TSRI Scientists Create First Stable Semisynthetic Organism

E. coli attached to intestines. Credit: ZEISS Microscopy

LA JOLLA, CA – January 23, 2017 – Life’s genetic code has only ever contained four natural bases. These bases pair up to form two “base pairs”—the rungs of the DNA ladder—and they have simply been rearranged to create bacteria and butterflies, penguins and people. Four bases make up all life as we know it.

Until now. Scientists at The Scripps Research Institute (TSRI) have announced the development of the first stable semisynthetic organism. Building on their 2014 study in which they synthesized a DNA base pair, the researchers created a new bacterium that uses the four natural bases (called A, T, C and G), which every living organism possesses, but that also holds as a pair two synthetic bases called X and Y in its genetic code.

TSRI Professor Floyd Romesberg and his colleagues have now shown that their single-celled organism can hold on indefinitely to the synthetic base pair as it divides. Their research was published January 23, 2017, online ahead of print in the journal Proceedings of the National Academy of Sciences.

“We’ve made this semisynthetic organism more life-like,” said Romesberg, senior author of the new study.

While applications for this kind of organism are still far in the future, the researchers say the work could be used to create new functions for single-celled organisms that play important roles in drug discovery and much more.

Building a Unique Organism

When Romesberg and his colleagues announced the development of X and Y in 2014, they also showed that modified E. coli bacteria could hold this synthetic base pair in their genetic code. What these E. coli couldn’t do, however, was keep the base pair in their code indefinitely as they divided. The X and Y base pair was dropped over time, limiting the ways the organism could use the additional information possessed in their DNA.

“Your genome isn’t just stable for a day,” said Romesberg. “Your genome has to be stable for the scale of your lifetime. If the semisynthetic organism is going to really be an organism, it has to be able to stably maintain that information.”

Romesberg compared this flawed organism to an infant. It had some learning to do before it was ready for real life.

In stepped TSRI Graduate Student Yorke Zhang and Brian Lamb, an American Cancer Society postdoctoral fellow in the Romesberg lab at the time of the study. Together, they helped develop the means for the single-celled organism to retain the artificial base pair.

First, Zhang and Lamb, co-first authors of the study, optimized a tool called a nucleotide transporter, which brings the materials necessary for the unnatural base pair to be copied across the cell membrane. “The transporter was used in the 2014 study, but it made the semisynthetic organism very sick,” Zhang explained. The researchers discovered a modification to the transporter that alleviated this problem, making it much easier for the organism to grow and divide while holding on to X and Y.

Next, the researchers optimized their previous version of Y. The new Y was a chemically different molecule that could be better recognized by the enzymes that synthesize DNA molecules during DNA replication. This made it easier for cells to copy the synthetic base pair.

A New Use for CRISPR-Cas9

Finally, the researchers set up a “spell check” system for the organism using CRISPR-Cas9, an increasingly popular tool in human genome editing experiments. But instead of editing a genome, the researchers took advantage of CRISPR-Cas9’s original role in bacteria.

The genetic tools in CRISPR-Cas9 (a DNA segment and an enzyme) originated in bacteria as a kind of immune response. When a bacterium encounters a threat, like a virus, it takes fragments of the invader genome and pastes them into its own genome—a bit like posting a “wanted” poster on the off chance it sees the invader again. Later, it can use those pasted genes to direct an enzyme to attack if the invader returns.

Knowing this, the researchers designed their organism to see a genetic sequence without X and Y as a foreign invader. A cell that dropped X and Y would be marked for destruction, leaving the scientists with an organism that could hold on to the new bases. It was like the organism was immune to unnatural base pair loss.

“We were able to address the problem at a fundamental level,” said Lamb, who now serves as a research scientist at Vertex Pharmaceuticals.

Their semisynthetic organism was thus able to keep X and Y in its genome after dividing 60 times, leading the researchers to believe it can hold on to the base pair indefinitely.

“We can now get the light of life to stay on,” said Romesberg. “That suggests that all of life’s processes can be subject to manipulation.”

A Foundation for Future Research

Romesberg emphasized that this work is only in single cells and is not meant to be used in more complex organisms. He added that the actual applications for this semisynthetic organism are “zero” at this point. So far, scientists can only get the organism to store genetic information.

Next, the researchers plan to study how their new genetic code can be transcribed into RNA, the molecule in cells needed to translate DNA into proteins. “This study lays the foundation for what we want to do going forward,” said Zhang.

Professor Floyd Romesberg (right) and Graduate Student Yorke Zhang led the new study at The Scripps Research Institute, along with Brian Lamb (not pictured).(Photo by Madeline McCurry-Schmidt.)

Additional authors of the study, “A semisynthetic organism engineered for the stable expansion of the genetic alphabet,” were Aaron W. Feldman and Anne Xiaozhou Zhou of TSRI; Thomas Lavergne of the University of Grenoble; and Lingjun Li of Henan Normal University.

The study was supported by the National Institutes of Health (grant GM060005), a National Science Foundation Graduate Research Fellowship (grant DGE-1346837), the National Natural Science Foundation of China (grant 21472036), a Labex ARCANE grant (ANR-11-LABX-0003-01), NanoBio-ICMG platforms (FR 2607) and a postdoctoral fellowship from the American Cancer Society, Illinois Division.

About The Scripps Research Institute

The Scripps Research Institute (TSRI) is one of the world's largest independent, not-for-profit organizations focusing on research in the biomedical sciences. TSRI is internationally recognized for its contributions to science and health, including its role in laying the foundation for new treatments for cancer, rheumatoid arthritis, hemophilia, and other diseases. An institution that evolved from the Scripps Metabolic Clinic founded by philanthropist Ellen Browning Scripps in 1924, the institute now employs more than 2,500 people on its campuses in La Jolla, CA, and Jupiter, FL, where its renowned scientists—including two Nobel laureates and 20 members of the National Academies of Science, Engineering or Medicine—work toward their next discoveries. The institute's graduate program, which awards PhD degrees in biology and chemistry, ranks among the top ten of its kind in the nation. In October 2016, TSRI announced a strategic affiliation with the California Institute for Biomedical Research (Calibr), representing a renewed commitment to the discovery and development of new medicines to address unmet medical needs. For more information, see www.scripps.edu.

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Can we synthetically engineer C4 photosynthesis?

“Photosynthesis as the engine for life on earth has high engineering potential, which has not yet been fully exploited…By step-wise identification of all the components needed for engineering, it will eventually become possible to employ this powerful machinery to increase yields for the future.”

Schuler, ML, Mantegazza, O & Weber, APM, 2016, ‘Engineering C4 photosynthesis into C3 chassis in the synthetic biology age’. The Plant Journal, vol. 87, pp. 62

These lines from the conclusion of the review we write about here are indicative of why so much effort is being put into understanding the more productive C4 photosynthetic system and working to increase important crop yields with it.

Schuler, Mantegazza and Weber’s article in the special issue of The Plant Journal on plant synthetic biology provides an excellent overview of the current status, significant hurdles and possible solutions to those problems of the current research aimed at bolstering rice yield by converting it from the common C3 photosynthesis system to the more efficient C4 system. We’ve previously written about C4 photosynthesis here and here.

C4 photosynthesis

C4 photosynthesis has evolved independently at least 66 times and is likely linked to a sudden drop in atmospheric CO2 levels sometime in the past. It is characterised by the concentration of CO2 around Rubisco (ribulose 1,5-bisphosphate carboxylase/oxygenase), the carbon-assimilating enzyme, reducing the competition that CO2 has with O2 to interact with the enzyme. More CO2 means greater growth and reduced photorespiration, an energy requiring process that is used to remove the O2 reaction products.

The concentration of CO2 in C4 photosynthesis is usually caused by a two-celled (but one-celled is possible) distribution of the process of fixing carbon and the process of reducing it. The two-celled system combines mesophyll (M) cells, which take up the CO2 from the leaf air space, and the bundle sheath (BS) cells, where the Rubisco enzymes reside, the final destination of CO2 for fixation and entry into the Calvin-Benson cycle. These two cells are arranged in concentric layers (called ‘Kranz Anatomy’) around leaf veins, maximising the contact between the two types of cells and increasing the transport of the molecules between them.

M cells convert CO2 to bicarbonate and then into the 4 carbon compound oxaloacetate via an enzyme that doesn’t react with oxygen. The modified compound is then passed to the BS cells where it is reformed into CO2 and fixed by Rubisco to enter the Calvin-Benson cycle.

Basically, by assimilating CO2 away from Rubisco, the plant reduces the ability of Rubisco to interact with O2 and instead it is steadily fed with CO2 from the M cells.

Of course, this description of the process is simplified and although most of the process and main enzymes that carry out the process are known, there are still gaps in our knowledge.

Recent Advances

The gathering of increasing amounts of genomic, trascriptomic and metabolimic data continue to improve our knowledge of C4 photosynthesis, how it evolved and how we might transition C3 crops to use the more efficient carbon fixation method.

Important C4 crop species have had their genomes sequenced and quantitative analysis of transcriptomes have begun to unravel the mystery behind the genes upregulated and downregulated, and the stage of development that these regulatory differences occur, that lead the formation of the Kranz anatomy. What we are finding is that many of the genes involved in C4 photosynthesis exist in C3 plants but are differently regulated at early stages to differentiate the BS and M cells, enable high throughput of metabolites between the cells and to increase the size of vascular tissue to support the increased activity.

Engineering C4 photosynthesis

Our initial attempts to engineer C4 photosynthesis relied on over-expressing one or more enzymes in C3 plants. However, given the enzymes involved in the C4 system are used in the C3 system in multiple alternative pathways, the effects of over-expression were multiple, varied and didn’t have the desire result. The compartmentalisation of reactions, whether in the single or two-celled reactions that make up the distinctive photosystem, is complex.

The notion of being able to engineer C4 photosynthesis is comforted by a number of factors:

1. The main enzymes are already present in C3 photosynthesis;
2. Characteristics such as the passing of metabolites between cells is seen in C3 species such as tobacco plants; and
3. Nature has done it herself in the past on multiple, independent occasions.

But the authors of the paper also note a number of engineering steps that need to be accomplished if we are re-enact evolution ourselves;

1. Higher order veins need to be initiated in plants (it previously being shown that such physical properties were already evolved in plants that subsequently evolved the Kranz anatomy);
2. The ratio of BS to M cells must be increased, ideally in a similar concentric organisation to Kranz anatomy;
3. Enlarging and enriching BS cells with additional chloroplasts;
4. Increasing the connection between M and BS cells;
5. Engineering the different morphologies of the chloroplasts to mimic the morphologies of chloroplasts found in M and BS cells;
6. Mirror the differing roles that M and BS cells take on in C4 photosynthesis so Rubisco reduction of CO2 occurs only in the BS cells with M cells feeding CO2 to the BS cells and excluding the oxidation of O2.

The tools we need

If we are to achieve success we still have some tools to develop and refine.

Chief among this list is a model plant that can be engineered and tested easily with speedy regeneration without requiring too much growing room. The authors point out that rice crops have some limitations in these criteria but identify Brachypodium distachyon as a model C3 plant with a small, annotated genome with quick flowering time, low growing space requirements and an efficient transformation protocol. A model such as this could hasten the engineering, testing and data gathering on conversion which can then be tested on important crop species.

A C4 model plant with similar characteristics is also required. Setaria viridis has previously been suggested as a possible model plant, as has the Fast Flowering Mini Maize.

The ability to drive and control expression of a transgene is also required. Cis-regulatory modules that promote gene expression are still under development in the wider plant synthetic biology area. This leaves a chasm between the tools we have to hand and the possibility that a large number of genes need to be differentially expressed in order to convert C3 photosynthesis to C4 photosynthesis.

Huge strides are being made with genetic manipulation, particularly with the discovery and modification of the CRISPR/Cas 9 system. But, according to the article, the maximum number of genes successfully introduced into a plant, at present, is 9. To induce C4 photosynthesis in a C3 plant, we may need the ability to stably transform a far larger number of genes plus regulatory elements, and do so without disrupting the remainder of the genome or the phenotype characteristics of our food crops.

Even when we do have these tools at the ready, we are still missing some vital information about the genes and regulatory elements that compose C4 photosynthesis. Increasing our knowledge of minutia of genetic composition and regulation of C4 systems compared to C3 systems is still a top priority. Identifying genera with the underlying predisposition that have allowed species within it to evolve from C3 to C4 for comparative analysis, particularly species displaying characteristics of a C3-C4 intermediate with sister taxa displaying C3 and C4 phenotypes, would be idyllic in assisting the study of the evolution. The authors highlight Morandia and Parthenium generas as possible true intermediates between C3 and C4 plants. Programs such as the Grass Phylogeny Working Group and the 1KP (1000 plants) project will greatly assist identifying and genotyping suitable candidates for understanding the genetics behind enhancing crop photosynthesis.

And some suggested means of pushing the research…

It is great to see that not only have the authors elucidated quite extensively the current knowledge and gaps within the field of C4 photosynthesis engineering, but have also suggested a couple of ways of advancing the research.

The first idea they suggested is synthetically replicating a simplified C4 photosynthetic system using known genetic components. The system replicates the targeting of specific enzymes to create a two-celled photosynthesis construct, limiting Rubisco to the BS cells using RNAi to interfere with its transcription in M cells. The article highlights specific transporters that can be used to transport the metabolites between the two cells.

A second suggested idea is using brute force to direct a speedy evolution of a C3 or C3-C4 intermediate species into a C4 plant. Identifying the minimum genetic requirements of a C4 plant in a candidate crop would then be followed by the repetitive growth under the selective pressure of a low CO2 atmosphere. By repeating genomic and transcription analysis of the evolving plant (if successful), a ‘mud-map’ of the road from C3 to C4 plants can be generated and be of enormous use to research seeking to synthetically install the same machinery.

Conclusion

Although its behind a pay-wall, get your hands on this article. Whether it be for a background in C4 photosynthesis or as a springboard for your own research, it is an area of immense potential that should be worthy of an X prize.

ORIGINAL: Legume Laboratory

September 6, 2016

C2c2: We can now 'cut and paste' RNA in addition to DNA, and it could disable viruses

Juan Gaertner/Shutterstock.com

Science gets smarter.

You've probably heard of CRISPR - the gene editing tool that essentially lets scientists cut and paste DNA, removing things like HIV and muscular dystrophyf rom our cells - and now scientists have discovered a way to edit RNA with just as much precision.

RNA is DNA's close biological cousin, responsible for translating messages from the nucleus to the rest of the cell, and being able to change it could open up all-new disease-fighting possibilities.

Just like CRISPR/Cas9 editing, the new procedure selectively cuts up RNA, which gives us microscopic control over genetic information, and the researchers behind it say it could open up the method could be used to block viruses and halt disease in its tracks.

"RNA is the blueprint through which genes regulate cellular processes," biologist Oliver Rackham at the University of Western Australia, who wasn't involved in the study, told Colin Barras from New Scientist. "The exciting thing about this study is that it now opens up the RNA world to the ease of experimental design afforded by CRISPR."

While DNA carries the genetic code for our bodies, RNA carries out those instructions and puts that code to work. Basically, DNA has the recipes but RNA reads that recipe and directs all the cooking.

And while scientists used to think the RNA was just a 'middleman', they've realised over the past few decades that it does so much more than that - it can control which proteins are produced in the cell.

In fact, the original CRISPR/Cas9 method uses a combination of DNA and RNA information to target viruses in the body, and identify genetic code that needs replacing. But right now, it focusses solely on DNA.

Now an international team of scientists, led by Feng Zhang from the Broad Institute of MIT and Harvard, has discovered another version of CRISPR called C2c2, and it can be used to slice up RNA molecules exclusively.

Just like CRISPR/Cas9, the C2c2 system was discovered inside bacteria, where it works like a pair of molecular scissors to 'cut out' any invading pathogen's RNA in order to disable an attack.

But the difference is that C2c2 only targets RNA, not DNA itself, which means scientists can use it to make changes and control what happens within a cell, without changing the genetic code itself.

Zhang's team tested the system by inserting C2c2 into E. coli bacteria, and showed they were able to use it to silence a gene simply by targeting RNA. This is because they attacked the RNA carrying the instructions from the gene, so the gene was effectively turned off, without the DNA being touched.

The researchers say C2c2 should also be able to bind to RNA without cutting it up, which would allow scientists to track the movement of RNA in a cell and monitor the effect that changing genetic information has on a single cell, or throughout the whole body.

Whereas DNA changes are permanent, RNA changes could be tweaked as treatment progresses, which gives a lot more opportunity for fighting disease.

"C2c2 opens the door to an entirely new frontier of powerful CRISPR tools," said Zhang. "There are an immense number of possibilities for C2c2, and we are excited to develop it into a platform for life science research and medicine."

While the technique did produce some collateral damage in this study - some RNA that looked similar to the target molecules was unintentionally attacked - it's still early days, and Rackham told New Scientist that the feature could actually be useful in some treatments.

"In a cancer treatment, for example, wiping out 'bystander' RNA as well as target RNA might offer a more effective way to fight a tumour," reports Barras.

This isn't the first time scientists have been able to target RNA instead of DNA through gene-editing techniques, but the difference is C2c2 uses a more natural and more versatile approach, and potentially has a range of different uses. We can't wait to see what scientists do with it.

The results of the study have been published in Science.

ORIGINAL: Science Alert
DAVID NIELD
3 JUN 2016

Bitter fight over CRISPR patent heats up

Unusual battle among academic institutions holds key to gene-editing tool’s future use.

Steve Jennings/Getty/Breakthrough Prize.Biologist Jennifer Doudna of the University of California, Berkeley, helped to develop the CRISPR gene-editing system.

A versatile technique for editing genomes has been called the biggest biotechnology advancesince the polymerase chain reaction (PCR), and the US Patent and Trademark Office (USPTO) is set to determine who will reap the rewards.

On 11 January, the USPTO granted a request to review a key patent awarded for the technique, known as CRISPR–Cas9. The outcome of the ensuing proceedings, called a patent interference, could be worth millions to the research institutions that are at war over the relevant patents. It might also influence who is allowed to use the technology — and under what terms.

“This is an absolutely humungous biotech patent dispute,” says legal scholar Jacob Sherkow of New York Law School. “We’re all waiting with bated breath.”

CRISPR–Cas9 is a bacterial defence system that uses the enzyme Cas9 to snip DNA at sites determined by the sequence of a ‘guide’ strand of RNA. Scientists can disable, replace or tweak genes by using the technique to rewrite snippets of DNA sequences. Use of the technology in research has exploded, thanks to CRISPR–Cas9’s relative simplicity and versatility compared to other gene-editing methods. Several companies have sprung up to harness the technique for generating improved crops, research reagents and therapies for human genetic diseases.

The roots of the CRISPR–Cas9 dispute date back to 2012, when researchers reported that they had reprogrammed the system to cut strands of isolated DNA at sites of their choosing1. The team, led by biologists Jennifer Doudna at the University of California, Berkeley, and Emmanuelle Charpentier, now at the Max Planck Institute for Infection Biology in Berlin and Umeå University in Sweden, filed a patent application on 15 March 2013.

By then, publications had emerged from other groups showing that the method works in human cells2–4 and bolstering dreams of CRISPR-based gene therapies — the basis for several companies that have sprung up to capitalize on the technique.

One of those groups, led by synthetic biologist Feng Zhang of the Broad Institute and the Massachusetts Institute of Technology, both in Cambridge, filed a patent application for the CRISPR–Cas9 technique in October 2013. The institutions filed the patent under a special expedited review programme, and it was granted in April 2014. Zhang has since been awarded additional patents on the technology. The original Doudna–Charpentier patent remains under review.

A pitched battle

In April 2015, the Berkeley team asked the USPTO to begin an interference proceeding to determine which team was the first to invent the technique. The proceedings will be much like a court case, with both sides presenting evidence culled from publications and laboratory notebooks. “Once the [USPTO] declares an interference, that’s really when the fur is going to fly,” Sherkow predicted in a June interview.

Nature special:CRISPR — the good, the bad and the unknown

The patent interference is also a testament to the high stakes involved: companies aiming to use CRISPR–Cas9 for gene therapy have raised hundreds of millions in venture capital and other funds in under three years. One company, Editas Medicine in Cambridge, Massachusetts, has already filed to go public.

Arti Rai, a legal scholar at Duke University in Durham, North Carolina, says that it is unusual for academic research institutions to battle so intensely over a patent. Instead, such institutions usually come to an agreement to share rights to the invention. “This seems more bitter than disputes I’ve heard of in the past,” she adds.

The two patents in question make broad claims to 'foundational' intellectual property thought to be necessary for most lucrative CRISPR–Cas9 applications. But many patents have been filed on CRISPR–Cas9 technologies, and there is still the chance that the winner of the interference will face additional challenges in court. Zhang's group has also reported another enzyme, called Cpf1, that could provide an alternative to Cas9. Researchers expect other alternatives to emerge with time.

As for the various CRISPR–Cas9 companies, Zhang remains involved in Editas, which was founded by both Zhang and Doudna, among others, in 2013. Doudna has since severed ties with Editas and thrown her support behind Intellia Therapeutics, also in Cambridge. Charpentier, meanwhile, co-founded CRISPR Therapeutics of Basel, Switzerland.

Licensing looms

For now, it is unclear how the dispute will affect researchers who use CRISPR–Cas9, if it does so at all. Academics who might use the technology for basic research make unattractive targets for patent lawsuits, says Rodney Sparks, a biotechnology patent counsel at the University of Virginia in Charlottesville. “Patent holders might send out a few cease-and-desist letters, but they probably won’t sue academic researchers,” he says. Doing so would take time and money with little reward: the spoils in a patent lawsuit are typically damages or a share of royalties from a marketed product. That leaves little to gain from suing academics who are not selling anything. But those who intend to use their research as the basis for a start-up company will need to be wary, Sparks says.

Some patent holders do ask that even scientists doing basic research take out a licence on a patented technology, typically for a fairly small fee. Such was the case for PCR, says Warren Woessner, a lawyer at Schwegman Lundberg and Woessner in Minneapolis, Minnesota. Woessner recalls how, during his previous career as a scientist, his institution decided to patent a method he developed. Officials at the institution later noticed that someone had published a paper that used the technique without a licence. “They sent the professor a little note,” recalls Woessner.“‘We have a patent on this. Pay up.’” The professor did.

The Broad Institute has noted on its website that it will continue to make CRISPR–Cas9 reagents available to the community, and has given no indication that it will pursue licensing fees from academics. But Sherkow warns against assuming that the spirit of academic camaraderie will prevail: licensing revenue has become increasingly important, particularly for major research institutions, he says. “We’re just living in a brave new world these days.”
Nature doi:10.1038/nature.2015.17961

ORIGINAL: Nature

Heidi Ledford

12 January 2016

Steven Pinker interview: case against bioethocrats & CRISPR germline ban

CRISPR-Cas9 gene editing technology is red-hot right now.

It has great power for research in the lab and there are hypothetical transformative clinical applications of CRISPR too. The latter efforts could include experimental attempts at reversal of disease-causing mutations in one-cell embryos with the hope that they then grow into full-fledged, healthy human beings. Hypothetically CRISPR could also be used for pursuing human enhancement via germline genetic modification.

As a tool CRISPR is exciting and my own lab is using it for genetic studies, but from a technical perspective it’s not perfect. It can introduce a range of types of errors into the genome, with largely unknown biological consequences. To date, the first and only report of CRISPR-based modification of human embryos was arguably most notable for the problems encountered including genetic errors. However, suboptimal CRISPR methods were used so better design would almost certainly reduce risks of errors.

CRISPR raises a number of questions and has sparked many discussions. 

• How should we handle a cutting edge biotechnology of this kind as a community of scientists? 
• What if anything should be the appropriate role of others in such considerations including bioethicists? }
• Should there be a temporary moratorium on clinical use of CRISPR? 

I tackled some of these issues in my own past piece Practical Plan for Managing Human Germline Genetic Modification. Others advocate for a more liberal perspective on the road to possible clinical use of CRISPR, focusing on the potential for great benefits.

Overall, this all might be summed up as follows: how do we balance the gas pedal and the brakes on CRISPR’s use in humans to aim for the greatest overall net benefit?

Steven Pinker

Professor Steven Pinker of Harvard has been one of the most outspoken advocates for more gas and less brakes here. Both in writing and in talks he has expressed the view that we should move forward without substantial impediments to CRISPR-Cas9. For instance, Pinker’s “get out of the way” editorial last week in The Boston Globe on CRISPR was very critical of bioethics and advocated an expeditious path forward for the research without constraints. It sparked wide-ranging discussions and even some anger from bioethicists. Update: see also this brief reply to Pinker’s interview from noted bioethicist, Art Caplan.

A few days ago I reached out to Dr. Pinker to do an interview to learn more of the specifics about his views with a goal toward increasing dialogue. For instance, I wondered if he really felt that strongly about the harms caused by bioethics that were suggested in his editorial. I want to thank him for taking the time to provide such detailed answers that make the full depth of his views on these issues far clearer here than in the past.

Knoepfler. 1. Related to your talk at BEINGS and your more recent editorial, what do you see as the appropriate role for bioethics and bioethicists in the life sciences? “Get out of the way” seems rather absolute. Can you help us understand the nuances there in your view of bioethics if any?

Pinker: There’s a difference between ethics, on the one hand, and “bioethics” and “bioethicists,” on the other. Of course everything a scientist does—everything a human being does—ought to be ethically guided. But bioethics has become a professional guild that all too often impedes sound ethical concerns rather than advancing them. Many moral philosophers—the scholars who specialize in evaluating the soundness of ethical arguments—believe that mainstream bioethics commonly trades in confused claims based on emotion and woolly thinking (see these articles by Julian Savulescu,Sally Satel, and me for examples).

Take the very foundation of ethics. You’d think it would be an obvious ethical principle that life is better than death, health is better than disease, and vigor is better than disability. But, astonishingly, so-called bioethicists have repeatedly denied these truisms, either explicitly (in the case of the country’s former bioethicist-in-chief, Leon Kass, who argued that the desire to extend life is a sign of shallowness and immaturity), or implicitly, by fetishizing sweeping rubrics such as dignity, equity, social justice, sacredness, privacy, and consent at the expense of the health and lives of actual people.

It’s not just that many bioethicists practice bad moral philosophy. It’s that they are entangled in a conflict of interest. Institutionalized bioethics has become an academic and bureaucratic industry, and they need to rationalize their existence. You hardly need a bioethicist to tell you that it’s wrong to inject typhus into twins or to withhold antibiotics from syphilis patients. But to come up with an abstruse argument as to why a parent should be prohibited from saving the life of her infant by donating a part of her liver—for that you need a “bioethicist.”

Regarding my advice to “get out of the way,” the nuances were stated, albeit tersely, in the article. The first is that a truly ethical bioethics must weigh the benefits of any restriction on research against the harm that will be caused to the vast number of people who would benefit if the research proceeded expeditiously. Savulescu puts it starkly: “To delay by 1 year the development of a treatment that cures a lethal disease that kills 100,000 people per year is to be responsible for the deaths of those 100,000 people, even if you never see them.”

The second is that a truly ethical bioethics should justify any restrictions on research with rigorous, defensible arguments about benefit and harm, not with moralistic grandstanding, science fiction dystopias, perverse analogies to Nazis and nuclear weapons, esoteric theories pulled out of the air, or freak-show scenarios like armies of cloned Hitlers, people selling their eyeballs on eBay, or warehouses of zombies to supply people with spare organs—all of which I’ve heard in these debates.

And as I wrote, no one questions the need to protect patients and research subjects from exploitation or harm. If there are flaws in the existing safeguards, as, for example, Alice Dreger argues, the safeguards should be fine-tuned or re-engineered. This is not the same as giving more power to the bioethocrats. A great deal of bioethical argumentation has nothing to do with protecting people. It rather cooks up reasons why consenting adults should be prohibited from doing things that help them or others while harming no one—a prominent example being recipient-solicited or incentivized organ and tissue donation. And establishment bioethics has caused preventable harm. Most infamous is the case of Jesse Gelsinger, the young man who died in a Phase-1 trial of gene therapy in 1999. Common sense would say that the experimental therapy should have been tested for safety on infants with a severe form of the disease who would have died anyway. But Arthur Caplan, the country’s most famous bioethicist, argued that the parents of such infants would be so consumed with grief that they could not truly give consent—the kind of paternalistic argument that is all too common in this field—and that an 18-year-old with a mild form of the disease, who technically could give consent, should be enrolled instead. A strained interpretation of the magic word “consent” was allowed to trump expected harm and benefit, and the result was tragedy.

Today mainstream bioethics gets in the way on a massive scale. The most obvious example is Institutional Review Boards. They are blatant abridgments of free speech, convenient weapons for fanatics to wield against people whose opinions they don’t like, and high-volume red-tape dispensers which bog down research while being unnecessary or even harmful to the protection of patients and research subjects. (See the Illinois White Paper and American Association of University Professors reports on IRB mission creep, David Hyman’s “The Pathologies of Institutional Review Boards,” and the new books The Ethics Police by Robert Klitzman and The Censor’s Hand by Carl Schneider). Regulations on confidentiality and consent to use data and tissues have also gone way overboard. The future of medicine hinges on the use of massive, open-access datasets to find signals in the noise. If every byte has to be multiply certified for consent and privacy, or even destroyed after a few years, no matter how inconsequential to the person who contributed it, then huge numbers of future patients will suffer or will fail to be helped by our faulty knowledge of the real effects of treatments.

There is, to be sure, an important role for bioethics. Satel puts it well: bioethicists at their best are “scholars who study the intellectual and social history of value controversies in medicine and biotechnology. They can teach us about the technical and cultural antecedents of modern debates and show us how to engage in disciplined moral inquiry. They are skilled at drawing conceptual maps of the dilemma at hand while enumerating various ways to resolve it.”

Knoepfler. 2. Forgetting bioethicists entirely for the moment, prominent scientists such as Jennifer Doudna, David Baltimore, and others have publicly called in unambiguous terms for at least a temporary moratorium on clinical applications of human germline editing technology. Do you disagree or agree? Why?

Pinker: Disagree. The specific harms they warn against, such as inducing cancer, mutations, or birth defects in the unborn child are already ruled out by a plethora of existing regulations and norms. Obviously we shouldn’t mess around with embryos in ways that have a significant probability of producing a sick or deformed child with no compensating benefit. But why do we need a new, across-the-board ban on an entire method to rule out what’s already ruled out on the uncontroversial grounds of protecting individuals against foreseeable harm? The authors seem to be acquiescing to the yuck-factor that surrounds the very idea of germline modification, if for no other reason than to draw a firewall around their own research programs, which are restricted to the genetic modification of somatic cells. But scientists should work to dismantle irrational taboos, not indulge them.

First, the idea that there is some sacrosanct entity called “the human germline,” such that deliberately manipulating it would violate this sanctity, or restrict the freedom of future generations, or alter the species in unprecedented and frightening ways, is biological nonsense. No two people, not even monozygotic twins, have the same germline. Each of us introduces dozens of random mutations into our germlines, often multiplied by voluntary choices such as exposing ourselves to mutagens like tobacco smoke or fathering a child in middle age. And we affect the genetic makeup of our offspring, and the species, every time we choose to have unprotected sex with one partner rather than another. So even if it did come to pass that some people edited out disease genes, or (far less likely—see below) edited in enhancement genes, it would be a droplet in the maelstrom of naturally churning genomes.

Second, a ban or moratorium would only reinforce the pernicious aura of dread that surrounds genomic modifications. This is the dread that incites across-the-board opposition to genetically modified organisms and that underpins the bogus moral arguments against cytoplasmic donation for mitochondrial disease (the so-called three-parent babies—another case in which so-called bioethical concerns increase rather than decrease death and suffering). And the spurious ideal of germline sacredness could compromise the treatment of disease in other ways. Though lots of things went wrong in the Gelsinger case, one complication was the decision to administer massive doses of the viral vector directly to his liver, with the risk of lethal inflammation, rather than systemically, out of the fear that (God forbid!) it might introduce the needed gene into his sperm-forming cells. That’s probably not what killed Gelsinger, but it did kill a monkey in a safety trial, and this germlinophobia could certainly endanger gene-therapy patients in the future.

Third, germline editing could have direct benefits in a number of scenarios:

• to parents with disease genes who don’t produce enough viable embryos for preimplantation genetic diagnosis (especially when more than one such gene is involved, which multiplies the number of necessary embryos); 
• to parents who both are homozygous for some recessive disease gene (not far-fetched given how often people meet each other through support groups); if future data were to show that PGD babies have compromised longevity or health; and 
• in other scenarios that perhaps we can’t imagine. 

For these reasons Savulescu, with Chris Gyngell, and Henry Miller with Drew Kershen argue that research on germline editing is not only morally permissible but morally imperative.

Knoepfler. 3. You appear relatively confident in future benefits of new biotechnology such as CRISPR to millions of people, but you seem very skeptical of the risk predictions that you described as “speculative harms”. What makes you so confident of benefit and at the same time so skeptical of risks? If us humans struggle generally at accurately predicting outcomes of biomedical science, why should there be a more accurate expectation of benefits as opposed to risks?

Pinker: No, this is wrong. Though it’s certain that the biomedical research enterprise as a whole will deliver benefits to billions of people, we can have no such confidence in particular technologies. That’s why we need a diversified research portfolio, without arbitrary bans. If you ban something, the probability that people will benefit is zero. If you don’t ban it, the probability is greater than zero.

As for the potential harms, they are far too nebulous to justify a ban or moratorium. Far from being confident in the power of gene editing, I’m on the record as being skeptical that we’ll ever see genetic enhancement of babies—the outcome that the prohibitionists and moratoristas dread and that many bioethicists blithely assume is inevitable. (For example, in 1999 Caplan announced that before the end of this century “We will see many children made by the artificial creation of embryos…This prediction is 100 percent certain.”) The prophesy of designer babies ought to be a relic of the early 1990s, when people thought there was “A Gene For” this or that talent. We now know that heritable psychological traits such as intelligence and personality are the product of hundreds or thousands of genes, each with a tiny effect, many of which may have harmful effects as well, such as an increased risk of neurological disease or cancer. With each enhancement gene providing a nugatory benefit and a non-negligible risk, and with the editing process itself imposing risks, it’s unlikely that today’s morbidly risk-averse helicopter parents will take a chance at enhancing a child—they won’t even feed their babies genetically modified applesauce! And that’s assuming that such a procedure ever got to the point of clearing conventional safety hurdles, which is far from likely. Add these risks to the fantastic expense and tribulation of IVF compared to good old-fashioned sex, and one should conclude that widespread genetic enhancement is too unlikely a possibility to worry about. And that’s assuming we should worry at all. There is, in addition, the argument (from Savulescu, the transhumanists, and others) that if enhancement were ever feasible it would be a good thing, not a bad thing—or at least a matter of individual freedom rather than government coercion.

Now, the story is different for editing out disease genes. There are more ways that a complex system can break down than that it can work better, and it’s easier to fix a defect than engineer in an improvement. Also, the benefits are very different for preventing death and disease (huge) than for implementing an enhancement (minor). So the possibility that germ-line editing might prevent disease in the future is well worth exploring.

Knoepfler. 4. Congress recently held a public hearing on human germline modification and is considering a legal provision to block editing of human embryos. What do you think of having such a hearing and the possibility of a restrictive legal provision? You said to bioethics, “get out of the way”. Should we scientists say the same thing to lawmakers? Why?

Knoepfler. 5. There is likely to be a NAS meeting sometime late this year on human germline modification by such technology as CRISPR-Cas9 and mitochondrial transfer (3-person IVF) in the spirit of the 1975 Asilomar meeting. Do you think this new meeting will achieve positive outcomes such as a white paper that appropriately has a vision for the future? What if the consensus is for a moratorium? Could you support that?

Pinker: I’ll answer these together. I think that scientists should reiterate the principle that no experiment should be permitted which imposes an unreasonable risk of an illness or birth defect on an individual. But no, I don’t think that scientists should support a ban or moratorium on germline genetic editing, for the reasons I set out in my answers to questions 2 and 3. Though the Asilomar recommendations have long been a source of self-congratulation among scientists, they were opposed by a number of geneticists at the time, who correctly argued that they were an overreaction which would needlessly encumber and delay important research. And the journalist Victor McElheny reminds us that the recommendations sowed a panic which came perilously close to shutting down some of the nation’s major laboratories, a danger he argues we are now in danger of repeating.

That having been said, I recognize that the political arena follows different rules than scientific and intellectual discourse. The scientists who lead major research institutions and deal with politicians and other public figures have to master the arts of compromise, tact, euphemism, and strategic deal-cutting. That’s how democracy works, and I’m grateful to the scientific leaders who carve out a space in which the rest of us can flourish. There are things they may believe but can’t say. But it’s important that someone says them, and that’s how I see my role in these debates.

ORIGINAL: Ipscell

August 10, 2015

28 Hot Plasmid Technologies from 2015

At Addgene we're continually impressed with the amazing plasmid technologies developed by our community of depositors. With over 40,000 plasmids avaliable in the repository, we can't give all of them attention they righlty deserve, but, in this post, we'll provide a small sampling of the many amazing new plasmid tools that have come through our doors in 2015. Do you have a favorite new plasmid tool from the past year? Let us know about it in the comments or shoot us an e-mail at blog@addgene.org and maybe we can write a blog post about it!

Split Cas9 System

The Cas9 protein is composed of an n-terminal DNA recognition domain and a mostly c-terminal nuclease domain. Feng Zhang's group utilized the bi-lobed architecture of Cas9 to engineer a series of "split" cas9 molecules that cannot function in isolation but form a fully functional Cas9 upon dimerization.

Splitting wild-type Cas9 into n-terminal (Cas9(N)-2xNLS) and c-terminal (Cas9(C)-2xNLS) fragments facilitates target DNA cleavage upon co-expression and spontaneous self-assembly. In an effort to obtain even more precise temporal control of gene knockout or activation, the c-terminal cas9 fragment was fused with FK 506 binding protein 12 (Cas9(C)-FKBP-2xNLS) and the n-terminal cas9 fragment with FKBP rapamycin binding domain of mTor (Cas9(N)-FRB-NES) resulting in a rapamycin-inducible Cas9 for genome editing. Without rapamycin treatment, the Cas9(N)-FRB-NES fragment is actively shuttled out of the nucleus due to the nuclear export sequence. Treatment with rapamycin induces Cas9(N)-FRB-NES and Cas9(C)-FKBP-2xNLS dimerization and net influx into the nucleus, where the functional Cas9 molecule can cleave the target DNA. The inducible split cas9 approach can also be used for activation of specific genes using dCas9-VP64 activator fragments (dCas9(C)-FKBP-2xNLS-VP64 and dCas9(N)-FRB-NES). This system provides users greater temporal control over CRISPR/Cas9 mediated genome modification and gene expression.

The plasmids associated with this article can be obtained through Addgene:

Rapamycin-inducible Cas9 sets (Addgene plasmids 62883 &62884; 62885 & 62886)

Rapamycin-inducible dCas9-VP64 activator set (Addgene plasmids 62887 & 62888)

Zetsche et al., Nat Biotechnol. 2015 Feb 2;33(2):139-42. doi: 10.1038/nbt.3149.

Light-Inducible CRISPR-Cas9 System for Endogenous Gene Activation

Optogenetics is a powerful tool that utilizes light to control and monitor individual living cells in order to understand how they work. Light activation allows scientists to spatially and temporally control which genes are turned on or off in a given area and the can do so in a very specific, precise manner. Previously, scientists have been successful in regulating gene transcription using DNA-guided optogenetic tools; however, targeting the necessary light-activated protein domains to the appropriate locus has been cumbersome.

To overcome many of the limitations intrinsic to DNA-guided systems such as TAL effectors or Zinc Finger Nucleases, the Gersbach lab has modified the RNA-guided CRISPR-Cas9 system to create a tool that is quick, versatile, and robust. Dubbed the LACE system for light-activated CRISPR-Cas9 effector, Polstein and Gersbach fused the light-inducible protein domains CibN and Cry2 to inactive dCas9 and VP64, respectively. CibN and Cry2 form a heterodimer in response to blue light, which will ultimately co-localize the VP64 transactivator with a dCas9 that has been targeted to a very specific site on the genome via the CRISPR system. The ease and flexibility of the LACE technology makes this system widely accessible for many potential applications.

Polstein LR & Gersbach CA, Nat Chem Biol 2015 Mar;11(3):198-200.

SunTag System for Single Molecule Imaging and Inducible Gene Expression

Brilliant technologies adopt basic biological concepts and employ them in creative and innovative ways. The principle of protein multimerization is seen in many important biological contexts, such as the enhancement of transcriptional activation via binding of multiple copies of a transcription factor to a promoter and localization of a protein via the presence of multiple copies of targeting sequences. Scientists have adopted and employed this concept in both imaging and inducible gene expression studies, which are evinced by the Tet system and visualization of single molecules via targeted fluorescent molecules. The Vale lab andthe Weissman lab have teamed up and advanced technology in the fields of single molecule imaging and inducible gene expression by creating the SunTag system.

The SunTag system, named after the "stellar explosion SUperNova", is a synthetic scaffold that recruits up to 24 copies of a protein to a target polypeptide. Multimerization in the SunTag system occurs via antibody-peptide labeling; specifically, cognate peptide epitopes fused to the protein of interest are recognized and thus fluorescently labeled by scFv antibodies fused to sfGFP. This system amplifies the intensity of fluorescence signal and enables tracking of single molecules within living cells without affecting protein function, thereby creating a single-molecule reporter of intracellular processes. Multimerization via the SunTag system also shows strong upregulation of gene activation when fused to dCas9. In the dCas9-SunTag-VP64 system, dCas9 is fused to a scaffold containing epitopes recognized by scFV antibodies fused to VP64 transcriptional activation domains. This system enabled the specific recruitment of multiple copies of VP64 to the sgRNA-targeted gene, resulting in increased activation of endogenous transcription of the target gene.

The brilliant SunTag system plasmids are available through Addgene and are already very popular. Use these plasmids to increase the brilliance of your fluorescent signal and endogenous activation of your target gene!

Tanenbaum et al., Cell. 2014 Oct 8. pii: S0092-8674(14)01227-6.

Multi-Bit Genetic Memory

One of the goals of synthetic biology is to engineer cells capable of recording permanent "memories" of molecular events. These memories would be recorded in the cell's DNA under the right conditions, and could initiate specified cellular processes or be observed at a later time via DNA sequencing or other readout. Cellular memory is accomplished by the use of sequence-specific enzymes (e.g. nucleases, recombinases) which irreversibly excise or invert a specific chunk of DNA. However, a limitation to this strategy is that each "bit" of information must be controlled independently; that is, the signal to record one bit of information must not act on any of the other bits in order to achieve accurate and reliable multi-bit memory.

To this end, Chris Voigt and colleagues at the MIT Synthetic Biology Center have deposited a set of 11 phage integrases which act irreversibly on their cognate attB/attP sites with no cross-talk. These pairs are closely related to the lambda phage integrase system which is the basis for Gateway cloning. The combination of all 11 att pairs in series with unique spacer sequences (pMemoryArray) gives a DNA sequence capable of recording 11 bits (1.375 bytes) of information. This leads to 2048 (211) possible combinations of states. Each of the integrases is supplied on its own expression plasmid or in various combinations, while the cognate att sites are supplied as individual reporter constructs or as the fully assembled pMemoryArray.

Yang et al., Nat Methods 2014 Dec;11(12):1261-6. doi: 10.1038/nmeth.3147.

Bioluminescent pH Sensor

Understanding intracellular pH regulation is important as pH regulation has roles in many cellular functions, from endosomal trafficking to signaling pathways to the cell cycle. Good tools for measuring intracellular pH are required in order to determine the relationships between pH changes and cellular events.

To overcome the issues with fluorescent pH sensors, a set of luciferase-fluorophore pH fusion reporters, pRSETb-pHlashand pcDNA3.1+-pHlash, that use Bioluminescence Resonance Energy Transfer (BRET) were developed by the Johnson Lab .

Characteristics of the pHlash reporter protein include:

• H+ specific response
• cytoplasmic retention
• insensitivity to other ions

In BRET, A mutant Renilla luciferase, catalyzes the oxidation of its substrate, luciferin, which releases energy as a photon. The photon energy is transferred to the acceptor fluorophore, a circularly permuted Venus. In the case of pHlash, the signal from the fluorophore is pH-dependent. Signal is simultaneously acquired from the the fluorescent and luminescent siganls and a ratio is calculated (the BRET ratio), which allows for compensation of varying fusion protein expression levels.

Zhang et al., PLoS One. 2012;7(8):e43072. doi: 10.1371/journal.pone.0043072.

Photoswitchable Tools for Spatial and Temporal Control of Cell Events

Three years ago, Brian Kuhlman’s lab designed photoswitchable dimers by fusing the SsrA peptide with the light-sensitive LOV2 domain of Avena Sativa phototropin 1 (AsLOV2). In the dark, this peptide is caged by the asLOV2 domain and has reduced affinity for its binding target, SspB. In the light, the Jα helix of the asLOV2 domain unfolds allowing it to bind to SspB. This original Light Inducible Dimer (oLID) could be used to modulate biological processes, but it did not show large changes in binding affinity with light stimulation. Indeed oLID showed only a two-fold change in affinity for SspB1.

In order to create a more powerful LID, the authors recently used computational protein design, phage display and high throughput binding assays, to engineer photoswitchable dimers which show over a 50 fold change in binding affinity with light stimulation2. The Kuhlman lab created two improved Light Inducible Dimers (iLID), iLID nano and iLID micro, which differ from each other by their light/dark affinity range (130nM to 4.7µM for iLID nano and 800nM to 47µM for iLID micro; Figure 1). With this great affinity range, these LIDs have been shown to be useful tools for light-mediated subcellular localization in mammalian cell culture and reversible control of signalling pathways.

These LIDs are generalizable, versatile and powerful tools which allow you to easily and reversibly switch off and on your favourite signalling pathways (Figure 2). They are all available now at Addgene.

1 Lungu et al., Chem Biol. 2012 Apr 20;19(4):507-17. 10.1016/j.chembiol.2012.02.006.

2 Guntas et al., Proc Natl Acad Sci USA 2015 Jan 6; 112(1):112-7. doi: 10.1073/pnas.

CRISPR-Cas9 Optogenetic Tool for Activating Endogenous Genes

Moritoshi Sato's Lab has engineered CRISPR-Cas9 to create a photoactivatable transcription system. The tool allows scientists to use light to spatially and temporally control endogenous gene activation.

This system works by bringing together an anchor protein and activator protein to drive transcription. The anchor consists of dCas9 fused to CIB1 and is targeted to specific genes via user-defined guide RNAs. Upon stimulation with blue light, the complex recruits a CIB1 binding protein (a portion of CRY2) fused to the p65 activation domain, which activates transcription of downstream genes.

Nihongaki et al describe the targeted activation of endogenous genes and detail the time course of gene activation using this system in the article cited below. Scientists can obtain dCas9-trCIB1, CRY2PHR-p65, and other plasmids from this article through Addgene.

Nihongaki et al., Chemistry and Biology. 2015 Feb 19; 22(2):169-74.

High Efficiency Cloning for Enzyme Screening

Dr. David Ackerley’s lab at the University of Wellington, Victoria, has developed an adaptable bacterial expression vector which gives close to 100% cloning efficiency and gene expression. The pUCXKT vectorensures that all screened plasmids contain a gene variant* and is amenable to both high and low throughput screening applications. The system can be readily modified to use different antibiotic resistances, restriction enzymes and plasmid backbones.

pUCXKT contains a nonfunctional truncated version of the kanamycin resistance gene (missing the first two codons). Two stop codons are inserted between the truncated gene and the promoter to prevent leaky expression of this resistance. Unlike previous systems, pUCXKT does not result in a translational fusion of the antibiotic cassette to the gene of interest. The gene to be expressed is amplified using a gene-specific forward primer containing the desired restriction site (the MCS has several options) and a pUCX-specific reverse primer containing the missing codons from the antibiotic resistance cassette, a ribosome binding site, SacI restriction site and a linker region. The PCR product is then ligated into the MCS/SacI site in pUCXKT, removing the stop codons during backbone digestion and providing the missing codons for functional antibiotic resistance. Colonies with an insert in pUCXKT will therefore have resistance to both kanamycin and ampicillin (from the pUC19 backbone).

*False positives can occur in this system if a small contaminating PCR product formed from primer dimers and containing the missing codons from the antibiotic resistance gene is inserted. It is important that primers be designed to minimize the possibility of dimer formation.

Prosser et al., Biotechnol Lett. 2015. 37:383-389.

Toolkit for Constructing Fusion Proteins with Customizable N- and C-terminal Modules

Thanks to the lab of Tomás Santalucía, you can now take any open reading frame (ORF) and with a simple recombination reaction, generate a protein with cell specific expression containing a variety of N- and C- terminal tags. TheSantalucía Lab has developed a cloning toolkit based on the Multisite Gateway® technology from Life Technologies.

The MultiSite Gateway Kit is a customizable Gateway®-based toolkit that allows scientists to generate fusion proteins from any ORF available as a standard entry clone. ORFs from an ORFeome Gateway library or any PCR-amplified ORF cloned into pDONR221 can be expressed either without a tag or as multiple fusions with different N- and C-terminal tags. Additionally, by using an adapted destination vector with a heterologous promoter outside of the Multisite Gateway® cassette, you can specifically express your new chimeric protein in a variety of model systems.

The MultiSite Gateway Kit contains a set of 12 promoter-less Gateway® entry clones to be used with other entry clones encoding the ORFs of interest:

Buj et al., BMC Molecular Biology. 2013 August 20th; 14(1).

TRICK: A Method for Visualization of the First Round of Translation in Living Cells

A novel technique developed in the labs of Jeffrey Chao, Robert Singer, and Anne Ephrussi allows researchers to observe when and where translation occurs in both live cells and whole animals. The system utilizes the bulldozer-like quality of the ribosome during translation, which knocks off all mRNA binding proteins as it travels along the mRNA to produce the protein product. The system, cleverly termed TRICK, for TranslationRNA Imaging by Coat protein Knock-off, is made up of three components:

• a GFP protein that binds to the coding region of a reporter mRNA via a PP7 coat protein (NLS-PCP-GFP)
• a RFP protein that binds to the 3’ UTR of the reporter mRNA via an MS2 coat protein (NLS-MS2-RFP)
• the reporter mRNA, which contains the tethering sites for the fluorescent proteins and an inducible promoter (Plasmid 64543or 64542)

When the reporter mRNA is first transcribed, it is bound by both the GFP and the RFP proteins in the nucleus. During the first round of translation, the GFP proteins that are bound to the coding region are displaced and do not rebind because they are transported back to the nucleus (thanks to a NLS on these proteins). The RFP proteins, however, remain bound to the mRNA since they are tethered AFTER the stop codon. Thus a non-translated mRNA will appear yellow, but a translated mRNA will be red. This system achieves great specificity and resolution by using 6-12 phage PP7 coat protein binding sites and 24 copies of the MS2 bacteriophage coat protein binding sites to tether many copies of the FP-coat protein fusions to their reporter mRNAs. Using the TRICK plasmids, Halstead et al demonstrated that 91% of reporter mRNAs were untranslated in the nucleus, confirming the hypothesis that most mRNAs are not translated until they are exported to the cytoplasm. Furthermore, they find that during stress, mRNAs in P-bodies are translationally repressed whereas the nonsequestered pool goes on to initiate translation. These TRICK plasmids provide valuable tools for studying translation in living cells.

Halstead, et al. Science. 2015 Mar 20;347(6228):1367-671. doi: 10.1126/science.aaa3380

MuLE Vectors for Customized Lentivirus Production

Finding a lentiviral vector backbone with just the right combination of promoter, cloning sites, fusion proteins or tags and reporter genes can be nearly impossible. Fortunately, a collection of plasmids comprising the multiple lentiviral expression (MuLE) system from the laboratory of Ian Frew simplifies the process of creating complex polycistronic lentiviruses.

The MuLE toolbox contains a series of building-block vectors that are combined in a specified order using Multisite Gateway cloning. A desired insert is cloned using traditional restriction enzyme cloning to create an Entry vector containing a promoter for constitutive (CMV, SV40, SFFV) or inducible (CMV-TO) cDNA expression, sgRNA (U6) expression, shRNA/shRNA-miR30 constitutive (H1, U6, 7SK) or inducible shRNA-miR30 (CMV-TO) expression, or a promoterless construct. Pre-constructed Entry vectors containing Cas9, EGFP, mCherry, iRFP, tdTomato, luciferase, LacZ, puromycin or CreERT2 can be combined with custom-cloned vectors in a Multisite Gateway reaction with a desired destination vector expressing a separate reporter gene (EGFP, iRFP, IFP1.4, puromycin, neomycin or luciferase) to create a final lentiviral vector containing up to 5 different elements.

Lentiviral vectors constructed with the MuLE toolbox have been used for a variety of experimental purposes including (1) single or multiple, constitutive or inducible, gene knockdown with shRNA or shRNA-miR30, (2) tamoxifen-inducible CreERT2 gene deletion, (3) simultaneous expression of cDNAs and shRNAs with a reporter gene, or (4) multiple sgRNAs with Cas9 for genome engineering. The versatility and ease of generating complex combinations of genetic elements in a single lentiviral vector provides a powerful tool to modify mammalian cells.

Multicolor CRISPR Labeling Chromosomal Loci in Genome

Thoru Pederson’s lab at the University of Massachusetts Medical School has recently developed a CRISPR-based multicolor tool enabling the imaging of multiple endogenous genomic loci simultaneously in live cells. This system allows the study of loci proximity and of the dynamic interactions between intra- and inter-chromosomal domains. The authors anticipate that this tool can also be used to visualize translocations, genome rearrangements and cancer-associated chromosome shattering.

This method is based on the recognition of genomic loci by three orthogonal nuclease-inactive Cas9's (dCas9) from S. pyogenes, N. meningitidis and S. thermophilus which have been used for gene editing in human cells without cross-talk in cognate sgRNA binding. By fusing these dCas9s with one of three fluorescent proteins (GFP, BFP and mCherry), the authors were able to visualize simultaneously several parts of the genome and to determine, for instance, the intra-nuclear distance between loci on different chromosomes.

The advantage of having this kind of system that works in live cells is that one can now follow movement of two or more targeted loci during the cell cycle or cell differentiation. This tool should delight anyone interested in understanding the spatio-temporal regulation of human genes. The plasmids are all available at Addgene ready to light up the genome of your favorite cells.

Ma et al., Proc Natl Acad Sci U S A. 2015 Mar 10;112(10):3002-7. doi: 10.1073/pnas.1420024112. Epub 2015 Feb 23.

A Novel dCas9 Activator Capable of Unprecedented Activation of Target Genes

A group of researchers headed by the Church Lab at the Wyss Institute have extended the transactivating capabilities of CRISPR/dCas9-fusion proteins and have made their dCas9 transactivators available through Addgene.

The development of dCas9 and dCas9-fusion proteins has dramatically increased the potential applications of CRISPR/Cas9 in genome-engineering. One particularly attractive application involves using dCas9-activator fusion proteins to induce endogenous gene expression. Several studies have utilized dCas9 fused to a VP64 activator domain and targeted toward endogenous promoters to activate transcription. Chavez et al. 2015extended upon these findings by screening for additional activation domains that increase the expression of target genes when fused to dCas9. One surprising finding was that, although three of the additional domains tested (VP64, p65, and Rta) were capable of increasing target gene expression, adding all three domains onto dCas9 to generate a dCas9-VP64-p65-Rta fusion protein (referred to as dCas9-VPR) had a synergistic effect on target gene activation.

The novel dCas9-VPR activator is capable of enhancing target gene expression anywhere from 87x to 20,000x over endogenous expression (depending on the specific gene) and induction routinely reaches levels 20-40x greater than the original dCas9-VP64 activator. They also revealed that dCas9-VPR mediated gene activation is capable of promoting induced-pluripotent stem cells (iPSCs) to take on a “neuron-like” phenotype when targeting genes involved in neurogenesis, which had proven challenging using the existing dCas9-VP64 activator. The dCas9-VPR activation plasmids have great potential and have been adapted for use in mammalian cells, yeast, and Drosophila.

Synchronization of Secretory Protein Traffic in Mammalian Cells

The journey of secretory proteins, from their synthesis in the ER to their arrival in their target compartment (e.g. the plasma membrane, the extracellular space, or the lysosomes) can take many paths. Understanding how a protein traffics through these pathways is key to understanding, and potentially perturbing, its function. In order to dissect protein traffic in different systems, Franck Perez’s lab, from the Institut Curie in Paris, has developed theretention using selecting hooks (RUSH) system.

How does it work? RUSH is an ingenious two-state assay based on the reversible interaction between a hook protein stably localized in the donor compartment (e.g. the ER) and a reporter protein of interest (Figure A). Franck Perez’s lab has engineered hook proteins fused to a streptavidin core that are able to retain reporter proteins fused to the streptavidin-binding peptide (SBP). The addition of biotin disrupts this interaction and thus triggers a synchronous release of the reporters which can then be tracked along the secretory pathway (Figure B). The RUSH system has already been used to study transport characteristics of various Golgi and plasma membrane reporters as well as secretory proteins or proteins targeted to membrane sub-domains. It has also been used to observe intra- and post-Golgi segregation of cargo during their transport. Finally, the system can be adapted to cellular screening to identify molecules that can perturb protein transport.

Franck Perez’s lab has developed a whole collection of hook and reporter proteins that can be used to test diverse secretory routes in various conditions. The plasmids and lentiviral vectors encoding these proteins can also be used with your own reporters to decipher their trafficking routes. This collection is now available at Addgene, so on your marks...get set...RUSH.

Boncompain et al., Nat Methods 2012. Mar 11;9(5):493-8.

MXS Cloning Platform for Imaging and Flow Cytometry Applications

Designing customized fluorescent reporters can be challenging enough, but incorporating multiple reporters into a single expression plasmid using traditional cloning strategies may seem preposterous. Luckily, the Pierre Neveu laboratory designed a new iterative, chaining-based cloning method that simplifies the process of constructing specialized fluorescent reporter plasmids with MXS-chaining.

Analogous to modular assembly methods, such as BioBricks, the MXS-chaining method uses specified restriction enzyme sites to combine two modules via compatible overhangs produced by the enzymes, such that the ligation regenerates the original restriction sites for further iterative cloning steps and the joined modules cannot be separated again by the same cloning enzymes. MXS (MluI-XhoI-SalI) chaining results in a translatable ligation scar (Val-Glu) between each module to permit easy construction of fusion proteins. These three enzymes were chosen because they cut human and mouse coding regions infrequently; any MluI, XhoI or SalI sites must be removed by site-directed mutagenesis in order to use the module as a building block for further chaining.

The MXS-chaining kit contains 5 empty chaining vectors plus a destination vector and a collection of building blocks including 14 different fluorescent proteins, 8 constitutive promoters, 2 includible promoters, 3 polyA terminators and various pieces for inducible expression, loxP sites, H2B, P2A or PEST2D sequences. Several pre-constructed promoter-selection marker or promoter-inducible expression related vectors are also ready for further chaining.

Plasmids constructed using MXS-chaining were used in a variety of experiments including subcellular visualization of 6 different organelles, a cell cycle indicator for mouse embryonic stem cells and a series of inducible promoter constructs. Up to 34 building blocks were used to construct 20kb long inserts. The easily reusable MXS-building blocks are particularly effective at generating plasmids with varying numbers of repeated sequences or inserts, which can be advantageous for imaging or flow cytometry experiments.

Sladitschek HL and Neveu PA. PLoS One. 2015. Apr 24;10(4):e0124958.

LOC8-ing Mitochondrial Fusion with a New Tri-chromatic Assay

Georg Ramm’s lab at Monash University has developed a unique tool for studying mitochondrial fusion using UV light to differentially photo-label multiple mitochondrial populations within the same cell. Mitochondrial fusion is typically assayed in living cells by using mitochondrially-targeted photoactivatable fluorescent proteins (FPs). A subpopulation of mitochondria is selectively labeled by irradiating the photoactivatable FP, then the overlap between labeled and unlabeled mitochondria, indicative of mitochondrial fusion, is quantified. This strategy relies on the current range of mitochondrially targeted photoactivatable and photoswitchable FPs that are activated by UV light (300-400nm) and that fluoresce in the green (500-550nm) and red (570-620nm) portions of the spectrum.

PhD student Benjamin S. Padman recently developed a mitochondrially-targeted variant of PSmOrange, a photoswitchable fluorescent protein which transitions from emitting orange (λ= 565nm) to far-red (λ= 662nm) fluorescence after exposure to blue-light (480nm). He also created an improved synthetic modular version of the COX8 leader sequence called Leader Of Cox8 Repeated (LOC8R) to more accurately target the mitochondria. The synthetic LOC8R sequence enhances mitochondrial targeting of PSmOrange, as well as other photoactivatable proteins like PAGFP. Co-expression of LOC8R-PSmOrange and LOC8R-PAGFP provides a novel strategy for assaying the dynamics of different mitochondrial subpopulations within the same cell; by photoswitching PSmOrange and photoactivating PAGFP in separate subpopulations of mitochondria, one can monitor how the two separate populations move and interact over time. A Ramm lab plasmid with an N1-vector-compatible LOC8R sequence is also available, N1-LOC8R. 

These plasmids are unpublished

Unnatural Amino Acid Incorporation in Mammalian Cells

Peter Schultz and colleagues at the Scripps Research Institute have deposited a pair of plasmids for the cotranslational incorporation of unnatural amino acids (UAAs) into proteins in mammalian cells. Each tRNA/aminoacyl-tRNA synthase pair is encoded on a single plasmid and facilitates the incorporation of a variety of bio-orthogonal UAAs via a re-assigned stop codon (TAG). This polyspecificity enables the site-specific introduction of many different functional groups, including reactive groups for conjugation reactions, fluorescent amino acids, posttranslationally modified amino acids, photoaffinity probes, and more, to your protein of interest using the same vector system. The tRNA/aaRS pairs are delivered via a pseudotyped baculovirus system, which is advantageous compared to other viral systems due to its large cargo capacity, minimal cytotoxicity, and broad host-tropism. These tools open up many powerful avenues through which one can examine molecular events with exceptional precision in a native context.

Chatterjee A et al. Proc Natl Acad Sci U S A. 2013 Jul 16;110(29):11803-8.

Plasmid Tools for All-optical Electrophysiology

A major challenge in the field of all-optical electrophysiology is designing the perfect voltage indicator and optogenetic actuator pair. Ideally, indicators should possess fast voltage sensing kinetics and have no spectral overlap with the optogenetic actuator. In 2012, Adam Cohen's lab presentedArchaerhodopsin 3 (Arch) as a new rhodopsin based voltage indicator for use in the detection of neuronal action potentials. The Cohen lab wished to improve on the Arch indicator and their collaborators in Robert Campbell’s lab carried out five rounds of random mutagenesis on a library of Arch mutants screening for improved brightness. The brightest mutants were then further mutagenized and screened for speed and voltage sensitivity. This hierarchical screen led to the generation of two enhanced voltage indicators: QuasAr1 and QuasAr2 (quality superior to Arch). Both QuasAr1 and QuasAr2 possess mutations for improved voltage sensitivity and speed, and an endoplasmic reticulum export motif and trafficking sequence for improved targeting to the plasma membrane.

To obtain a more sensitive channelrhodopsin actuator for triggering action potentials at low light intensities, the Cohen lab further modified sdChr- a blue-shifted channelrhodpson from freshwater green alga identified by Ed Boyden’s lab. The enhanced sdChr construct, named CheRiff, exhibits high light sensitivity and fast kinetics. By combining both the voltage indicator, QuasAr, and actuator, CheRiff, in the bicistronic Optopatch vector, the Cohen lab has provided the field with an improved tool for electrophysiology, minus the use of electrodes. The Cohen lab has made the original Arch indicator (pJMK004), as well as the CheRiff, QuasAr, and Optopatchvectors available to the scientific community through Addgene.

Hochbaum, et al. Nat Methods. 2014.

Kralj, et al. Nat Methods. 2011.

High-Throughput Luciferase Reporter System for Detection of pre-mRNA Splicing

The Gideon Dreyfuss lab has developed rapid-response luciferase (firefly P. pyralis) reporter plasmids for use in high-throughput screening of pre-mRNA splicing. This reporter system can be used for identifying previously unknown factors and pathways involved in pre-mRNA splicing in a compound screen.

Their reporter system is comprised of two plasmids: Luc (intronless; CMV-LUC2CP/ARE and Luc-I (intron-containing; CMV-LUC2CP/intron/ARE). The Luc-I intron is a chimeric β-globin/immunoglobulin intron that has been commonly used in constitutive splicing studies and has been optimized for high efficiency splicing. If intron splicing does not occur in Luc-I, the produced luciferase is truncated and lacks enzymatic activity. The Luc plasmid (intronless) is useful for counter-screening, i.e. to filter out any compounds that are affecting other global cell processes, like translation.

This luciferase reporter system was also designed for a short screening time (<4 hours) in order to avoid complicating effects from global toxicity due to loss of splicing. To this end, destabilizing sequences were added to shorten the half-life of both the luciferase mRNA (3’UTR AUUUA [5 consecutive]) and protein (C-terminal CL1 & PEST). These modifications ensure that any signal from full-length luciferase produced prior to the start of the screen is quickly removed. This system should help researchers further elucidate the many factors that affect alternative splicing in mammalian cells.

Younis, et al. Mol Cell Biol. 2010 Apr;30(7):1718-28.

New & Improved Light-Switchable Bacterial TCSs

Jeffrey Tabor's lab has developed new synthetic biology tools by engineering light-switchable sensors developed from bacterial two-component signal transduction systems (TCSs). TCSs are utilized by bacteria to sense and respond to their environment - typically through a histidine kinase, which triggers phosphorylation of the response regulator and downstream transcription activation or repression. The TCS light sensors described in ACS Synthetic Biology (2014) represent the newest versions of both a red- and a green-light switchable bacterial TCS system that has been improved through engineering efforts over the years. The authors optimized these systems by: 1) decreasing the number of plasmids required for each input/output system; 2) removing all chemically inducible promoters; 3) improving the strength of the promoters used; and 4) optimizing expression of both the light sensor and the response regulator. These changes reduced system leakiness and increased the dynamic range, resulting in more user-friendly and tunable tools for controlling gene expression.

For more details about these light-switchable TCSs, read our blog post: Synthetic Photobiology: Optogenetics for E. coli.

Schmidl, et al. ACS Synth Biol. 2014 Nov 21;3(11):820-31.

Inncredible New Tools for Epitope Tagging: Inntags

When fused to your protein of interest (POI), epitope tags give you the ability to monitor POI expression via western blot, to purify your POI, to perform pull down experiments like co-IPs and ChIP, and more - all without the need to develop custom antibodies. Unfortunately, while there are many types of epitope tags (eg. HA, MYC, Flag), they can be deleterious and cause aberrant protein folding, expression, or even affect cell viability. In theirrecent work in Nature Methods, Georgieva et al describe the rational production of a new series of epitope tags, inntags (innocuous tags), that they believe will have fewer effects when fused to your POI.

Georgieva et al developed the inntags via a bioinformatics screen for proteins with known structure and characteristics favorable for innocuous tags including high solubility, limited affinity for other proteins, ordered structures, and accessibility to an antibody. After identifying possible protein domains containing these properties, they generated monoclonal antibodies against them, fused them to GFP, and compared them to popularly used epitopes tags such as HA and MYC in vivo. The authors found that fusions to one particular tag, the IT6 tag, interacted with fewer other proteins, had fewer effects on yeast cell viability, had less of an effect on GFP diffusion, properly localized in mammalian cells without aggregation, and were successfully used in such techniques as western blots, immunoprecipitation, and immunofluorescence.

Marti Aldea's lab has graciously made IT6 and other inntags available through Addgene so you can try these incredible new tools yourself! The tags can be found in the pBluescript II SK+ vector here.

For more information on inntags, read our recent blog post.

Georgieva, et al. Nature Methods. 2015 Aug 31.

CRISPR-Display: A Novel CRISPR technology for Targeting RNA Cargos to Specific DNA Loci

John Rinn’s lab is interested in understanding the role of long noncoding RNAs (lncRNAs) in human health and disease. A primary question in their quest to understand lncRNAs: What can lncRNAs do by themselves? The Rinn lab has developed a novel technique, called CRISPR-Display, that may help answer this difficult question. CRISPR-Display uses the dCas9 enzyme to deploy large RNA cargos to specific DNA loci. In CRISPR-Display, your RNA of interest is fused to an sgRNA used to direct Cas9 to a specific genomic locus. When fused to the sgRNA and co-expressed with dCas9 – the dCas9, sgRNA/RNA cargo complex localizes to the sequence targeted by the sgRNA. This allows scientists to target RNA-based protein-binding cassettes, artificial aptamers, pools of random RNA sequences, and natural lncRNAs to specific locations in the genome. Furthermore, these different RNA-based functions can be multiplexed using a shared pool of dCas9.

To show that their sgRNA-RNA fusions were functional, Shechner et al used the constructs in their study to target various RNA molecules - such as MS2 stem loops, PPL stem loops, the Spinach2 aptamer, and more - to sequences on a Gaussia luciferase reporter vector. dCas9 or dCas9-VP64 (a control that is known to enhance gene expression) were then cotransfected with these constructs, providing the machinery that brings the sgRNA/cargo RNA to the appropriate DNA locus. The sgRNA/RNA fusions retained their appropriate functions affecting luciferase reporter expression as anticipated. For instance, an aptamer designed to bind a transcriptional activator activated gene expression from the reporter construct. The sgRNA targeting Gaussia luciferase can be easily replaced with an sgRNA targeting your sequence of choice. Many of the available sgRNA-cargo constructs utilize a U6 promoter and contain the RNA cargo cassette inserted within the sgRNA core (ie an INT construct), as the authors found that, in many situations, this was the ideal promoter/location combination. However, other constructs which contain different promoters and/or allow for insertion at the 5′ or 3′ end of the sgRNA (i.e. TOP1 or TOP2 constructs; see Supplementary Note 3 in the article for more info) are available. Furthermore, there is a general purpose cloning vector pU6_(Gluc)_INT(GenPurpClon) which can accept a novel RNA sequence of interest within the INT location. One plasmid to make note of is the INT construct bearing the "Bunch of Baby Spinach"aptamer, which is a brighter version of Spinach2, an RNA aptamer that fluoresces upon binding of a cell-permeable dye. Bunch of Baby Spinach, or BoBS, consists of three tandem copies of the Baby Spinach core embedded in a single, extended stem-loop, contiguous with the sgRNA core (see image above).

Shechner, et al. Nat Methods. 2015. PubMed PMID: 26030444

Fluorescent Fusion Proteins for Understanding Cellular Dynamics

The Voeltz Lab has deposited several new fluorescent protein fusions that can be used for monitoring cellular dynamics. These fusion proteins have been used as subcellular markers to visualize the dynamics and interactions between the endoplasmic reticulum (ER), mitochondria, cytoskeleton, and endosomes. These fusion proteins can act as markers for visualization of subcellular structure, for performing live-cell imaging experiments (e.g. fluorescence recovery after photobleaching [FRAP]) for studying membrane dynamics, for monitoring cargo trafficking, and for discovering organelle contact sites.

This suite of deposited fluorescent fusion proteins can visualize:

Cell Structure	Plasmids:	Cell Structure	Plasmids:
ER	mCherry-Sec61 beta BFP-Sec61 beta BFP-KDEL Rtn4a-GFP (tubular ER)	Microtubules	mCherry-alpha-tubulin
Early Endosome	Vacuolar Compartment mCherry-Rab5 BFP-Rab5 GFP-Rab5B Vacuolar & Budding Compartment GFP-Rab4B	Mitochondria	mito-BFP mCherry-Drp1 (GTPase in division) GFP-Mff (outer membrane protein)
Late Endosome	mCherry-Rab7A GFP-Rab7A

Friedman, et al. J Cell Biol. 2010. PubMed PMID: 20696706

Friedman, et al. Science. 2011. PubMed PMID: 21885730

Friedman, et al. Mol Biol Cell. 2013. PubMed PMID: 23389631

Rowland, et al. Cell. 2014. PubMed PMID: 25416943

Shibata, et al. J Biol Chem. 2008. PubMed PMID: 18442980

Zurek, et al. Traffic. 2011. PubMed PMID: 20955502

CRISPR Tagging Techniques for Studying Protein Interactions in Their Endogenous Context

Researchers commonly use tandem affinity purification (TAP) followed by mass spectroscopy to determine interaction partners for a protein of interest (POI). This technique requires the production and expression of fusions between the POI and a TAP tag (a tag with two epitopes that can be used to sequentially purify the POI along with any proteins bound to it). Classically it has been easier to overexpress the POI-tag fusion from a plasmid or from a random genomic location than to express the tagged protein from a defined genomic location or from its endogenous locus. In their recent Cell Reports paper, researchers from the Doyon Lab show that, with the many nuclease based genome editing techniques out there (and particularly CRISPR), it is now relatively easy to design constructs and repair templates to insert TAP tag fusions directly into the genome. In this work, they use AAVS1_Puro_PGK1_3xFLAG_Twin_Strep and nuclease driven recombination to insert TAP-tagged proteins into a well-defined “safe harbor” site in the genomic locus of the PPP1R12C gene in single copy. They show that these fusions can be purified, re-capitulate interactions observed in previous studies, and provide new insights into the make-up of multiprotein complexes like the EPC1 complex.. Going one step further, they also show that fusions can be targeted to their endogenous loci using validated gRNAs from Addgene and well designed homology directed repair templates (see the supplementary info from the paper for details on repair template design). Because these fusions are regulated in their endogenous context, they should be expressed at similar levels and under similar conditions as their wild-type counterparts - a real boon to those trying to understand physiological interactions and functions of their POIs.

Dalvai, et al. Cell reports. 2015. PubMed PMID: 26456817.

Nano-lanterns for Multicolor Luminescence Imaging 

Fluorescent imaging techniques have become indispensable tools for molecular and cell biologists over the last two decades, but their use is sometimes limited by the drawbacks of autofluoresence and photobleaching, as well as the need for external light activation. In order to overcome these limitations, Takeharu Nagai and colleagues at Osaka University have developed two new color variants of their Nano-lantern technology, first developed in 2012. The new cyan and orange Nano-lantern fusions are designed to complement the original yellow variant, with all 3 having distinct emission spectra suitable for single cell, multi-color imaging. The Nano-lanterns rely on the principle of bioluminescence resonance energy transfer (BRET), whereby photons emitted by a Renilla luciferase variant are used to excite a fused fluorescent protein, eliminating the need for an excitation light source. Instead, coelenterazine is supplied to the culture media as a chemical substrate for Renilla luciferase. This approach is especially suited to complement the use of optogenetic tools, where light used for excitation can cause unintended activation of the optogenetic system. In their 2015 PNAS manuscript, Takai et al validate the Nano-lantern approach as a tool for monitoring multi-gene expression and adapt it for multi-color calcium sensing (plasmids coming soon!).

Takai, et al. PNAS. 2015. PubMed PMID: 25831507.

Saito, et al. Nat Commun. 2012. PubMed PMID: 23232392

Testing Your Cell-free Expression System: There’s a Kit for That!

Cell-free expression systems have been improved for both eukaryotic and prokaryotic expression in recent years, with many different systems now available. However, it can be difficult to compare their performance, as most translation initiation sequences are optimised for a particular system. The Alexandrov lab at the University of Queensland’s Institute for Molecular Bioscience has provided a solution for this problem by developing a general translation initiation sequence known as the Species-Independent Translational Sequence (SITS). This sequence bypasses the 5′ mRNA cap required for eukaryotic systems and instead directly engages the ribosome for translational complex assembly. PhD student Dejan Gagoski then used SITS to create the pCell-Free vectors, Gateway-compatible backbones which enable cell-free expression of proteins in both prokaryotic and eukaryotic cell extracts (Gagoski et al 2015). He has also constructed a library of eGFP-tagged human ORF clones to allow testing and comparison of different cell-free expression systems; the Cell-free expression test kit represents a set of 88 clones in a pCellFree vector that enables protein expression in any in vitro translation system. The gene set was carefully chosen to perform statistically relevant benchmarking of cell-free expression systems, and tested for product integrity, expression level, and aggregation propensity in four expression systems: E. coli, wheat germ, HeLa, and Leishmania (Gagoski, et al 2015[2]). The proteins encoded in this set range in size from 4 to 156 kDa, enabling the user to characterise the efficiency of their cell-free system in correlation to the size of the product. Analysis of protein size and expression level can be conveniently performed through the N-terminal eGFP tag carried on all constructs.

pXPG: An Alternative to the Widely Used Luciferase Reporter Gene Plasmid pGL3

pGL3 luciferase reporter gene plasmids have been used extensively by many labs to study promoter regulation in live cells since their introduction by Sherf and Woods in the nineties. However, reliance on pGL3 has not been without problems due to the high rate of read-through transcription originating from cryptic promoters in the plasmid backbone (Giannaks et al 2003, Bert et al 2000). In 2000, Peter Cockerill and his lab developed an improved luciferase reporter gene plasmid, pXPG, which shows lower background expression than pGL3, enabling its use for the study of promoters with either weak or high activities (Bert, et al. 2000).

To design pXPG, the authors incorporated a high-copy origin of replication and a modified luciferase gene into apXP1-derived vector that more efficiently blocks read-through transcription in eukaryotic cells. pXPG contains the Luc+ luciferase gene derived from pGL3 but has a distinct advantage over the latter plasmid as it contains a duplicated SV40 polyadenylation region instead of the synthetic polyadenylation signal present in pGL3. This appears to contribute to more efficient blocking of read-through transcription for pXPG. The other advantage of pXPG over pXP1 is that its new origin of replication increases plasmid copy number in E. coli. The authors used pXPG to study the human GM-CSF promoter and enhancer, a finely regulated promoter controlled by a mixture of transcription factors and chromatin remodeling events, demonstrating the utility of the pXPG construct in understanding complex gene regulation (Bert et al 2000, Johnson et al 2004).

So if you are looking for a luciferase reporter plasmid that provides a more sensitive means of studying promoter function, pXPG is an excellent choice. And guess what? pXPG is now available at Addgene.

Sherf, et al. 1994. Promega Notes Mag.

Giannakis, et al. BioTechniques. 2003. PubMed PMID: 12951775

Bert, et al. Plasmid. 2000. PubMed PMID: 10964627

Johnson, et al. Mol Cell Biol. 2004. PubMed PMID: 15340054

Imaging RNA in Bacterial and Mammalian Cells with Broccoli

Vegetable tags, including Spinach, Spinach2, andBroccoli, can benefit your RNA experiments in several ways. The laboratory of Samie Jaffrey describes using these tags in an in-gel RNA visualization technique with RNA derived from bacterial and mammalian cells. Broccoli is an RNA aptamer that acts as a GFP mimetic and induces fluorescence in the presence of DHFBI, a small molecule fluorophore. When fused to an RNA of interest, Broccoli-DHFBI fluorescence can be used to visualize the fusion in vitro or in vivo. The protocolinvolves isolating total RNA, resolving by PAGE and then staining the gel with DHFBI to selectively visualize the Broccoli-tagged RNA. This technique is faster and less labor intensive than northern blotting.

To improve folding of the Broccoli apatamer, Filonov and colleagues engineered a new scaffold, F30, based on the Phi29 viral RNA junction motif, that increases RNA stability and avoids processing of the scaffold in mammalian cells into unexpected cleavage products. This scaffold allows insertion of two dimeric Broccoli tags (2xdBroccoli) to further enhance fluorescence of the tagged RNA. Bacterial and mammalian expression vectors for the F30-2xdBroccoli are available, as well as 5S RNA F30-2xdBroccoli fusion and pET28c-F30-Broccolicontrols.

Don’t just eat more broccoli, use Broccoli tags in your experiments too!

Filonov, et al. Chem Biol. 2015. PubMed PMID: 26000751

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ORIGINAL: Addgene

by Tyler Ford

Dec 23, 2015