Principe de la méthode: La mutagenèse dirigée de plasmides entière expliqué dans cette vidéo est une méthode de mutagenèse qui vous permet de modifier un gène cible clonée par substitution, délétion ou d&#39;insertion de quelques bases directement dans un plasmide. Il agit en amplifiant le plasmide entier, dans une réaction thermocyclage PCR non. Lors de la réaction amorces mutagènes, portant la mutation désirée sous forme d&#39;inadéquation au plasmide d&#39;origine, sont intégrés dans le plasmide nouvellement synthétisé. Après l&#39;élimination du plasmide d&#39;origine de la réaction du plasmide muté est transformé en E. coli. Les étapes suivantes sont à des fins de dépistage seulement, car l&#39;efficacité de la mutation de cette méthode n&#39;est pas 100%. La procédure complète prend trois jours, mais la partie principale peut être fait en une journée. 1.0 La première journée: 1.1 réaction thermocyclage: D&#39;origine plasmidique: Cette mutagenèse dirigée protocole qui fonctionne le mieux avec des plasmides à 10ko. Agrandir Les plasmides sont un peu difficiles à muter avec cette méthode et peut prendre un peu de patience et d&#39;adaptation des conditions de thermocyclage et / ou de cellules compétentes. En outre, le plasmide qui vous travaillez avec doit être isolé à partir d&#39;un barrage + souche de bactéries. Pour la réaction thermocyclage vous aurez besoin de 10 à 60 END du plasmide vous voulez muter. Primaires: Avant que vous pouvez configurer votre réaction thermocyclage vous devez avoir vos amorces à portée de main. Vous avez besoin d&#39;environ 150 ng de chaque primer mutagène. Il est ok, si vous prenez 1,5 pi d&#39;une dilution 1:10 100 actions pM. Il ya quelques règles simples que vous devez tenir compte lorsque vous concevez votre amorces mutagènes: <ul><li> Les amorces devraient être complémentaires les unes aux autres </li><li> Les amorces doivent être entre 25 et 45 nucléotides de long </li><li> Les mutations dans la forme de l&#39;inadéquation de la plasmide d&#39;origine doivent être contenues dans les deux amorces </li><li> Les décalages doivent être centrées dans l&#39;amorce et flanqué d&#39;au moins 8 nucléotides de chaque côté </li><li> Les amorces doivent avoir un contenu en GC d&#39;au moins 40% </li><li> Les amorces devraient extrémité 5 premiers et 3 premiers avec un ou plusieurs Gs ou Cs </li><li> Les amorces ne doivent pas être phosphorylée, et ils n&#39;ont pas à être FPLC ou PAGE purifiée, simplement dessalée qu&#39;ils devraient être. </li><li> Pour le calcul de Tm vous n&#39;avez pas besoin d&#39;aucune formule, mais Tm sur le certificat d&#39;expédition doit être supérieure à 60 ° C. </li><li> À des fins de dépistage, il est pratique pour insérer ou supprimer un site de restriction avec votre mutation. En raison de la dégénérescence du code génétique il ya beaucoup de possibilités pour insérer un site de restriction avec votre mutation désirée. Le site du New England Biolabs fournit un outil pour trouver un tel site de restriction approprié. Entrez simplement votre séquence de l&#39;amorce codant pour la séquence d&#39;acides aminés que vous désirez, en utilisant le code d&#39;ambiguïté pour l&#39;ADN. L&#39;outil de recherche d&#39;enzymes vous dira quels sites de restriction peuvent être introduits en parallèle à votre mutation désirée. Si vous ne pouvez pas trouver tout site de restriction possible ou pratique de cette façon, vous pouvez insérer un nouveau site de restriction concernant les sans code génétique sur le site de mutation. Par la suite, vous pouvez utiliser ce plasmide comme matrice pour une série de mutations dans lequel ce site de restriction sera supprimée par l&#39;insertion de la nouvelle amorces mutagènes. Cette approche peut être très pratique si vous avez l&#39;intention de faire une série de mutations sur le même site du plasmide. </li></ul> ADN-polymérase: Pour cette méthode, vous avez besoin d&#39;une ADN-polymérase thermostable qui présente une activité exonucléase 3&#39;-5 &#39;et crée des extrémités franches. Nous utilisons toujours recombinante Pfu-polymérase du Fermentas. Si vous avez des problèmes avec les étapes d&#39;amplification, vous pouvez essayer une polymérase de qualité supérieure, mais pour la plupart des fins de la Pfu standard sera suffisant. Remplissez la réaction avec dNTP, polymérase tampon et d&#39;eau. 1,2 conditions thermocyclage Le thermocycleur doit être mis en place de la façon suivante. La phase initiale de dénaturation et récurrent est réglé sur 30 sec à 95 ° C. La température de recuit des amorces ne doivent pas être calculées avec des formules compliquées. En général, nous réglez la température de recuit à 55 ° C et le temps de recuit à une minute. Pour amorces de 25 à 30 nucléotides qui vous allez utiliser pour la plupart, cela fonctionne pour nous 95% du temps. Quoi qu&#39;il en soit, si cela ne devrait pas travailler, essayez simplement varier la température de recuit entre 50 - et 60 ° C. La température dépend de l&#39;élongation de la polymérase vous utilisez. Dans cette démonstration, nous utilisons la Pfu polymérase du Fermentas, qui appelle à une température d&#39;élongation de 72 ° C. Le temps d&#39;élongation varie en fonction de la taille du plasmide. Nous calculons toujours 1 min par ko, et ajouter une minute supplémentaire pour que lestemps. Par exemple, pour un plasmide 9kb nous choisirions 10min que le temps d&#39;élongation. 18 cycles sont suffisantes pour créer suffisamment muté plasmide pour l&#39;utilisation ultérieure. Maintenir le nombre de cycles de basse, vous fait également gagner du temps. <img src="/files/ftp_upload/1135/figure1.png" alt="Figure 1" width="600" height="287" /> 1,3 Gelcheck après réaction thermocyclage L&#39;amplification de succès devrait être vérifiée en effectuant une électrophorèse. Directement après le vélo est terminée, charge de 5 ul de la réaction sur un gel à 1% d&#39;agarose TAE. Si l&#39;amplification a été un succès, vous devriez voir une bande distincte. Toutefois, si l&#39;ADN nouvellement synthétisé n&#39;est pas clairement visible sur le gel, vous pouvez essayer de précipiter toute la réaction et l&#39;utiliser pour la transformation à l&#39;étape suivante. Pour nous, cela a rarement travaillé, mais il vaut la peine de donner un essai. La meilleure chose à faire si la réaction ne fonctionne pas est d&#39;ajuster la température de recuit. 1,4 digestion Dpnl: Avant la transformation du plasmide original qui a servi de modèle doit être retiré de la réaction pour éviter de solides antécédents. Cela se fait par digestion de restriction avec Dpnl. Cette endonucléase de restriction ne coupe que méthylé plasmide. Sa reconnaissance et site de restriction est la séquence GATC alors A doit être méthylées. Lorsque la digestion avec Dpnl seul le plasmide d&#39;origine non muté qui a été isolée à partir d&#39;un barrage + souche est ensuite coupée, le plasmide muté nouvellement synthétisées qui n&#39;est pas méthylé n&#39;est pas affectée par Dpnl. Il suffit d&#39;ajouter 1-2 ul Dpnl à la réaction et incuber c&#39;est au moins une heure à 37 °. Lorsque vous utilisez le Fast digérer Dpnl le temps d&#39;incubation peut être réduit à environ 15 minutes. La qualité de votre Dpnl et le temps d&#39;incubation de cette digestion de restriction détermine grandement la force de vos antécédents avec le plasmide non muté sera. 1.5 Transformation: Après digestion du plasmide Dpnl est prêt pour la transformation en l&#39;autorité compétente E. cellules coli. Il suffit d&#39;ajouter 5 ul de la réaction de digestion Dpnl dans les cellules compétentes et de réaliser la transformation tel que recommandé dans votre laboratoire. Dans notre cas, nous utilisons la procédure de choc thermique en incubant le mélange de bactéries plasmide sur la glace pendant 30 min puis choc thermique c&#39;est à 42 ° C pendant 90 sec. Après l&#39;ajout de 200 ul SOC-solution, les bactéries sont incubées, secouer vigoureusement, à 37 ° C pendant 1 h. Après 1 heure du bactéries sont étalées sur la sélection d&#39;agar. Jour deux 2,0 2.1 Dépistage d&#39;une partie des clones 1: Sélection des clones Parce que l&#39;efficacité mutation n&#39;est pas à 100% vous avez besoin de l&#39;écran de votre mutants. Nous faisons cela par digestion de restriction dans lequel nous vérifier la présence ou l&#39;absence de site de restriction auquel nous avons ajouté ou supprimé par l&#39;intégration d&#39;apprêt. A cet effet, nous avons l&#39;habitude ramasser huit colonies et les cultiver pendant la nuit pour la préparation de plasmide sur le lendemain. 3.0 La troisième journée 3.1 Dépistage d&#39;une partie des clones 2: digestion de restriction avec l&#39;enzyme marqueur Le troisième jour, vous effectuez une préparation Mini et la digestion de restriction ultérieurs avec votre marqueur enzymatique. Sur le gel vous pouvez alors voir lequel de vos clones porteurs de la mutation désirée et ceux qui pas. La méthode de dépistage à l&#39;aide la digestion de restriction fonctionne très bien. Néanmoins, vous devez confirmer votre mutation réussie par séquençage.

<ol>
	<li>Papworth, C., Bauer, J. C., Braman, J., Wright, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=QuikChange+site-directed+mutagenesis.">QuikChange site-directed mutagenesis.</a> Strategies. 9, 3-4 (1996).</li></ol>

The authors have nothing to disclose.

Mutagenèse dirigée est une méthode qui mutagenèse fournit un moyen rapide de muter un gène porté par un plasmide. La réaction peut être fait tout en une seule journée. Avec cette méthode, il suffit d&#39;une paire d&#39;amorces complémentaires portant les mutations souhaitées et une polymérase Pfu relecture tels que-polymérase. Le plasmide nouvellement synthétisés peuvent être séparés du plasmide parental en digérant la réaction avec l&#39;enzyme de restriction Dpnl. Cette enzyme digère l&#39;ADN m...

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		<th>Catalogue Number</th>
		<th>Comment</th>
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<td>Pfu polymerase (recombinant or native)</td>
<td>&nbsp;</td>
<td>fermentas</td>
<td>EP0571 or EP0501</td>
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<td>dNTP mix 10 mM </td>
<td>&nbsp;</td>
<td>fermentas</td>
<td>R0191</td>
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<td>DpnI or DpnI Fast digest</td>
<td>&nbsp;</td>
<td>fermentas</td>
<td>ER1701</td>
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<td>primers</td>
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homemade site directed mutagenesis of whole plasmids

Mutagenèse dirigée de plasmides ensemble est un moyen simple de créer des variations légèrement différentes d&#39;un plasmide d&#39;origine. Avec cette méthode, le gène cible clonés peuvent être modifiés par substitution, délétion ou d&#39;insertion de quelques bases directement dans un plasmide. Il fonctionne tout simplement en amplifiant le plasmide entier, dans une réaction thermocyclage PCR non. Lors de la réaction amorces mutagènes, porteurs de la mutation souhaitée, sont intégrés dans le plasmide nouvellement synthétisé. Dans ce tutoriel vidéo, nous démontrons une manière facile et rentable d&#39;introduire des substitutions de bases dans un plasmide. Le protocole fonctionne avec des réactifs standard et est indépendant de kits commerciaux, qui sont souvent très coûteux. L&#39;application de ce protocole peut réduire le coût total d&#39;une réaction à un huitième de ce qu&#39;il en coûte en utilisant certains des kits commerciaux. Dans cette vidéo, nous avons également des commentaires sur les étapes essentielles du processus et donner des instructions détaillées sur la façon de concevoir les amorces mutagènes.

Watch this Scientific Journal Video about Homemade Site Directed Mutagenesis of Whole Plasmids at JoVE.com

Homemade Site Directed Mutagenesis of Whole Plasmids

Mutagenèse dirigée de plasmides ensemble est un moyen simple de créer des variations légèrement différentes d&#39;un plasmide d&#39;origine. Ici nous démontrons une manière facile et rentable d&#39;introduire des substitutions de bases dans un plasmide standard en utilisant des réactifs.

Mutagenèse dirigée maison des plasmides entiers

Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. With this method the cloned ...

homemade-site-directed-mutagenesis-of-whole-plasmids

Research

JoVE Journal

Biology

33.0K vues.  Johannes Gutenberg-University Mainz, Germany. Mutagenèse dirigée de plasmides ensemble est un moyen simple de créer des variations légèrement différentes d'un plasmide d'origine. Ici nous démontrons une manière facile et rentable d'introduire des substitutions de bases dans un plasmide standard en utilisant des réactifs.

Vidéo: Homemade Site Directed Mutagenesis of Whole Plasmids

Principles of Site-Specific Recombinase (SSR) Technology

Site-specific recombinase (SSR) technology allows the manipulation of gene structure to explore gene function and has become an integral tool of molecular biology. Site-specific recombinases are proteins that bind to distinct DNA target sequences. The Cre/lox system was first described in bacteriophages during the 1980's. Cre recombinase is a Type I topoisomerase that catalyzes site-specific recombination of DNA between two loxP (locus of X-over P1) sites. The Cre/lox system does not require any cofactors. LoxP sequences contain distinct binding sites for Cre recombinases that surround a directional core sequence where recombination and rearrangement takes place. When cells contain loxP sites and express the Cre recombinase, a recombination event occurs. Double-stranded DNA is cut at both loxP sites by the Cre recombinase, rearranged, and ligated ("scissors and glue"). Products of the recombination event depend on the relative orientation of the asymmetric sequences.
SSR technology is frequently used as a tool to explore gene function. Here the gene of interest is flanked with Cre target sites loxP ("floxed"). Animals are then crossed with animals expressing the Cre recombinase under the control of a tissue-specific promoter. In tissues that express the Cre recombinase it binds to target sequences and excises the floxed gene. Controlled gene deletion allows the investigation of gene function in specific tissues and at distinct time points. Analysis of gene function employing SSR technology --- conditional mutagenesis -- has significant advantages over traditional knock-outs where gene deletion is frequently lethal.

Principles of Site-Specific Recombinase SSR Technology

The advent of site-specific recombinase (SSR) technology and the Cre/lox system has led to numerous advances in molecular biology, and has proven itself as a valuable tool for assessing gene function in transgenic animals. This interview discusses the mechanism of site specific recombination by Cyclization recombinase (Cre) and how the use of this enzyme has led to the development of conditional mutagenesis, which has significant advantages over traditional knock out strategies.

Principes de la recombinase site-spécifique (SSR) Technologie

Site-specific recombinase (SSR) technology allows the manipulation of gene structure to explore gene function and has become an integral tool of molecular ...

Recherche

Biologie

Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying potassium (referred to as GIRK) channels, which are important for regulating the excitability of neurons. There are four different mammalian GIRK channel subunits (GIRK1-GIRK4) - we focus on GIRK2 because it forms a homotetramer. Stimulation of different types of G protein-coupled receptors (GPCRs), such as the muscarinic receptor (M2R), leads to activation of GIRK channels. Alcohol also directly activates GIRK channels. We will show how to mutate one amino acid by specifically changing one or more nucleotides in the cDNA for the GIRK channel. This mutated cDNA sequence will be amplified in bacteria, purified, and the presence of the point mutation will be confirmed by DNA sequencing. The cDNAs for the mutated and wild-type GIRK channels will be transfected into human embryonic kidney HEK293T cells cultured in vitro. Lastly, whole-cell patch-clamp electrophysiology will be used to study the macroscopic potassium currents through the ectopically expressed wild-type or mutated GIRK channels. In this experiment, we will examine the effect of a L257W mutation in GIRK2 channels on M2R-dependent and alcohol-dependent activation.

We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

Mutagenèse et analyse fonctionnelle des canaux ioniques hétérologue exprimée dans les cellules de mammifères

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying ...

An Introduction to Worm Lab: from Culturing Worms to Mutagenesis

This protocol describes procedures to maintain nematodes in the laboratory and how to mutagenize them using two alternative methods: ethyl methane sulfonate (EMS) and 4, 5', 8-trimethylpsoralen combined with ultraviolet light (TMP/UV). Nematodes are powerful biological systems for genetics studies because of their simple body plan and mating system, which is composed of self-fertilizing hermaphrodites and males that can generate hundreds of progeny per animal. Nematodes are maintained in agar plates containing a lawn of bacteria and can be easily transferred from one plate to another using a pick. EMS is an alkylating agent commonly used to induce point mutations and small deletions, while TMP/UV mainly induces deletions. Depending on the species of nematode being used, concentrations of EMS and TMP will have to be optimized. To isolate recessive mutations of the nematode Pristionchus pacificus, animals of the F2 generation were visually screened for phenotypes. To illustrate these methods, we mutagenized worms and looked for Uncoordinated (Unc), Dumpy (Dpy) and Transformer (Tra) mutants.

Screening for mutants with phenotypic defects is a straightforward method for identifying genes that function in a given biological process. In this article we describe how to culture free living worms (e.g., Pristionchus pacificus) in the laboratory and show two different mutagenesis methods, EMS and TMP/UV.

Une introduction au laboratoire Worm: La culture de Worms à la mutagenèse

This protocol describes procedures to maintain nematodes in the laboratory and how to mutagenize them using two alternative methods: ethyl methane ...

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular signatures, or to evolve proteins in the laboratory. Here, we present a protocol that allows the generation of large (>1011) mutant libraries for a given target sequence. This method is based on replication of a ColE1 plasmid encoding the desired sequence by a low-fidelity variant of DNA polymerase I (LF-Pol I). The target plasmid is transformed into a mutator strain of E. coli and plated on solid media, yielding between 0.2 and 1 mutations/kb, depending on the location of the target gene. Higher mutation frequencies are achieved by iterating this process of mutagenesis. Compared to alternative methods of mutagenesis, our protocol stands out for its simplicity, as no cloning or PCR are involved. Thus, our method is ideal for mutational labeling of plasmids or other Pol I templates or to explore large sections of sequence space for the evolution of activities not present in the original target. The tight spatial control that PCR or randomized oligonucleotide-based methods offer can also be achieved through subsequent cloning of specific sections of the library. Here we provide protocols showing how to create a random mutant library and how to establish drug-based selections in E. coli to identify mutants exhibiting new biochemical activities.

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

Here we demonstrate a simple protocol to create a random mutant library for a given target sequence. We show how this method, which is performed in vivo in Escherichia coli, can be coupled with functional selections to evolve new enzymatic activities.

Protocoles de sélection mutagenèse et fonctionnelles pour l&#39;évolution dirigée de protéines dans les E. coli</em

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

L&#39;examen de la dynamique conformationnelle des protéines membranaires In situ Avec site-dirigé marquage fluorescent

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Analyzing the Function of Small GTPases by Microinjection of Plasmids into Polarized Epithelial Cells

Epithelial cells polarize their plasma membrane into biochemically and functionally distinct apical and basolateral domains where the apical domain faces the 'free' surfaces and the basolateral membrane is in contact with the substrate and neighboring cells. Both membrane domains are separated by tight junctions, which form a diffusion barrier. Apical-basolateral polarization can be recapitulated successfully in culture when epithelial cells such as Madin-Darby Canine Kidney (MDCK) cells are seeded at high density on polycarbonate filters and cultured for several days 1 2. Establishment and maintenance of cell polarity is regulated by an array of small GTPases of the Ras superfamily such as RalA, Cdc42, Rab8, Rab10 and Rab13 3 4 5 6 7. Like all GTPases these proteins cycle between an inactive GDP-bound state and an active GTP-bound state. Specific mutations in the nucleotide binding regions interfere with this cycling 8. For example, Rab13T22N is permanently locked in the GDP-form and thus dubbed 'dominant negative', whereas Rab13Q67L can no longer hydrolyze GTP and is thus locked in a 'dominant active' state 7. To analyze their function in cells both dominant negative and dominant active alleles of GTPases are typically expressed at high levels to interfere with the function of the endogenous proteins 9. An elegant way to achieve high levels of overexpression in a short amount of time is to introduce the plasmids encoding the relevant proteins directly into the nuclei of polarized cells grown on filter supports using microinjection technique. This is often combined with the co-injection of reporter plasmids that encode plasma membrane receptors that are specifically sorted to the apical or basolateral domain. A cargo frequently used to analyze cargo sorting to the basolateral domain is a temperature sensitive allele of the vesicular stomatitis virus glycoprotein (VSVGts045) 10. This protein cannot fold properly at 39&deg;C and will thus be retained in the endoplasmic reticulum (ER) while the regulatory protein of interest is assembled in the cytosol. A shift to 31&deg;C will then allow VSVGts045 to fold properly, leave the ER and travel to the plasma membrane 11. This chase is typically performed in the presence of cycloheximide to prevent further protein synthesis leading to cleaner results. Here we describe in detail the procedure of microinjecting plasmids into polarized cells and subsequent incubations including temperature shifts that allow a comprehensive analysis of regulatory proteins involved in basolateral sorting.

This article details the procedures involved in overexpression and analysis of small GTPases in polarized epithelial cells using microinjection technique.

Analyse de la fonction des petites GTPases par micro-injection de plasmides dans les cellules épithéliales polarisées

Epithelial cells polarize their plasma membrane into biochemically and functionally distinct apical and basolateral domains where the apical domain faces ...

Single Oocyte Bisulfite Mutagenesis

Epigenetics encompasses all heritable and reversible modifications to chromatin that alter gene accessibility, and thus are the primary mechanisms for regulating gene transcription1. DNA methylation is an epigenetic modification that acts predominantly as a repressive mark. Through the covalent addition of a methyl group onto cytosines in CpG dinucleotides, it can recruit additional repressive proteins and histone modifications to initiate processes involved in condensing chromatin and silencing genes2. DNA methylation is essential for normal development as it plays a critical role in developmental programming, cell differentiation, repression of retroviral elements, X-chromosome inactivation and genomic imprinting.
One of the most powerful methods for DNA methylation analysis is bisulfite mutagenesis. Sodium bisulfite is a DNA mutagen that deaminates cytosines into uracils. Following PCR amplification and sequencing, these conversion events are detected as thymines. Methylated cytosines are protected from deamination and thus remain as cytosines, enabling identification of DNA methylation at the individual nucleotide level3. Development of the bisulfite mutagenesis assay has advanced from those originally reported4-6 towards ones that are more sensitive and reproducible7. One key advancement was embedding smaller amounts of DNA in an agarose bead, thereby protecting DNA from the harsh bisulfite treatment8. This enabled methylation analysis to be performed on pools of oocytes and blastocyst-stage embryos9. The most sophisticated bisulfite mutagenesis protocol to date is for individual blastocyst-stage embryos10. However, since blastocysts have on average 64 cells (containing 120-720 pg of genomic DNA), this method is not efficacious for methylation studies on individual oocytes or cleavage-stage embryos.
Taking clues from agarose embedding of minute DNA amounts including oocytes11, here we present a method whereby oocytes are directly embedded in an agarose and lysis solution bead immediately following retrieval and removal of the zona pellucida from the oocyte. This enables us to bypass the two main challenges of single oocyte bisulfite mutagenesis: protecting a minute amount of DNA from degradation, and subsequent loss during the numerous protocol steps. Importantly, as data are obtained from single oocytes, the issue of PCR bias within pools is eliminated. Furthermore, inadvertent cumulus cell contamination is detectable by this method since any sample with more than one methylation pattern may be excluded from analysis12. This protocol provides an improved method for successful and reproducible analyses of DNA methylation at the single-cell level and is ideally suited for individual oocytes as well as cleavage-stage embryos.

Bisulfite mutagenesis is the gold standard for analyzing DNA methylation. Our modified protocol allows for DNA methylation analysis at the single-cell level and was specifically designed for individual oocytes. It can also be used for cleavage-stage embryos.

Simple mutagenèse bisulfite d&#39;ovocytes

Epigenetics encompasses all heritable and reversible modifications to chromatin that alter gene accessibility, and thus are the primary mechanisms for ...

High-throughput Functional Screening using a Homemade Dual-glow Luciferase Assay

We present a rapid and inexpensive high-throughput screening protocol to identify transcriptional regulators of alpha-synuclein, a gene associated with Parkinson&#39;s disease. 293T cells are transiently transfected with plasmids from an arrayed ORF expression library, together with luciferase reporter plasmids, in a one-gene-per-well microplate format. Firefly luciferase activity is assayed after 48 hr to determine the effects of each library gene upon alpha-synuclein transcription, normalized to expression from an internal control construct (a hCMV promoter directing Renilla luciferase). This protocol is facilitated by a bench-top robot enclosed in a biosafety cabinet, which performs aseptic liquid handling in 96-well format. Our automated transfection protocol is readily adaptable to high-throughput lentiviral library production or other functional screening protocols requiring triple-transfections of large numbers of unique library plasmids in conjunction with a common set of helper plasmids. We also present an inexpensive and validated alternative to commercially-available, dual luciferase reagents which employs PTC124, EDTA, and pyrophosphate to suppress firefly luciferase activity prior to measurement of Renilla luciferase. Using these methods, we screened 7,670 human genes and identified 68 regulators of alpha-synuclein. This protocol is easily modifiable to target other genes of interest.

We present a rapid and inexpensive screening method for identifying transcriptional regulators using high-throughput robotic transfections and a homemade dual-glow luciferase assay. This protocol rapidly generates direct side-by-side functional data for thousands of genes and is easily modifiable to target any gene of interest.

Criblage à haut débit fonctionnelle utilisant une maison double lueur de dosage de luciférase

We present a rapid and inexpensive high-throughput screening protocol to identify transcriptional regulators of alpha-synuclein, a gene associated with ...

Generation of Enterobacter sp. YSU Auxotrophs Using Transposon Mutagenesis

Prototrophic bacteria grow on M-9 minimal salts medium supplemented with glucose (M-9 medium), which is used as a carbon and energy source. Auxotrophs can be generated using a transposome. The commercially available, Tn5-derived transposome used in this protocol consists of a linear segment of DNA containing an R6K&#947; replication origin, a gene for kanamycin resistance and two mosaic sequence ends, which serve as transposase binding sites. The transposome, provided as a DNA/transposase protein complex, is introduced by electroporation into the prototrophic strain, Enterobacter sp. YSU, and randomly incorporates itself into this host&#8217;s genome. Transformants are replica plated onto Luria-Bertani agar plates containing kanamycin, (LB-kan) and onto M-9 medium agar plates containing kanamycin (M-9-kan). The transformants that grow on LB-kan plates but not on M-9-kan plates are considered to be auxotrophs. Purified genomic DNA from an auxotroph is partially digested, ligated and transformed into a pir+ Escherichia coli (E. coli) strain. The R6K&#947; replication origin allows the plasmid to replicate in pir+ E. coli strains, and the kanamycin resistance marker allows for plasmid selection. Each transformant possesses a new plasmid containing the transposon flanked by the interrupted chromosomal region. Sanger sequencing and the Basic Local Alignment Search Tool (BLAST) suggest a putative identity of the interrupted gene. There are three advantages to using this transposome mutagenesis strategy. First, it does not rely on the expression of a transposase gene by the host. Second, the transposome is introduced into the target host by electroporation, rather than by conjugation or by transduction and therefore is more efficient. Third, the R6K&#947; replication origin makes it easy to identify the mutated gene which is partially recovered in a recombinant plasmid. This technique can be used to investigate the genes involved in other characteristics of Enterobacter sp. YSU or of a wider variety of bacterial strains.

Generation of Enterobacter sp. YSU Auxotrophs Using Transposon Mutagenesis

Enterobacter sp. YSU grows in glucose minimal salts medium. Auxotrophs are generated by transforming it with a transposome which randomly inserts itself into the host genome. Mutants are found by replica plating from complex medium to minimal medium. Interrupted genes are identified by gene rescue and sequencing.

Génération de<em&gt; Enterobacter</em&gt; Sp. UEE auxotrophes utilisant mutagénèse par transposon

Prototrophic bacteria grow on M-9 minimal salts medium supplemented with glucose (M-9 medium), which is used as a carbon and energy source. Auxotrophs can ...

A Protocol for Functional Assessment of Whole-Protein Saturation Mutagenesis Libraries Utilizing High-Throughput Sequencing

Site-directed mutagenesis has long been used as a method to interrogate protein structure, function and evolution. Recent advances in massively-parallel sequencing technology have opened up the possibility of assessing the functional or fitness effects of large numbers of mutations simultaneously. Here, we present a protocol for experimentally determining the effects of all possible single amino acid mutations in a protein of interest utilizing high-throughput sequencing technology, using the 263 amino acid antibiotic resistance enzyme TEM-1 &#946;-lactamase as an example. In this approach, a whole-protein saturation mutagenesis library is constructed by site-directed mutagenic PCR, randomizing each position individually to all possible amino acids. The library is then transformed into bacteria, and selected for the ability to confer resistance to &#946;-lactam antibiotics. The fitness effect of each mutation is then determined by deep sequencing of the library before and after selection. Importantly, this protocol introduces methods which maximize sequencing read depth and permit the simultaneous selection of the entire mutation library, by mixing adjacent positions into groups of length accommodated by high-throughput sequencing read length and utilizing orthogonal primers to barcode each group. Representative results using this protocol are provided by assessing the fitness effects of all single amino acid mutations in TEM-1 at a clinically relevant dosage of ampicillin. The method should be easily extendable to other proteins for which a high-throughput selection assay is in place.

We present a protocol for the functional assessment of comprehensive single-site saturation mutagenesis libraries of proteins utilizing high-throughput sequencing. Importantly, this approach uses orthogonal primer pairs to multiplex library construction and sequencing. Representative results using TEM-1 &#946;-lactamase selected at a clinically relevant dosage of ampicillin are provided.

Un protocole pour l&#39;évaluation fonctionnelle de Whole-Protein Saturation mutagenèse Bibliothèques Utilisant à haut débit de séquençage

Site-directed mutagenesis has long been used as a method to interrogate protein structure, function and evolution. Recent advances in massively-parallel ...