Synthetic biology

Cellular Control – at the Flick of a Light Switch

What if you could turn on an enzyme inside a living cell—or release a cellular factor from its anchor—with the flick of a light switch?

Researchers at the University of Alberta’s Department of Chemistry have developed a new tool for manipulating biochemical processes within cells using light. By applying the unique properties of a photoconvertible fluorescent protein called mMaple, the team created such a light switch, a photocleavable protein called PhoCl (pronounced “focal”).

mMaple, whose name was inspired by the green-to-red color change of maple leaves as seasons transition, undergoes a light-dependent conformational change. Dr. Robert E. Campbell’s team engineered PhoCl to cleave into two pieces when exposed to light.

This novel optogenetic tool is especially useful for applications that involve manipulating cellular processes. For example, PhoCl can be used to create “caged” proteins that will not become activated until exposed to light. Researchers link one terminus of PhoCl to a cellular enzyme and the other terminus to an inhibitor, “caging” the enzyme and preventing it from performing its function. Upon exposure to violet light, PhoCl is cleaved to separate the inhibitor from the enzyme, thus activating the enzyme at the user’s command.

The cleavage mechanism of PhoCl is particularly useful for the activation of proteins within a specific location of a cell. Because intact PhoCl is fluorescent, researchers can visualize its location and movement within the cell and have control over when it cleaves. Upon cleavage, the fluorescence is quenched, enabling users to visually determine where the event took place.

As Allele Biotechnology & Pharmaceuticals is a licensed distributor of plasmids containing the gene for mMaple, the development of PhoCl is particularly exciting news to us and our customers. Interested readers can learn more about PhoCl in their paper published in Nature Methods.

Wednesday, March 22nd, 2017 Fluorescent proteins, Synthetic biology No Comments

Visualizing Endogenous Synaptic Proteins in Living Neurons

The recently published method is based on the generation of disulfide-free “intrabodies”, a structure from the 10th fibronectin type III domain known as FingRs. These affinity molecules were fused to GFP for direct fluorescence miscroscopy. The FingRs do not need di-sulfite bonds and are therefore better folders in mammalian cells. Specifically, a library was screened with in vitro display to identify FingRs that bind two synaptic proteins, Gephyrin and PSD95. After the initial selection, the researchers from USC secondarily screened binders using a cellular localization assay to identify potential FingRs that bind at high affinity in an intracellular environment. As it turned out, only 10-20% of the original positive clones bind well inside the cells, suggesting this type of further screening was a critical step.

The expression of intrabody is transcriptionally regulated by the target protein through a ZFN-repressor fusion. This transcriptional control system matches the expression of the intrabody to that of the target protein regardless of the target’s expression level. This design virtually eliminates unbound FingR, resulting in very low background that allows unobstructed visualization of the target proteins. As result, the FingRs presented in this study enabled live cell visualization of excitatory and inhibitory synapses, and apparently without affecting neuronal function.

Technically, the reason to use in vitro mRNA display was required by the need to use a large library (>10exp12, beyond the limit of the more commonly used phase display) to find good binders. A similar visualization system can be established using more potent affinity domains such as the VHH single-domain antibodies that have only one, sometimes dispensable, di-sulfite bond. The VHH domain nanobodies can be more easily isolated from camelid animals. Another improvement to the visualization system can be made by using stronger, superresolution-ready FPs such as mNeonGreen or mMaple to enable single molecule imaging, which is particularly interesting for studying synapses and applied to the BRAIN initiative.

Gross et al. Neuron, June 2013,

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Genome Modification—a Practical Approach

The ability to modify genomes has always been fervidly sought after by molecular, developmental biologists and geneticists as it would provide them with the means for finding out what a particular piece of the genome may do in the biological process they are studying. The discovery of naturally existing P-element helped a generation of Drosophila geneticists and made the fruit fly a prime model system for gene function studies in the 80’s and 90’s. But P-elements inserted at uncontrolled sites, making it essentially a gene transfer vehicle without much control. The introduction of prokaryotic recombination systems, e.g. LoxP and Cre, provided researchers with tools to obtain more control of the inserted genes in a host chromosome during a biological process such as development. Transposons like Sleeping Beauty, Piggybac, or Tol2 made similar experiments possible in mammalian cells.

Still, the randomness of transposon-type elements’ insertion, much like retrovirus or lentivirus, could cause trouble if they land in an undesirable spot. Methods of inserting transgenes only in well-known, harmless, and transcriptionally active regions, so called “safe harbors”, were subjects of interest of researchers and NIH grant topics in the past couple of years under “directed genome editing”. Gene knock-out or knock-in can be achieved through vector-mediated homologous recombination such as the rAAV genome engineering system and the “TARGATT” system, which are commercially available as kits or services.

However, instead of inserting an exogenous gene, it is often highly desirable to modify an endogenous genome sequence, which requires the modification apparatus to first recognize the target sequence. ZFN and TALEN both recognize DNA targets through specific nucleotide binding protein domains, with TALEN having more flexibility if assembled in a “Lego”-like format because each domain can specifically recognize a “C”, “G”, “T”, or “A” base. The description of using CRISPR/cas system in a recent burst of publications opened up new ways of binding to specific DNA sequences and nicking or severing the dsDNA. This system does not require engineeredDNA binding domain assembly; instead, it uses a guide RNA to find the target DNA sequence to direct endonuclease, in a sense quite like RNAi. However, the enthusiasm about CRISPR/cas was somewhat dampened by a report last month in Nature Biotechnology that reported off-target effects of CRISPR/cas was much higher than ZFN and TALEN. Particularly, if mismatches are located in the 5’ portion of the guide RNA targeting sequence, they can be well tolerated up to 3 or 4, even 5 mismatches. Unfortunately this is also similar to the tolerance of the RNAi matching region outside the core 12-base region. The difference is: for RNAi, the off-target damage is temporary and ignorable if the extent is insignificant compared to the effects on the intended target while for CRISPR/cas, an off-target cut on the chromosome is permanent.

On the positive side, in an even more recent publication in Nature Methods, mutant strains of C. elegans were obtained using the CRISPR/cas system and no evidence was found for off-target changes, at least not in an overwhelming fashion. Much value of the estimates of off-target effects relates to the methods used for analysis. Currently, most of the studies looked at potential off-target sides by searching for partial matches. In the future, whole genome sequencing will be increasingly required for submitting such publications.

On a practical note, if you intend to take a dive and try to use any one of these methods, your number one problem will be that none of the methods will result in 100% modification even if you can ignore the off-target problems for now. Therefore, many of our customers ask about a screening strategy. One could use traditional drug selection and fluorescent protein (FP)-based sorting, but these can only help you find cells that are successfully transfected with the ZFN, TALEN, or CRISPR/cas expressing DNA molecules, not necessarily having the genome modification result. We have formulated the idea of inserting the target site into an FP-bearing plasmid as a surrogate target cutting indicator, and use another FP to track transfection of the TALEN plasmid. Nonetheless, in the end, PCR-amplifying the target region of the chromosome and doing either an enzymatic mismatch detection assay (e.g. T7 endonuclease) or sequencing is the only way to know for sure whether genome editing has occurred.

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Wednesday, July 10th, 2013 Synthetic biology, Viruses and cells 1 Comment

The Power of Cas

Precise engineering of the genomes of higher eukaryotes can enable a variety of biological and medical applications. Targeted gene disruption, editing, and insertion can translate into the much desired freedom to generate cells or organisms bearing a desired genetic change. Recent developments in the stem cell field have created even more excitement for genetically modifying genomes because it enables delivering more beneficial stem cell-derived therapeutic cells to patients. For instance, by correcting a gene mutation known to be critical to Parkinson’s disease, LRRK2 G2019S, in patient-specific iPSCs (induced pluripotent stem cells), researchers were able to rescue neurodegenerative phenotypes [1].

Cumbersome reagent development and high costs have been major barriers to targeted genome modification using the current technologies, which include the zinc finger nuclease (ZFN) and transcription activator-like effector nuclease (TALEN). Unlike the ZFN and TALEN systems, CRISPR/cas does not require assembly of DNA pieces that encode the functional proteins every time a new sequence is to be targeted. Instead, it uses a guide RNA to direct the traffic of a nuclease complex. Five recent publications of modifying eukaryotic chromosomes showed the importance of the CRISPR/cas system [2-6], they also hinted at the ease of adapting this system in eukaryotes given that the functions of cas and the small guide RNA were described in bacteria merely few months ago [7].

The concern that the bacterial CRISPR/cas system would not access the chromatin structures of eukaryotic genome was muted as a result of recent publications; it also seems that the cas9 protein is as powerful an enzyme as one could have hoped in an endonuclease. As a matter of fact, cas9 from S. pyogenes contains 2 different single-stranded DNAse domains independent of each other, and can be mutated to change from a double-stranded DNA endonuclease to a single-strand cutter, or a non-cutting block. That’s not all, a more recent Nature publication further showed that cas9 (from another species, F. novicida), can bind to yet another small RNA and, instead of cutting chromosomal DNA, it degrades RNA, apparently through a direct cas9/RNA binding mechanism [8]. It may be chromosomal modification and RNAi rolled in one (cas9 from different genera are quite different though). One has to admire the powerful cas!

1. Reinhardt, P., et al., Genetic Correction of a LRRK2 Mutation in Human iPSCs Links Parkinsonian Neurodegeneration to ERK-Dependent Changes in Gene Expression. Cell Stem Cell, 2013. 12(3): p. 354-67.
2. Qi, L.S., et al., Repurposing CRISPR as an RNA-Guided Platform for Sequence-Specific Control of Gene Expression. Cell, 2013. 152(5): p. 1173-83.
3. Mali, P., et al., RNA-guided human genome engineering via Cas9. Science, 2013. 339(6121): p. 823-6.
4. Cong, L., et al., Multiplex genome engineering using CRISPR/Cas systems. Science, 2013. 339(6121): p. 819-23.
5. Cho, S.W., S. Kim, J.M. Kim, and J.S. Kim, Targeted genome engineering in human cells with the Cas9 RNA-guided endonuclease. Nat Biotechnol, 2013. 31(3): p. 230-2.
6. Hwang, W.Y., et al., Efficient genome editing in zebrafish using a CRISPR-Cas system. Nat Biotechnol, 2013. 31(3): p. 227-9.
7. Jinek, M., et al., A Programmable Dual-RNA-Guided DNA Endonuclease in Adaptive Bacterial Immunity. Science, 2012.
8. Sampson, T.R., et al., A CRISPR/Cas system mediates bacterial innate immune evasion and virulence. Nature, 2013.

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Solving the world’s problems with new biotechnology

The ability to isolate, create, synthesize, or artificially evolve living organisms towards desirable phenotypes may be increasingly important for solving many of the problems the world is facing. Such problems may include creating renewable energy using biowaste, finding biocontrol products that kill food-spoiling fungi “organically”, or assaying pathogens in the field using synthetic biological detection systems. With the arrival of synthetic biology, “it is possible to design and assemble chromosomes, genes and gene pathways, and even whole genomes”, according to the J. Craig Venter Institute. That is, if you know which genes or gene pathways you would need to put into the synthetic genome that would lead to the desired traits. So far, most published synthetic biology work involves bringing in transcription factors from a non-host source to set up an artificial network like circadian oscillators, showing that it can be done and it is interesting.

Through the process of evolution biological systems aptly self-engineer favorable traits in order to survive, but these changes require millions of years to manifest. However, there are quicker adaptations to environmental cues, such as developing antibiotic resistance, which can be achieved through a small number of mutations in hundreds or even dozens of generations. The question is how to harness this kind of adaptation for new strains that can be used as products with defined purposes? As a first requirement, you must have an assay for identifying the wanted mutants or method for augmenting their subpopulation, which is not necessarily easy and normally takes some clever designs to establish. Since evolutionary success in nature results from continuous “rounds” of gene mutagenesis, expression and selection, an evolution in the lab should ideally proceed with continuity. Previously, each round of mutation and selection takes a few days to complete. Recently, Esvelt et al. in David Liu’s lab at Harvard demonstrated one way of doing in vitro continuous evolution, by creating a lagoon of mixed E. coli and phages. By continuous dilution of the phage population through outflow, those phages that remain in the pool with properties that help them propagate in the host bacteria will have a better chance to regenerate and accumulate mutations towards the design of the assay [1].

Another aspect of natural evolution is that it occurs in a heterogeneous environment separated into niches of subpopulations with uneven stress levels. Although most evolutions with human intervention were conducted in a homologous population under the same stress and selection, a spatially complex environment may speed up evolution. This may not be easy to imagine, but if a mutant acquires some level of resistance to its environmental stress level and has a chance to move to join a population under higher stress, its relative fitness will likely increase. In addition, in a smaller population in the niche under higher stress, the mutant with marginally beneficial properties acquired under lower pressure can take over more quickly. This was demonstrated by Zhang et al. who showed that with a gradient of antibiotics applied to an array of microwells interconnected through tiny channels, new resistant strains can evolve in less than a day. Without the gradient, or separate the interconnected niches into discrete wells, no resistant populations could be obtained [2].

With more understandings like these and equipped with large scale gene synthesis, chromosome assembly, and deep sequencing technologies, we should see increasing numbers of human-made organisms serving special needs for food, health, energy, and the environment. Synthetic biology or artificial evolution won’t solve all the world’s problems, but if applied effectively and diligently, they can certainly help with many critical aspects as the technology “coevolves” with the environment.

[1] Kevin M. Esvelt, Jacob C. Carlson, & David R. Liu. “A system for the continuous directed evolution of biomolecules” Nature 499, 2011.
Qiucen Zhang, Guillaume Lambert, David Liao, Hyunsung Kim, Kristelle Robin, Chih-kuan Tung, Nader Pourmand, Robert H. Austin. “Acceleration of Emergence of Bacterial Antibiotic Resistance in Connected Microenvironments” Science 333, 2011.

New Products of the week: Modified UTP (Pseudouridine-5´-triphosphate), and Modified CTP (Methylcytidine-5´-triphosphate) for in vitro transcription of mRNA.

Promotion of the week: Friday special this week, buy 2 GFP-Trap get 1 free. Email the code “2+1GFPTrap” after placing your order of 2 GFP-Trap beads (0.25ml or 0.5ml scales only).

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