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.



I am aware that it has been a while since I updated new posts on fluorescent microscopy. I blame life. And work.
Anyway, I have several papers piled up, waiting to get blogged.

In other news, I wrote a blog post for Addgene blog. Addgene, for those of you who do not know, is a non-profit organization that helps biologists share plasmids. Briefly, researchers can send Addgene their plasmids (which were featured in their publications) and other researches can order these plasmids from Addgene at a relatively small fee (relative to the amount invested in cloning a plasmid). My post is about how to choose the best suited fluorescent protein, something I wrote about in this blog sporadically. It should get published soon.

On May 22 I’m going to present my work at the annual meeting of the Israeli Society for Cancer Research.

On July 12-13 I’m going to present my work at the Gordon-Merck Research Seminar on Post-Transcriptional Gene Regulation  and the very next day at the Gordon Research conference on the same topic. You can still apply (for poster presentation only) for both of these.

On the publications’s front, a revised version of a paper I co-authored (which continues my work on the decay-transcription story) was recently sent back to the editors. We’re hoping for good news soon.

Am co-authoring review paper on mRNA localization for Nature Reviews Molecular & Cell Biology. Hope to submit within a couple of weeks.

And, the research I’ve been working on for the past two years is finally starting to take shape as a paper. Hope to submit within a month or so.

Also, for the first time, I’m now an official reviewer of a peer-reviewed journal ( RNA journal ).

The next evolutionary step

Human have always tried to improve on nature, from domestication of plants & animals through directed evolution in the test tube and GMO and up to Craig Venter’s synthetic bacteria and the expansion of the genetic code.
Today, another step was taken towards creating completely artificial life.

The natural genome is composed of four bases, which form two pairs: A-T and G-C. This pairing allows the double-helical DNA to maintain the information upon replication (the  semi-conservative replication).

The group of Floyd Romesberg from the Scripps institute now published a paper in Nature titled “A semi-synthetic organism with an expanded genetic alphabet”. I think that the title is a bit misleading, since its not really semi-synthetic, but is still awesome.

Roemsberg have previously published several papers on a synthetic deoxyribonucleotide pair: d5SICS and dNaM (I have failed to discover what these abbreviations stand for). These bases create an unnatural base-pair, yet it is still recognized, in vitro, by DNA and RNA polymerases and is maintained upon semiconservative replication (e.g. by PCR).

Now, they incorporated this pair into the bacteria E. coli to see if this pair can be maintained in vivo.

Unnatural (d5SICS-dNaM) and natural Watson–Crick (dC-dG) base pairs. Source: Malyshev et al. (2012) PNAS vol 109:12005-12010.

Chemical structures and the unnatural (d5SICS-dNaM) and natural Watson–Crick (dC-dG) base pairs. Source: Malyshev et al. (2012) PNAS vol 109:12005-12010.

The main problem going in vivo is how to get these bases into the bacteria, and how to get achieve the tri-phosphate (tri-P) form that can be incorporated by the DNA polymerase. They decided to use synthetic tri-P forms and screen to a transported that can import them into the cell. Having found a suitable nucleotide triphosphate transporters (NTT) from Phaeodactylum tricornutum and solved a problem of de-phosphorylation of the tri-P bases before they enter the cell, they set out to test if the bases-pair can be maintained in the E. coli cell.

To this end, they synthetically created a plasmid that harbors one d5SIC-dNaM base pair at a specific location. After transformation, they show that this base-pair is maintained at 95-97% of the plasmids for 15hr (24 generations). If they let the bacteria grow to stationary phase without supplementing them with fresh media (with fresh synthetic bases) then they slowly lose this pair to an A-T pair. but even after 6 days, ~10% of plasmids maintain this pair. This suggests that the replacement is not due to repair but due to the polymerase errors, and depletion of the bases in the media (though they don’t actually prove its not repair).

At this point, the paper ends. So, first of all, having one base pair is not semi-synthetic. I was disappointment :-)

Second, I hope these experiments can be repeated by other labs, and we will not get another “arsenic life” scandal.  But by the looks of it, it seems more reliable.

Third, this could be a platform for real synthetic biology, with a 6-letter code, instead of the 4 we have now, (that’s 216 3-letter codons, instead of the 64 we naturally have.  If we go to 4-letter codons, that’s 1296 codons to use.)

These could be used to create synthetic, unnatural proteins to produce new products; ribozymes and other RNA structures; imagine gene therapy that can only be maintained by ingesting these synthetic bases and I’m sure there are many options I haven’t thought of. In terms of this blog – maybe fluorescent proteins or RNA aptamers with interesting or useful properties (e.g. fast folding, very bright, etc..).

And last, this is the first proof that a DNA molecule does not have to be composed by the bases we’re used to. Aliens from other planets could have DNA molecules with similar double-helix Watson-Crick structure, semiconservative replication, but made out of other ribo nucleotide bases.

At the very least, we can revive dinosaurs without fearing they will run off the island and get their Lysine contingency supplemented elsewhere.

ResearchBlogging.orgMalyshev, D., Dhami, K., Lavergne, T., Chen, T., Dai, N., Foster, J., Corrêa, I., & Romesberg, F. (2014). A semi-synthetic organism with an expanded genetic alphabet Nature DOI: 10.1038/nature13314

iBiology Microscopy Course

Originally posted on Optical Nanoscopy Blog:

Light microscopy has become one of the most useful tools in the life sciences. Following the traditions of great courses on light microscopy, such as those offered by the Marine Biological Laboratory, EMBO, and the NCBS in Bangalore, this free online comprehensive course begins with the basics of optics, proceeds through transmitted light microscopy, covers the various methods of imaging fluorescent samples, describes how cameras work and image processing, and concludes with some of the latest advances in light microscopy. In addition to lectures, they also provide labs (filmed at a microscope) and short tips, so as to cover pragmatics of how to use microscopes. Assessments are provided for each lecture. Enjoy learning microscopy!


View original

sequencing localized RNA in single cells by FISH

To celebrate the 2-year anniversary of this blog, lets talk about the new Science paper in which the authors claim to performs in situ single cell, single molecule  RNA sequencing.

So what’s the big deal?

Well, RNA sequencing (RNA-Seq) has become a very common method to study gene expression. In many cases, RNA-Seq uses cell extraction from an entire cell population -thus averaging the RNA content of each individual cells. In recent years, single-cell RNA-Seq is becoming more feasible (for example). In this case, cells are sorted via flow cytometry so that one can sort individual cells into designated wells in a multi-well plate. Thus, RNA from a single cell can be sequenced. Though this process is becoming both efficient and accurate, you loose information about the cellular localization of the RNA.

FISH is a method that enables to determine the accurate localization of the target mRNA. However, FISH is limited to only a small number of mRNAs. Using color barcoding of the FISH probes can increase the complexity that is achieved (i.e. – one can simultaneously detect multiple types of RNAs) but these are still only a few compared to the entire transcriptome.

This new paper in Science combines FISH with RNA sequencing to give Fluorescent in situ sequencing (FISSEQ). How did they do that?

First, they generated cDNA by performing reverse-transcription in fixed cells with random primers. The cDNA was then circularized and then amplified. One of the nucleotides has a reactive group so that the cDNA is cross-linked to the surrounding macromolecules. Thus, the cDNA is localized to the same location as the RNA.  The primer has an adapter sequence that can be used as template for sequencing or for FISH. They show some 3D FISH images of cells and tissues using a probe for this adapter. Pretty pictures, but not much info there.

From here on, it gets trickier.

For the sequencing, they used the SOLiD method. One problem was to reduce the spot density so that it will be sufficiently low to distinguish single molecules. If I understand them correctly, they modified the SOLiD sequencing probes so that the sequencing primers have mismatches that reduce the efficiency.

The second problem was to identify auto fluorescence and other fluorescent artifacts in their images. Rather than fine-tuning a specific detection threshold, they relied on the fact that they are monitoring sequences. That means that the fluorescence at each spot should account for a known sequence (i.e. the colors pattern should change based on the sequence), whereas auto fluorescence should remain stable. So, they actually used no threshold at all.

The sequencing reaction cycles and images of the first 15 cycles in primary fibroblasts. Source: Lee JH et. al. (2014) Science 343:1360.

The sequencing reaction cycles and images of the first 15 cycles in primary fibroblasts. Source: Lee JH et. al. (2014) Science 343:1360.

They them showed a few applications of this method. One interesting feature was to show that the nucleus is enriched for non-coding and antisense RNAs compared to the cytoplasm that is enriched with mRNAs.

Unfortunately, the nuclear/cytoplasm dichotomy was the only “localization” aspect in their paper. It would have been much more interesting to show mitochondrially-localized vs ER localized vs plasma membrane localized etc…

Or, look at highly polarized cells like neurons and use FISSEQ to look at somatic vs dendritic or axonic RNAs.

Another experiment they did was to look at the gene expression changes in response to a wound healing model. They found the expected increase in genes related to cell migration, and with some genes differentially expressed only in the migrating cells (at the “wound”) compared to contact inhibited cells that are close to them.  Again, It would have been even better if they could show the subcellular of these mRNAs – are they at focal adhesion sites or other unique sites in the migrating cells?

Their results on the whole seem convincing, but I didn’t check all the little technical details.  One thing which is missing is a high resolution image showing the sequencing of just one RNA, compared to an auto fluorescence spot. All their images show one or many cells with multiple spots in different colors – but that is it.

The authors say upfront that this is just a demonstration of their method, and that they expect it to improve in coming years, just like what happened with next-generation sequencing.

We’ll see…
ResearchBlogging.orgLee JH, Daugharthy ER, Scheiman J, Kalhor R, Yang JL, Ferrante TC, Terry R, Jeanty SS, Li C, Amamoto R, Peters DT, Turczyk BM, Marblestone AH, Inverso SA, Bernard A, Mali P, Rios X, Aach J, & Church GM (2014). Highly multiplexed subcellular RNA sequencing in situ. Science (New York, N.Y.), 343 (6177), 1360-3 PMID: 24578530


[post-pub note: this post was also published at the RNA-Seq blog, here].


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