In order to target a single protein to multiple subcellular organelles, plants typically duplicate the relevant genes, and express each gene separately using complex regulatory strategies including differential promoters and/or signal sequences. Metabolic engineers and synthetic biologists interested in targeting enzymes to a particular organelle are faced with a challenge: For a protein that is to be localized to more than one organelle, the engineer must clone the same gene multiple times. This work presents a solution to this strategy: harnessing alternative splicing of mRNA. This technology takes advantage of established chloroplast and peroxisome targeting sequences and combines them into a single mRNA that is alternatively spliced. Some splice variants are sent to the chloroplast, some to the peroxisome, and some to the cytosol. Here the system is designed for multiple-organelle targeting with alternative splicing. In this work, GFP was expected to be expressed in the chloroplast, cytosol, and peroxisome by a series of rationally designed 5’ mRNA tags. These tags have the potential to reduce the amount of cloning required when heterologous genes need to be expressed in multiple subcellular organelles. The constructs were designed in previous work11, and were cloned using Gibson assembly, a ligation independent cloning method that does not require restriction enzymes. The resultant plasmids were introduced into Nicotiana benthamiana epidermal leaf cells with a modified Gene Gun protocol. Finally, transformed leaves were observed with confocal microscopy.
21 Related JoVE Articles!
Protein-protein Interactions Visualized by Bimolecular Fluorescence Complementation in Tobacco Protoplasts and Leaves
Institutions: Ludwig-Maximilians-Universität, München.
Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular fluorescence complementation (BiFC) is an in vivo
method to monitor such interactions in plant cells. In the presented protocol the investigated candidate proteins are fused to complementary halves of fluorescent proteins and the respective constructs are introduced into plant cells via agrobacterium-mediated transformation. Subsequently, the proteins are transiently expressed in tobacco leaves and the restored fluorescent signals can be detected with a confocal laser scanning microscope in the intact cells. This allows not only visualization of the interaction itself, but also the subcellular localization of the protein complexes can be determined. For this purpose, marker genes containing a fluorescent tag can be coexpressed along with the BiFC constructs, thus visualizing cellular structures such as the endoplasmic reticulum, mitochondria, the Golgi apparatus or the plasma membrane. The fluorescent signal can be monitored either directly in epidermal leaf cells or in single protoplasts, which can be easily isolated from the transformed tobacco leaves. BiFC is ideally suited to study protein-protein interactions in their natural surroundings within the living cell. However, it has to be considered that the expression has to be driven by strong promoters and that the interaction partners are modified due to fusion of the relatively large fluorescence tags, which might interfere with the interaction mechanism. Nevertheless, BiFC is an excellent complementary approach to other commonly applied methods investigating protein-protein interactions, such as coimmunoprecipitation, in vitro
pull-down assays or yeast-two-hybrid experiments.
Plant Biology, Issue 85, Tetratricopeptide repeat domain, chaperone, chloroplasts, endoplasmic reticulum, HSP90, Toc complex, Sec translocon, BiFC
Monitoring Cell-autonomous Circadian Clock Rhythms of Gene Expression Using Luciferase Bioluminescence Reporters
Institutions: The University of Memphis.
In mammals, many aspects of behavior and physiology such as sleep-wake cycles and liver metabolism are regulated by endogenous circadian clocks (reviewed1,2
). The circadian time-keeping system is a hierarchical multi-oscillator network, with the central clock located in the suprachiasmatic nucleus (SCN) synchronizing and coordinating extra-SCN and peripheral clocks elsewhere1,2
. Individual cells are the functional units for generation and maintenance of circadian rhythms3,4
, and these oscillators of different tissue types in the organism share a remarkably similar biochemical negative feedback mechanism. However, due to interactions at the neuronal network level in the SCN and through rhythmic, systemic cues at the organismal level, circadian rhythms at the organismal level are not necessarily cell-autonomous5-7
. Compared to traditional studies of locomotor activity in vivo
and SCN explants ex vivo
, cell-based in vitro
assays allow for discovery of cell-autonomous circadian defects5,8
. Strategically, cell-based models are more experimentally tractable for phenotypic characterization and rapid discovery of basic clock mechanisms5,8-13
Because circadian rhythms are dynamic, longitudinal measurements with high temporal resolution are needed to assess clock function. In recent years, real-time bioluminescence recording using firefly luciferase
as a reporter has become a common technique for studying circadian rhythms in mammals14,15
, as it allows for examination of the persistence and dynamics of molecular rhythms. To monitor cell-autonomous circadian rhythms of gene expression, luciferase reporters can be introduced into cells via transient transfection13,16,17
or stable transduction5,10,18,19
. Here we describe a stable transduction protocol using lentivirus-mediated gene delivery. The lentiviral vector system is superior to traditional methods such as transient transfection and germline transmission because of its efficiency and versatility: it permits efficient delivery and stable integration into the host genome of both dividing and non-dividing cells20
. Once a reporter cell line is established, the dynamics of clock function can be examined through bioluminescence recording. We first describe the generation of P(Per2
reporter lines, and then present data from this and other circadian reporters. In these assays, 3T3 mouse fibroblasts and U2OS human osteosarcoma cells are used as cellular models. We also discuss various ways of using these clock models in circadian studies. Methods described here can be applied to a great variety of cell types to study the cellular and molecular basis of circadian clocks, and may prove useful in tackling problems in other biological systems.
Genetics, Issue 67, Molecular Biology, Cellular Biology, Chemical Biology, Circadian clock, firefly luciferase, real-time bioluminescence technology, cell-autonomous model, lentiviral vector, RNA interference (RNAi), high-throughput screening (HTS)
Genome Editing with CompoZr Custom Zinc Finger Nucleases (ZFNs)
Institutions: Sigma Life Science.
Genome editing is a powerful technique that can be used to elucidate gene function and the genetic basis of disease. Traditional gene editing methods such as chemical-based mutagenesis or random integration of DNA sequences confer indiscriminate genetic changes in an overall inefficient manner and require incorporation of undesirable synthetic sequences or use of aberrant culture conditions, potentially confusing biological study. By contrast, transient ZFN expression in a cell can facilitate precise, heritable gene editing in a highly efficient manner without the need for administration of chemicals or integration of synthetic transgenes.
Zinc finger nucleases (ZFNs) are enzymes which bind and cut distinct sequences of double-stranded DNA (dsDNA). A functional CompoZr ZFN unit consists of two individual monomeric proteins that bind a DNA "half-site" of approximately 15-18 nucleotides (see Figure 1
). When two ZFN monomers "home" to their adjacent target sites the DNA-cleavage domains dimerize and create a double-strand break (DSB) in the DNA.1
Introduction of ZFN-mediated DSBs in the genome lays a foundation for highly efficient genome editing. Imperfect repair of DSBs in a cell via the non-homologous end-joining (NHEJ) DNA repair pathway can result in small insertions and deletions (indels). Creation of indels within the gene coding sequence of a cell can result in frameshift and subsequent functional knockout of a gene locus at high efficiency.2
While this protocol describes the use of ZFNs to create a gene knockout, integration of transgenes may also be conducted via homology-directed repair at the ZFN cut site.
The CompoZr Custom ZFN Service represents a systematic, comprehensive, and well-characterized approach to targeted gene editing for the scientific community with ZFN technology. Sigma scientists work closely with investigators to 1) perform due diligence analysis including analysis of relevant gene structure, biology, and model system pursuant to the project goals, 2) apply this knowledge to develop a sound targeting strategy, 3) then design, build, and functionally validate ZFNs for activity in a relevant cell line. The investigator receives positive control genomic DNA and primers, and ready-to-use ZFN reagents supplied in both plasmid DNA and in-vitro transcribed mRNA format. These reagents may then be delivered for transient expression in the investigator’s cell line or cell type of choice. Samples are then tested for gene editing at the locus of interest by standard molecular biology techniques including PCR amplification, enzymatic digest, and electrophoresis. After positive signal for gene editing is detected in the initial population, cells are single-cell cloned and genotyped for identification of mutant clones/alleles.
Genetics, Issue 64, Molecular Biology, Zinc Finger Nuclease, Genome Engineering, Genomic Editing, Gene Modification, Gene Knockout, Gene Integration, non-homologous end joining, homologous recombination, targeted genome editing
Site-specific Bacterial Chromosome Engineering: ΦC31 Integrase Mediated Cassette Exchange (IMCE)
Institutions: University of Waterloo.
The bacterial chromosome may be used to stably maintain foreign DNA in the mega-base range1
. Integration into the chromosome circumvents issues such as plasmid replication, plasmid stability, plasmid incompatibility, and plasmid copy number variance. This method uses the site-specific integrase from the Streptomyces
phage (Φ) C312,3
. The ΦC31 integrase catalyzes a direct recombination between two specific DNA sites: attB
(34 and 39 bp, respectively)4
. This recombination is stable and does not revert5
. A "landing pad" (LP) sequence consisting of a spectinomycin- resistance gene, aadA
), and the E. coli
ß-glucuronidase gene (uidA
) flanked by attP
sites has been integrated into the chromosomes of Sinorhizobium meliloti, Ochrobactrum anthropi,
and Agrobacterium tumefaciens
in an intergenic region, the ampC
locus, and the tetA
locus, respectively. S. meliloti
is used in this protocol. Mobilizable donor vectors containing attB
sites flanking a stuffer red fluorescent protein (rfp)
gene and an antibiotic resistance gene have also been constructed. In this example the gentamicin resistant plasmid pJH110 is used. The rfp
may be replaced with a desired construct using Sph
I and Pst
I. Alternatively a synthetic construct flanked by attB
sites may be sub-cloned into a mobilizable vector such as pK19mob7
. The expression of the ΦC31 integrase gene (cloned from pHS628
) is driven by the lac
promoter, on a mobilizable broad host range plasmid pRK78139
A tetraparental mating protocol is used to transfer the donor cassette into the LP strain thereby replacing the markers in the LP sequence with the donor cassette. These cells are trans-integrants. Trans-integrants are formed with a typical efficiency of 0.5%. Trans-integrants are typically found within the first 500-1,000 colonies screened by antibiotic sensitivity or blue-white screening using 5-bromo-4-chloro-3-indolyl-beta-D-glucuronic acid (X-gluc). This protocol contains the mating and selection procedures for creating and isolating trans-integrants.
Bioengineering, Issue 61, ΦC31 Integrase, Rhizobiales, Chromosome Engineering, bacterial genetics
Detection of Protein Interactions in Plant using a Gateway Compatible Bimolecular Fluorescence Complementation (BiFC) System
Institutions: University of Western Ontario, Agriculture and Agri-Food Canada.
We have developed a BiFC technique to test the interaction between two proteins in vivo
. This is accomplished by splitting a yellow fluorescent protein (YFP) into two non-overlapping fragments. Each fragment is cloned in-frame to a gene of interest. These constructs can then be co-transformed into Nicotiana benthamiana
mediated transformation, allowing the transit expression of fusion proteins. The reconstitution of YFP signal only occurs when the inquest proteins interact 1-7
. To test and validate the protein-protein interactions, BiFC can be used together with yeast two hybrid (Y2H) assay. This may detect indirect interactions which can be overlooked in the Y2H. Gateway technology is a universal platform that enables researchers to shuttle the gene of interest (GOI) into as many expression and functional analysis systems as possible8,9
. Both the orientation and reading frame can be maintained without using restriction enzymes or ligation to make expression-ready clones. As a result, one can eliminate all the re-sequencing steps to ensure consistent results throughout the experiments. We have created a series of Gateway compatible BiFC and Y2H vectors which provide researchers with easy-to-use tools to perform both BiFC and Y2H assays10
. Here, we demonstrate the ease of using our BiFC system to test protein-protein interactions in N. benthamiana
Plant Biology, Issue 55, protein interaction, Gateway, Bimolecular fluorescence complementation, Confocal microscope, Agrobacterium, Nicotiana benthamiana, Arabidopsis
Massively Parallel Reporter Assays in Cultured Mammalian Cells
Institutions: Broad Institute.
The genetic reporter assay is a well-established and powerful tool for dissecting the relationship between DNA sequences and their gene regulatory activities. The potential throughput of this assay has, however, been limited by the need to individually clone and assay the activity of each sequence on interest using protein fluorescence or enzymatic activity as a proxy for regulatory activity. Advances in high-throughput DNA synthesis and sequencing technologies have recently made it possible to overcome these limitations by multiplexing the construction and interrogation of large libraries of reporter constructs. This protocol describes implementation of a Massively Parallel Reporter Assay (MPRA) that allows direct comparison of hundreds of thousands of putative regulatory sequences in a single cell culture dish.
Genetics, Issue 90, gene regulation, transcriptional regulation, sequence-activity mapping, reporter assay, library cloning, transfection, tag sequencing, mammalian cells
Characterization of Complex Systems Using the Design of Experiments Approach: Transient Protein Expression in Tobacco as a Case Study
Institutions: RWTH Aachen University, Fraunhofer Gesellschaft.
Plants provide multiple benefits for the production of biopharmaceuticals including low costs, scalability, and safety. Transient expression offers the additional advantage of short development and production times, but expression levels can vary significantly between batches thus giving rise to regulatory concerns in the context of good manufacturing practice. We used a design of experiments (DoE) approach to determine the impact of major factors such as regulatory elements in the expression construct, plant growth and development parameters, and the incubation conditions during expression, on the variability of expression between batches. We tested plants expressing a model anti-HIV monoclonal antibody (2G12) and a fluorescent marker protein (DsRed). We discuss the rationale for selecting certain properties of the model and identify its potential limitations. The general approach can easily be transferred to other problems because the principles of the model are broadly applicable: knowledge-based parameter selection, complexity reduction by splitting the initial problem into smaller modules, software-guided setup of optimal experiment combinations and step-wise design augmentation. Therefore, the methodology is not only useful for characterizing protein expression in plants but also for the investigation of other complex systems lacking a mechanistic description. The predictive equations describing the interconnectivity between parameters can be used to establish mechanistic models for other complex systems.
Bioengineering, Issue 83, design of experiments (DoE), transient protein expression, plant-derived biopharmaceuticals, promoter, 5'UTR, fluorescent reporter protein, model building, incubation conditions, monoclonal antibody
Polymerase Chain Reaction: Basic Protocol Plus Troubleshooting and Optimization Strategies
Institutions: University of California, Los Angeles .
In the biological sciences there have been technological advances that catapult the discipline into golden ages of discovery. For example, the field of microbiology was transformed with the advent of Anton van Leeuwenhoek's microscope, which allowed scientists to visualize prokaryotes for the first time. The development of the polymerase chain reaction (PCR) is one of those innovations that changed the course of molecular science with its impact spanning countless subdisciplines in biology. The theoretical process was outlined by Keppe and coworkers in 1971; however, it was another 14 years until the complete PCR procedure was described and experimentally applied by Kary Mullis while at Cetus Corporation in 1985. Automation and refinement of this technique progressed with the introduction of a thermal stable DNA polymerase from the bacterium Thermus aquaticus
, consequently the name Taq
PCR is a powerful amplification technique that can generate an ample supply of a specific segment of DNA (i.e., an amplicon) from only a small amount of starting material (i.e., DNA template or target sequence). While straightforward and generally trouble-free, there are pitfalls that complicate the reaction producing spurious results. When PCR fails it can lead to many non-specific DNA products of varying sizes that appear as a ladder or smear of bands on agarose gels. Sometimes no products form at all. Another potential problem occurs when mutations are unintentionally introduced in the amplicons, resulting in a heterogeneous population of PCR products. PCR failures can become frustrating unless patience and careful troubleshooting are employed to sort out and solve the problem(s). This protocol outlines the basic principles of PCR, provides a methodology that will result in amplification of most target sequences, and presents strategies for optimizing a reaction. By following this PCR guide, students should be able to:
● Set up reactions and thermal cycling conditions for a conventional PCR experiment
● Understand the function of various reaction components and their overall effect on a PCR experiment
● Design and optimize a PCR experiment for any DNA template
● Troubleshoot failed PCR experiments
Basic Protocols, Issue 63, PCR, optimization, primer design, melting temperature, Tm, troubleshooting, additives, enhancers, template DNA quantification, thermal cycler, molecular biology, genetics
The Production of C. elegans Transgenes via Recombineering with the galK Selectable Marker
Institutions: Beth Israel Deaconess Medical Center, Harvard Medical School, University of Pittsburgh.
The creation of transgenic animals is widely utilized in C. elegans
research including the use of GFP fusion proteins to study the regulation and expression pattern of genes of interest or generation of tandem affinity purification (TAP) tagged versions of specific genes to facilitate their purification. Typically transgenes are generated by placing a promoter upstream of a GFP reporter gene or cDNA of interest, and this often produces a representative expression pattern. However, critical elements of gene regulation, such as control elements in the 3' untranslated region or alternative promoters, could be missed by this approach. Further only a single splice variant can be usually studied by this means. In contrast, the use of worm genomic DNA carried by fosmid DNA clones likely includes most if not all elements involved in gene regulation in vivo
which permits the greater ability to capture the genuine expression pattern and timing. To facilitate the generation of transgenes using fosmid DNA, we describe an E. coli
based recombineering procedure to insert GFP, a TAP-tag, or other sequences of interest into any location in the gene. The procedure uses the galK
gene as the selection marker for both the positive and negative selection steps in recombineering which results in obtaining the desired modification with high efficiency. Further, plasmids containing the galK
gene flanked by homology arms to commonly used GFP and TAP fusion genes are available which reduce the cost of oligos by 50% when generating a GFP or TAP fusion protein. These plasmids use the R6K replication origin which precludes the need for extensive PCR product purification. Finally, we also demonstrate a technique to integrate the unc-119
marker on to the fosmid backbone which allows the fosmid to be directly injected or bombarded into worms to generate transgenic animals. This video demonstrates the procedures involved in generating a transgene via recombineering using this method.
Genetics, Issue 47, C. elegans, transgenes, fosmid clone, galK, recombineering, homologous recombination, E. coli
High Throughput Quantitative Expression Screening and Purification Applied to Recombinant Disulfide-rich Venom Proteins Produced in E. coli
Institutions: Aix-Marseille Université, Commissariat à l'énergie atomique et aux énergies alternatives (CEA) Saclay, France.
Escherichia coli (E. coli)
is the most widely used expression system for the production of recombinant proteins for structural and functional studies. However, purifying proteins is sometimes challenging since many proteins are expressed in an insoluble form. When working with difficult or multiple targets it is therefore recommended to use high throughput (HTP) protein expression screening on a small scale (1-4 ml cultures) to quickly identify conditions for soluble expression. To cope with the various structural genomics programs of the lab, a quantitative (within a range of 0.1-100 mg/L culture of recombinant protein) and HTP protein expression screening protocol was implemented and validated on thousands of proteins. The protocols were automated with the use of a liquid handling robot but can also be performed manually without specialized equipment.
Disulfide-rich venom proteins are gaining increasing recognition for their potential as therapeutic drug leads. They can be highly potent and selective, but their complex disulfide bond networks make them challenging to produce. As a member of the FP7 European Venomics project (www.venomics.eu), our challenge is to develop successful production strategies with the aim of producing thousands of novel venom proteins for functional characterization. Aided by the redox properties of disulfide bond isomerase DsbC, we adapted our HTP production pipeline for the expression of oxidized, functional venom peptides in the E. coli
cytoplasm. The protocols are also applicable to the production of diverse disulfide-rich proteins. Here we demonstrate our pipeline applied to the production of animal venom proteins. With the protocols described herein it is likely that soluble disulfide-rich proteins will be obtained in as little as a week. Even from a small scale, there is the potential to use the purified proteins for validating the oxidation state by mass spectrometry, for characterization in pilot studies, or for sensitive micro-assays.
Bioengineering, Issue 89, E. coli, expression, recombinant, high throughput (HTP), purification, auto-induction, immobilized metal affinity chromatography (IMAC), tobacco etch virus protease (TEV) cleavage, disulfide bond isomerase C (DsbC) fusion, disulfide bonds, animal venom proteins/peptides
A Restriction Enzyme Based Cloning Method to Assess the In vitro Replication Capacity of HIV-1 Subtype C Gag-MJ4 Chimeric Viruses
Institutions: Emory University, Emory University.
The protective effect of many HLA class I alleles on HIV-1 pathogenesis and disease progression is, in part, attributed to their ability to target conserved portions of the HIV-1 genome that escape with difficulty. Sequence changes attributed to cellular immune pressure arise across the genome during infection, and if found within conserved regions of the genome such as Gag, can affect the ability of the virus to replicate in vitro
. Transmission of HLA-linked polymorphisms in Gag to HLA-mismatched recipients has been associated with reduced set point viral loads. We hypothesized this may be due to a reduced replication capacity of the virus. Here we present a novel method for assessing the in vitro
replication of HIV-1 as influenced by the gag
gene isolated from acute time points from subtype C infected Zambians. This method uses restriction enzyme based cloning to insert the gag
gene into a common subtype C HIV-1 proviral backbone, MJ4. This makes it more appropriate to the study of subtype C sequences than previous recombination based methods that have assessed the in vitro
replication of chronically derived gag-pro
sequences. Nevertheless, the protocol could be readily modified for studies of viruses from other subtypes. Moreover, this protocol details a robust and reproducible method for assessing the replication capacity of the Gag-MJ4 chimeric viruses on a CEM-based T cell line. This method was utilized for the study of Gag-MJ4 chimeric viruses derived from 149 subtype C acutely infected Zambians, and has allowed for the identification of residues in Gag that affect replication. More importantly, the implementation of this technique has facilitated a deeper understanding of how viral replication defines parameters of early HIV-1 pathogenesis such as set point viral load and longitudinal CD4+ T cell decline.
Infectious Diseases, Issue 90, HIV-1, Gag, viral replication, replication capacity, viral fitness, MJ4, CEM, GXR25
Mouse Genome Engineering Using Designer Nucleases
Institutions: University of Zurich, University of Minnesota.
Transgenic mice carrying site-specific genome modifications (knockout, knock-in) are of vital importance for dissecting complex biological systems as well as for modeling human diseases and testing therapeutic strategies. Recent advances in the use of designer nucleases such as zinc finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and the clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system for site-specific genome engineering open the possibility to perform rapid targeted genome modification in virtually any laboratory species without the need to rely on embryonic stem (ES) cell technology. A genome editing experiment typically starts with identification of designer nuclease target sites within a gene of interest followed by construction of custom DNA-binding domains to direct nuclease activity to the investigator-defined genomic locus. Designer nuclease plasmids are in vitro
transcribed to generate mRNA for microinjection of fertilized mouse oocytes. Here, we provide a protocol for achieving targeted genome modification by direct injection of TALEN mRNA into fertilized mouse oocytes.
Genetics, Issue 86, Oocyte microinjection, Designer nucleases, ZFN, TALEN, Genome Engineering
Recombineering Homologous Recombination Constructs in Drosophila
Institutions: University of Texas Southwestern Medical Center at Dallas, University of Texas Southwestern Medical Center at Dallas, University of Texas Southwestern Medical Center at Dallas.
The continued development of techniques for fast, large-scale manipulation of endogenous gene loci will broaden the use of Drosophila melanogaster
as a genetic model organism for human-disease related research. Recent years have seen technical advancements like homologous recombination and recombineering. However, generating unequivocal null mutations or tagging endogenous proteins remains a substantial effort for most genes. Here, we describe and demonstrate techniques for using recombineering-based cloning methods to generate vectors that can be used to target and manipulate endogenous loci in vivo.
Specifically, we have established a combination of three technologies: (1) BAC transgenesis/recombineering, (2) ends-out homologous recombination and (3) Gateway technology to provide a robust, efficient and flexible method for manipulating endogenous genomic loci. In this protocol, we provide step-by-step details about how to (1) design individual vectors, (2) how to clone large fragments of genomic DNA into the homologous recombination vector using gap repair, and (3) how to replace or tag genes of interest within these vectors using a second round of recombineering. Finally, we will also provide a protocol for how to mobilize these cassettes in vivo
to generate a knockout, or a tagged gene via knock-in. These methods can easily be adopted for multiple targets in parallel and provide a means for manipulating the Drosophila
genome in a timely and efficient manner.
Genetics, Issue 77, Bioengineering, Molecular Biology, Biomedical Engineering, Physiology, Drosophila melanogaster, genetics (animal and plant), Recombineering, Drosophila, Homologous Recombination, Knock-out, recombination, genetic engineering, gene targeting, gene, genes, DNA, PCR, Primers, sequencing, animal model
Investigating Protein-protein Interactions in Live Cells Using Bioluminescence Resonance Energy Transfer
Institutions: Max Planck Institute for Psycholinguistics, Donders Institute for Brain, Cognition and Behaviour.
Assays based on Bioluminescence Resonance Energy Transfer (BRET) provide a sensitive and reliable means to monitor protein-protein interactions in live cells. BRET is the non-radiative transfer of energy from a 'donor' luciferase enzyme to an 'acceptor' fluorescent protein. In the most common configuration of this assay, the donor is Renilla reniformis
luciferase and the acceptor is Yellow Fluorescent Protein (YFP). Because the efficiency of energy transfer is strongly distance-dependent, observation of the BRET phenomenon requires that the donor and acceptor be in close proximity. To test for an interaction between two proteins of interest in cultured mammalian cells, one protein is expressed as a fusion with luciferase and the second as a fusion with YFP. An interaction between the two proteins of interest may bring the donor and acceptor sufficiently close for energy transfer to occur. Compared to other techniques for investigating protein-protein interactions, the BRET assay is sensitive, requires little hands-on time and few reagents, and is able to detect interactions which are weak, transient, or dependent on the biochemical environment found within a live cell. It is therefore an ideal approach for confirming putative interactions suggested by yeast two-hybrid or mass spectrometry proteomics studies, and in addition it is well-suited for mapping interacting regions, assessing the effect of post-translational modifications on protein-protein interactions, and evaluating the impact of mutations identified in patient DNA.
Cellular Biology, Issue 87, Protein-protein interactions, Bioluminescence Resonance Energy Transfer, Live cell, Transfection, Luciferase, Yellow Fluorescent Protein, Mutations
Genetic Manipulation in Δku80 Strains for Functional Genomic Analysis of Toxoplasma gondii
Institutions: The Geisel School of Medicine at Dartmouth.
Targeted genetic manipulation using homologous recombination is the method of choice for functional genomic analysis to obtain a detailed view of gene function and phenotype(s). The development of mutant strains with targeted gene deletions, targeted mutations, complemented gene function, and/or tagged genes provides powerful strategies to address gene function, particularly if these genetic manipulations can be efficiently targeted to the gene locus of interest using integration mediated by double cross over homologous recombination.
Due to very high rates of nonhomologous recombination, functional genomic analysis of Toxoplasma gondii
has been previously limited by the absence of efficient methods for targeting gene deletions and gene replacements to specific genetic loci. Recently, we abolished the major pathway of nonhomologous recombination in type I and type II strains of T. gondii
by deleting the gene encoding the KU80 protein1,2
. The Δku80
strains behave normally during tachyzoite (acute) and bradyzoite (chronic) stages in vitro
and in vivo
and exhibit essentially a 100% frequency of homologous recombination. The Δku80
strains make functional genomic studies feasible on the single gene as well as on the genome scale1-4
Here, we report methods for using type I and type II Δku80Δhxgprt
strains to advance gene targeting approaches in T. gondii
. We outline efficient methods for generating gene deletions, gene replacements, and tagged genes by targeted insertion or deletion of the hypoxanthine-xanthine-guanine phosphoribosyltransferase (HXGPRT
) selectable marker. The described gene targeting protocol can be used in a variety of ways in Δku80
strains to advance functional analysis of the parasite genome and to develop single strains that carry multiple targeted genetic manipulations. The application of this genetic method and subsequent phenotypic assays will reveal fundamental and unique aspects of the biology of T. gondii
and related significant human pathogens that cause malaria (Plasmodium
sp.) and cryptosporidiosis (Cryptosporidium
Infectious Diseases, Issue 77, Genetics, Microbiology, Infection, Medicine, Immunology, Molecular Biology, Cellular Biology, Biomedical Engineering, Bioengineering, Genomics, Parasitology, Pathology, Apicomplexa, Coccidia, Toxoplasma, Genetic Techniques, Gene Targeting, Eukaryota, Toxoplasma gondii, genetic manipulation, gene targeting, gene deletion, gene replacement, gene tagging, homologous recombination, DNA, sequencing
Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos
Institutions: University of Wisconsin-Madison, University of Wisconsin-Madison, University of Wisconsin-Madison, University of Wisconsin-Madison.
The zebrafish is an ideal model for imaging cell behaviors during development in vivo
. Zebrafish embryos are externally fertilized and thus easily accessible at all stages of development. Moreover, their optical clarity allows high resolution imaging of cell and molecular dynamics in the natural environment of the intact embryo. We are using a live imaging approach to analyze cell behaviors during neural crest cell migration and the outgrowth and guidance of neuronal axons.
Live imaging is particularly useful for understanding mechanisms that regulate cell motility processes. To visualize details of cell motility, such as protrusive activity and molecular dynamics, it is advantageous to label individual cells. In zebrafish, plasmid DNA injection yields a transient mosaic expression pattern and offers distinct benefits over other cell labeling methods. For example, transgenic lines often label entire cell populations and thus may obscure visualization of the fine protrusions (or changes in molecular distribution) in a single cell. In addition, injection of DNA at the one-cell stage is less invasive and more precise than dye injections at later stages.
Here we describe a method for labeling individual developing neurons or neural crest cells and imaging their behavior in vivo
. We inject plasmid DNA into 1-cell stage embryos, which results in mosaic transgene expression. The vectors contain cell-specific promoters that drive expression of a gene of interest in a subset of sensory neurons or neural crest cells. We provide examples of cells labeled with membrane targeted GFP or with a biosensor probe that allows visualization of F-actin in living cells1
Erica Andersen, Namrata Asuri, and Matthew Clay contributed equally to this work.
Developmental Biology, Issue 36, zebrafish, axon guidance, neural crest, cell behavior, actin, microinjection, embryos
Generation of Stable Transgenic C. elegans Using Microinjection
Institutions: University of Alabama.
Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1
. The plasmid DNA rearranges to form extrachromosomal concatamers that are stably inherited, though not with the same efficiency as actual chromosomes 2
. A gene of interest is co-injected with an obvious phenotypic marker, such as rol-6 or GFP, to allow selection of transgenic animals under a dissecting microscope. The exogenous gene may be expressed from its native promoter for cellular localization studies. Alternatively, the transgene can be driven by a different tissue-specific promoter to assess the role of the gene product in that particular cell or tissue. This technique efficiently drives gene expression in all tissues of C. elegans except for the germline or early embryo 3
. Creation of transgenic animals is widely utilized for a range of experimental paradigms. This video demonstrates the microinjection procedure to generate transgenic worms. Furthermore, selection and maintenance of stable transgenic C. elegans lines is described.
Developmental Biology, Issue 18, C. elegans, microinjection, transgenic, stable lines
Electroeluting DNA Fragments
Institutions: Research and Advanced Studies Center of National Polytechnic Institute.
Purified DNA fragments are used for different purposes in Molecular Biology and they can be prepared by several procedures. Most of them require a previous electrophoresis of the DNA fragments in order to separate the band of interest. Then, this band is excised out from an agarose or acrylamide gel and purified by using either: binding and elution from glass or silica particles, DEAE-cellulose membranes, "crush and soak method", electroelution or very often expensive commercial purification kits. Thus, selecting a method will depend mostly of what is available in the laboratory. The electroelution procedure allows one to purify very clean DNA to be used in a large number of applications (sequencing, radiolabeling, enzymatic restriction, enzymatic modification, cloning etc). This procedure consists in placing DNA band-containing agarose or acrylamide slices into sample wells of the electroeluter, then applying current will make the DNA fragment to leave the agarose and thus be trapped in a cushion salt to be recovered later by ethanol precipitation.
Basic Protocols, Issue 43, electroelution procedure, nucleic acids, agarose, acrylamide
Transformation of Plasmid DNA into E. coli Using the Heat Shock Method
Institutions: University of California, Irvine (UCI).
Transformation of plasmid DNA into E. coli using the heat shock method is a basic technique of molecular biology. It consists of inserting a foreign plasmid or ligation product into bacteria. This video protocol describes the traditional method of transformation using commercially available chemically competent bacteria from Genlantis. After a short incubation in ice, a mixture of chemically competent bacteria and DNA is placed at 42°C for 45 seconds (heat shock) and then placed back in ice. SOC media is added and the transformed cells are incubated at 37°C for 30 min with agitation. To be assured of isolating colonies irrespective of transformation efficiency, two quantities of transformed bacteria are plated. This traditional protocol can be used successfully to transform most commercially available competent bacteria. The turbocells from Genlantis can also be used in a novel 3-minute transformation protocol, described in the instruction manual.
Issue 6, Basic Protocols, DNA, transformation, plasmid, cloning
Purifying Plasmid DNA from Bacterial Colonies Using the Qiagen Miniprep Kit
Institutions: University of California, Irvine (UCI).
Plasmid DNA purification from E. coli is a core technique for molecular cloning. Small scale purification (miniprep) from less than 5 ml of bacterial culture is a quick way for clone verification or DNA isolation, followed by further enzymatic reactions (polymerase chain reaction and restriction enzyme digestion). Here, we video-recorded the general procedures of miniprep through the QIAGEN's QIAprep 8 Miniprep Kit, aiming to introducing this highly efficient technique to the general beginners for molecular biology techniques. The whole procedure is based on alkaline lysis of E. coli cells followed by adsorption of DNA onto silica in the presence of high salt. It consists of three steps: 1) preparation and clearing of a bacterial lysate, 2) adsorption of DNA onto the QIAprep membrane, 3) washing and elution of plasmid DNA. All steps are performed without the use of phenol, chloroform, CsCl, ethidium bromide, and without alcohol precipitation. It usually takes less than 2 hours to finish the entire procedure.
Issue 6, Basic Protocols, plasmid, DNA, purification, Qiagen
Mosaic Zebrafish Transgenesis for Evaluating Enhancer Sequences
Institutions: University of Pennsylvania .
The completion of the human genome sequence, along with that of many other species, has highlighted the challenge of ascribing specific function to non coding sequences. One prominent function carried out by the non coding fraction of the genome is to regulate gene transcription; however, there are no effective methods to broadly predict cis-regulatory elements from primary DNA sequence. We have developed an efficient protocol to functionally evaluate potential cis-regulatory elements through zebrafish transgenesis. Our approach offers significant advantages over cell-culture based techniques for developmentally important genes, since it provides information on spatial and temporal gene regulation. Conversely, it is faster and less expensive than similar experiments in transgenic mice, and we routinely apply it to sequences isolated from the human genome. Here we demonstrate our approach to selecting elements for testing based on sequence conservation and our protocol for cloning sequences and microinjecting them into zebrafish embryos.
Cellular Biology, Issue 41, zebrafish, transgenesis, microinjection, GFP, enhancers, transposon