mutated binding sites. The numbers obtained by this method were very consistent with the in vivo results.

One of the cooler stuff described in this paper is the multi-focal microscope. This microscope has the capacity to simultaneously take images in 9 focal planes, thus allowing really fast single molecule 3D imaging.

Diagram of the basic principles of the aberration corrected multifocal microscope. Image from the JRC site.

[I heard a talk about the construction of this microscope. Wasn’t easy… The grating had to be lithographed very precisely, at the 100 micron accuracy, i think. This was a one-color microscope, but I know they are developing a two color microscope. because of the difference in wave length of green and red, the grating for the green is not suitable for the red. Anyway, very cool microscope]

Using this method, they developed a kinetic model for Sox2 diffusion, search and binding to the DNA. Based on their results, their model suggests that Sox2 “samples” the DNA on average for 84 times before finding a target binding site. The search duration is ~377 seconds. This means it take a Sox2 transcription factor molecule ~6 minutes to find its target, but it only remains bound for several seconds , presumably to stimulate transcription.

Biochemical results suggest there are ~7000 binding sites for Sox2. They used FCS measurements to calculate how many Sox2 molecules there are in an ES cell nucleus. Their calculations suggest 115,000 molecules!  Thus, on average, every Sox2 binding site is sampled every 24 second by a Sox2 molecule, meaning that the TF binding sites are occupied at least half the time.

In summary – this paper used several imaging techniques to get new insights on the behavior of TFs in live cells. Their results are really interesting and shed light on the dynamics of TF behavior, on the effect of TF expression level and diffusion rates and other aspects I did not mention here.
ResearchBlogging.orgChen J, Zhang Z, Li L, Chen BC, Revyakin A, Hajj B, Legant W, Dahan M, Lionnet T, Betzig E, Tjian R, & Liu Z (2014). Single-molecule dynamics of enhanceosome assembly in embryonic stem cells. Cell, 156 (6), 1274-85 PMID: 24630727

SmartFlare live RNA detection

I recently came across a booklet regarding SmartFlare for live cell RNA level detection. The basic idea, if i understand it correctly, is the use of gold nanoparticles that are conjugated to fluorescently tagged short double stranded probes. The “capture” strand is bound to the gold particle, is non-fluorescent and complementary to the target mRNA. The “reporter” strand is complementary to the capture strand but shorter. It is fluorescently labeled, but the gold is quenching the fluorescence.

This particle can enter the cells and if the target RNA is present, it replaces the “reporter” strand which is set free to the cytoplasm and fluoresce. This fluorescent can then be detected by microscopy or any other fluorescent detection.

SmartFlare concept. Source: Merck Millipore website

I haven’t used it yet because it clearly isn’t suited for single-molecule analysis. Furthermore, is does not give any data on RNA localization since the fluorescence appears only after the target RNA is bound and the “reporter” is a short oligo which probably diffuses very fast.

Furthermore, I’m concerned about the biological effects. Although they claim in their FAQ section that at the concentrations used it does not cause RNA knockdown and is non-toxic, I am concerned first about the short reported oligo which presumably will hybridize with antisense to the target RNA, if present and second, that capturing the target RNA will affect its gene expression (probably prevent its translation, maybe mis-localize it and can promote its decay thus actually affect its transcription rate as well).

Particularly I’m concerned due to the long incubation time they suggest in their protocol – 18 hours. Why so long? Shouldn’t it work at shorter time-scales?

This system can be a good reporter for relative expression levels for whole cell populations or single-cell analysis, but I’m hesitant about the use of these same cells for downstream assays.

Also, their illustrations always show complementary of the capture strand to the 5′ end of the target RNA. Does it have to be the 5′ end?

I am considering the use of this system as a reporter for my experimental purposes. Any thoughts? Did anyone use that and can share their experience?

Nobel prize in chemistry 2014

The Nobel prize in chemistry 2014 was awarded to Eric Betzig, Stefan W. Hell & William E. Moerner for the development of Super resolution fluorescent microscopy!

See here for details on the history of this discovery.

As readers of this blog know, Super-resolution microscopy has made a revolution in the field of fluorescent microscopy in a very short time.

This is a justified award.