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.
Buxbaum, 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

This month’s Nature methods (part 2): optogenetics

Optogenetic tools are light-sensitive genetically encoded proteins that, upon light activation, affect a molecular change in the cells. In the previous post I described an optogenetic system to induce transcription. However, the most common use is of channelrhodopsin (ChR) molecules, that alter ion homeostasis upon illumination, and when expressed in neurons, can affect neuronal activity and – in live animals – a change in behavior.

Though optogenetics were used extensively in neurons in culture or in mice, its use was limited in flies. The reason was that the most common ChRs are activated by blue light, which does not penetrate well into the brains of live flies. Therefore, theremogenetic control has been used. In thermogenetics, a thermosensitive channel is expressed in the neurons. However, theremogenetics is far less accurate than optogenetics in terms of temporal resolution and intensity. On top of that, one needs to consider the effect of changes in heat on behavior.

To overcome that, the group of David Anderson decided to use a newly developed red-shifter ChR, ReaChr.

Using ReaChr, but not other Chrs, the researchers were able to activate specific neurons and alter fly behavior. Here’s one of their videos. Using this tool, they learned about the sexual behavior of the male fly and came to the conclusion that there are two sets of neurons which can be separated, with social experience affecting only one set of neurons.

The second optogenetic paper in this issue is more technical. The group of Edward Boyden decided to screen for new candidate ChRs in over 100 species of alga. They sequenced transcriptomes of 127 alga species and isolated 61 candidate ChRs which they tested for light-induced currents using electrophysiology methods. They chose 20 candidates and tested them in a range of wavelength excitations looking at current maxima, kinetics and more parameters.

characteristics of selected channelrhodopsins from different alga species. A, B, C, D – currents amplitude at different excitation wavelengths in HEK293 (A-C) and neurons (D). E – Chrimson is the most red-shifted ChR. F – off kinetics. G – on-kinetics. H – recovery kinetics. Source: Klapoetke et. al. (2014) Nat. Meth. 11:338-346.

Of these, they selected two unique ChR: Chronos (with very high activity after blue or green light excitation) and Chrimson (the most red-shifted ChR – 45nm more red-shifted than ReaChR). After further characterization they showed that Chrimson is a good optogenetic tool for live flies and showed that they can stimulate neurons in fly brains.

Last, they created a two-color system. One of the limitations of the current ChRs is that all of them can be stimulated to some extent by blue light. Although Chrimson can be activate by blue light, the kinetics of Chronos are 10 fold faster. They did a very detailed work in finding the best conditions for a two-color system, based on the excitation pulse and power as well as expression level of the different ChRs.

All in all, a very nice work.

 

 
Klapoetke NC, Murata Y, Kim SS, Pulver SR, Birdsey-Benson A, Cho YK, Morimoto TK, Chuong AS, Carpenter EJ, Tian Z, Wang J, Xie Y, Yan Z, Zhang Y, Chow BY, Surek B, Melkonian M, Jayaraman V, Constantine-Paton M, Wong GK, & Boyden ES (2014). Independent optical excitation of distinct neural populations. Nature methods, 11 (3), 338-46 PMID: 24509633
Inagaki HK, Jung Y, Hoopfer ED, Wong AM, Mishra N, Lin JY, Tsien RY, & Anderson DJ (2014). Optogenetic control of Drosophila using a red-shifted channelrhodopsin reveals experience-dependent influences on courtship. Nature methods, 11 (3), 325-32 PMID: 24363022
 

 

Open source microscope & software: openSPIM & Fiji

Just hears a great talk today by Pavel Tomancak from Max-Planck institute. He’s doing amazing work in systematic imaging of fly RNAome & proteome during development. Check out his website (click his name above).

He also talked about Fiji, which an open source software that is just like ImageJ, but with supporting community that develops new scripts & applications.

A most unusual “open source” development that he is very proud about is the openSPIM.

SPIM – Selective Plane Illumination Microscopy – is a microscopy method in which a laser beam sends a narrow light sheet to the specimen, and the objective is at 90 degree from the light sheet (there are SPIM developments with up to 4 objectives that can image from all 4 sides. See here). This type of microscopy is good for imaging thick live specimens (e.g. whole worm, fly or fish embryo etc..). Due to the thickness of the sample, wide field imaging will cause too much background.

So, there are good SPIM microscopes that you can buy from companies like Zeiss. but he developed the openSPIM, which is a build-it-yourself  SPIM microscope, that actually fits into a suitcase (he took it to South-Africa with him to show college & high-school kids). He claims that non-specialist can construct it in one hour. All the details (parts, assembly instructions (“IKEA/Lego style”) are found at the website. The cost, he estimates, is ~40,000$ (with the camera being about half the cost). He claims that the openSPIM is comparable, in quality of images, to commercial SPIM microscopes from 5 years ago.  Pretty good for standard imaging.

I like the idea.

Nature methods special issue: focus on bioimage informatics (Part III)

Continuing with the Brief communications section:
Rapid, accurate particle tracking by calculation of radial symmetry centers

Tracking single particles is a major challenge, since in many cases the particles are smaller than the pixel size. Several image analysis methods have been developed to analyze subpixel localization of particles. Here, Raghuveer Parthasarathy describes a new approach to calculate subpixel localization of particles, using radial symmetry analysis. Although the accuracy of his algorithm is similar to that of other algorithms (Gaussian fittings such as Non-Linear Least Square minimization – NLLS and Maximum likelihood estimations –MLE), his calculations are ~100 faster than other algorithms.

For those of you who do single particle analysis and super-resolution microscopy, this algorithm may be very helpful.

Rational design of true monomeric and bright photoactivatable fluorescent proteins

As already mentioned in the previous post, photoactivated localization microscopy (PALM) and Stochastic optical reconstruction microscopy (STORM) are two of the techniques used in super-resolution microscopy. In many cases PALM/STORM use photoactivatable fluorescent proteins (PA-FPs). The efficiency of super-resolution microscopy relies on the properties of these PA-FPs, such as brightness, photostability, pH stability, oligomeric state, maturation rate, photoswitching/activation yields etc…

EoSFP, which was cloned in 2004 from the scleractinian coral Lobophyllia hemprichii and furhter engineered, is a green-to-red photoswitchable protein with the highest photon output of all PA-FPs. Upon UV irradiation, it permanently switches its emission peak from 516 to 581 (excitation is at 505).  Monomeric form, mEoSFP and mEos2 were developed (mEoSFP is less used, since its chromofore does not maturate at 37°C, limiting the use to non-mammalian cells).

Here, the authors claim than mEoS2 forms oligomers at high concentrations, which may limit the use of this protein as a fusion partner to the studied protein, and can also skew super-resolution analysis that assumes only monomeric mEoS2 forms. Therefore, the authors solved the crystal structure of mEoS2. Based on the structure, they developed improved, true monomeric variants (mEoS3.1 & mEoS3.2), which are also brighter and mature faster.

Confocal images of HEK293 cells transiently transfected with plasmids encoding indicated fusions and imaged at the middle layer (top) or near the plasma membrane (bottom). See the differnces between mEoS2 and mEoS3! Source: Zhang et al. Nature Methods 9,727–729(2012)

In the supplementary data of this paper you will find a lot of data on the different mEoS variants (not only 2, 3.1 and 3.2 but others as well).

The lesson to be learned here – the properties of the fluorescent protein that you are using to tag your protein of interest may affect the properties of the studied fusion protein/organelle/cell and these factors should always be taken into consideration.

Tracking mitochondria dynamics in live HeLa cells. The large box: Mitochondria in HeLa cells tagged with mEoS3.2-mito prior to photoswitching. Rectangle – area of UV illumination to switch color. boxes on left: time-lapse of mEoS3.2-mito only in the activated region. Source: Zhang et al. Nature Methods 9,727–729(2012)

Multiview light-sheet microscope for rapid in toto imaging

Embryogenesis and morphogenesis are highly dynamic processes that are difficult to image since it involves multicellular samples in the millimeter range. In such cases, it is difficult to image subcellular processes on the one hand, and get a clear 3-D view of the entire sample (which need to be properly rotated). Some techniques that allow sample rotation exist, and an emerging method called selective plane illumination microscopy (SPIM), are helpful in following such processes on whole embryos. However, samples are required to rotate in several angles, often not keeping with the same axis, and the time resolution required for each rotation sometimes exceeds the biological dynamics. Here, research from the lab of Lars Hufnagel developed a new microscopy system, which they term MuVi-SPIM, consisting of four arms (with objectives) that can perform as illuminating or detecting objective. This allows rapid four 3D-view imaging of the sample. Very nifty!

3D image reconstruction of a Drosophila embryo expressing the membrane marker Gap43-mCherry in cycle 14. Alternating green and magenta colors correspond to the image contributions from the eight different views. Inset shows a close-up view of the image fusion on the boundary between two different views. Source: Krzic et al. Nature Methods 9,730–733(2012)

The next parts will review the four articles in this issue.

Parthasarathy R (2012). Rapid, accurate particle tracking by calculation of radial symmetry centers. Nature methods PMID: 22688415
Zhang M, Chang H, Zhang Y, Yu J, Wu L, Ji W, Chen J, Liu B, Lu J, Liu Y, Zhang J, Xu P, & Xu T (2012). Rational design of true monomeric and bright photoactivatable fluorescent proteins. Nature methods, 9 (7), 727-729 PMID: 22581370
Krzic U, Gunther S, Saunders TE, Streichan SJ, & Hufnagel L (2012). Multiview light-sheet microscope for rapid in toto imaging. Nature methods, 9 (7), 730-733 PMID: 22660739