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.

Eliminating mutated mitochondria during in-vitro fertilization

There are several genetic diseases which originate not from mutations in the nuclear genome but mutations in the mitochondrial genome. In humans, the threshold for disease occurrence is if 60% of the mitochondria has mutated mitochondrial DNA (mtDNA) (a mixed mitochondrial origin is called heteroplasmy). There is currently no cure, and no good way to prevent these genetic diseases.

Since the source of the mitochondria is solely from the oocyte, a lot of effort is invested in trying to get rid of mutated mitochondria by in-vitro fertilization (IVF) procedures – thus preventing the disease in the offspring. Two methods have been tried so far – pronuclear transfer (PNT) and spindle-chromosome transfer (ST).  The idea is to extract the nuclear genetic material of the oocyte or zygote and transfer it to an enucleated oocyte or zygote with healthy mitochondria. However, in both cases, there is still carry-over of some mutated mitochondria to levels that can be as high as 44%.

In this new paper published in Cell, a group of researchers from China suggest a different approach. During the oocyte, and later zygote development, a unique germ cell that is extruded. These are called polar body 1 (PB1), a diploid germ cell, which is extruded from the oocyte before ovulation and PB2, a haploid  germ cell, which is extruded from the zygote after fertilization. The authors reasoned that the nuclear genetic material in PB1 and PB2 is identical to the spindle chromosome and female pronuclear, respectively.  But, PB1 and PB2 are very small cells and therefore may contain very few mitochondria.

Levels of mitochondria in the different nuclei. Red - MitoTracker. Blue: Hoecht (stains DNA).

Levels of mitochondria in the different nuclei. Red – MitoTracker. Blue: Hoecht (stains DNA). Source: Wang T. et. al. (2014) Cell 157:1591-1604.

Indeed, staining with MitoTracker (a dye specific for mitochondria) shows only a small amount of mitochondria in PB1 and PB2.  So, after several tested to see if the PB1 and PB2 has characteristics similar to the corresponding oocyte/zygote nuclei (using immunofluorescence against specific nuclear and epigenetic markers) they compared the four different methods (PNT, ST and PB1 or PB2 transfer) to prevent mitochondrial transfer in IVF.

IF staining with antibody against Lamin B1 (green) shows the nuclear envelope of the different nuclie at distinct stages of zygote development. Red- propidium iodide (PI) stains DNA. Source: Wang T. et. al. (2014) Cell 157:1591-1604

IF staining with antibody against Lamin B1 (green) shows the nuclear envelope of the different nuclie at distinct stages of zygote development. Red- propidium iodide (PI) stains DNA. Source: Wang T. et. al. (2014) Cell 157:1591-1604

Their results are pretty amazing.

All methods have similar success rates of the IVF procedure and give similar number of offspring, with normal development. However, PNT still maintains high levels of heteroplasmy (20-60% in 2nd generation (F2) mice). ST maintains moderate-low heteroplasmy (5-20% in F2). PB2T has even lower heteroplasmy rates (<5% in F2). But PB1T has no heteroplasmy (at least, below the detection level).

This work, done in mice, is a real breakthrough , because if this also works in humans then we can have genetic screens for heteroplasmy and women with high levels of heteroplasmy (or already suffering from an inherited mitochondrial disease) could use PB1T-IVF to give birth to healthy children.

Pretty awesome.
ResearchBlogging.orgWang, T., Sha, H., Ji, D., Zhang, H., Chen, D., Cao, Y., & Zhu, J. (2014). Polar Body Genome Transfer for Preventing the Transmission of Inherited Mitochondrial Diseases Cell, 157 (7), 1591-1604 DOI: 10.1016/j.cell.2014.04.042

An excelent new tool for comparing fluorescent protein properties

Screenshot of the new tool

Screenshot of the new tool

Two very useful tools for visualizations of many fluorescent and photoswitchable proteins have been developed by Talley Lambert and Kurt Thorn (UCSF).

With these tools you can compare the properties of the different FPs by changing the X & Y axes parameters & range on the left. Play with it. Its fun!

If you want exact numbers, there are extensive tables and references below the graph.

Thanks to Optical Nanoscopy Blog for pointing this out.