Newsletter Hot Plasmids
July 2017: New Optogenetic Tool PhoCI, LSSmCherry1 & RDSmCherry1, CRISPR Coselection, New Base Editors, & PSU DNA LaddersArticle Contributors
PhoCl can be used for light-dependent protein activation by a) sterically uncaging a protein or b) releasing an inhibitory domain. Image courtesy of Zhang W, et al. 2017.
Increasingly popular Optogenetics tools use light to alter cell physiology and molecular processes via genetically encoded proteins. PhoCl is a new member of this class that works via a different mechanism - the PhoCl protein is irreversibly cleaved when exposed to light.
Robert Campbell’s lab illustrates the utility of PhoCl in a variety of settings. For instance, inserting PhoCl in between a protein and a tag, such as an NLS or NES, enables control over localization. Inserting PhoCl between an enzyme and an inhibitor domain similarly allows for control of enzymatic activity. In addition, proteins can be “caged” by fusion to specific domains thereby rendering them inactive in the cytoplasm. The authors use PhoCl in combination with this caging technique to beautifully demonstrate light-dependent Gal4 transcriptional activation and Cre recombination.
Find PhoCl plasmids at Addgene.
Zhang W, et al. Nature Methods. 2017. PubMed PMID: 28288123
The toolbox of fluorescent proteins (FP) for cellular imaging is constantly expanding. Case in point - check out the growing list of popular plasmids in Addgene’s FP collection. Just last month, Robert Campbell’s lab added two new mCherry variants to the repository. These variants can be used as tools to learn about the influence of structure on an FP’s properties. pBAD-LSSmCherry1 is a long Stokes shift variant, which could be compatible with two-photon microscopy using Ti-Sapphire lasers. pBAD-RDSmCherry1 is a red-shifted variant which, with further development, could be better suited for deep-tissue in vivo imaging. These variants were developed as part of a larger strategy to engineer improved red fluorescent proteins (RFPs) and, while the authors note that there are other RFP variants available that may be preferred for their intended uses, they provide a well-defined framework for the production of additional variants. Read the group’s PLOS ONE paper for more on the directed evolution of mCherry and the spectral properties of various RFPs.
For tips on how to choose the right FP for your next experiment, check out our recent guest blog series: A Practical Approach to Choosing the B(right)est Fluorescent Protein.
Or sign-up to receive Addgene’s Fluorescent Proteins 101 eBook (Coming soon! Summer 2017).
Shen Y, et al. PLoS ONE. 2017. PubMed PMID: 28241009
Blue flame depositor Yannick Doyon’s lab has developed a new method to increase CRISPR editing efficiency in mammalian cells. Their marker-free coselection, analogous to methods previously used in C. elegans, targets the sodium/potassium pump ATP1A1 in addition to a researcher’s locus of interest. NHEJ- or HDR-induced mutations in ATP1A1 can render cells resistant to the small molecule ouabain - and editing at the ATP1A1 locus increases the probability that the locus of interest has also been edited! Thus, ouabain selection enriches the population for CRISPR edits and improves editing efficiency. Importantly, the point mutations created by Agudelo et al. prevent ouabain binding to ATP1A1 but do not compromise the ion pump’s function or cause cellular growth delays.
With eSpCas9(1.1) and coselection, Agudelo et al. achieved editing rates as high as 83% with NHEJ and 40-50% with HDR. Coselection efficiency is so high that some applications may permit working with edited “pools” of cells, eliminating the time-consuming step of single-cell isolation. Agudelo et al. also showed high editing efficiencies with AsCpf1, indicating that ouabain coselection is a robust and flexible way to increase CRISPR editing efficiency across multiple CRISPR enzymes.
Plasmids for CRISPR coselection are available at Addgene, and the Doyon lab has also provided a detailed protocol at Protocol Exchange. Plasmid maps can be found at Addgene and in Figure S12 of the publication.
Ouabain coselection increases the fraction of correctly edited cells. Image from Agudelo D, et al. 2017.
Agudelo D, et al. Nature Methods. 2017. PubMed PMID: 28417998
The popular CRISPR base editing technique uses dCas9/Cas9 nickase fused to a cytidine deaminase for targeted conversion of cytosine to thymine without a double stranded break. However, the method was initially limited by off-target effects, including the conversion of other cytosines near the target and in predicted Cas9 off-target sites. Two recent papers from blue flame depositor David Liu’s lab increase both the range and specificity of base editing.
Kim et al. used natural and engineered Cas9 variants to develop five new base editors with distinct PAM sequences, expanding the number of available target sites for base editing. For each base editor, they observed editing activity with a minimum efficiency of ~50% and confirmed that the fusion protein retained the PAM properties of the individual Cas9. Kim et al. also mutagenized the cytidine deaminase portion of the base editor to create SpCas9 base editors with editing windows as small as 1-2 nucleotides!
To reduce off-target effects associated with base editing, Rees et al. created HF-BE3, a base editor containing high fidelity Cas9 variant HF-Cas9. They subsequently tested the original BE3 and HF-BE3 using a gRNA with high off-target cutting, and they found that HF-BE3 showed 37-fold less off-target editing than BE3, with only a slight reduction in on-target editing efficiency. To further improve specificity, they purified HF-BE3 protein for delivery in ribonucleoprotein particles (RNPs) to both zebrafish embryos and the mouse inner ear. These exciting papers further show the potential of base editing for precise genome modification.
Schematic of the high-fidelity base editor (HF-BE3) described in Rees et al. bound to target DNA. HF-BE3 contains HF-Cas9 nickase, a uracil DNA glycosylase inhibitor (UGI) and a cytidine deaminase. Image from Rees H, et al. 2017.
|Plasmid ID||Plasmid Name||Cas9 Variant (PAM)||Special Features|
|85171||pBK-VQR-BE3||VQR-Cas9 (NGA)||Lower off-target activity than BE3|
|85172||pBK-EQR-BE3||EQR-Cas9 (NGAG)||Lower off-target activity than BE3|
|85174||pBK-YE1-BE3||SpCas9 (NGG)||Editing window ~2 nt|
|85175||pBK-EE-BE3||SpCas9 (NGG)||Editing window ~2 nt|
|85176||pBK-YE2-BE3||SpCas9 (NGG)||Editing window ~2 nt|
|85177||pBK-YEE-BE3||SpCas9 (NGG)||Lower on-target activity than BE3; editing window ~1-2 nt|
|87438||pET42b-HF-BE3||HF-Cas9 (NGG)||Very low off-target activity; protein purification vector|
|87439||pCMV-HF-BE3||HF-Cas9 (NGG)||Very low off-target activity; mammalian expression vector|
The Tan lab at Penn State - set out to “rule” Molecular Biology. Under the supervision of Dr. Tan, a team of undergraduates created DNA ladders (aka DNA molecular weight markers) that can be used to measure the size of DNA fragments. Depending on which restriction enzyme is used to cut the plasmids encoding the ladders, either 100bp or 1kb fragments are produced (see figure above).
3 Reasons The PSU Ladders Are Awesome Tools:
- They are comprised of 2 plasmids, pPSU1 and pPSU2, that can be amplified affordably in bacteria.
- They are compatible with both agarose and polyacrylamide gels.
- They are shared through Addgene without licensing restrictions.
Use these instructions to prepare the PSU ladders!
Find more plasmids from the Tan Lab!
Henrici R, et al. Scientific Reports. 2017. PubMed PMID: 28550309
March 2017: Tol2 Gateway-Compatible Toolbox, Polycistronic PiggyBac Cassettes, GABA-ergic Interneuron AAV Vectors, mCyRFP, & mScarletArticle Contributors
An example of a transgenic zebrafish created with our toolbox to study the relationship between neurodegenerative disease and motor neuron degeneration. This fish was generated from the entry vector elements collated in this toolbox and the original Tol2kit: pTol2pA2-bactin2-Hsa.TDP-43-EGFP and pTol2pA2--3mnx1:mCherry. Each expression cassette has a three prime SV40 polyadenylation signal (pA) in either the 3’ vector or backbone to aid expression.
The Cole lab recently deposited the Tol2 Gateway-Compatible Toolobox, which will allow scientists to quickly and easily generate zebrafish transgenic lines to study neurobiology and neuorodgenerative diseases.
The Tol2 Gateway-Compatible Toolbox is based on the original Tol2kit generated by the Chi-Bin Chien lab (Kwan et al., 2007) and includes four promoters, six fluorophores with nonoverlapping emission spectrums (N- and C-terminal tags for mTagBFP, TagRFPt, EGFP, mVenus, mCerulean3, mKOFP2) and empty vectors that have standard cloning sites or gateway compatible cloning sites for easy cloning of your genes of interest. In order to improve the study of specific neuronal cell types involved in neurodegenerative diseases, the Cole Lab selected promoters for cell types directly linked to disease, including motor neurons (-3mnx1/Hb9), pan neuronal (elavl3/HuC), microglial/macrophage (mpeg1.1), and astrocytic (gfap).This toolbox adds new neuronal tools to the expanding gateway compatible vectors currently shared amongst the zebrafish community.
Don E, et al. Zebrafish. 2017. PubMed PMID: 27631880
Kwan K, et al. Dev Dyn. 2007. PubMed PMID: 17937395
Deriving iPSCs (induced pluripotent stem cells) through the expression of transcription factors OCT3/4, SOX2, KLF4, and c-MYC is the established model for reprogramming somatic cells. A common method for generating iPSCs is expressing the transcription factors from a polycistronic cassette. However, it was unknown whether variations in gene expression from these polycistronic cassettes could lead to comparable experimental results.
Using PiggyBac transposons, the Woltjen Lab compared different variants of the polycistronic cassettes and discovered reprogramming discrepancies, which they traced to a 9 amino acid N-terminal variation in the Klf4 isoform (KLF4s or KLF4L). Polycistronic cassettes that contained KLF4s had overall diminished KLF4 expression levels; this altered stoichiometry impacted reprogramming and global gene expression patterns. Polycistronic cassettes that contained the KLF4L isoform had more robust initiation of reprogramming (evaluated by an alkaline phosphatase-positive assay) as well as stabilization (Nanog-GFP reporter activation and silencing of factor-linked mCherry). They additionally discovered that monocistronic expression of either variation of Klf4 did not lead to the observed reprogramming discrepancies.
The polycistronic piggyBac transposon vectors tested in this work are available at Addgene.
Graphical Abstract from Kim et al.
Kim, et al. Stem Cell Reports. 2015. PubMed PMID: 25772473
This interneuron in the adult mouse cortex was infected with AAV-Dlx-HA-Gq-DREADD-P2A-nls-dTomato ( one week post injection, staining with HA-tagged Gq-DREADD in green, nuclear dTomato in red, Dapi nuclei in Blue). Image from Dimidschstein J et al.
Genetically modified organisms like transgenic mice are important tools for understanding the function of cell types in the nervous system. While certain cell types, such as principal excitatory cells and glia, have been successfully targeted via viral particles, an effective approach to target specific neuronal subtypes was not available until now.
In order to target specific neuronal subtypes such as inhibitory GABA-ergic interneurons, Gordon Fishell’s lab has developed a new AAV approach using regulatory elements that restrict expression to this cell type. To this end, they constructed AAV plasmids bearing one of the distal-less homeobox 5 and 6 (Dlx5/6) enhancer elements and combined it with a variety of reporters (i.e. eGFP) and effectors (i.e. DREADDs). The team demonstrated that viral expression is robust and specific to interneurons in mice as well as other organisms, opening the possibility of using these tools in virtually any vertebrate.
This novel AAV approach allows you to target and functionally manipulate GABA-ergic interneurons across species and will undoubtedly constitute a valuable tool for the research community.
You can find the AAV vectors for infecting GABA-ergic interneurons here.
Dimidschstein J, et al. Nature Neuroscience. 2016. PubMed PMID: 27798629
mCyRFP labeled actin in HeLa cells. Image adapted from Supplementary Figure 4 of Laviv et al.
Researchers in the labs of Michael Lin and Ryohei Yasuda have teamed up to develop a novel monomeric cyan-excitable red fluorescent protein (mCyRFP1). Derived from cyan-excitable organge fluorescent protein (CyOFP1), some of the exciting properties of mCyRFP1 include a redshifted-emission spectrum compared to that of CyOFP1, a large stokes shift, long fluorescence lifetime, and a stable monomeric form as determined by the organized smooth endoplasmic reticulum (OSER) assay. A major advantage of mCyRFP1 is its ability to be coexcited along with EGFP while retaining an emission spectrum that is easily separable from that of EGFP. This makes mCyRFP1 a powerful tool for use as a donor in FRET (fluorescence resonance energy transfer) and FLIM (Fluorescence Lifetime Imaging Microscopy) applications. To demonstrate the power of mCyFP1, the Lin and Yasuda groups use both mCyFP1 and EGFP as FRET donors in experiments simultaneously measuring the activities of CaMKII and RhoA during induction of structural plasticity in single dendritic spines. The set of CyRFP constructs available at Addgene includes bacterial and mammalian mCyRFP1 or CyRFP1 expression plasmids, FRET constructs, and CaMKII and RhoA sensors.
Laviv, et al. Nat Methods. 2016. PubMed PMID: 27798609
mScarlet-I used to label F-actin in U-2 OS cells. Image adapted from Bindels et al Supplementary Figure 13.
The Dorus Gadella lab has recently developed a synthetic red fluorescent protein variant they call mScarlet. Bindels et al used a combination of deliberate and random mutagenesis to isolate the best RFP. They targeted known residues on the outer surface to break dimerization interfaces (affecting many of the RFPs developed from natural ancestors), and screened through libraries of new RFP variants for increased fluorescence, brightness, and proper maturation. Three new RFPs were isolated and characterized: mScarlet (fluorescent lifetime of 3.9 ns, quantum yield of 0.70), mScarlet-I with T74I (accelerated maturation in cells compared to mScarlet, fluorescent lifetime of 3.1 ns, quantum yield of 0.54) and mScarlet-H with M164H (2-fold improvement in photostability compared to mScarlet, fluorescent lifetime of 1.3 ns, quantum yield of 0.20). All three mScarlet variants have shown great performance in cellular functional imaging and in protein fusions. The Gadella lab has deposited all mScarlet constructs with Addgene.
Bindels DS, et al. Nat. Methods. 2017. PubMed PMID: 27869816
Dec 2016: Yeast Prototrophy Plasmids, T. gondii CRISPR Library, CRISPR-X, Magneto 2.0, & the Human Kinase Domain Constructs KitArticle Contributors
Complement your auxotrophic strains with this elegantly designed set of plasmids--a must-have toolbox for every yeast lab. The kit's minichromosomal plasmids contain HIS3, LEU2, URA3, MET17, LYS2, and combinations thereof. Image source: Mülleder et al 2016.
Though auxotrophic markers are used frequently for selection in yeast, Addgene's own Plasmids 101: Yeast Vectors blog post concedes that there are some drawbacks with this technique. Great care needs to be taken with common S288c strains since failing to fully complement strains with multiple auxotrophies or using different auxotrophic backgrounds can lead to disparate physiological effects and complicate any metabolic study. The Ralser lab has created a unique tool with their Yeast Prototrophy Kit, a set of plasmids designed to complement unused auxotrophies in Saccharomyces strains by compensating for histidine (HIS3), leucine (LEU2), uracil (URA3), methionine (MET17), and lysine (LYS2) deficiencies, and combinations thereof. The 23 plasmids in this kit are derived from the Ralser lab's own pHLUM minichromosomal vector, and besides being used for restoring strain prototrophy, can also aid in designing self-establishing metabolically cooperating (SeMeCo) communities. The uniform multiple cloning site in the plasmid series also allows for protein expression under a range of auxotrophic markers. Read more about the Ralser lab's important new tool for the yeast community in their open access article.
Mülleder M, et al. F1000Res. PubMedPMID: 27830062
- Listen to Our Yeast Prototrophy Kit Podcast Segment
Parasites from the apicomplexan phylum cause severe human and livestock diseases such as malaria and toxoplasmosis. Despite their importance to global health, most apicomplexan genes remain uncharacterized. Recent studies carried out by Sebastian Lourido and colleagues constitute an important step toward understanding the biology of these parasites. In this work, they used the CRISPR/Cas9 technology to measure the contribution of each gene from the model apicomplexan organism Toxoplasma gondii to the infection of human fibroblasts.
Toxoplasma mutants were generated by transforming parasites that constitutively express Cas9 with a library of sgRNA expression vectors (containing ten guides against each of the 8,158 predicted T. gondii protein-coding genes). Dr. Lourido’s studies demonstrate that genome-scale genetic screening is an efficient approach to identify genes involved in drug-resistance. Remarkably, this method also enabled Lourido’s group to define ~200 previously uncharacterized genes that are conserved among apicomplexans and important for T. gondii fitness. Sixteen of these genes were investigated for functions during infection of human cells, and one of them called “claudin-like apicomplexan microneme protein” (CLAMP) was shown to be important for the initiation of the T. gondii infection, and necessary for the asexual cycle of malaria parasites.
Fluorescence microscopy image showing the localization (green) of select genes in Toxoplasma. Image courtesy of the Lourido Lab.
Sidik and Huet, et al. Cell. 2016. PubMedPMID: 27594426
Directed evolution is an important method for protein engineering, but it can be difficult to generate enough variation in a native genomic context to find useful mutants. CRISPR-X, a new system from Michael Bassik’s lab, overcomes this obstacle. With CRISPR-X, dCas9 is used to recruit an sgRNA with MS2 hairpin binding sites, which then recruit an MS2-fused hyperactive variant of activation-induced cytidine deaminase (AID).
AID normally mediates somatic hypermutation to generate diverse antibodies; in the CRISPR-X system, AID*Δ similarly produces single base changes at each gRNA-specified target locus. Within the editing window -50 to +50 bp from the PAM site, CRISPR-X produces point mutations at a rate of up to 1 per 500-1000 bp. In comparison, the DNA replication error rate is 1 per 109 bp.
Unlike wt Cas9, CRISPR-X produces few indels, so it maintains reading frame and instead creates diverse, localized point mutations. As a proof of concept, Hess et al. successfully evolved wild type GFP to EGFP using CRISPR-X and subsequent FACS sorting. They also mutagenized proteasome subunit PSMB5 to find mechanisms of resistance to a proteasome inhibitor; CRISPR-X identified both previously characterized and novel mutants. Hess et al. envision CRISPR-X as not only a protein engineering tool, but also as a way to characterize/engineer promoters, enhancers, and other regulatory elements.
In the fields of opto- and chemogenetics, scientists are always on the hunt for new actuators that are non-invasive and can rapidly and reversibly stimulate neurons. Recent studies have designed multi-component actuators that are sensitive to radio waves or allow for magnetogenetic control, but these actuators were not optimally designed for neuronal systems. To develop a new single-component tool for magnetogenetic control of neurons, the Guler Lab engineered an actuator in which two subunits of the paramagnetic ferritin protein were tethered to the C-terminus of TRPV4 (a pressure-sensitive channel). The Guler lab optimized the actuator’s subcellular localization using trafficking signals and named it Magneto 2.0.
Using Magneto2.0-p2A-mCherry constructs, the lab verified that Magneto2.0 was magnetically sensitive and could manipulate cell activity in vitro. The Guler lab went on to test Magneto2.0 in the mammalian brain after delivering it via AAV injection, finding that it could increase the activity of neurons from brain slice preparations. Magneto2.0 was also tested in vivo by expression in zebrafish and mice. These in vivo studies revealed that Magneto 2.0 allows for magnetic remote control of neuronal activation. Find Magneto 2.0 plasmids by following the links below.
A magnetic field increases the activity of neurons expressing Magneto 2.0 and thereby induces behavioral responses in animals. Image courtesy of the Guler lab.
|Plasmid ID||Plasmid Name||Plasmid Type|
The misregulation of human kinases has been linked to several diseases, especially cancer. A great deal of chemical biology and drug discovery is focused on developing selective kinase inhibitors to probe these signaling pathways or develop potential therapeutics, but expressing and purifying human kinases remains challenging. Small molecule kinase inhibitors usually target soluble kinase domains, but these domains often do not express well in E. coli without their partner regulatory domains.
The Chodera Lab recently compiled the Human Kinase Domain Constructs Kit, a library of human kinase domain constructs developed through a large-scale expression screen that can be used to generate His-tagged human kinase constructs that express well in a simple bacterial expression system (Parton et. al. 2016). The key to obtaining high expression of the catalytic domains of each kinase is to express them with an appropriate phosphatase, a technique originally pioneered by Markus Seeliger for the high-yield expression of Src and Abl (Seeliger et. al. 2005). In this kit, Lambda phosphatase (Plasmid 79748) enhances bacterial expression of Serine/Threonine kinases while YopH (residues 164-468, Plasmid 79749) enhances expression of Tyrosine kinases. All sequences and expression data were kindly provided online by the Chodera Lab and the plasmids are now available through Addgene. All kinases in this library have crystallographic structures available in the PDB.
Parton DL, Hanson SM, Rodríguez-Laureano L, Albanese SK, Gradia S, Jeans C, Seeliger MA, Levinson NM, Chodera JD. bioRxiv preprint. 2016.Full Text.
Seeliger MA, Young M, Henderso MN, Pellicena P, King DS, Falick AM, and Kuriyan J. Protein Sci. 2005. PubMedPMID: 16260764
- Listen to Our Human Kinase Domain Constructs Kit Podcast Segment
Phylogenetic tree representing kinase domain expression. Dark green circles represent kinases with expression above 50 g/mL culture yield. Light green circles represent expression between 50 and 12 g/mL. Yellow circles represent expression between 12 and 7 g/mL. Yellow circles represent kinases with any expression (even below 2 g/mL) up to 7 g/mL yield. Figure courtesy of the Chodera lab. Authors note that the image was created using KinMap.
Sept 2016: Zika Plasmids, DULIP, CRISPR-AID Tagging, TRIBE, PiggyBac CRISPR Activators, & the pDGE Dicot Genome Editing KitArticle Contributors
The Vaithi Arumugaswami lab at Cedars-Sinai Medical Center has kindly constructed and deposited an unpublished set of Zika gene plasmids to Addgene’s repository. The genes, from the Zika virus Asian genotype PRVABC59, were obtained from the CDC and cloned into a third generation lentiviral transfer vector. Each gene has been FLAG-tagged for convenient experimental use. The plasmids were validated by both Western blotting and immunohistochemistry, and are summarized in the table below:
Aedes egypti mosquito (left) photo by jentavery via Flickr (Creative Commons 2.0 Generic License). Zika virus capsid (right) by Manuel Almagro Rivas (Own work) (CC BY-SA 4.0), via Wikimedia Commons.
DULIP schema. Plasmids expressing fusions of ProteinA-Renilla luciferase-Protein X and Firefly luciferase-Protein Y are co-transfected and expressed in a mammalian cell. If the two proteins interact, both Renilla and Firefly luciferase activities will be detected after co-immunoprecipitation by Protein A.
Proteins rarely act alone. Molecular processes from signal transduction to transport rely on protein-protein interactions and therefore identifying and studying interaction networks is important to detemine how these systems work. Researchers screening protein-protein interactions in mammalian systems have many assays to choose from, however, these methods are not without limitations. Some, such as the well-known yeast two-hybrid assay are not optimal for studying native mammalian protein interactions, whereas others may be too cumbersome to use in a high throughput manner or make it difficult to quantify interaction strengths. Erich Wanker’s group at the Max-Delbrueck Center for Molecular Medicine has recently developed a new method called DULIP (dual luminescence-based co-immunoprecipitation) to overcome the limitations of other methods and allow researchers to robustly identify and quantify protein-protein interactions in mammalian systems.
The DULIP system is comprised of Gateway-compatible plasmids in which “bait” proteins are tagged with Protein A-Renilla luciferase and the “prey” proteins are tagged with Firefly luciferase. The bait and prey plasmids are co-transfected and proteins are immunoprecipitated from the cells using the Protein A tag. The luciferase activities can then be measured and normalized to assess interactions. Higher relative firefly luciferase activity in the Co-IP compared of a control Co-IP indicates stronger interaction. The system has been demonstrated to detect high and low affinity interactions, identify changes in interaction strength due to point mutations, and is suitable for high throughput applications.
Trepte P, et al. J. Mol. Biol. 2015. PubMedPMID: 26264872
Seven years ago, Masato Kanemaki's lab successfully used the plant-based auxin-inducible degron (AID) technology to conditionally knock down proteins in a variety of model organisms. As long as it was expressed in a cell that also expressed auxin-sensitive TIR1, any protein fused with their AID tag could be targeted for degredation via ubiquitination. The key feature of auxin-induced degredation is the speed at which it works--typical half lives of AID-tagged proteins after the introduction of auxin range from 10-20 minutes, making AID an appealing tool for studying essential proteins.
Until recently, it was challenging to apply the AID-tagging system to essential genes in human cells and mouse ES cells because it was difficult to fuse the AID tag to endogenous proteins. As described in their recent Cell Reports paper, the Kanemaki lab overcame this hurdle using CRISPR technology. First, they used CRISPR to create OsTIR1-expressing cell lines by inserting OsTIR1 into the "safe harbor" AAVS1 locus (a tet-inducible OsTIR1 plasmid is also available). Second, they co-transfected short-arm donor vectors containing mAID and either Neo or Hygro resistance markers with CRISPR/Cas vectors targeting their genes of interest to generate mAID fusions to endogenous genes. Cell lines with the appropriate fusions could then be selected for using the indicated antibiotics. In addition, some versions of the mAID vectors come with mCherry2 or mClover fluorescent proteins allowing you to screen for fusions. This combination of CRISPR and AID technologies provides a clever means of investigating hard-to-study genes in hard-to-study cell types.
Adding the mAID tag to an endogenous gene in mammalian cells using CRISPR. Once tagged, the addition of auxin to the culture medium induces target protein ubiquitination and subsequent degradation. Image adapted from Natsume et al., 2016.
Identifying targets of RNA-binding proteins (RBPs) has traditionally been difficult, as techniques like CLIP require a specific antibody and a large number of cells to analyze. A new technique from Michael Rosbash’s lab, TRIBE, solves both of these problems by fusing an RBP to the catalytic domain of RNA editing enzyme, ADAR. This RBP-ADARcd fusion protein irreversibly edits adenine residues in the region of mRNA binding, and these sequence changes are subsequently identified through transcriptome sequencing. Importantly, results obtained using TRIBE compared favorably to those of CLIP, the gold standard for characterizing RBP targets.
One key advantage of TRIBE is the low input required - McMahon et al., report using as few as 150 fly neurons as their starting material! Thus, TRIBE can be used to analyze a particular cell population within a given tissue rather than a mixed sample. TRIBE was developed in Drosophila, and further work is needed to assess its function in mammalian cells or other animal models, but this technique has laid the groundwork for better analysis of RBP function.
McMahon A, et al. Cell. 2016. PubMedPMID: 27040499
TRIBE schematic courtesy of McMahon et al., 2016.
VPR uses multiple activation domains to increase expression of a gene of interest targeted by a gRNA and can now be delivered to mammalian cells using the PiggyBac system.
We previously reported on a novel Cas9-based activator developed by the Church lab, termed “dCas9-VPR”, which facilitates robust activation of target genes (20-40x greater than standard dCas9-VP64 activators) when introduced into mammalian cells.
Although the VPR activation domain is an incredible tool for researchers trying to activate target genes, the relatively large size of dCas9-VPR limits one’s ability to package it into lentivirus and later create stable cell lines expressing it. The Church lab fixed this issue by creating a new dCas9-VPR expression plasmid that uses the piggybac transposase to integrate dCas9-VPR into the genome of target cells. The piggybac system has a much larger cargo capacity than lentivirus and is capable of efficiently creating stable cell lines using direct transfection precluding the need for virus. Furthermore, dCAS9-VPR expression is under the control of the TRE promoter, which allows for doxycycline-inducible expression. The end result is genomic integration of dCas9-VPR and controlled expression of the dCas9-VPR transgene by doxycycline in your target cells.
This plasmid adds to the growing collection of dCas9-VPR plasmids, which already includes dSaCas9-VPR in an AAV transfer vector for in-vivo expression and plasmids optimized for dCas9-VPR expression in drosophila and yeast.
Chavez A, et al. Nat. Methods 2015. PubMedPMID: 25730490
The Stuttmann lab generated the vectors in this kit to make it easy to apply a variety of CRISPR tools to dicotyledonous plants. The pDGE Dicot Genome Editing Kit contains Cas9, Cas9 nickase, and Cas9 activator expressing plasmids that can co-express 1-8 gRNAs. Ordon et al., 2016 tested the Cas9 constructs for their ability to generate both small (< 100 bp) and large (up to 120 kb) deletions in N. benthamiana and Arabidopsis. The authors found that deletion efficiency was dependent upon 1) Cas9 and gRNA dose (both of which can be modulated using the different promoters in the kit and by altering the number of tandemly expressed gRNAs respectively) and 2) the size of the deletion. Smaller deletions generally occurred at higher frequency and could be detected by PCR while large deletions were relatively rare. The vectors are compatible with Golden Gate Cloning and Agrobacterium-mediated delivery making them easy to customize for targeting your gene of interest. Try them today!
Ordon J, et al. Plant J. 2016. PubMedPMID: 27579989
The pDGE Dicot Genome Editing Kit makes it easy to do generate chromosomal deletions in dicotyledonous plants like Arabidopsis (right). Arabidopsis image by Marie-Lan Nguyen (CC-BY 2.5)
June 2016: CRISPR Epigenetic Tools, BioID2, FusX TALEN Assembly System, Snoop Catcher, Easy Clone 2.0, NgAgo & SapTrapArticle Contributors
The traditional approach to studying epigenetic modifications has been to use inhibitors of epigenetic modifiers like DNA methyltransferases or histone deacetylases. Unfortunately, these inhibitors are often non-selective, affecting the entire genome and not specific loci. To overcome this flaw, the Zoldos lab has developed CRISPR tools for epigenome editing that enable the precise study of particular epigenetic modifications and their effects on gene regulation.
Professor Zoldos and colleagues from the Department of the Biology, Division of Molecular Biology at the University of Zagreb, constructed a CRISPR-Cas9-based tool for targeting CpG methylation by fusing catalytically inactive Cas9 (dCas9) with the catalytic domain from DNA methyltransferase, DNMT3A. Co-expression of this modified Cas9 along with a gRNA targeting a specific genomic locus causes that locus to be methylated. This powerful tool has been used to direct methylation to and lower expression from both the IL6ST and BACH2 promoters.
Vojta A, et al. Nucl. Acids Res. 2016. PubMedPMID: 26969735
CRISPR linked to DNMT3A methylates a target site specified by a gRNA. Image courtesy of Vojta A, et al. 2016.
All the vectors used in this study are now available at Addgene and we’re excited to see how you use them in your own work!
The Roux Lab has deposited a smaller version of the promiscuous biotin ligase used in their popular biotinylation method for identifying protein-protein associations. This method uses a bait protein fused to a biotin ligase to biotinylate proximate proteins in the cell. These candidate proteins can be enriched via biotin pull-down methods and identified using mass spectrometry.
The Roux Lab’s new and improved version of BioID, called BioID2, is significantly smaller, allows for more selective targeting of fusion proteins, requires less biotin, and shows enhanced labeling of proximate proteins. Overall BioID2 improves the efficiency of screening for protein–protein interactions.
Kim, et al. Mol Biol Cell. 2016. PubMedPMID: 26912792
TALEs as nucleases (TALENs) are genome-editing tools that are deployed in both in vitro cell systems and diverse model organisms. Addgene’s newest TALEN kit, the FusX assembly system from Stephen Ekker’s lab, is a modified version of the Golden Gate TALEN kit (GGT) that streamlines the assembly of TALE repeats into a single-tube 3 day process by utilizing pre-assembled trimers to reduce the number of cloning steps. The FusX system was verified in zebrafish and was shown to offer high activity and unparalleled specificity in genomic targeting design flexibility. The FusX kit is backward compatible with all TAL effector scaffolds previously constructed for the Golden Gate TALEN and TAL Effector Kit 2.0.
Ma, et al. Hum Gene Ther. 2015. PubMedPMID: 26854857
Examples of FusX-Compatible Destination Vections Available at Addgene:
|Dan Carlson||RCIscript-GoldyTALEN, pC-GoldyTALEN|
|Tom Ellis||pTAL5-BB, pTAL6-BB|
|David Grunwald||pCS2TAL3-DD, pCS2TAL3-RR|
|Pawel Pelczar||pCAG-T7-TALEN(Sangamo)-Destination series, pCAG-Golden-Gate-Esp3I-Destination|
|Takashi Yamamoto||pcDNA-TAL-NC2, pCAGGS-TAL-NC2|
|Charles Gersbach||pcDNA3.1-GoldenGate, pcDNA3.1-GoldenGate-VP64|
|Maria-Elena Torres-Padilla||pTALYM3, pTALYM4|
|Boris Greber||pTAL7a, pTAL7b|
|Nathan Lawson||pJDS Series|
The development of recombinant DNA technology has allowed molecular biologists to fuse fragments of DNA together in a desired order and express fusion proteins in a target cell. In addition to nucleotide-based linking strategies, post translational protein fusion would be advantageous when individual protein expression is preferred or required, or when it is difficult for a cell to synthesize or fold a protein because of its length. Mark Howarth’s lab recently engineered a new peptide/protein pair, SnoopTag/SnoopCatcher, that can be used to irreversibly link desired proteins together via a spontaneous isopeptide bond. When used in combination with their previously described SpyTag/SpyCatcher pair, multiple protein fusions can be synthesized in an iterative process by alternating the use of the SnoopTag/SnoopCatcher and SpyTag/SpyCatcher pairs.
"Polyproteam" production using Snoop/Spy tags and Snoop/Spy catcher. Image courtesy of Veggiani et al 2016.
In order to improve the ease and convenience of assembling these multi-protein fusions or “polyproteams”, the Howarth lab created the MBPx-SpyCatcher construct. This construct is used to produce a maltose binding protein/SpyCatcher fusion that can be anchored to amylose resin for solid-phase synthesis of larger fusions through SpyTag/SpyCatcher and SnoopTag/SnoopCatcher linkages. Once all of the desired sequential additions have been made, the final polyproteam can be eluted using maltose. This procedure allows for stable and directional protein fusion, with a SpyCatcher-SnoopCatcher linker remaining between each protein unit, and provides a new method for generating protein chains or clusters.
Veggiani, et al. Proc Natl Acad Sci U S A. 2016. PubMedPMID: 26787909
Many industries, such as the food and beverage industry, the bioethanol industry, and the pharmaceutical industry, depend on the yeast, Saccharomyces cerevisiae, as a key host for the production of their products, including fermented beer and wine, fuels, pharmaceutical ingredients, and recombinant proteins. In order to keep production yields high and costs low, and to deal with the increasing variety of new compounds being generated, there is a need for genetic optimization of industrial yeast strains. However, industrial yeast strains are more difficult to genetically modify than common lab yeast strains since they are prototrophic, usually have low levels of homologous recombination, and can be difficult to transform.
To increase the genetic toolbox for industrial yeast strains, the Borodina Lab has recently designed the EasyClone 2.0 toolkit. EasyClone 2.0 consists of 25 integrative vectors containing long homology arms and both auxotrophic and dominant selection markers, allowing for use with both laboratory and prototrophic strains. The vectors were based on the original EasyClone vectors (Jensen et al 2013) and allow for cloning of up to two genes with a bidirectional promoter, integrate at 11 specific chromosomal locations, and stably express the integrated gene. These vectors contain uracil-excision based (USER) sites, flanked by ADH1 and CYC terminators for easy cloning of genes and promoters. In addition to USER cloning, this toolkit is also compatible with systems such as in-fusion, Gibson, and MoClo. The EasyClone 2.0 kit has been made publically available through Addgene.
Schematic of EasyClone 2.0 vectors containing various selection markers. Image courtesy of the Borodina lab.
Cas9’s ability to cleave target DNA at user defined genomic targets is a hallmark of CRISPR and contributes to its growing popularity as a genome engineering tool. However, novel endonucleases may be necessary to circumvent inherent limitations of Cas9. Among these limitations is Cas9’s requirement for a short PAM sequence adjacent to any putative cut site. Chunyu Han’s laboratory at the University of Science and Technology in Shijiazhuang, China characterized a novel Argonaute gene from Natronobacterium gregoryi (so called “NgAgo”) that, like Cas9, can be guided to cut specific DNA sequences using a short nucleotide-based guide, but doesn’t require a PAM. This and several other features may make NgAgo more useful than Cas9 for specific applications.
Many of the basic principles of Cas9-mediated genome engineering hold true for NgAgo with some important differences. Because NgAgo doesn’t require a PAM, it can theoretically modify any genomic target. Furthermore, NgAgo uses a ~24 nucleotide 5’ phosphorylated guide DNA (“gDNA”) to guide NgAgo to target DNA, as opposed to the 18-20 nucleotide guide RNA used by Cas9. Like Cas9, NgAgo is capable of generating double-strand breaks in genomic DNA targets in mammalian cells which can be repaired by error-prone non-homologous end joining resulting in knock outs or homology directed repair resulting in specific edits when delivered with a repair template. Target cleavage via NgAgo is sensitive to mismatches between the gDNA and the genomic target, which ensures specificity of target binding and cleavage. Furthermore, NgAgo is roughly ⅔ the size of SpCas9, meaning it may be more amenable to packaging into lentivirus or AAV, a useful means of delivery for difficult to transfect cell types or in-vivo applications, respectively.
NgAgo does not require a PAM sequence in order to cut at its target sites. Image Courtesy of Joel McDade and Tyler Ford.
It remains to be determined whether NgAgo will surpass Cas9 in terms of efficiency, specificity, and versatility, but the availability of NgAgo expands the toolbox of reagents available to genome engineers beyond Cas9.
A mammalian expression vector containing NgAgo can be obtained through Addgene: plasmid number 78253.
Gao, et al. Natl Biotechnol. 2016. PubMedPMID: 27136078
CRISPR/Cas9 technology has revolutionized genome editing in multiple organisms thanks to its simple, easily modifiable RNA-guided targeting mechanism. However, the laborious construction of repair templates and guide RNA constructs, as well as the difficult screens required to identify correctly modified organisms has limited routine use of this technology. To streamline CRISPR/Cas9-based genetic tag insertion in the C. elegans genome, Erik Jorgensen’s lab from the University of Utah developed SapTrap.
The SapTrap system uses golden gate assembly to produce single plasmid targeting vectors that encode both a guide RNA transcript and a repair template for an individual tagging event. Commonly used sequences, including fluorescent tags, a floxed Cbr-unc-119 selectable marker, and “connectors” that link tags to the targeted gene are supplied to the reaction from a prebuilt donor plasmid library, available from Addgene. Site-specific sequences for homology arms and the guide RNA are supplied as annealed synthetic oligos - eliminating the need for PCR or additional molecular cloning steps during plasmid assembly.
This toolkit reduces the expense and workload necessary to produce vectors for genome editing in worms to the point that high-throughput tagging projects can be performed.
Schwartz, et al. Genetics. 2016. PubMedPMID: 26837755
March 2016: STARR-seq, CIDAR MoClo Parts Kit, Ras Pathway, pORTMAGE, CRISPR Libraries & MoreArticle Contributors
Although many technologies have been developed for the study of enhancer function, strength, and binding (e.g. DHS-seq, FAIRE-seq, and CHIP-seq), genome-wide identification and quantification of enhancer activity has remained challenging due to the variable availability of enhancer sequences in different chromatin states. Alexander Stark and colleagues at the Research Institute of Molecular Pathology have developed an alternative method, called STARR-seq (self-transcribing active regulatory region sequencing) that tests enhancer sequences outside of their endogenous chromatin environment and relies on the fact that enhancers function independently of their position relative to a transcription start site. In STARR-seq, randomly sheared genomic libraries are cloned downstream of a minimal promoter sequence. Should one of these random stretches of DNA contain an enhancer, it will activate its own transcription from the minimal promoter and its strength can be determined from its enrichment over input levels using next gen sequencing. The STARR-seq backbone is available from Addgene in both human and fly versions.
Arnold, et al. Science. 2013. PubMedPMID: 23328393
The Wu lab developed a pooled lentiviral gRNA library to screen for human genes that confer resistance to West Nile Virus induced cell death, which revealed a novel role for genes involved in ER-assisted Protein Degredation (ERAD). This library adds to the growing number of CRISPR knockout libraries available through Addgene. Image designed by Joel McDade.
CRISPR pooled libraries enable researchers to screen the entire genome for genes that regulate a wide variety of phenotypes, including but not limited to cell growth and drug resistance. Recently, the Wu Lab at Texas Tech developed a pooled CRISPR knockout library and used this library to screen for genes involved in West Nile Virus (WNV) induced cell death.
The Wu lab library consists of 77,406 individual gRNAs targeting a total of 20,121 genes within the human genome (the identity of the genes and target sequences can be found here). To use the library, the authors packaged it into lentivirus and used the resulting lentiviral library to generate a population of HEK293FT cells expressing a single gRNA targeting a single gene. The gRNA expressing cells were then transfected with a Cas9-expressing plasmid to generate a population of mutant cells. The mutant cells were treated with WNV at a dose and duration sufficient to kill all control cells and surviving “mutant” colonies were deep sequenced to identify the gRNAs that confer protection to WNV. From this screen the authors were able to identify 7 previously unsuspected genes with known roles in ER-associated degradation (ERAD) that positively regulate the ability of WNV to induce cell death in infected cells (SEL1L, UBE2J1, EMC3, EMC2, DERL2, UBE2G2 and HRD1).
The Wu lab knockout library is available alongside a growing number of CRISPR libraries from Addgene’s repository that can be used to perform screens in human cells. General information regarding how to use a CRISPR pooled library as well as criteria of a good screening experiment can be found on our blog post entitled “Genome-wide Screening Using CRISPR/Cas9”. Detailed information regarding the Wu Lab library can be found in the associated publication.
Ma, et al. Cell Rep. 2015. PubMedPMID: 26190106
Nearly one third of all medications target G-protein coupled receptors (GPCRs) yet one third of the GPCRs in the human genome are orphan receptors meaning their ligands are unknown. Identifying new ligands for orphan GPCRs is of significant medical interest, however efforts to do so have been hindered by a lack of tools, compounds, and assays to monitor activation of these receptors. Bryan Roth's lab has a newly available PRESTO-TANGO kit which allows researchers to interrogate the druggable human GPCR-ome using a quick, affordable, and ubiquitous reporter assay. In their recent publication, Kroeze et al. describe their enhanced TANGO arrestin recruitment assay incorporating 314 codon-optimized GPCR sequences for human cell line expression. In a twist on the classical TANGO assay, Kroeze et al. developed a new approach termed Parallel Receptor-ome Expression and Screening via Transcriptional Output-TANGO (PRESTO-TANGO) to screen the NCC-1 library of approved drugs against the entire kit; a parallel analysis that successfully identified new, highly specific agonists for orphan GPCRs demonstrating the utility of this platform for drug discovery. Researchers can use plasmids from the PRESTO-TANGO kit directly for drug screening or easily shuttle the optimized, expression validated GPCR sequences to any desired backbone.
Synthetic and basic biologists alike often need to test multiple expression systems before they achieve the appropriate expression level for their gene of interest. Although the parts involved in altering gene expression are relatively well known, particularly in bacteria, the process of cloning, combining, and testing these different parts can be quite arduous. The cloning process alone involves much literature searching, DNA synthesis, and plasmid assembly. Luckily, researchers from Douglas Densmore’s lab at Boston University have greatly simplified the process of generating multiple E. coli gene expression plasmids with the Cross-disciplinary Integration of Design Automation Research lab Moduclar Cloning (CIDAR MoClo) Parts Kit.
This Kit consists of a variety of plasmids containing many different promoters, RBS’, coding sequences (CDS), and terminators of varying strengths. These can be easily combined with your gene of interest and cloned into provided destination vectors using low-cost, one-pot golden gate cloning that results in the creation of many E. coli constructs with greatly varied levels of expression. The 93 plasmids in the CIDAR MoClo Parts Kit are available as glycerol stocks in a single plate with every plasmid or as individual plasmids shipped as agar stabs.
The CIDAR MoClo Parts Kit contains plasmids with various promoters, RBS’, coding sequences (CDS), and terminators that can easily be ligated into provided expression vectors using golden gate cloning. Image courtesy of Iverson et al 2016.
Iverson, et al. ACS Synth Bio. 2015. PubMedPMID: 26479688.
Ras proteins are small GTPases involved in cell signaling pathways that control many cellular processes such as differentiation, proliferation, and apoptosis. In humans, mutations that permanently activate Ras are found in one third of all cancers, including a high percentage (up to 95%) of pancreatic cancers. Ras oncogenes have been difficult to target therapeutically due in no small part to the central role Ras proteins play in cell signaling. Understanding how molecules in the Ras signaling pathway interact will allow scientists to develop better strategies for combating cancers. To facilitate innovative cancer research, the Reference Reagents Group of the NCI RAS Initiative, led by Dominic Esposito, has recently released a collection of 360 Ras pathway plasmids to the scientific community.
This collection consists of plasmids containing genes from a crowd-sourced map of the RAS signaling pathway and includes 180 unique transcripts. Each gene was chosen to represent the primary splice variant in 38 types of human cancer. Many of the transcripts are not available from other existing sources and are provided in both open (no stop codon) and closed (including a stop codon) formats. The collection is compatible with both standard Gateway cloning and the FNLCR Combinatorial Cloning Platform (CCP); the latter of which greatly enhances the utility of this collection as it can be used to easily create protein expression constructs fused with epitopes or fluorophores, generate expression vectors with various promoters suitable for in vivo expression, and/or produce vectors to make mutant cell lines or transgenic animals.
Wall, et al. Methods Mol Biol. 2014. PubMedPMID: 24395366
The Ras signaling pathway with major kinases and other players indicated. Signaling through Ras ultimately results in the alteration of gene expression and is important to a variety of cellular processes including differentiation, proliferation, and apoptosis. Image attribution: JWSchmidt at the English language Wikipedia.
Addgene has had the privilege of distributing several human and murine CRISPR pooled libraries, but thanks to a deposit from the laboratory of Dr. Ji-Long Liu at Oxford University, we are now providing a powerful CRISPR-based screening source for the Drosophila community as well. The pooled library, designed to be used with cultured Drosophila cells, targets 13,501 fly genes with at least three independent gRNAs, each chosen strategically to maximize the likelihood of creating a functional knock-out via frameshift and minimize off-target effects. The library was cloned into the Liu lab’s own pAc-sgRNA-Cas9 insect expression backbone, which expresses the gRNA from a Drosophila U6:2 promoter and Cas9 from the actin 5C promoter. Addgene’s fly library web page offers more details and protocols, and the Liu lab’s OXfCRISPR page provides many other useful resources for fly labs who wish to harness this powerful genome engineering tool.
Bassett, et al. J Genet Genomics. 2015. PubMedPMID: 26165496
Gibson assembly-based modular assembly platform (GMAP). a) A variety of promoters, genes, and backbones from the GMAP collection can be used to generate DNA constructs in one step. b) 5 common 30bp overlap sequences (Sites #1-5) allow for multiple orderings of genes and promoters. C) Example of a GMAP retroviral backbone used to generate new vectors with a variety of different promoters. (Image adapted from Akama-Garren et. al. Sci Rep. (2016)).
Have you ever wished you could design a gene knockdown or overexpression experiment, generate the necessary constructs, and complete screening in just 3 days? Thanks to Tyler Jacks’ lab, this 3-day goal is is now attainable. The Jacks lab has recently introduced a new assembly cloning platform called GMAP (Gibson assembly-based modular assembly platform). GMAP is based on the Gibson Assembly method, which allows for easy assembly of DNA fragments by utilizing regions of sequence homology (usually 30-40bp overlaps). To expand the capabilities of Gibson assembly and make a more modular platform, the Jacks lab generated 5 common 30bp overlap sequences (Sites #1-5). Each overlap site encodes a unique restriction enzyme site and unique primer sites for more rapid screening by restriction digest and Sanger sequencing, respectively. The speed and simplicity of GMAP allows scientists to easily design, generate, and test constructs composed of complex genetic elements.
To demonstrate the usefulness of GAMP, the Jacks Lab constructed several GMAP-compatible vectors for a wide variety of biological applications. First, GMAP-compatible backbones for lentivirus LV 1-5 and retrovirus RV 2-5 were constructed. The authors then used GMAP assembly to establish a collection of over 30 promoters and 140 genes, including constructs with unique tissue-specific promoters expressing GFP, tetracycline response elements, and shRNAs many of which are available at Addgene. Since GMAP assembly utilizes 5 different overlap sites, this technique also allows for easy and rapid targeting of complex elements to specific genomic sites. For instance, the authors generated a GMAP-compatible backbone containing Rosa26 homology arms. The Rosa26 locus is commonly used to drive ubiquitous expression of a target gene in mice. The Jacks Lab used this backbone to target a CAG-driven loxP-stop-loxP cassette into the Rosa26 locus, allowing for Cre-dependent expression of a GMAP inserted gene. The applications for such a modular system are endless and, as scientists use GMAP, there will be a continuous expansion of compatible backbones, promoters, and genes available to the community. If you plan to use GMAP for your experiments, think about depositing your new plasmids with Addgene to help expand the collection!
The Jacks lab has made several GMAP-compatible plasmids available to the scientific community through Addgene.
Akama-Garren, et al. J Genet Genomics. 2016. PubMedPMID: 26887506
Genotyping using HRM (High Resolution Melting) analysis. The chart displays a normalized melting curve used for HRM analysis to detect small sequence differences between two DNAs. Image courtey of the Pál lab.
Recent discoveries in the fields of genome engineering and synthetic biology have transformed the way scientists create new traits in bacteria and enabled the in-depth study of many biological processes. While these methods can be used to easily modify an individual locus within the genome, they tend to show their limits when it comes to simultaneously modifying multiple loci (multiplexing).
Currently, the only genome engineering method in bacteria that enables rapid, automated, and high-throughput genome editing is multiplex automated genome engineering (MAGE) (1). MAGE uses recombineering (2) to simultaneously incorporate multiple single-strand DNA (ssDNA) oligonucleotides (oligos), and thereby rapidly create desired allele combinations and combinatorial genomic libraries. MAGE has allowed genome-engineering endeavours of unparalleled complexity in Escherichia coli, like the construction of a so-called “genomically recoded organism” (3) but its portability to other bacterial strains remains seriously limited, as prior optimizations are required for each individual target species. Indeed in order to use MAGE, the λ Red recombinase enzymes need to be expressed and the native methyl-directed mismatch repair (MMR) system needs to be repressed in the host strain.
To address this problem, Csaba Pál’s lab has created a set of vectors that allows one to use the MAGE method in unmodified bacterial species (4). This set of plasmids (dubbed pORTMAGE) expresses the λ Red recombinase enzymes, as well as a dominant-negative mutator allele of the E. coli MMR protein MutL, all under the control of the cI857 temperature-sensitive repressor. The temperature-controlled expression of the MutL mutant enables transient suppression of DNA repair during oligonucleotide integration, allowing MAGE in otherwise unmodified bacterial strains (5). MutL is highly conserved in distant relatives of E. coli so it can be used in a broad range of strains. Thus, pORTMAGE simultaneously allows genome editing and mutant library generation in several biotechnologically and clinically relevant bacterial species.
pORTMAGE vectors open new horizons to modify genomes in a broad range of bacterial hosts. This fantastic tool is now available at Addgene for you to be the “MAGE” that will create new traits in bacteria.
The Moffat lab used the TKO CRISPR library to test genes for essentiality across a variety of different contexts. Those that were essential across all contexts were considered core-fitness genes. Image courtesy of the Moffat Lab.
In order to identify both core and context-dependent human fitness genes, the laboratory of Jason Moffat at the University of Toronto has created the Toronto KnockOut (TKO) CRISPR Library. This complex second-generation CRISPR lentiviral library targets nearly all human protein-coding genes, and is unique in that it is composed of two sub-pools. The first is a base “90k library” that contains ~90,000 gRNAs that target ~15,000 genes with 6 gRNAs per gene. The second is a "supplementary" library that contains an additional 6 gRNAs per gene generated using slightly relaxed gRNA design parameters.
To demonstrate the effectiveness of their library design, Moffat laboratory members conducted multiple screens in different human cell models and identified ~4-5-fold more fitness-related genes than have been found in previous screens of this type, and they were able to functionally characterize positive hits. This work has been published in Cell.
The Moffat laboratory maintains a website for the TKO library that contains up-to-date information and data-sets, as well as an interactive gRNA viewer. The Moffat lab has also deposited their Cas9 and gRNA backbone plasmids with Addgene. Additional control plasmids for the TKO library will be available at Addgene soon.
Hart, et al. Cell. 2015. PubMedPMID: 26627737
A pH-sensitive fluorescent biosensor-based assay system was recently deposited with Addgene by Dr. Mark Prescott. Dr. Prescott and Dr. Rod Devenish (both at Monash University) originally developed the system for monitoring and analysing autophagy of cytosol and organelles in yeast cells. Named after the brightly-coloured Australian parrot, the dual colour-emission biosensor Rosella consists of DsRed connected to a pH-sensitive variant of GFP (SEP) by a 9 amino-acid linker (see Figure 1). The key to the biosensor lies in pH: DsRed is relatively pH-insensitive, while SEP fluoresces at cellular pH (7.0) but not at vacuolar pH (6.2). The colour of Rosella therefore varies with cellular location: it is both red and green through much of the cell, but turns more red in lower (e.g. vacuolar) pH environments that inactivate SEP and leave only DsRed to fluoresce (1). With variants that can be targeted to specific cellular compartments (cytosol, Addgene catalog #71245; mitochondria, Addgene catalog #71247) Rosella provides an easy way to follow uptake of cellular material into the yeast vacuole, as well as to examine the mechanisms behind autophagic pathways.
Rosado, et al. Autophagy. 2008. PubMedPMID: 18094608
Left: Australian parrot from which the Rosella biosensor gets its name. Right: The Rosella biosensor. From the N-terminal end (N): targeting sequence for subcellular localisation (not present in the cytosolic version of Rosella) - blue circle; DsRed.T3 - red box; purple bar - 9 amino acid linker; GFP-variant (SEP) - green box. Image designed by Jessica Welch.
December 2015: CRISPR-Display, FP Fusions, Broccoli, Spinach, Nano-laterns, & MoreArticle Contributors
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.
Shechner, et al. Nat Methods. 2015. PubMedPMID: 26030444
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:|
Friedman, et al. J Cell Biol. 2010. PubMedPMID: 20696706
Friedman, et al. Science. 2011. PubMedPMID: 21885730
Friedman, et al. Mol Biol Cell. 2013. PubMedPMID: 23389631
Rowland, et al. Cell. 2014. PubMedPMID: 25416943
Shibata, et al. J Biol Chem. 2008. PubMedPMID: 18442980
Zurek, et al. Traffic. 2011. PubMedPMID: 20955502
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 a randomn genomic location than to express the tagged protein from the a defined site in the genome 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 previously observed interactions, 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.
Graphical Abstract from Dalvai et al 2015. Technique used to endogenously TAP-tag a protein of interest.
Dalvai, et al. Cell reports. 2015. PubMedPMID: 26456817.
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!).
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). 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.
Example pCellFree vector from the Alexandrov lab Cell-Free Protein Expression Kit, showing the position of the Species-Independent Translational Sequence (SITS), N-terminal EGFP tag, ORF with Gateway cloning sites, and other features.
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 a pXP1-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.
Vegetable tags, including Spinach, Spinach2, and Broccoli, 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 protocol involves 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.
Broccoli for your research. Image attribution: David Monniaux.
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-Broccoli controls.
Don’t just eat more broccoli, use Broccoli tags in your experiments too!
Filonov, et al. Chem Biol. 2015. PubMedPMID: 26000751
September 2015: Customized FP Reporters, Optogenetics, A New Epitope Tag & MoreArticle Contributors
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 the retention 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.
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.
A "Colorful Cell" generated using a MXS-chaining vector containing 6 expression cassettes. Image courtesy of Pierre Neveu.
Sladitschek HL and Neveu PA. PLoS One. 2015. Apr 24;10(4):e0124958.
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.
The mitochondrial fusion assay using differential photo-tagging of mitochondrial LOC8R-PAGFP & LOC8R-PSmOrange. a) Before photoactivation b) 488 nm PSmOrange photoswitching of mitochondria in the center of the cell c) 405 nm PAGFP photoactivation of mitochondria surrounding the photoswitched region. Image courtesy of Benjamin S. Padman.
These plasmids are unpublished.
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 to examine molecular events with exceptional precision in a native context.
Plasmid map of pAcBac2.tR4-OMeYRS/GFP, for baculovirus-based delivery of E. coli tyrosyl-tRNA synthetase and 4 copies of the Tyr tRNA cassette to mammalian cells.
Chatterjee A et al. Proc Natl Acad Sci U S A. 2013 Jul 16;110(29):11803-8.
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 presented Archaerhodopsin 3 (Arch) as a new rhodopsin based voltage indicator for use in 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 Optopatch vectors available to the scientific community through Addgene.
Hochbaum, et al.Nat Methods.2014.
Kralj, et al.Nat Methods.2011.
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.
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.
June 2015: Optogenetics, dCas9 Transactivators, Fusion Protein Cloning Kit, Visualizing Translation, & MoreArticle Contributors
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 the anchor 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.
The article describes the targeted activation of endogenous genes and details the time course of gene activation using this system. 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.
Nihongaki et al.,Chemistry and Biology 2015 Feb 19; 22(2):169-74.
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 vector ensures 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.
Summary of cloning mechanism of pUCXKT vector.
With kind permission from Springer Science+Business Media: Biotechnology Letters ‘A gain-of-function positive-selection expression plasmid that enables high-efficiency cloning’; 2015, 37:383-389; Prosser GA, Williams EM, Sissons JA, Walmsley KE, Parker MR, Ackerley DF; Fig 1.
Prosser et al., Biotechnol Lett. 2015. 37:383-389.
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. The Santalucía 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).
Buj et al., BMC Molecular Biology. 2013 August 20th; 14(1).
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 Translation RNA 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 (Plasmid64543or64542)
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. The great specificity and resolution of this system are achieved 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, the scientists 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
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 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 (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 all the people 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 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. 2015 extended 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.
Chavez et al., Nat Methods. 2015 12(4): 326-328.
March 2015: LACE System, Split Cas9 systems, Multi-bit Genetic Memory & MoreArticle Contributors
The Cas9 protein is composed of an n-terminal DNA recognition domain and 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 plasmids62883&62884;62885&62886)
- Rapamycin-inducible dCas9-VP64 activator set (Addgene plasmids62887&62888)
Zetsche et al.,Nat Biotechnol.2015 Feb 2;33(2):139-42. doi: 10.1038/nbt.3149.
Zetsche et al.,Nat Biotechnol. 2015 Feb 2;33(2):139-42.
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.
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 and the 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 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.
Tanenbaum et al.,Cell. 2014 Oct 8.
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.
pMemoryArray with position 9 inverted, allowing for detection by PCR.
Yang et al., Nat Methods 2014 Dec;11(12):1261-6. doi: 10.1038/nmeth.3147.
Understanding intracellular pH regulation is important as pH regulation has many roles in cellular function, 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-pHlash and pcDNA3.1+-pHlash, were developed by the Johnson Lab that use Bioluminescence Resonance Energy Transfer (BRET).
Characteristics of the pHlash reporter protein include:
- H+ specific response
- cytoplasmic retention
- insensitivity to other ions
A mutant Renilla luciferase, acting as the donor, catalyzes the oxidation of its substrate, luciferin, which releases energy as a photon. The energy is transferred to the acceptor fluorophore, a circularly permuted Venus. Luminescence is simultaneously acquired at two wavelengths and a ratio is calculated (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.
Zhang et al.,PLoS One. 2012;7(8):e43072.
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 the binding of the target to the peptide. 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 Dimer (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, for your service.
Figure 1: Comparison of light and dark affinities between the original heterodimer pairs and the two new pairs, iLID nano and iLID micro.
Figure 2: Engineering of an improved Light Inducible Dimer to control cell signalling
December 2014: Mini-transposon Vectors, Golden GATEway Cloning Kit, & MoreArticle Contributors
Updated Mini-transposon Vector for Bacterial Mutagenesis or Gene Targeting
Victor de Lorenzo's lab has engineered a modular mini-Tn5 vector that can be used to generate random mutagenesis libraries or to insert heterologous genes, reporters, or other markers into a target genome. They did this by selecting the important elements from existing transposon and vector systems and creating an all-synthetic vector that included only the elements needed for function.
The lab validated this vector, called pBAM1, by conducting random mutagenesis in the soil bacterium Pseudomonas putida and demonstrate that they can successfully create GFP fusion proteins with a variety of genes across the genome. Although this tool was published in 2011, it was only recently made available through Addgene and we want to highlight it for use in your research.
Martínez-García et al., BMC Microbiology 2011 Feb 22;11:38.
Martínez-García et al., BMC Microbiology 2011 Feb 22;11:38.
APEX2 for Proteomic Mapping and Electron Microscopy
A thorough understanding of complex biological systems requires both a catalog of molecular players and contextualized knowledge of their roles inside cells. The original APEX (enhanced ascorbate peroxidase) reporter from Alice Ting’s lab has enabled progress on both fronts – as a genetic electron microscopy (EM) tag providing superior subcellular localization of tagged proteins compared to traditional fluorescence microscopy, and as a targeted tool for spatial proteomic mapping in living cells. Despite these advances, the Ting lab’s first-generation APEX suffered from inconsistent activity levels and limited sensitivity under the stringently low expression conditions necessary to avoid biological perturbation in a variety of contexts. In order to overcome these limitations, they evolved and characterized an improved peroxidase reporter, APEX2, through yeast display. APEX2 has been shown to exhibit superior performance over APEX for both EM imaging and proximity-dependent proteomic mapping with far greater sensitivity at lower expression levels. APEX2 is a monomeric, 27 kD genetic tag which can be fused to proteins of interest for EM imaging, or targeted to subcellular regions or protein complexes for proteomic mapping in live cells.
Lam et al., Nat Methods 2014 Nov 24. doi: 10.1038/nmeth.3179.
Martell et al., Nat Biotechnol 2012 Nov;30(11):1143-8.
Activity comparison between APEX2 and APEX. HEK-293T cells were transfected with cytosolically-targeted APEX or APEX2 (Addgene ID: 49386) constructs. Separate experiments compared enzymatic activity with the substrates utilized for proteomics (biotin-phenol, left) and EM (diaminobenzidine (DAB), right). Left, extent of APEX/APEX2 labeling with biotin-phenol corresponds to intensity of streptavidin-fluorophore signal. Right, APEX/APEX2 polymerizes DAB to form a light-absorbing precipitate that can be visualized by bright-field microscopy.
Image courtesy of Stephanie Lam.
Highly Bright and Extremely Dim GFPs
The availability of the three dimensional structure of GFP, its brighter mutant S65T-GFP, and the subsequent biochemical characterization, has enabled scientists to engineer a wide variety of GFPs with diversity in fluorescence brightness, intensity, excitation and emission spectra. The lab of Dimitri Deheyn recently identified a family of GFP proteins in the cephalochordate Branchiostoma floridae, named bfloGFPs. They went on to characterize the structural and spectral properties of two of the family members – one with an unprecedentedly-high brightness, bfloGFPa1, and the other with extremely-dim brightness, bfloGFPc1. The plasmids used for determining the structures are now available from Addgene - bfloGFPa1 and bfloGFPc1.
Bomati et al., Sci Rep 2014 Jun 27;4:5469.
Images courtesy of Dimitri Deheyn, Scripps Institution of Oceanography, UCSD
MitoTimer Plasmids to Report Mitochondrial Turnover
Mitochondrial health, transport and turnover are important features to be monitored to study mitochondrial dysfunction. Easy-to-use tools to monitor these features were lacking until Roberta Gottlieb and Zhen Yan labs developed MitoTimer plasmids.
MitoTimer proteins are derived from the Timer protein - a mutant of the fluorescent protein dsRed that changes irreversibly its color from green to red when it is oxidized. The Timer tool has already been used by several groups to monitor protein and cell ageing, as the spectrum shift occurs during Timer protein life due to a form of oxidation (dehydrogenation). MitoTimers are composed of a Timer protein fused to the mitochondrial targeting sequence of cytochrome c oxidase subunit VIII, so that they can be imported to the mitochondrial matrix. Gottlieb’s lab has used an inducible MitoTimer reporter using a Tet-on system (pTRE-Tight-MitoTimer) to show its usefulness in cell culture to report changes in mitochondrial turnover and transport. pMitoTimer from Yan’s lab, which is constitutively expressed, reports on the balance of biogenesis and mitochondrial degradation in vitro and in vivo.
Merged green and red channel fluorescence images of C2C12 cells stably expressing pTRE-Tight-MitoTimer and rtTA. The cells were pulsed with 2µg/ml of Doxycycline for 2 hours, and imaged on the Keyence BZ-9000 fluorescence microscope 8 hours post-pulse. The distinct differences of red-to-green fluorescence in individual mitochondria in the networks marks the differences in import of newly synthesized (green) protein and organelles with older (yellow to orange) protein. Image courtesy of Roberta Gottlieb.
A Quicker Method for Protein Inactivation
Your protein of interest does x, but it also participates in y. Depletion of your protein may upregulate an alternative pathway or induce other compensatory mechanisms. How can protein function be dissected while minimizing confounding secondary effects?
An approach described as “knocksideways” (a British idiom for “taking by surprise”) by Margaret Robinson’s lab, and more recently used by Stephen Royle’s lab, acutely depleted proteins from their compartment of action via sequestration.
The strategy relies on several observations:
- Rapamycin is a membrane-permeable pharmacological agent that can rapidly induce heterodimerization of proteins that contain rapamycin-binding domains, such as FKBP and FRB.
- The cell does not appear to mind extraneous proteins bound to its outer mitochondrial membrane.
Sequestration is accomplished by knocking down your endogenous protein of interest and co-expressing its replacement recombinant protein with a FKBP domain and a mitochondrial outer membrane protein with a FRB domain (MitoTRAP). Rapamycin binds your recombinant protein (via FKBP) and then sequesters the protein at the mitochondria (binding FRB), allowing for a rapid and inducible inactivation. Find theRobinson plasmidsor theRoyle plasmidsand get to work!
Image courtesy of Stephen Royle.
Assemble Multiple Component Plasmids from Building Blocks via Golden GATEway Cloning
Frustrated by the complexity of assembling recombination of transgenesis constructs? A new cloning system combining the advantages of Golden Gate and Multisite Gateway cloning methods developed by the lab of Joachim Wittbrodt might be just the set of tools that you need.
The Golden GATEway cloning kit simplifies the cloning process for complex DNA constructs, particular for those involving recombination elements such as FRT or Lox sites, and provides a modular system for easier exchange and re-use of existing elements. DNA sequences for a desired component are cloned into Golden Gate entry vectors via traditional restriction enzyme cloning, TA cloning, or annealing of oligonucleotides. These entry vectors are used in a Golden Gate cloning step to assemble the individual components into a destination vector in a predefined order. Finally, the destination vectors are used with Multisite Gateway cloning to generate the final construct.
Up to eight entry vectors can be used for each Multisite Gateway compatible destination vector for a maximum of 24 elements assembled into a final transgenesis construct. Plasmids constructed using Golden GATEway cloning have been utilized to create recombination template vectors, to perform multiple site mutagenesis and create complex fusion or recombination vectors. The efficient and flexible cloning process provides improvements to classical cloning methods, particularly for complex transgenesis constructs.
Kirchmaier et al., PLOS One 2013 Oct 7;8(10):e76117.
September 2014: New Channelrhodopsins, Tools for CRISPR gRNA Validation, & More
Shining a Light on Channelrhodopsins – Chronos & Chrimson
Through de novo-sequencing of 127 algal transcriptomes, as well as further optimization through engineering, Ed Boyden’s lab has identified a variety of new channelrhodopsins for use as optogenetics tools. These tools include a pair of channelrhodopsins, Chronos and Chrimson, which have enabled the group to perform independent two-color optical excitation of neurons.
The paper describes five previously unknown channelrhodopsins from different species: Stigleoclonium helveticum (ShChR), Chlamydomonas noctigama (CnChR1), Chloromonas subdivisa (CsChR), Chloromonas oogama (CoChR), and Scherffelia dubia (SdChR). ShChR, nicknamed Chronos, is blue and green light-excitable and has faster kinetics than those of other channelrhodopsins. CnChR1, nicknamed Chrimson, is the first reported yellow-peaked channelrhodopsin with a spectral peak at 590 nm, which is 45 nm more red-shifted compared to other variants. The group further optimized Chrimson through mutagenesis (K176R) to improve the otherwise slow off-kinetics (15.8 ± 0.4ms from 21.4 ± 1.1 ms); this new variant was named ChrimsonR. Plasmids containing Chronos, Chrimson, ChrimsonR, CsChR, and CoChR have been deposited to Addgene, including lentiviral and AAV expression vectors for certain variants.
Klapoetke et al., Nat Methods 2014 Mar 11;(3):338-46.
For more information and descriptions of various optogenetics plasmid tools, visit Addgene’s optogenetics guide.
More Optogenetics Tools – Light-activated Receptor Tyrosine Kinases
Receptor tyrosine kinases (RTKs) are a large class of cell-surface receptors which play a critical role in development and are often implicated in disease progression. One of the major challenges in signaling research is the inability to replicate the spatio-temporal precision with which signaling events occur in a physiological setting. Genetic techniques typically rely on overexpression or knock-down, where signaling dynamics are influenced by the relatively long life-cycle of a protein from expression to degradation. Chemical approaches may require expensive peptides or small molecules and exceed a physiologically relevant exposure in intensity and/or duration. Both of these strategies carry the risk of off-target effects.
Harald Janovjak and his team at the Institute of Science and Technology Austria, decided to take a novel approach to control for these effects, drawing on the rapidly growing field of optogenetics. Using a rational protein engineering approach, they designed Opto-RTKs, which activate signaling cascades on exposure to low-intensity blue light. This approach relies on the incorporation of light-oxygen-voltage (LOV) sensing domains from algae into chimeric Opto-RTKs. The LOV domains bind flavin cofactors and dimerize on exposure to light, bringing the intracellular kinase domains into contact and initiating signaling. Grusch et al. demonstrate that Opto-RTKs have a similar level of background activity and activation as their wild-type counterparts.
The Janovjak lab has deposited their 3 Opto-RTK constructs – Opto-mFGFR1, Opto-hEGFR, and Opto-hRET – as well as various LOV-domain-mVenus constructs used in the study for those who would like to extend this work into their RTK of interest.
Grusch, M. et al., EMBO J 2014 Aug 1;33(15):1713-26.
RUSH & CRUSH – Rapid & Conditional Gene Silencing in RNAi Transgenic Mice
The Jackson-Grusby lab has designed two vectors for use in conditional and reversible gene silencing in RNAi transgenic mouse models and embryonic stem (ES) cells. These vectors, termed RUSH (For ROSA26 U6 short hairpin) and CRUSH (Conditional RUSH) use Cre-mediated recombination to turn on or off the expression of shRNA. This system requires less work than using established methods and minimizes some of the technical issues associated with high levels of shRNA expression. The capability to turn RUSH and CRUSH alleles off or on enables this method to rapidly address questions of tissue-specificity and cell autonomy of gene function in development. To validate the use of the CRUSH vector in transgenic mice, the lab established a dual-color RNAi “sensor” mouse strain in which Cre expression causes the induction of both DsRed fluorescence and GFP shRNA. Mice carrying the R26DsRedR; CRUSH-GFP; and Nestin-Cre alleles showed efficient GFP knockdown in eyes and clonogenic neural stem cells concomitant with activated DsRed2 expression.
Brown et al., Genesis 2014 Jan;52(1):39-48.
A New Tool for CRISPR gRNA Validation
Selecting a gRNA sequence that effectively targets your gene/region of interest is a key step for CRISPR/Cas9 gene editing. Dr. Masahito Ikawa has created a GFP reporter plasmid for scientists to validate the efficacy of their gRNAs.
The first step is to clone the target sequence in between two fragments of EGFP in pCAG-EGxxFP. The EGFP fragments contain 482bp of overlapping sequence that direct Homologous Recombination (HR) or Single Strand Annealing (SSA) in the event of a DNA break. Next, the gRNA being tested is expressed along with Cas9. If the gRNA effectively cuts the target sequence, the plasmid undergoes HR or SSA to reconstitute functional GFP, and the cells will turn green.
Mashiko et al., Sci Rep 2013 Nov 27; 3:3355.
For more CRISPR plasmids and resources, visit Addgene's frequently updated CRISPR/Cas9 guide.
Irreversible Peptide-Peptide Ligation Using SpyLigase
Building off their SpyTag/SpyCatcher system for protein-peptide locking, Mark Howarth’s lab has developed a new tool for peptide-peptide locking. The new technology is known as SpyLigase and is a protein domain that promotes the formation of an isopeptide bond between 2 peptide tags, SpyTag and KTag. The group demonstrated the use of the SpyLigase peptide-peptide interaction to link affibodies or antibodies against common tumor markers to enhance cancer cell capture.
For further reading about SpyLigase technology, read the Addgene interview with Mark Howarth.
Rinehart lab reagents for improved expression of recombinant phosphoproteins
Protein phosphorylation is one of the most abundant forms of posttranslational modifications in cells and research into its many roles in protein function and signaling networks continues to expand. The labs of Jesse Rinehart and Dieter Söll at Yale University previously changed the way researchers can explore important questions surrounding serine phosphorylation by adding this phosphorylated amino acid to the genetic code of E. coli (Park et al., Science 2011).
The Rinehart lab has now made improvements to this system by engineering cells that lack release factor one (RF-1; Bacterial strain EcAR7) and minimizing the set of plasmids required to make singly or multiply phosphorylated proteins (B40 OTS and pCRT7-GFP). To demonstrate the improvements of the system, the Rinehart lab synthesized the activated form of human mitogen-activated ERK activating kinase 1 (MEK1) with either one or two phosphoserine residues cotranslationally inserted in their canonical positions (SP218, SP222) using the original or improved phosphoprotein synthesis reagents. One phosphoserine (SP218) insertion was moderately enhanced while two phosphoserine insertions (SP218/SP222) was dramatically enhanced with the improved system. This MEK vector is also available at Addgene and can be used as a control for your experiments. For more information, please see Addgene’s information page for the Rinehart phosphoprotein system, which includes detailed protocols provided by the Rinehart lab.
pOSIP and the Clonetegration
Classic genetic engineering methods enabling chromosomal integration of sequences in bacteria are time-consuming and involve many steps. The Drew Endy and Keith Shearwin labs have developed a new, streamlined approach to genetic engineering which drastically reduces the time and effort needed to insert new genes into bacteria. They designed the pOSIP (one-step integration plasmid) series of plasmids, vectors that convey both the sequence to be integrated and a removable integrase cassette. They validated this methodology in two differents types of bacteria (E. coli and Salmonella typhimurium) by integrating DNA sequences either sequentially or simultaneously.
This method, called by the authors the “Clonetegration”, is quick and easy to do. Clonetegration could become a “valuable technique facilitating genetic engineering with difficult-to-clone sequences and rapid construction of synthetic biological systems” as they predict. The pOSIP plasmid kit can be found at Addgene, so what are you waiting for? Start building up your own designer bacteria.
St-Pierre et al., ACS Synth Biol 2013 Sep 20;2(9):537-41.
June 2014: MoClo & Platinum Gate Kits, Davidson Lab FPs, CRISPRs, & More
DREADD-based Chemogenetic Technologies
After several years of distributing his Designer Receptors Exclusively Activated by Designer Drugs (DREADD) plasmids on his own, UNC-Chapel Hill's Bryan Roth has now deposited many of these constructs with Addgene. These G-protein coupled receptors have been engineered by the Roth lab to be activated by otherwise pharmacologically inert drug-like small molecules, allowing labs to precisely and non-invasively control neuronal signaling.
For more information on DREADDs, please see the Roth lab's DREADD users blog.
Michael Davidson Lab Fluorescent Protein Collection
Michael Davidson and his lab from Florida State University have contributed their comprehensive collection of ORFs tagged with a variety of fluorescent proteins. In addition to this collection of ORFs, over 100 empty backbones are available from the Davidson lab for tagging your gene of interest.
The Davidson plasmid collection includes excitation/emission, localization, sequence, and supplemental information for many of the plasmids so you can easily find what you need.
We currently have ~ 300 plasmids (empty backbones and mEmerald tagged ORFs) available for request, with more plasmids becoming available everyday. Browse the collection here.
For more information on these tools, visit Davidson's Molecular Expressions website for additional images, tutorials, optical microscopy protocols, and many more resources.
Looking for other fluorescent proteins? Find more at Addgene’s Fluorescent Protein Guide.
New CRISPR Plasmids Available!
New Lentiviral CRISPR activator and repressor plasmids from Scot Wolfe's lab. These include Tet-inducible CRISPR activators and repressor plasmids. (Kearns et al., Development. 2014..)
CRISPRs for Xenopus! From the lab of Yonglong Chen, pCS2-3xFLAG-NLS-SpCas9-NLS is a Cas9 expression plasmid that was used by Chen and colleagues for genome editing in Xenopus tropicalis. (Guo et al., Development. 2014..)
A new, higher specificity genome editing system that combines TALENs and CRISPRS. Developed by David Liu and colleagues, FokI-dCas9 expresses Fok1 nuclease domain fused to catalytically inactive Cas9 DNA-binding domain in mammalian cells. (Guilinger et al., Nat Biotechnol. 2014..)
Our CRISPR-Cas collection of plasmids updates frequently, so visit our CRISPR pages often to find the most recently deposited tools, resources, protocols, and more.
MoClo Modular Cloning System
Click on image to see full figure and more information about the MoClo Kits available at Addgene. Image from Weber et al., PLoS One. 2011 Feb 18;6(2):e16765.
Synthetic biologists have developed a modular cloning strategy, MoClo, which uses the Type IIS restriction enzymes BsaI and BpiI/BbsI to efficiently assemble up to six DNA fragments at a time. This method (based on the Golden Gate technology) exploits the ability of Type IIS enzymes to cut outside their recognition site, and permits DNA fragments with compatible overhangs to be efficiently assembled. Scientists can engineer unique enzyme recognition sites that flank a DNA module in an inverse orientation, so that multiple DNA components can directionally assemble in a single reaction, while retaining only a defined 4bp fusion site in between.
The MoClo system is comprised of three sets of cloning vectors (Level 0, 1, or 2) which can be utilized in three successive assembly steps. Before beginning, scientists can insert fragments of DNA containing basic parts (promoters, UTRs, coding sequences, terminators, etc) into individual Level 0 plasmids, or choose from a growing number of libraries containing pre-constructed standardized modules. In the first assembly step, compatible Level 0 vectors are directionally assembled into a Level 1 vector creating a single transcriptional unit (Ex: a promoter, 5’UTR, coding region, and terminator). Next, up to six Level 1 modules can be similarly assembled into a Level 2 vector, thus forming a functional genetic circuit. Flexibility has been built into the Level 2 vectors to allow for additional iterations of Level 1 assembly if necessary. Combining multiple Level 2 vectors in the final assembly step permits the creation of more complex constructs constrained only by the ability of E. coli to maintain the final plasmid after transformation.
Addgene depositors Sylvestre Marillonnet and Nicola Patron have assembled two collections of standardized genetic modules compatible with the MoClo system. The MoClo Toolkit provided by the Marillonnet Lab can be used to assembly general eukaryotic multigene constructs, while the Patron Lab MoClo Plant Parts kit contains modules specific for plant transformation.
Weber et al., PLoS One . 2011. Feb 18;6(2):e16765.
Engler et al., ACS Synth Biol 2014. Feb 5 (Web); DOI: 10.1021/sb4001504.
For further reading about Nicola Patron's MoClo kit and her plant synbio research, read the Addgene interview.
Engineering TALENs containing variable-repeats using Platinum Gate system
Interested in optimizing TALEN assembly and activity? The laboratory of Takashi Yamamoto has created a complete TALEN assembly system after systematically analyzing the effect of both the TALE scaffold and module on TALEN activity in a single-strand annealing (SSA) assay.
This new Platinum Gate TALEN Kit utilizes a 4-module assembly system, which reduces the number of individual repeat-variable di-residue (RVD) module plasmids and simultaneously increases the success rate of module assembly in the first Golden Gate reaction. While the DNA-binding specificity is imparted by the RVD at residues 12 and 13 in the TALE repeat, other naturally-occurring variations in the TALE repeat, referred to as “non-RVD variations”, were found to improve TALEN activity. Each positional group of module plasmids (ex. p1HD, p1NG, p1NI & p1NN) in the Platinum Gate system contains an identical variable repeat (VR) at the 4th and 32nd residues of the TALE repeat and the VR differs between the positional groups. Eight final destination vectors, consisting of all four final RVD modules in each of two different TALE scaffolds are provided, as the optimal scaffold can depend on the length of the spacer region between the target sequences. These updates to TALEN genome engineering demonstrate improved efficiency over previous reports.
Click on image to see full figure and more information about the Platinum Gate TALEN Kit. Figure from Sakuma et al., Sci Rep (2013).
For additional information on using the Platinum Gate TALEN Kit, please see the Yamamoto lab’s protocol for TALEN construction.
- Sakuma et al., Sci Rep . 2013. Nov 29;3:3379.
Interested in more genome engineering technologies? Browse others on Addgene's Genome Engineering Guide.
March 2014: New Neuronal Imaging Tools, GreenGate Cloning System, & More
Fire Up Those Neurons: mGRASP
A Nature 2012 article by Jinny Kim and colleagues describes their efforts to map the location and distribution of synapses in the mouse brain. Kim et al is utilizing a mammalian GRASP (GFP reconstitution across synaptic partners) technique based on functional complementation between two non-fluorescent split GFP fragments. When the two fragments, expressed in the presynaptic region of one neuron and postsynaptic region of a different neuron, come into proximity in the synaptic cleft, functional fluorescent GFP is reconstituted in vivo. Currently available from the Kim lab are 2 presynaptic and 2 postsynaptic targeting mGRASP plasmids. Additionally, the lab recently described the use of another set of mGRASP plasmids in their Neuron 2014 paper.
Next-Gen Brainbow Toolkit for Neuronal Imaging
Joshua Sanes and his team at the Center for Brain Science at Harvard University have developed a next-generation Brainbow toolkit for high-resolution fluorescent imaging of individual neurons. The technology generates a unique spectral identity for each cell in a population by expressing a randomly generated mix of fluorescent proteins, determined by competing recombination events at the genetic level. Upon Cre/loxP recombination, each transgene expresses one of three possible fluorescent proteins, chosen for minimal spectral overlap, minimal protein aggregation, and high photostabilty. When multiple cassettes are integrated, each recombines independently, generating tens or hundreds of possible combinations (depending on the number of copies). This facilitates the distinction of neighboring cells in imaging applications and the mapping of neuronal projections to their associated cell bodies.
The first versions of the Brainbow system were reported in 2007. In order to extend the utility of the system, the researchers developed three new versions. Flipbow employs the Flp recombinase/FRT system in place of Brainbow’s Cre/loxP, allowing for simultaneous use of the two systems in different tissues of the same animal. Flipbow additionally incorporates SUMO tags in the FP sequence for separation from Cre-based Brainbow-expressing cells. Autobow plasmids are all-in-one versions which express self-excising Cre recombinase from the same transgene as the Brainbow cassette, simplifying experiments when additional cross-breeding steps are undesired or infeasible. Finally, an adeno-associated viral (AAV) system enables greater spatio-temporal control over expression and increases the number of species in which Brainbow may be used. This system uses two plasmids in tandem, with Cre-dependent inversion determining between 0 and 2 FPs expressed from each copy. Brainbow, Flipbow, and Autobow systems are available with either the Thy1 or CAG promoter, while Brainbow AAV is under control of the EF1a promoter.
pCoofy Vectors for Optimizing Protein Expression
Under the direction of Sabine Suppmann, the Recombinant Protein Production group at Max-Planck Institute of Biochemistry has developed a number of expression vectors for use with Sequence and Ligation Independent Cloning (SLIC). The pCoofy series of plasmids contain a variety of N- and C-terminal tags (including His, S-tag, OneStrep, CBP, Trx, GST, Halo, MBP, NusA and SUMO) for optimizing expression, solubilization and purification and have been tested in bacterial, insect and mammalian cells. These vectors were designed for parallel testing and screening of constructs in multiple host cells in order to optimize expression. The expression plasmids for a given species are based on the same backbone to permit expression levels to be directly compared amongst the different tags
To clone a sequence or gene of interest into the pCoofy vectors, select the appropriate pair of vector and gene primers from Table 2 of the associated publication which contain regions of sequence homology between the vector and gene primer for SLIC cloning. Amplify the pCoofy vector and the sequence of gene of interest in separate PCR amplication reactions for recombination in SLIC cloning to form the desired vector. The pCoofy vectors contain the ccdB cassette, which is not copied during the PCR amplification step, so that only vectors with the desired sequence of interest are retained after SLIC cloning. The general cloning strategy described in the associated publication can be used to generate any combination of tags for optimizing protein expression and purification in a fast, efficient and affordable way.
Scholz et al.., BMC Biotechnology. 2013. Feb 14;13:12.
Image from Scholz et al.., BMC Biotechnology. 2013. Feb 14;13:12.
GreenGate Cloning System for Plant Transgenesis
Developed by Jan Lohmann and colleagues, GreenGate is a cloning system for the rapid assembly of plant transformation constructs. As the name suggests, GreenGate is based on the Golden Gate cloning method, but has been modified specifically to improve plant transgenesis. The GreenGate kit available at Addgene includes six individual types of pre-cloned insert modules (plant promoter, N-terminal tag, coding sequence of the gene of interest, C-terminal tag, plant terminator, and plant resistance cassette) in pUC19 based entry vectors, as well as the pGreen-IIS based destination vectors.
To learn more about the GreenGate cloning system, see the detailed plasmid kit page or read our blog post: Quick, Versatile Plant Transgenesis with GreenGate Plasmids.
Lampropoulos et al., PLoS One . 2013. Dec 20;8(12):e83043.
Lentiviral CRISPR Libraries for Knockout Screening
New systems have been developed and deposited with Addgene which allow scientists to use CRISPR-Cas technology to perform genome-wide knockout screens. These vectors and sgRNA libraries expand upon the CRISPR family of plasmids by offering a lentivirus-based mechanism for sgRNA delivery and providing a means for large scale functional screens.
For more information on these new CRISPR screening tools, see our CRISPR/Cas Plasmids: Pooled Libraries webpage or read our blog post, Lentiviral CRISPR Libraries Enable Genome-Scale, Knockout Screening.
Also, check out our updated CRISPR-Cas resources at www.addgene.org/CRISPR/ to browse CRISPR plasmids, watch informational videos, download protocols, and more! Looking for backgroubnd information on CRISPR technology? See our improved CRISPR-Cas Guide.
December 2013: Light Controlled Genome Editing, Hydrogen Peroxide Sensor, CRISPRs, & More
Affinity and Fluorescent Protein Tagged Bacterial Expression Vectors
Looking for bacterial expression vectors with affinity tags for purification or fluorescent reporter gene fusions? The laboratory of Thorben Dammeyer constructed a set of plasmids with several combinations of affinity tags and fluorescent YFP-fusion proteins for periplasmic and cytoplasmic expression.
The plasmids in the pTD series share a broad host range RK2 origin of replication and a strong, IPTG-inducible lacIq-Ptrc promoter. Variations in the presence and location of Strep, Twin-Strep, and His tags allow for affinity purifications or co-purifications, while an optional pelB signal sequence utilizes the secretory pathway for export to the periplasmic space. A synthetically engineered EYFP variant was created to permit expression of a mature, active EYFP in the periplasm of Gram negative bacteria such as E. coli and P. putida, as standard EGFP and EYFP are unable to mature in the periplasm after export by the secretory pathway. These plasmids are based on the Standard European Vector Architecture (SEVA) platform to permit exchange of the origin of replication, promoter/MCS and antibiotic resistance modules with other SEVA compatible modules.
Dammeyer et al., Microb Cell Fact . 2013. May 20;12(1):49.
Light Controlled Genome Editing: LITE
Optogenetics meets genome editing in the newest tools developed by the lab of Feng Zhang. These light-inducible transcriptional effectors (LITEs) are designed to bind specific genes and turn them on or off in response to light. These LITEs have been packaged in viral vectors and can be targeted to specific cell populations. Konermann et al. demonstrated the use of these tools to control gene expression in mouse neurons and in the brains of living mice.
For more about these new optogenetic tools, check out our blog post: Let There Be LITE Plasmids.
Konermann et al., Nature . 2013. Aug 22; 500: 472–476.
HyPer3: Fluorescent Protein Sensor for Reactive Oxygen Species
The lab of Vsevolod Belousov has deposited their latest hydrogen peroxide sensor, HyPer3. In the presence of reactive oxygen species (ROS), the excitation properties of HyPer3 change such that the intensity of the emitted light (516 nm) from 500 nm excitation increases relative to that emitted by 420 nm excitation (increase in 500/420 ratio). The new generation sensor has greater dynamic range than HyPer1, faster response times than HyPer2, and is available for either mammalian or bacterial expression. Use it to track ROS changes in real time by fluorescence imaging.
Bilan et al., ACS Chem Biol . 2013. Mar 15; 8(3):535-42.
Latest CRISPR-Cas9 Plasmids
The genome engineering technology known as CRISPR/Cas has recently been utilized in exciting new ways.
Scientists have devised ways to harness Cas9 nuclease to activate or repress genes. This was accomplished by fusing known transcriptional activator proteins (example VP64) or repressor proteins (example KRAB domain) to a catalytically inactive Cas9 nuclease. When targeted to promoter regions by a specific gRNA, these activator or repressor proteins have been shown to up or down regulate gene expression.
Additionally, multiple research groups have recently identified and utilized the type II CRISPR/Cas systems from several different bacterial species. For reference, the original discovery and application of CRISPR genome engineering technology utilized the type II CRISPR/Cas system from Streptococcus pyogenes. The key difference between these new CRISPR systems is the unique PAM (Protospacer-Adjacent Motif) sequence recognized by the Cas9 nuclease in each species. Different PAM sequences allow for an increase in potential target sites for gRNAs (a gRNA can be targeted to any sequence in the genome that ends with an appropriate PAM sequence). Different PAM sequences also allow for multiple, simultaneous CRISPR/Cas driven genome manipulations. For example, a cutting Cas9 from S. pyogenes and an activating Cas9 from N. meningitidis can function within the same cell, at the same time, without interfering with one another. These advances add increased functionality to the already versatile system that is CRISPR.
Interested in reading about the history of CRISPR-Cas technology? Checkout our blog post: History of CRISPR Cas - A tale of survival and evolution.
pDusk and pDawn: Light Regulated Bacterial Expression Plasmids
Building on advances in optogenetics, Andreas Mӧglich's lab has built pDusk and pDawn, two complementary plasmids for light regulated expression of recombinant proteins in E. Coli. These plasmids rely on the engineered two-component regulatory system YF1/FixJ. YF1 is a synthetic, photosensitive kinase, which uses the ubiquitous flavin mononucleotide as its chromophore, and phosphorylates the transcriptional activator FixJ in the absence of blue light. Phosphorylated FixJ is able to drive high levels of gene expression from the FixK2 promoter. These elements are the basis for pDusk, which allows for insertion of your gene of interest directly downstream of the FixK2 promoter. Genes cloned into the multiple-cloning site (MCS) of pDusk will be expressed in the absence of blue or ambient (white) light, and expression levels can be varied with light intensity.
Ohlendorf et al., J Mol Biol. 2012. Mar 2, 416(4):534-42.
The complementary plasmid pDawn is a light-activated expression system, and may be used in combination with pDusk in experiments where alternating expression of different genes is desired. pDawn contains all of the elements of pDusk, except that phosphorylated FixJ now drives expression of the λ phage repressor cI, which in turn represses gene expression from the λ promoter pR, located upstream of the MCS. Because of the strength of both the λ phage repressor and promoter, pDawn has both lower background expression and better induction than pDusk, making it the better choice for preparative expression. After transformation, cultures can be grown to the desired density in the dark, and induced for expression by exposure to blue or ambient light. This obviates the need for chemical inducers such as IPTG, saving money and reducing potential exposures to contaminants. A big advantage of light vs. chemical induction is the ability to turn off expression by removing the light source.
Ohlendorf et al., J Mol Biol . 2012. Mar 2, 416(4):534-42.
Looking for other plasmids for optogenetics research? Check out Addgene’s Optogenetics Guide.
September 2013: Tools for Proteomic Mapping, NIR Fluorescent Probes, Newest TALEN Kit, & More
Tool for Proteomic Mapping of Mitochondria in Living Cells
Alice Ting's lab has designed a new technology for creating a spatially and temporally resolved proteomic map of large numbers of proteins in living cells. The method works by targeting ascorbate peroxidase (APEX) within the cell, resulting in biotinylation of nearby proteins. The biotinylated proteins are then analyzed by mass spectrometry, providing a readout of colocalized proteins from live cells.
The Ting Lab validated this technique by localizing APEX to the mitochondrial matrix with the plasmid pcDNA3-mito-APEX. This resulted in the identification of 495 proteins within the human mitochondrial matrix, including 31 which were not previously linked to the mitochondria. Browse the relevant plasmids.
Rhee et al., Science . 2013. Mar 15; 339(6125):1328-31.
Gateway-compatible Cloning and Expression Vectors
Have you ever wished that your favorite empty vector or tag was available in a Gateway-compatible version? We may have exactly what you are looking for already in our repository.
Image from Dubin et al., Plant Methods (Biomed Central). 2008. Jan 22; 4:3.
The lab of Giovanna Benvenuto created a series of twelve Gateway Entry vectors with six commonly used tags for either N- or C-terminal fusions. The available tags (STREP, HA, MYC, GST, ECFP and EYFP) are present within the attachment sites, avoiding any extra linker amino acids between the tag and insert. Traditional restriction enzyme cloning is used to insert a gene of interest into these Entry vectors, followed by recombination with a Gateway-compatible Destination vector of your choice for bacterial, insect, mammalian, plant or yeast expression. All of the vectors utilize the same cloning sites and contain a stop codon before the final attachment site in the entry cassette.
Converting existing vectors to Gateway-compatible vectors that can be used with recombination-based cloning can be a tedious process. Fortunately, the Yu-Zhu Zhang laboratory has developed a method using site-specific recombination to convert non-Gateway based vectors to Gateway-compatible vectors. This process was then used to create eleven Gateway-compatible Destination vectors from commonly used conventional empty vectors containing His, hemoglobin, EGFP, Flag, Myc-His tags or from other empty vectors used in adenoviral, bacterial, mammalian or yeast systems. An Entry vector containing your gene of interest can be recombined with one of these Gateway-compatible Destination vectors to generate an expression-ready construct.
More empty backbones can be found at Addgene’s Empty Backbones Guide.
New NIR Fluorescent Probes: iRFPs, PAiRFPs, and iSplit
Several types of new near-infrared fluorescent proteins derived from bacterial phytochrome photoreceptors (BphPs) have been developed by Vladislav Verkhusha’s lab and can be used for deep-tissue optical in vivo imaging. They fluoresce in mammalian cells and tissues without adding exogenous biliverdin. The four new spectrally distinct permanently fluorescent iRFP variants (iRFP670, iRFP682, iRFP702, and iRFP720) described by Shcherbakova et al., along with the group’s original iRFP (iRFP713), were shown to have high effective brightness and allowed multicolor imaging. Next, Piatkevich et al. described the engineering of photo-activatable iRFPs (PAiRFP1 and PAiRFP2), which can be ‘turned on’ by non-phototoxic far-red light and used for spatially selective imaging of tissues in living animals. Most recently, further development of the original iRFP resulted in a split fluorescence complementation probe, iSplit, by Filonov et al. iSplit was tested both in vitro and in vivo as a biomolecular fluorescence complementation (BiFC) reporter to detect protein-protein interactions.
Shcherbakova et al., Nat Methods . 2013. Jun 16; 10(8):751-4.
Piatkevich et al., Nat Commun . 2013. Jul 10; 4:2153.
Filonov et al., Chem Biol . 2013. Aug 22; 20(8):1078-86.
Looking for other fluorescent proteins? Check out Addgene’s Fluorescent Protein Guide.
The Open Source Wnt Project
Wnt signalling pathways play essential roles in embryonic development as well as tissue homoeostasis in adults, and their aberrant regulation has been linked to diseases in man including diabetes, neurodegeneration and cancer. In order to allow direct side by side comparison of the various mammalian Wnts and their function, the labs of Marian Waterman and David Virshup have developed a standardized set of Wnt expression plasmids. The kit contains the ORFs of all 19 human Wnts in the same expression backbone. Each ORF is cloned into 2 entry backbones, pENTR/D-TOPO with and without a STOP codon, and 2 mammalian expression backbones, with and without a C-terminal V5 tag. In addition, the Xi He lab has contributed a fifth set containing a modified version of each tagged ORF that enables epitope tagging without loss of Wnt signaling activity.
Najdi et al., Differentiation . 2012. Sep; 84(2):203-13.
Learn more about how the Open Source Wnt Kit was developed using crowd-sourcing in our interview with Dr. Marian Waterman.
Genome Engineering in hPSCs: New Musunuru/Cowan TALEN Kit
Developed by the labs of Kiran Musunuru and Chad Cowan, the newest TALEN kit allows for the quick and easy delivery of TALENs into human pluripotent stem cells and other difficult-to-transfect mammalian cell types. TALEN construction can be completed in 1-2 days without PCR amplification.
- Gene knockout by indel mutation induction
- Reporter line generation
- Correcting causal mutation in iPSC lines
- And more!
Ding et al., Cell Stem Cell . 2013. Feb 7;12(2):238-51.
Check out the Latest CRISPR Plasmids
Do you want to edit plant, fly, worm, or fish genomes? We now have CRISPRs for that! Are you interested in activating your gene of interest? New CRISPR technology lets you selectively activate your gene of choice!
Kamoun Lab: Using CRISPRs to modify plant genomes.
“Targeted mutagenesis in the model plant Nicotiana benthamiana using Cas9 RNA-guided endonuclease.” Nat Biotechnol . 2013.
Joung Lab: A new use of the CRISPR/Cas9 system to target and activate specific genes.
“CRISPR RNA-guided activation of endogenous human genes.” Nat Methods . 2013.
Chen and Wente Labs: Using CRISPRs to modify the zebrafish genome.
“Efficient multiplex biallelic zebrafish genome editing using a CRISPR nuclease system.” PNAS . 2013.
Calarco Lab: Using CRISPRs to modify the C. elegans genome.
“Heritable genome editing in C. elegans via a CRISPR-Cas9 system.” Nat Methods . 2013.
O’Connor-Giles, Wildonger, and Harrison Labs: Using CRISPRs to modify Drosophila genome.
“Genome Engineering of Drosophila with the CRISPR RNA-Guided Cas9 Nuclease.” Genetics . 2013.
Goldstein Lab: Using CRISPRs to modify the C. elegans genome.
“Engineering the Caenorhabditis elegans genome using Cas9-triggered homologous recombination.” Nat Methods . 2013.
Don't see the CRISPR/Cas system you're looking for here? Find more CRISPR/Cas9 plasmids at Addgene.
June 2013: Fluorescent Protein Kit, CRISPRi, New Vectors for Use with Golden Gate TALEN Kit
New Plasmid Kit: Fluorescent Proteins
Organizing your fluorophore combinations at the start of a research project can be a tricky task. Many times, having access to additional colors or combinations can be a big asset. The Hamdoun lab constructed a number of fluorescent plasmids for their recent JBC publication, and the fluorescent protein- containing empty vectors have wide applications for use in Zebrafish, Sea urchin, Xenopus, and C. elegans. The following 9 plasmids have been bundled together to provide a useful “starter kit” for screening fluorescent protein fusion expression in organisms or cells in which an exogenous mRNA can be injected and expressed. The utility of this kit is that it enables the user to generate N and C terminal fusions to mCherry, Cerulean, mCitrine, or EGFP, and could also be employed for many types of multicolor experiments or assays with fluorescent small organic molecules.
Gokirmak et al., J Biol Chem . 2012. Dec 21; 287(52):43876-83E.
CRISPRi: Repurposing of the CRISPR/Cas Genome-Editing Technology for Transcriptional Silencing
The CRISPR/Cas system is quickly becoming well known as an effective genome-editing technology, and a simple alternative to TALENs. With only two components, CRISPR systems utilize Cas9 nuclease to cleave DNA and chimeric guide RNA (gRNA) to target the Cas9 to a specific region of the genome. The system has been optimized for DNA editing in a variety of different species and cell types. The current technology utilizes Cas9’s nuclease abilities to destroy a specific locus in DNA through directed cleavage followed by non-homologous end joining (NHEJ). Alternatively, a modified Cas9 can be made to nick the DNA, cutting only one of the DNA strands to facilitate DNA replacement by homologous recombination. This requires the transfection of an additional plasmid that contains a complimentary sequence.
The Stanley Qi lab has created a new function of the CRISPR/Cas system called CRISPR Interference. CRISPRi is the latest tool available to scientists looking to manipulate the genome of their favorite organism. CRISPRi utilizes a catalytically inactive Cas9 nuclease in complex with a gRNA to interfere with transcription of the DNA downstream of its binding site. The mechanism of the interference is believed to involve physically impairing the RNA polymerase progression past the Cas9:gRNA complex. These CRISPRi plasmids are optimized for use in human cells and an additional set of CRISPRi plasmids are optimized for use in bacteria.
Qi et al., Cell . 2013. Feb 28;152(5):1173-83.
Alternative TALEN Assembly and Validation Vectors for Golden Gate TALEN Kit
Recent advances in genome editing technology, such as transcription activator-like effector nuclease (TALEN) systems, have reduced the barrier to studying gene function. With a focus on efficiency, the laboratory of Takashi Yamamoto has developed optimized array and destination vectors for use in combination with original vectors from the Golden Gate TALEN and TAL Effector kit, deposited by the labs of Dan Voytas and Adam Bogdanove. The modified pFUS array vectors are designed for six-module assembly to improve the assembly success rate and efficiency of the first Golden Gate cloning step, while reducing the number of necessary RVD module vectors. An optional ligation with pre-digested RVD and array vectors for the first module assembly step is reportedly more robust than the typical Golden Gate assembly method. The mammalian destination vectors offer the choice of a CMV/T7 or CAG promoter with codon-optimized FokI and Flag tag, ready for use in mammalian cells without additional cloning.
A novel TALEN evaluation system utilizing the pGL4-SSA vector included in the Yamamoto Lab TALEN Accessory Pack allows for validation of TALEN plasmids in mammalian cells using a luciferase reporter containing custom oligonucleotides corresponding to the TALEN target sequence in a single-strand annealing (SSA) assay. The Yamamoto lab has provided a step-by-step protocol describing the use of their array and destination vectors, as well as this universal TALEN validation assay in mammalian cells.
Sakuma et al., Genes Cells . 2013. Apr;18(4):315-26.
FREQ-Seq: a rapid method to determine specific allele frequencies from mixed populations.
To help scientists track the frequencies of specific alleles in microbial populations through time Christopher Marx's lab has engineered and validated a new method called FREQ-Seq. This strategy allows scientists to construct barcoded, locus-spe cific libraries compatible with Illumina next generation sequencing in order to study evolutionary dynamics. By counting DNA sequence reads, FREQ-Seq can quantitatively determine allele frequencies across timepoints or populations. This kit consists of a 48 plasmid adaptor library, with each plasmid carrying a unique barcoded Illumina-M13F bridging primer. These bridging primers are amplified and used to generate Illumina sequencing libraries using a 2-step PCR-based protocol and are compatible with single-end or paired-end read flow cells. FREQ-Seq is an open source platform and the libraries can be generated in a cost-effective manner with minimal bias.
Chubiz et al., PLoS One . 2012.; 7(10): e47959. doi:10.1371/journal.pone.0047959.
Linearly Tunable Gene Expression Systems
Experimental dose–response data for htetR::NLS::eGFP and mCherry expression in the two gene mammalian linearizer system plotted on log-log scale after background subtraction (defined as the fluorescence at 0 ng/mL of doxycycline).
Gábor Balázsi’s lab has developed a series of systems that allow for tunable gene expression. The group first developed the system in yeast by creating a synthetic linearizer gene circuit that controlled gene expression with a linear dependency on the extracellular concentration of an inducer (anhydrotetracycline). The use of a negative feedback circuit also resulted in gene expression that was homogenous across the cell population. Plasmids for the various reporter and regulator parts of the circuits are available at Addgene.
Using computational modeling as a guide, the circuit was further adapted for use in mammalian cells. Enhancements to the mammalian gene expression system were sequentially made by identifying additions to the circuit that would improve transcription, translation, and nuclear localization (including addition of an intron, use of a nuclear localization sequence, generation of new TetR-repressible promoters, and more). Gene circuits were successfully developed that showed a linearly tunable expression response to the doxycycline inducer when tested in MCF-7 cells, both for a one- and two-gene linearizer system. The related constructs can be found here.
PCR-mediated Modification of Chromosomal Genes in Yeast
PCR-mediated modification and deletion of chromosomal genes is a tried-and-tested technique for analyzing gene function in S. cerevisiae.
Peter Philippsen's and John Pringle's laboratories have created a set of plasmids that serve as templates for the PCR synthesis of fragments used for a variety of gene modifications. The modifications, based on the groundbreaking work by Baudin et al., include gene deletion, gene overexpression (using the regulatable GAL1 promoter), and tagging with a variety of epitopes and fusion proteins. Browse a table of these plasmids here.
Tim Formosa's lab has constructed 36 plasmids that can also be used to generate linear PCR products. The PCR products can easily be fused to the 3' end of an ORF in the yeast genome, thereby adding a variety of tags preceded by a TEV or PreScission protease site. Browse these plasmids here.
Longtine et al., Yeast . 1998. Jul;14(10):953-61.
March 2013: Fluorescent Biosensors/Markers, Improved Reporter for HTS, New Lentiviral Vectors
New Optogenetic Tool Derived from Box Jellyfish - JellyOp
Optogenetics, a recent technological breakthrough in neuroscience, combines the fields of optics and genetics to allow for precise spatial and temporal control of individual neurons. Photons stimulate cells expressing microbial light-gated ion channels, such as channelrhodopsin-2 and halorhodopsin, to modulate neuronal firing. The laboratory of Robert Lucas designed a new optogenetic tool incorporating an opsin from the box jellyfish to achieve improved sensitivity and reproducibility in signaling. JellyOp is more bleach resistant than existing variants of mammalian rod opsin and allows for sustained signaling under conditions of repeated light exposure. The present JellyOp construct is ideally suited for mimicking the activity of Gs-coupled G protein coupled receptors, while further structural modifications could permit coupling of JellyOp to additional signaling pathways.
Bailes et al., PLoS ONE . 7.(1): e30774. doi:10.1371/journal.pone.0030774.
Additional plasmids for optogenetics research can be found on Addgene's Optogenetics Guide.
Optimized Glutamate-sensing Fluorescent Reporter
Glutamate signaling is important in many species and biosensors are allowing scientists to study this process with greater ease and resolution. The Looger lab has created a new intensity-based glutamate sensing fluorescent reporter and has validated it in a wide variety of neurological systems. To learn more and browse these plasmids, click here.
Marvin et al., Nat Methods . 2013. Feb; 10(2):162-70.
FLuc-P2A-RLuc - A New Reporter System for High Throughput Screening
High throughput screening (HTS) with a single reporter-gene is a convenient method for rapidly assaying diverse compounds (such as small molecules) to identify those that may modulate a specific biomolecular pathway. Reporter-gene HTS identifies the ‘active’ compounds by measuring the amount of the reporter-gene activity compared to controls and thus provides a great starting point for downstream experiments. Unfortunately, direct interactions between compounds and the reporter-gene itself can cause misleading, false-positive results that complicate data interpretation.
To overcome this problem, James Inglese’s lab has devised a “coincidence reporter biocircuit” system that expresses two unique bioluminescent genes (firefly and renilla lucifereases) at equivalent levels in order to efficiently distinguish compounds with active biological activity from those that are interfering with the reporter-gene itself. Because the two luciferase genes are nonhomologous, any compound that shows activity from both reporters has higher probability of being biologically relevant.
Cheng KC, et al. Nat Methods . 2012. Oct;9(10):937.
RGB-Marking with LeGO Vectors
The lab of Boris Fehse initially described their lentiviral "gene ontology" (LeGO) vectors as a system "that allows simultaneous expressing and/or suppression of several genes in a single cell to facilitate the analysis of gene networks." These 3rd-generation lentivectors consist of various combinations of fluorescent markers, promoters/enhancers, and shRNA expression cassettes, all of which can be used in conjunction with each other to visualize complex systems.
The Fehse lab recently published a new application for the LeGO vectors: Red, Green, Blue (RGB) Marking. The fluorescent protein marking system is based on the same principle as TV screens, which combine red, green, and blue beams of light at different intensities to make all colors (see the layman's explanation here). The applications go beyond making pretty images--the ability to produce an almost limitless number of colors allows researchers to identify and track single cell clones. For example, all pink cells in a pink cell cluster are derived from a single pink cell.
This unique marker system gives scientists the means to follow tumor clonalities and to investigate the development of monoclonal or polyclonal metastases. Also, in a setting of organ regeneration after transplantation of stem cells, RGB marking enables the visualization of what individual cells are doing after engraftment. The number of cell clusters with different colors shows the number of engrafted cells, and the size of each cell cluster shows how often the cell has divided after engraftment. There are multiple LeGO vectors available, but all you need to get started with RGB marking is a standard fluorescence microscope and the following three LeGO vectors: LeGO-Cer2, LeGO-C2, and LeGO-V2.
Newest pLX Lentiviral Expression Vectors
From the laboratory of David Root, these newest '300' series pLX vectors (pLEX) are similar in function to the previous 300s, but with new promoters and selectable markers. For example, while pLX301 - 4 utilize the CMV promoter, which is known to be silenced in ES cells, pLEX_307 contains the EF1a promoter which is strongly expressed in ES cells.
The '400' series provides all-in-one doxycycline inducibility. These vectors show almost no leakiness in the off-state, allowing the user to titrate expression by varying the levels of doxycycline. This is ideal for avoiding spurious phenotypes caused by expression far above endogenous levels, such as when performing RNAi rescue experiments or when trying to compare activities of gene variants. Available new pLX plasmids include:
Constitutive Lentiviral Expression
- pLEX_305: SV40-puro; PGK-gateway-no tag
- pLEX_306: SV40-puro; PGK-gateway-V5 tag
- pLEX_307: SV40-puro; EF1a-gateway-V5 tag
Inducible Lentiviral Expression
December 2012: New Tools for Imaging, Golden Gate TALEN Add-Ons, & More
New TALEN Destination Vectors – pc-GOLDYTALEN & RCIscript-GoldyTALEN
Transcription activator-like effector nucleases (TALENs) consist of assembled DNA binding motifs coupled to FokI nuclease monomers that can dimerize and introduce a DNA double strand break. In the past year TALENs have become the tool for genome editing primarily due to their simple and straightforward design and assembly strategies. The laboratory of Dan Carlson and Stephen Ekker designed a new and improved TALEN scaffold, GoldyTALEN, truncated at both the N and C terminus and inducing higher mutation rates than the parental pTAL vector. Dan Carlson deposited 2 destination vectors containing the GoldyTALEN compatible with the Voytas lab Golden Gate TALEN kit. pC-GoldyTALEN directs expression of TALENs from a truncated CAGs promoter. RCIscript-GoldyTALEN is designed for in vitro synthesis of TALEN mRNAs. Both 5’ and 3’ Xenopus β-globin UTRs are included in the vector to enhance expression of the message.
Carlson et al., PNAS . 2012. Oct; 109(43):17382-7.
Improved Genetically Encoded Calcium Indicators - GCaMP6 Variants
The capacity to image and measure neuronal activity in vivo has been improved by the development of various calcium (Ca2+) indicators. Such indicators bind Ca2+ and induce a change in fluorescence signal, allowing scientists to measure action potentials and other receptor activation events which trigger Ca2+ fluxes. Genetically encoded calcium indicators (GECIs), such as GCaMP, express indicators in specific tissues or cells.
Douglas Kim’s lab at Janelia Farm recently developed and deposited novel GCaMP6 variants. pGP-CMV-GCaMP6s, pGP-CMV-GCaMP6m, and pGP-CMV-GCaMP6f have increased ΔF/F0 and faster kinetics compared to previous GCaMP3 and GCaMP5G.
Imaging neuronal calcium responses with novel GCaMP6 sensor variants
(A) GCaMP5G (Akerboom et al., 2012) basal fluorescence in rat neurons transfected in culture. Scale bar: 100 µm. (B) GCaMP6s (manuscript submitted) basal fluorescence. (C) Peak GCaMP5G response to 1 action potential stimulus. (D) Peak GCaMP6s response. Fluorescence change (ΔF/F0) is shown in color. (E) Averaged fluorescence traces of neurons after 1 action potential stimulation (arrow) comparing GCaMP6s, 6m, and 6f with GCaMP3 (Tian et al., 2009) and GCaMP5G. GCaMP6 variants were named based on their slow, medium, and fast response kinetics. Data from Tsai-Wen Chen, Trevor J. Wardill, Eric R. Schreiter, Rex A. Kerr, Vivek Jayaraman, Loren L. Looger, Karel Svoboda, Douglas S. Kim; Genetically-Encoded Neuronal Indicator and Effector Project, Janelia Farm Research Campus, Howard Hughes Medical Institute, www.janelia.org/genie.
Live Visualization of Single mRNAs with MS2 and PP7 Systems
Fluorescent in situ hybridization (FISH) has been considered to be the gold standard for labeling nucleic acids in their native environment; however, the technique lacks spatiotemporal resolution available in living cells. An alternative imaging technique has been developed in the laboratory of Robert Singer that permits live detection of single mRNAs by tagging mRNAs of interest with a repeated MS2- or PP7-derived nucleotide sequence. These binding site sequences form hairpin loops that can subsequently bind to the corresponding MS2 or PP7 coat proteins tagged with a fluorescent protein. Modified MS2 and PP7 coat protein constructs, tdMCP and tdPCP respectively, improve labeling and imaging of target mRNAs by reducing fluorescent background to allow for more intricate investigation of mRNA processing.
Wu et al., Biophys J . 2012. Jun 20;102(12):2936-44.
Enhanced FRET Pairing using Clover and mRuby 2
Förster (or Fluorescence) Resonance Energy Transfer (FRET) is an important tool for determining whether two fluorophores are within a certain distance of each other, and is widely utilized to observe and quantify dynamic biological processes. Historically, CFP and YFP have been the most common FRET fluorophore duo; however, limitations such as emissions overlap between the donor-acceptor pair, sub-optimal FRET efficiency/dynamic range, and low photostability make constructing improved fluorophores desirable.
Michael Lin’s group at Stanford University has recently engineered the novel Clover-mRuby2 FRET pair which shows not only the brightest fluorescence for their respective colors (green and red), but also improves FRET efficiency, dynamic range, and photostability while limiting emissions overlap. The Clover-mRuby2 couple was tested in 4 established FRET reporters (Camuiα-CR, AKAR2-CR, VSFP-CR, and Raichu-RhoA-CR) with noticeable improvement, making these updated sensors more useful in detecting rapid cellular processes in real-time. This new FRET pair has great potential not only for enhancing existing sensors, but also for the construction of new, more sensitive FRET reporters.
Lam et al., Nat Methods . 2012. Oct; 9(10):1005-12.
A New Split GFP for Studying in vivo Protein-Protein Interaction – spGFP
A new superpositive split GFP (spGFP) construct for detecting protein-protein interactions in vivo at room temperature was develop by Brian McNaughton's lab. The superpositive split GFP has a greater reassembly speed than previous split GFP constructs. The increased positive charge significantly reduces protein aggregates. This new design also shows robust signal at room temperature making this ideal for studying protein-protein interactions in vivo. The pET11a-Z-NspGFP and pMRBad-Z-CspGFP plasmids are available through Addgene.
Blakeley et al., Mol Biosyst . 2012. Aug; 8(8):2036-40.
EM Imaging in all Cellular Compartments Using APEX
A key component of creating high-resolution electron microscopy (EM) images is contrast. Existing genetic tags designed to increase EM contrast, such as horseradish peroxidase (HRP), are helpful in some cellular locales, but have restrictions. In their recent Nature Biotechnology article, the MIT laboratory of Alice Ting describes the engineering of ascorbate peroxidase (APX) to make a new genetic tag that overcomes the shortcomings of previous EM reporters. The Ting lab introduced a series of mutations to APX to change it from monomeric to dimeric, as well as more highly active towards DAB, thus creating enhanced APX (APEX).
APEX offers the following advantages as a genetic tag EM reporter:
- Fixation and staining of cells does not require a detergent, allowing ultrastructure to be maintained
- APEX does not require light, is easy to use and should have applicability to tissue samples
- The nature of APEX staining makes it a useful tool for 3-D EM applications
- APEX can be fused to fluorescent proteins, allowing for correlative studies using both light microscopy and EM
- The DAB stain generated by APEX is tightly localized, giving spatial resolution in EM on the order of 10 nm
- APEX works in all cell compartments tested, including the cytosol, nucleus, mitochondria, and endoplasmic reticulum
COS7 cell in the metaphase of mitosis
The chromosomes are aligned at the center of the cell. The cell is expressing APEX-Histone2B, which causes APEX to be incorporated throughout chromatin structures. This cell was fixed and stained with 3,3'-diaminobenzidine, resulting in a dark reaction product that highlights chromatin and is visible under a conventional light microscope (top left). The cell was subsequently processed for electron microscopy and imaged with much higher resolution under the electron microscope (low magnification EM at top right; high magnification EM at bottom). Image courtesy of the Ting lab.
APEX plasmids are now available at Addgene.
Martell et al., Nat Biotechnol . 2012. Nov; 30(11):1143-8.
Improved Mos-1-mediated Transgenesis Reagents for C. elegans
Erik Jorgensen's lab had previously created a set of plasmids for targeted transgene insertions (Mos1-mediated single-copy transgene insertions; MosSCI) and targeted deletions (Mos1-mediated deletions; MosDEL) in C. Elegans. In their 2012 Nature Methods paper, they add to this collection, making it even easier to use.
First, they have created a Mos-1 expression vector under the eft-3 promoter that significantly increases the insertion and deletion frequencies. Second, they introduce a set of plasmids to facilitate selection of transgenic strains with antibiotic markers and additional transgene insertion sites. Plasmids from the paper are available from Addgene.
Frokjaer-Jensen et al., Nat Methods . 2012.; 9(2):117-8.
September 2012: Add-Ons for the Golden Gate TALEN Kit, Brighter ECFPs, and More
A Brighter ECFP Variant – mTurquoise2
Cyan based fluorescent proteins suffer from low quantum yield and hence are mostly used as the acceptor, rather than donor, in FRET assays. Using site-directed mutagenesis and fluorescent lifetime-based screening, the Dorus Gadella lab identified mTurquoise2, a variant of ECFP (enhanced cyan fluorescent protein). mTurquoise2 has the highest quantum yield of any monomeric fluorescent protein. In vivo studies in mammalian cells show a 20% gain of brightness, high photostability and better performance in FRET studies.
Addgene distributes several mTurquoise2 vectors with various targeting sequences - mitochondria, nucleus, ER, etc.
Goedhart et al., Nat Commun . 2012.; 3: 751.
Drosophila phiC31 Transgenesis Vectors – pBID and pMartini Gate
Advanced genetic tools available for examining gene function in Drosophila have strongly contributed to its widespread use as a model organism in the genomics era. Integration of transgenes into the Drosophila genome via phiC31 integrase permits efficient and site-specific targeting.
The laboratory of Brian McCabe at Columbia University has recently improved upon phiC31 transgenesis by creating a series of vectors for expressing transgenes in Drosophila. The pBID series of plasmids incorporates several design enhancements, including: (1) gypsy insulator sequences to permit uniform expression levels independent of genomic integration site; (2) backwards compatibility with pUAST cloning sites; (3) Gateway cloning compatibility; (4) DSCP promoter with 10 UAS binding sites to improve expression while eliminating leaky expression in the absence of GAL4; and (5) a variety of fluorescent protein or epitope tagged constructs. The pMartini-Gateway series of plasmids, a set of intermediate vectors, was generated concurrently with the pBID series to assist in Gateway cloning and in developing novel destination vectors.
Wang et al., PLoS ONE . 2012.; 7(7): e42102.
Drosophila transcription factor ORFs
Drosophila melanogaster, one of the best-known model organisms, has been used to advance our knowledge of genetics and developmental biology since first employed over 100 years ago. The complete sequence of the fly genome, published back in 2000, opened many doors to investigating not only the 15,000+ genes, but also the >60% of functional, non-coding DNA found within it. Over the course of the last decade, many methodologies have been employed to identify some of the regulatory elements found within the non-coding DNA; however, the specific functions of these have not been deeply explored as this requires not only identifying the elements, but also the transcription factors that bind to them.
Bart Deplancke’s group from the Laboratory of Systems Biology and Genetics at the Swiss Federal Institute of Technology has recently deposited a nearly complete collection of Drosophila transcription factor open reading frames (ORFs) comprised of 692 plasmids. These ORFs were cloned open-ended into Gateway compatible Entry vectors, permitting easy and efficient subcloning for a variety of downstream applications. Using this clone library, the authors developed and validated a gene-centered, high-throughput yeast 1-hybrid system by which they identified some previously uncharacterized direct interactions between transcription factors and Drosophila cis-regulatory elements. This ORF library will be a significant resource for many scientists studying the biological importance of specific DNA-protein interactions within the Drosophila regulatory gene network.
Browse this whole collection of Drosophila transcription factor ORFs available through Addgene.
Hens et al., Nat Methods . 2011.; 8(12): 1065-70.
Add-Ons for the Golden Gate TALEN Kit
The Golden Gate TALEN kit, deposited by the Voytas and Bogdanove labs, has proven to be Addgene's most popular kit. Due to its popularity, a number of labs designed new plasmids to be used in conjunction with this powerful tool. In the summer of 2012, three new Addgene depositors contributed destination vectors compatible with this kit:
pCS2TAL3-DD and pCS2TAL3-RR
Developed in the lab of David Grunwald, pCS2TAL3-DD and pCS2TAL3-RR are next generation TALEN backbone vectors in place of pTAL1, 2, 3, or 4 from the Golden Gate TALEN kit. For both plasmids, sequence positions 1214–2210 of pTAL3 were cloned into a pCS2 expression vector resulting in shorter N- and C-terminal tal protein segments (136AA and 63AA, respectfully). This next generation architecture has been shown to increase mutation induction when using TALENs. The FokI domains (DD, RR) used are obligate heterodimers that require cloning of left and right TALEN monomer proteins into opposite vectors.
Dahlem et al., PLoS Genet . 2012.; 8(8): e1002861.
pTAL5-BB and pTAL6-BB
Plasmids pTAL5-BB and pTAL6-BB were created in Tom Ellis’ lab and function as alternative destination vectors to generate TAL Orthongal Repressors (TALORs). TALORs can be used to custom repress gene expression in yeast. They consist of the DNA-binding domain of a TALE and strong nuclear localization tags. TALORs repress transcription initiation when targeted to DNA sequences within core promoter regions. pTAL5-BB contains the GAL1 promoter, placing TALORs built into this vector under galactose-inducible expression. pTAL6-BB contains the TEF1 promoter, resulting in constitutive expression of TALORs built into this vector.
Blount et al., PLoS One . 2012.; 7(3): e33279.
pCAG-T7-TALEN(Sangamo)-Destination with homo- and heterodimeric FokI domains
pCAG-T7-TALEN(Sangamo)-Destination constructs were designed by Pawel Pelczar’s lab for the purpose of optimal mammalian expression of Voytas Golden Gate-assembled TALENs, both in microinjected embryos and transfected cells. TALEN expression is driven by the strong CAG promoter or can be achieved by in vitro mRNA synthesis from the T7 promoter. Truncations were introduced to the N- and C-terminus of the pTAL3 TALEN backbone, which were initially published by Sangamo BioSciences (N153AA, C63AA) and showed robust cleavage activity in several later studies. pCAG-T7-TALEN(Sangamo)-Destination vectors are available with homodimeric or enhanced heterodimeric (ELD, KKR mutations) FokI domains.
June 2012: Promiscuous Biotin Ligases, QB3 MacroLabs Expression Vectors, GCaMP5, and more
Promiscuous Biotin Ligases
In recent years there has been a significant focus on finding a protein’s “interactome”, identifying neighboring and potentially interacting partners of your protein of interest. Current means of identifying one’s network are limited and have several drawbacks, for example biochemical approaches encounter problems with insolubility of expressed proteins and in Y2H systems the interactions are tested outside of the natural environment.
In his recent paper, Kyle Roux developed a simply and quick technique for identifying interacting proteins. The system relies on promiscuous biotin protein ligase fused to a protein of interest; once the culture media is supplemented with biotin, the ligase will biotinylate proteins that are in close proximity. Biotinylated proteins can then be captured by affinity purification and identified using mass spec.
Specifically, Roux et al utilize a mutated form of the E. coli DNA-binding biotin protein ligase, BirA. BirA* (with R118G mutation) lacks the specificity of BirA and has been shown to promiscuously biotinylate proteins in a proximity dependent fashion. Roux et al test their system on lamin-A, a structural element of the nuclear envelope, thus identifying proteins that interact and/or are in close proximity to lamin-A.
Roux et al., J Cell Biol . 2012. Mar 19;196(6):801-10. Epub 2012 Mar 12.
Expression Vectors from the QB3 MacroLab
Finding the right empty vector for your purification technique of choice is often challenging. Finding a vector with the right tags and the right fluorescent fusion protein is even more challenging. What are the chances that someone has a backbone with just the right combination of tags and fusion proteins?
Actually, if you look at Addgene's collection of backbones from QB3 MacroLab, your chances are pretty good. This core facility for the California Institute for Quantitative Biosciences (QB3) designed these backbones to offer different combinations of the following TEV-cleavable N-terminal tags: His6, MBP, FLAG, NusA, Mocr, proteingG, StrepII, Sumo, gCrystallin, N10, and Biotin. The N-terminal tags can be found in various combinations with the C-terminal fusions that include mCherry, mOrange, mCitrine, msfGFP, and mCerulean. Most of the vectors were designed for bacterial expression, but there are also baculovirus and mammalian expression backbones.
The vectors were designed for ligation independent cloning (LIC), for which there are convenient protocols on the MacroLab website. You can also download a full expression vector list in Excel format from their website.
GCaMP5: Out With the Old and In With the New
The very popular calcium sensor GCaMP3 has been updated. GCaMP5G, a.k.a. GCaMP3-T302L R303P D380Y, has improved deltaF/F0 and lower F0 as compared to GCaMP3. Addgene now offers pCMV-GCaMP5G from Loren Looger's lab or the membrane targeted version, Lck-GCaMP5G, from Baljit Khakh's lab.
Zinc Finger Arrays Targeting Endogenous Zebrafish Genes
Gene targeting is a powerful technique that induces specific DNA stand breaks followed by stand rejoining to modify genes of interest. Engineered zinc-finger nucleases (ZFNs) are widely used for targeted genome modification in Drosophila, C. elegans, D. rerio, plants and humans. ZFNs function as dimmers and consist of a DNA-binding zinc finger domain that is covalently linked to a non-specific DNA cleavage domain of a restriction endonuclease, such as FokI. When the two zinc finger DNA binding domains bind to the target sites, the cleavage domains are able to dimerize and cleave at the desired target site. In order to provide researchers with easy-to-use zinc finger arrays, the laboratories of Keith Joung, Randall Peterson and Joanna Yeh at Massachusetts General Hospital have produced 49 pairs of zinc finger arrays targeting a variety of zebrafish genes.
Methods for engineering zinc finger domains:
New Retroviral Vectors for Cellular Reprogramming
Alzheimer’s disease is a neurodegenerative disorder for which there is no known cure. The disease most often arises sporadically, but familial forms can also be genetically inherited. The majority of research to date has focused on the less prevalent familial form, as this is more easily modeled in the lab. A new technology used to reprogram primary cells into induced pluripotent stem cells (iPSCs) has been successfully utilized in the study of some neurological diseases; however, it is not known whether this approach would be effective for studying both the sporadic and familial forms Alzheimer’s disease. Steven Dowdy and Larry Goldstein at UCSD have developed retroviral plasmids to generate iPSCs from the primary cells of Alzheimer’s patients. The resulting data provides evidence that iPSCs are effective for studying pathogenesis at the early stages of sporadic and familial Alzheimer’s disease, and can be successfully used in patient-specific cells. This technology not only provides a means to study the mechanisms of disease pathogenesis, but may also prove to be a useful tool for Alzheimer’s disease diagnosis.
Israel et al., Nature . 2012. Jan 25;482(7384):216-20.
March 2012: O-phosphoserine, Genome Wide Knockdown, Brain MiniPromoters, and Cyclin Reporters
Engineering Phosphoproteins in E. Coli
Protein kinases post-translationally phosphorylate specific residues as a means of regulating cell-signaling cascades. O-phosphoserine (Sep) being, by far, the most common phosphorylation modification. Phosphorylation events often require specific stimuli or conditions that are difficult to experimentally replicate, limiting investigations into the functional roles of these phosphoamino acids. Jesse Rinehart’s group at Yale has devised a system to incorporate Sep directly into the genetic code of E. coli using a re-engineered tRNA (tRNAsep). This process requires not only the orthogonal tRNAsep:Sep-tRNA synthetase pair, but also a modified EF-Tu, which allows tRNAsep to be incorporated during protein synthesis. This study elegantly deduces the minimal requirements for genetic code expansion as well as provides a unique tool for protein engineering and research.
Park et al., Science . 2011. Aug 26;333(6046):1151-4.
See details on the Rinehart & Söll Phosphoprotein Synthesis Kit here.
Genome-wide shRNA Knockdown Screens Made Easier with DECIPHER
In 2011, Addgene began collaborating with Dr. Alex Chenchik of Cellecta, Inc. and Dr. Gus Frangou of the Fred Hutchinson Cancer Research Center to distribute Cellecta's DECIPHER pooled lentiviral shRNA libraries. The DECIPHER Project was designed to make high-throughput RNAi genetic screening accessible to academia.
Each pooled shRNA library (or module) targets approximately 5,000 genes/transcripts, with 5 to 6 bar-coded shRNAs per target gene to maximize screening efficiency. The library modules were constructed as a series with non-overlapping sets of target genes. There are currently three modules for human and two modules for mouse available which target 15,000 and 10,000 genes, respectively. The first two library modules for human and mouse target mostly well characterized pathway-associated and disease-associated genes. The third human module targets cell surface and DNA-binding proteins, and other conserved genes. Just like Cellecta's custom libraries, the DECIPHER Project libraries are constructed with shRNA expression cassette oligonucleotides containing unique sequence tags ("bar-codes") synthesized on Agilent's array-based platform. Cellecta also offers free bar-code analyzer/deconvoluter software to assist scientists in processing Illumina HT Sequencing data generated from their RNAi screens. For more information about this powerful new screening tool, visit http://www.addgene.org/decipher/.
MiniPromoters for Brain-Specific Gene Expression
Region and cell-specific expression of a gene of interest is a critical experimental consideration, especially when studying the human brain which by some estimates contains 100 billion cells and possibly 100 trillion cellular connections. In order to address the lack of tools available for brain-region specific gene expression, a team of investigators joined forces to create the Pleiades Promoter Project (www.pleiades.org). Using a novel design pipeline, they produced MiniPromoters derived from endogenous human gene promoter sequences and combined them with reporter genes such as EGFP, EGFP/Cre or LacZ in constructs designed for knock-in insertion of transgenes to yield region specific expression as verified by in vivo experiments in mouse brain. Some MiniPromoters produced neuron or glia specific expression, as expected based on the gene the promoter was taken from, while others produced interesting, but unrelated expression patterns. The MiniPromoters available in the Pleiades Plasmids offer researches a more refined tool to obtain localized gene expression in the brain than previously available.
Portales-Casamar et al., Proc Natl Acad Sci USA . 2010. Sep 21; 107(38):16589-94.
Cyclin D1 Reporters
Regulated progression through the cell cycle requires sequential expression of a family of proteins called cyclins. Frank McCormick's group at UCSF recently deposited a series of human cyclin D1 promoter constructs from his Nature 1999 publication with Addgene. Cyclin D1 is over-expressed in many colon carcinomas and has been identified as a target of β-catenin mediated transcription via the core TCF/LEF-binding consensus sites in the promoter region. The McCormick group demonstrates this regulation using Cyclin D1 luciferase reporter constructs containing mutated TCF-binding sites.
Tetsu O, McCormick F. Nature . 1999l. Apr 1;398(6726):422-6.
December 2011: GECOs, iMNs, Zinc Fingers, and Multicistronic Drosophila Vectors
New Flavors for Genetically Encoded Ca2+ Indicators (GECOs)
Fluorescent indicators have long been used to measure intracellular Ca2+ levels, an important process in many signaling activities. In a recent Science paper, Robert Campbell’s laboratory at the University of Alberta describe an improved screening method for detecting changes in Ca2+ dependent fluorescence that they used to develop improved genetically encoded Ca2+ indicators, dubbed GECOs. The green (G-GECOs) demonstrate a two-fold improvement in Ca2+-dependent change in fluorescence over earlier fluorescent protein Ca2+ indicators. Blue-shifted (B-GECO) and red-shifted (R-GECO) Ca2+ indicators fill a void in the spectrum of available colors for genetically encoded Ca2+ indicators. By recombining these newly created constructs, additional ratiometric GECOs, GEM-GECO and GEX-GECO, were developed. The GECO plasmids offer researchers additional choice and flexibility when selecting fluorescent Ca2+ indicators.
Zhao et al., Science . 2011. Sep 30;333(6051):1888-91.
Transcription Factor Mediated Reprogramming of Fibroblasts into Motor Neurons (iMNs)
Studying cellular subtypes of the central nervous system (CNS) has proven to be challenging. For one, isolating subtypes of the human CNS can be strategically limiting in terms of supply and attainability. Moreover, differentiating embryonic stem cells (ESCs) into neuronal sub-populations has been fairly unsuccessful to date. Kevin Eggan's group at Harvard recently showed that they could use transcription factor-mediated expression to induce mouse and human fibroblasts into spinal motor neurons. This particular subset of neurons controls the contraction of muscle fibers involved in movement. Characterization of these reprogrammed cells showed they exhibit a distinct motor neuron identity . The complete set of reprogramming transcription factors are available at Addgene cloned into a retroviral backbone.
Son EY et al., Cell Stem Cell . 2011. Sep 2;9(3):205-18.
Zinc Finger Nuclease Assembly in Zebrafish
Zinc Finger Nucleases (ZFNs), chimeric fusions between a zinc-finger protein and the nuclease domain of FokI, are used in a wide range of model organisms for gene inactivation. Gene disruption occurs by imprecise repair of a ZFN-induced double-strand break within the coding sequence of a target gene. Using the modular assembly-based approach to ZFNs construction, the Lawson and Wolfe labs from UMass Worcester created a library of over 70 different zinc-finger cassettes. In parallel, the labs assembled a new publicly available database of zebrafish genes that can be targeted by ZFNs. The Lawson and Wolfe labs also demonstrate that ZFNs are a viable tool for creating an heritable gene inactivation in vertebrates by creating several germline mutations in zebrafish.
Zhu et al., Development . 2011. Oct;138(20):4555-64.
Vectors for Simplified Multicistronic Expression in Drosophila
Techniques to produce stable mammalian cell lines have been around for years; however, the ability to stably transform insect cells is especially limited. Because of these limitations, cell-culture based approaches are generally disregarded when attempting to validate chemical/RNAi library screens or produce recombinant protein. The Sutherland Lab has developed two Droshophila vectors, pAc5-STABLE1-Neo and pAc5-STABLE2-Neo, which facilitate the multicistronic expression of proteins without relying on co-transfection with a second vector (for antibiotic selection) or the need for dual-promoters. Both of these versatile vectors allow for either N- or C-terminal tagging with a fluorescent protein and pAc5-STABLE2-Neo gives the additional option of co-expressing a separate fluorescent tag, that can by used for FACS analysis.
Gonzalez et al., Scientific Reports . 2011. Aug; 1, Article: 75.
September 2011: TALENS, Photoswitchable Proteins, a Novel iPSC Factor and More
PSmOrange- a Novel Photoswitchable Protein
Photoconvertible proteins are widely used in tracking the migration, fate, and dynamics of cells, organelles, and proteins. Photoconvertible proteins can be divided into those that are photoactivatable- that can be turned on or off; or those that are photoswitchable- that can switch from one color to another. In a recent Nature Methods publication by Vladislav Verkhusha’s group at Albert Einstein, a novel photoswitchable protein, PSmOrange was characterized. PSmOrange is initially Orange (excitation, 548 nm; emission, 565 nm) but becomes far-red (excitation, 636 nm; emission, 662 nm) after irradiation with blue-green light. The photoconverted or the far-red version of PSmOrange is brighter than conventional far-red fluorescent proteins and can be imaged in deep tissues. PSmOrange is in an easy to clone vector and has been used to generate numerous fusion proteins.
Subach et al., Nat Methods . 2011. Jul 31;8(9):771-7.
Golden Gate TALEN and TAL Effector Kit
TAL effector nucleases (TALENs) are fusion proteins containing transcription activator-like (TAL) DNA binding domains fused to the FokI nuclease. TALENs have shown to be robust tools for DNA targeting in a variety of species. Binding specificity is determined by customizable arrays of polymorphic amino acid repeats in the TAL effectors. The Voytas laboratory at the University of Minnesota recently published a system for custom TALEN design and assembly. The Golden Gate TALEN and TAL Effector Kit and accompanying documentation allow one to efficiently assemble TALEN constructs with custom repeat arrays, containing anywhere between 12 and 31 of these repeats. The reagents include a plasmid construct for making custom TAL effectors and one for TAL effector fusions to additional proteins of interest.
Cermac et al., Nucleic Acids Res . 2011. Jul;39(12):e82.
GLI Transcription Factor Improves iPSC Generation
Generating induced pluripotent stem cells (iPSCs) from somatic cells involves finding the “right” combination of factors. The transcription factor Myc has been used successfully in producing iPSCs (in combination with other proteins)- although these mice have been shown to develop cancer. The Yamanaka lab at Kyoto University recently performed a screen to find a transcription factor that could improve efficiency of iPSC generation and potentially improve the viability of iPSC mice. Glis1 (Glis family zinc finger 1) was shown to promote iPSC generation in combination with Oct3/4, Sox2, and Klf4 in both mouse and human fibroblasts- replacing the requirement of Myc in generating iPSCs. Since the beginning of the summer, Glis1 has been requested by over 60 laboratories and continues to be a gene of interest for many researchers. Additional reprogramming factors are also available at Addgene.
Maekawa et al., Nature . 2011. Jun 8;474(7350):225-9.
New Recombinases for Animal Genomes
Site-specific recombination is a robust tool used for controlling gene expression. Site-specific recombinases can be used in animal models to turn expression of a gene on or off, create conditional mutants, or introduce novel genetic material. Gerald Rubin’s lab at the Janelia Farm Research Campus has characterized a set of novel recombinases from yeast- KD, B2, B3, and R. All recombinases were shown to be active in Drosophila and do not cross-react. In addition, KD and B3 were shown to be non-toxic and active in mice. Increasing the number of functional, specific recombinases allows for greater flexibility when performing experiments that involve multiple recombination events.
Nern et al., Proc Natl Acad Sci U S A . 2011. Aug 23;108(34):14198-203.
RNAi vectors for improved detection and inducible expression
To improve the efficiency of shRNA delivery and expression, specifically in targeting genes involved in proliferation and survival, Scott Lowe’s laboratory at Cold Spring Harbor has designed a set of plasmids that can be used for tracking and induction of retroviral-mediated shRNA expression. The retroviral plasmids express two transcripts. The first transcript, driven by a tetracycline response element (TRE)-encodes for a dsRed tagged miR-embedded shRNA, while the second transcript encodes for Venus and a mammalian antibiotic resistance cassette or Tet-inducible transactivator (rtTA3). The dual fluorescent system adds sensitivity to detection and the TRE allows for targeted knockdown, helping mimic in vivo therapeutic strategies. The TRMPV vectors, as they’re called, come in a variety of flavors- with different antibiotic resistance cassettes- and make up part of Addgene’s collection of inducible shRNA plasmids.
Zuber et al., Nat Biotechnol . 2011. Jan;29(1):79-83.
STAG2 and Aneuploidy
Todd Waldman’s lab at Georgetown recently showed that mutations in STAG2 can result in aneuploidy. Aneuploidy-- an abnormal number of chromosomes-- is often seen in cancer cells. STAG2 is a member of the cohesin complex, which regulates the separation of sister chromatids during cell division. In the Waldman lab’s recent Science paper (Solomon et al., 2011)- they identify specific STAG2 mutations found in cancer cells- that result in chromosomal disruption. STAG2 plasmids, wild-type and these aneuploidy-associated mutant forms, are now available through Addgene; along with a set of shRNAs targeting STAG2.
Solomon et al., Science . 2011. Aug 19;333(6045):1039-43.
Hot Article Contributors:
Addgene's Hot Article page is a compilation of summaries describing new plasmids available at Addgene and their related journal articles. These summaries were written by: Susanna Bachle, Andy Baltus, Chari Cortez, Michelle Cronin, Melina Fan, Matthew Ferenc, Tyler Ford, Mary Gearing, Benoit Giquel, Melanie Herscovitch, Max Juchheim, Caroline LaManna, Joel McDade, Jason Niehaus, Marcy Patrick, Eric Perkins, Brook Pyhtila, Maria Soriano, Lianna Swanson, Julian Taylor-Parker, Nicole Waxmonsky, and Jessica Welch.