Tag Archives: neurons

Imaging translation of single mRNAs in live cells

Translating the information encoded in mRNAs into proteins is one of the most basic processes in biology. The mechanism requires a machinery (i.e. ribosomes) and components (mRNA template, charged tRNAs, regulatory factors, energy) that are shared by all organisms on Earth. We’ve learned a great deal about translation over the last century. We know how it works, how it is being regulated at many levels and under varuious conditions. We know the structures of the components. We have drugs that can inhibit translation. With the emergance of next-gen sequencing, we can now perform ribosome profiling and determine exatly which mRNAs are being translated, how many ribosomes occupay each mRNA species and where these ribosomes “sit” on the mRNA, on average. New biochemical approaches like SILAC and PUNCH-P can quantifiy newly synthesized proteins & peptides. Yet, all of that information comes from population studies, typically whole cell populations. Rarely, whole transcriptome/ribosome analysis of a single cell is performed. Still, there is no dynamic information of translation, since cells are fixed and/or lysed. And there is no spatial information regarding where in the cell translation occurs (poor spatial information can be determined if cell fractionation is performed, which is never a perfect separation of organelles/regions and we are still not at the stage of single organelle sequencing).

Imaging translation in single cells is intended to provide both spatial and dynamic information on translation at the single cell and, hopefully, single mRNA molecule resolution. Recently, four papers were published (on the same day!) providing information on translation of single mRNAs. Here is a summary of these papers.

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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.

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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

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.

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.

 

 
ResearchBlogging.orgKlapoetke 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
 

 

Looking at single mRNAs in neurons hints at memory formation

It is postulated that learning and memory are modulated by synaptic plasticity – molecular changes  that result in changes in the synapse morphology and signaling capacity. Local protein translation is considered important for synaptic plasticity. Two works from our lab were published last month (back to back!) in Science. Both papers deal with how beta-actin mRNA localization and dynamics in neurons may account for local protein translation upon stimulation, and hence, may supply insight into memory formation.

The first paper by Adina Buxbaum shows that beta-actin mRNAs in dendrites are “unmasked” upon activation of the dendrites. Using single molecule FISH, She noticed that the average number of probes bound to the mRNAs in dendrites (but not in adjacent glia cells) was lower than expects, and this number increased upon stimulation. Not only that, there were more mRNAs in the stimulated dendrites. This indicated masking by a protein “coating” that prevented FISH probe binding in the unstimulated cells. A modified FISH protocol which included a protease digestion step prior to probe hybridization showed that indeed the mRNAs were masked by proteins.

single molecule FISH for beta-actin mRNA in dendrites shows that mRNAs in unstimulated neurons are masked. A) Unstimulated neuron. B) stimulated neuron showing increased number of spots. C) Unstimulated neuron, in which the fixed cells were digested with protease prior to FISH probe hybridization. Source: Buxbaum, Wu & Singer (2014). Science Vol. 343  pp. 419-422

single molecule FISH for beta-actin mRNA in dendrites shows that mRNAs in unstimulated neurons are masked. A) Unstimulated neuron. B) stimulated neuron showing increased number of spots. C) Unstimulated neuron, in which the fixed cells were digested with protease prior to FISH probe hybridization. Source: Buxbaum, Wu & Singer (2014). Science Vol. 343 pp. 419-422

 

She further showed that this masking relates to other mRNAs, as well as to ribosomes, and that this is due to a metabolic process resulting from stimulation. Thus, this unmasking process may be a way to “activate” localized mRNAs for translation.

Apart from being a very neat paper technically and biologically, I think it was exceptionally entertaining to begin her paper by quoting an 1894 work by Cajal, the father of neuroscience.

The second paper by Hye-Yoon Park follows the dynamics of single molecule endogenous beta-actin mRNAs in neurons by live imaging, using the MS2 system. She shows movement of mRNAs along dendrites, as well as some events of merging or splitting – suggesting that some mRNAs are packed together in larger granules – which may regulate local translation. She also looked at brain slices, visualizing beta actin transcription dynamics. This is an important achievement since it is much harder to look at mRNA dynamics in tissue slices than in single cells on plate, due to background fluorescence. Though some biological insight is derived here, this is more of a “new technology” report.

Live imaging of beta-actin mRNAs in dendrites (movie. Source: Park HY et al. (2014) Science Vol. 343 pp. 422-424)

These papers are just the beginning of a long-term story of how mRNA localization and local translation are regulated in neurons.  A lot of cool experiments are being done in our lab in this regard and I’ll report more as they are published.

ResearchBlogging.orgBuxbaum AR, Wu B, & Singer RH (2014). Single β-actin mRNA detection in neurons reveals a mechanism for regulating its translatability. Science (New York, N.Y.), 343 (6169), 419-22 PMID: 24458642
Park HY, Lim H, Yoon YJ, Follenzi A, Nwokafor C, Lopez-Jones M, Meng X, & Singer RH (2014). Visualization of dynamics of single endogenous mRNA labeled in live mouse. Science (New York, N.Y.), 343 (6169), 422-4 PMID: 24458643

Watching Neurons in action

Fluorescent sensors are important tools that can allow real-time, live, single molecule imaging of microscopic millisecond scale events. It is even better if these sensors are genetically encoded sensors (i.e. fluorescent proteins). We have already encountered the pH sensors pHluorin and pHTomato and the Ca2+ sensor GCaMP. There have been a few others, such as HyPer that detects H2O2 or ArcLight and ElectrikPk which are voltage sensors.

Now, the group of Loren Looger from HHMI Janelia  developed a sensor for a very important molecule: L-glutamate. Continue reading

Sensing pH in neurons

A recent paper in Nature Neuroscience demonstrates the usefulness of the pH sensitivity of fluorescent proteins.

I have briefly mentioned the importance of pH when I discussed mKeima. Here I will describe the work from Richard Tsien’s lab which utilizes the effect of pH on FP in order to study synaptic activity.

One avenue of communication between synapse is vesicle exocytosis, i.e. the release of neurotransmitter-containing vesicles from the pre-synaptic cell. A surge of calcium ions in the postsynaptic cell follows vesicle exocytosis. Interestingly, when a vesicle is exocytosed, the lumen of the vesicle changes its pH from 5.5 to 7.4.

A pH sensitive GFP-based protein called pHluorin was developed as early as 1998. pHluorin was fused to a vesicle protein called VAMP2. Thus, a pH change from 5.5 to 7.5 increase the emission intensity (i.e.the protein becomes brighter when excited).

Although this is a good tool, the fact that it is GFP based limits the use of dual-color microscopy, particularly with other types of fluorescent sensors since most of them are GFP, CFP or YFP based, which makes spectral distinction difficult. Therefore, the authors set out to develop a RFP based pH sensor. After several rounds of mutagenesis and shuffling of mRFP and mStrawberry, they identified a bright pH sensor which they named pHTomato since its ex/em peaks (550nm/580nm) are similar to those of dTomato. Importantly, increasing the pH has increase the brightness of the protein, without affecting the ex/em peaks (unlike mKeima which I mentioned above). Furthermore, fusion to VAMP2 did not change its characteristics; the pH dependence is reversible (i.e. once pH is acidic again, the intensity decreases).

Since VAMP2 gives them a high background expression on the membrane, the authors decided to fuse pHTomato to a protein called synaptophysin (sy), which is more vesicle specific. sypHTomato was then compared to sypHluorin in the same neuronal cell. The authors show (by graph) that both proteins perform similarly. However, it would be nice to also show an image.

Once the authors establish that pHTomato is a good pH-sensor, they turn to the real exciting work of dual color sensing. As mentioned above, following exocytosis, there is a Ca2+ increase at the post-synapse. Electrical stimuli also cause a Ca2+ surge in the presynaptic cell. The authors then used a Ca2+ sensor called GCaMP3. Briefly, GCaMP3 is a GFP-based protein that was modified such that that the two part of the protein are fused to Calmodulin (CaM) and a peptide called M13. In the presence of Ca2+, CaM binds M13, thus bringing the two parts of the GFP together, which results in fluorescence (ex/em 489/509 – similar to EGFP).

Expressing both sensors in the same neurons allowed them to visualize the spike in Ca2+ (green spike) concurrent with an increase in red signal, that deteriorates slowly, upon stimuli to the neurons. I am not a neuroscientist, so I cannot evaluate their system regards to choice of cells/stimuli, but the fluorescence response following the different stimuli they give seems distinct and impressive. However, it is very difficult to see it in the snapshot images of the cells.

Expressing each sensor in a different cell allowed the authors to distinctly visualized pre and post synaptic cells at the same synapse. What they looked at are pre synaptic sypHTomato-positives cells, in contact with post-synaptic GCaMP3 positive cells. I think that the image is beautiful:

Figure 3: Dual-color imaging of synaptic connection by sypHTomato and GCaMP3. (partial figure)

Now that the technical issues have been address, it was time to ask some meaningful biological questions. The first question which they asked is whether the vesicle content at presynaptic termini (called boutons) of the same cell is the same in all synapses of the same cell, or are there differences based on the target, postsynaptic cell (obviously a pre-synaptic cell can create synapses with multiple post-synaptic cells).

They used stimuli to measure the volume of the readily-releasable vesicles, and ammonium chloride (NH4Cl), a strong base, to measure the total volume of all vesicles in each synapse. They found that synapses that were targeted to the same postsynaptic neuron had reduced variability in total and readily-releasable vesicle volume, compared to the variability of synapses targeted to different cells. This indicates that there is some reverse feedback from the post-synaptic cell to the pre-synapses.

It would have been interesting to see if pre-synaptic boutons from different cells also have less variability if they are targeted to the same post-synaptic cell. However, the authors did not measure that.

They then looked what happens when you stimulate the cells electrically. As ecpected, there is an immediate increase in sypHTomato signal (vesicle exocytosis) with an almost immediate increase in GCaMP3 signal (Ca2+surge) in the post synaptic cell. This Ca2+ surge begins at the synapse but propagates along the dendrites, indicating opening of voltage-gated Ca2+ channels with the propagation of the ation potential. They confirmed that the Ca2+ surge is due to the vesicle release by adding inhibitors for the neurotransmitter receptors on the post-synaptic cell, and detecting little or no Ca2+ increase (The inhibitors did not affect vesicle release). This approach, then, was capable to image neuronal activity at single synapse resolution.

But the authors went one step further, to get achieve an all-optical system.

The experiment described above was performed by electrically stimulating the cells. However, such methods are invasive and affect the entire cell. An alternative strategy to activate cells is by chemical application of agonists or antagonists. However, addition of drugs or neurotransmitters to the cell culture media will affect all cells in the media. Here comes optogenetics – a system that allows optical activation of a single cell, or part of it. I do not want to discuss optogenetics here; this should get a post or two of its own. The basic idea is to use genetically encoded light-responsive ion channels. Once you shine light (at a specific wavelength) on the part of the cell you want to activate, the ion channel opens and you get action potential originating from the part of the cell you shined on. So here comes the cool stuff:

The authors co-expressed channelrhodopsin2 (ChR2) with a pHluorin-tagged vesicular glutamate transporter (vGluTpH). ChR2 gets to the plasma membrane, whereas vGluTpH to the vesicles. When they shined blue light (to activate ChR2 and excite pHluorin) they get an increase in the green signal, which is abolished by a drug that inhibits action potential. Similarly, co-expression of VChR1 (another optogenetic tool) with stpHTomato and shining green light (to activate VChR1 and excite pHTomato) led to an increase in red emission, that was abolished by the drug.

Better yet, co-expressing ChR2 with sypHTomato showed an increase in red signal only when cells were illuminated by blue light (that activates ChR2) and not green light (Which doesn’t). This figure should have been a main figure. I don’t know why they put it in the supplementary.

Fig S6(b) Tests of ChR2 in combination with sypHTomato as proof-of-principle of all-optical yet independent monitoring and stimulation. Left, interrogation of sypHTomato with Green light (546 -566nm) excitation alone without activation of ChR2-driven vesicular turnover. Right, positive control with blue light (457-482 nm), showing robust ChR2-driven sypHTomato transient (right).

And there you have it – an all optical system to study vesicle release in neurons.

So what can we do with this system?

The authors state several exciting possibilities:

  1. Perform dual color experiments with other green sensors.
  2. Deciphering pre and post synaptic strength in different scenarios.
  3. Probing neuronal circuits: combining optogenetic photostimulation of two spectrally distinct channlerhodopsins with two spectrally distinct sensors (pH sensors in this case) will allows us to follow the neural pathway when we activate a distinct neuron with a specific color.

But I could think of other options. For instance, the authors did not discuss at all the growing use of caged molecules. In brief, caged molecules are biologically relevant molecules that are in an inactive state. These molecules can be activated by shining light of a specific waveband. Thus, if you have a caged neurotransmitter and you shine the light at a specific location, synapses can be activated only if the uncaged molecule is at high enough concentration (i.e. where you shined your light).

The authors kind of ignored it, but there seems to be a difference in the sypHTomato intensity, and signal decay, when comparing electrical and optical stimulation. This difference could be biologically meaningful and could be further explored.

All in all, I think it is a very nice paper, describing a new system to study neurons.

Further reading:

Yulong Li & Richard W Tsien (2012) pHTomato, a red, genetically encoded indicator that enables multiplex interrogation of synaptic activity. Nature Neuroscience 15, 1047–1053.

A good post on GCaMP at “brain Windows” blog.

Optogenetics resource center

Info on chemical synapses, from Wikipedia

ResearchBlogging.org
Li Y, & Tsien RW (2012). pHTomato, a red, genetically encoded indicator that enables multiplex interrogation of synaptic activity. Nature neuroscience, 15 (7), 1047-53 PMID: 22634730