Principio del Metodo: Il sito mutagenesi di plasmidi tutto spiegato in questo video è un metodo di mutagenesi che consente di modificare un gene bersaglio clonato attraverso la sostituzione, cancellazione o l&#39;inserimento di alcune basi direttamente in un plasmide. Funziona, amplificando il plasmide intero, in modo non basati sulla PCR reazione termici. Durante la reazione di primer mutagenico, portando la mutazione desiderata in forma di mismatch al plasmide originale, sono integrati nel plasmide di nuova sintesi. Dopo la rimozione del plasmide originale dalla reazione del plasmide mutato si trasforma in E. coli. seguenti passaggi sono solo a scopo di screening, perché l&#39;efficienza mutazione di questo metodo non è al 100%. L&#39;intera procedura dura tre giorni, ma la parte principale può essere fatto in un giorno. 1,0 ° giorno: 1,1 reazione termici: Plasmidi originale: Questo sito mutagenesi protocollo funziona meglio con i plasmidi fino a 10kb. Grandi plasmidi sono un po &#39;difficile da mutare con questo metodo e può prendere po&#39; di pazienza e la regolazione delle condizioni termici e / o cellule competenti. Inoltre il plasmide che si lavora con deve essere isolato da una diga + ceppo di batteri. Per la reazione termici è necessario 10-60 NGS del plasmide si vuole mutare. Primer: Prima di poter impostare la vostra reazione termici devi avere la tua primer a portata di mano. Avete bisogno di circa 150 ng di ogni fondo mutageno. E &#39;ok se si prende 1,5 ml di una soluzione 1:10 diluito al 100 pM magazzino. Ci sono alcune semplici linee guida è necessario considerare quando si progetta il vostro primer mutagenico: <ul><li> I primer devono essere complementari l&#39;uno all&#39;altro </li><li> Il primer deve essere compreso tra 25 e 45 nucleotidi di lunghezza </li><li> Le mutazioni nella forma dei disallineamenti al plasmide originale deve essere contenuta in entrambi i primers </li><li> La non corrispondenza dovrebbe essere al centro del fondo e fiancheggiata da almeno 8 nucleotidi su ogni lato </li><li> I primer dovrebbe avere un GC-Content di almeno il 40% </li><li> I primer dovrebbe finire 5 Prime e 3 primi con uno o più G o Cs </li><li> Il primer non devono essere fosforilata, né devono essere FPLC o PAGINA purificata, semplicemente dissalati dovrebbero essere. </li><li> Per il calcolo della Tm non hai bisogno di alcuna formula ma Tm sul certificato di spedizione deve essere superiore a 60 ° C. </li><li> Ai fini dello screening è pratico di inserire o eliminare un sito di restrizione con il tuo mutazione. A causa della degenerazione del codice genetico ci sono molte possibilità di inserire un sito di restrizione con la tua mutazione desiderata. Il sito web del New England Biolabs fornisce uno strumento per trovare un simile sito appropriato restrizione. Basta inserire la sequenza di innesco che codifica per la sequenza di aminoacidi che desideri, utilizzando il codice ambiguità del DNA. L&#39;enzima strumento di ricerca vi dirà quali siti di restrizione può essere introdotto parallelamente al mutazione desiderata. Se non riesci a trovare alcun sito di restrizione possibile o pratico in questo modo, è possibile inserire un qualsiasi nuovo sito senza restrizioni riguardanti il ​​codice genetico nel sito di mutazione. Successivamente è possibile utilizzare questo plasmide come modello per una serie di mutazioni in cui questo sito di restrizione sarà cancellato dal inserimento del nuovo primer mutagenico. Questo approccio può essere molto conveniente se hai intenzione di fare una serie di mutazioni nello stesso sito del plasmide. </li></ul> DNA polimerasi: Per questo metodo hai bisogno di una DNA polimerasi termostabile che presenta 3&#39;-5 &#39;attività esonucleasi e crea le estremità smussata. Usiamo sempre ricombinante Pfu-Polymerase da Fermentas. Se hai problemi con la procedura di amplificazione è possibile provare una polimerasi di qualità superiore, ma per la maggior parte degli scopi del Pfu standard sarà sufficiente. Completa la reazione con tampone polimerasi dNTPs, e l&#39;acqua. 1.2 Condizioni termici Il termociclatore dovrebbe essere istituito nel seguente modo. La fase iniziale di denaturazione e ricorrenti è impostato per 30 sec a 95 ° C. La temperatura di annealing del primer non deve essere calcolato con formule complicate. Di solito impostare la temperatura di annealing a 55 ° C e il tempo di ricottura a un minuto. Per primer da 25 a 30 nucleotidi, che si prevede di utilizzare per lo più, questo funziona per noi il 95% del tempo. Comunque se questo non dovrebbe funzionare, basta provare variando la temperatura di ricottura tra i 50 - e 60 ° C. La temperatura di estensione dipende dalla polimerasi in uso. In questa dimostrazione si usa il Pfu-Polymerase da Fermentas, che prevede un allungamento temperatura di 72 ° C. Il tempo allungamento varia a seconda delle dimensioni del plasmide. Abbiamo sempre calcolare 1 minuto per ogni kb, e aggiungere un minuto supplementare a quelladel tempo. Ad esempio, per un plasmide 9kb che sceglieremmo 10 minuti il ​​tempo di allungamento. 18 cicli sono sufficienti a creare abbastanza mutato plasmide per l&#39;ulteriore utilizzo. Mantenere il numero di cicli a bassa, anche il tempo. <img src="/files/ftp_upload/1135/figure1.png" alt="Figura 1" width="600" height="287" /> 1,3 Gelcheck dopo la reazione termici L&#39;amplificazione di successo dovrebbe essere controllato eseguendo una elettroforesi. Subito dopo il ciclismo è finito, carico 5 l di reazione su un 1% gel di agarosio al TAE. Se l&#39;amplificazione è stata eseguita correttamente, si dovrebbe vedere un gruppo distinto. Tuttavia, se il DNA di nuova sintesi non è chiaramente visibile sul gel, si può tentare di accelerare la reazione di tutto e usarlo per la trasformazione nella fase successiva. Per noi questo ha funzionato raramente, ma vale la pena di fare un tentativo. La cosa migliore da fare se la reazione non funziona è quello di regolare la temperatura di ricottura. 1,4 DpnI digestione: Prima della trasformazione il plasmide originale che servì come modello deve essere rimosso dalla reazione per evitare forte background. Questo viene fatto con la digestione di restrizione DpnI. Questa endonucleasi di restrizione taglia solo metilati plasmide. Il suo riconoscimento e sito di restrizione è la sequenza GATC, mentre A deve essere denaturato. Durante la digestione con DpnI solo il plasmide originale immutata, che è stato isolato da una diga + ceppo viene tagliato, il plasmide mutato di nuova sintesi che non è metilato non è influenzata dal DpnI. Basta aggiungere 1-2 ml di DpnI alla reazione e incubare che almeno un&#39;ora a 37 °. Quando si utilizza il veloce digerire DpnI il tempo di incubazione può essere ridotto a circa 15 minuti. La qualità della vostra DpnI e il tempo di incubazione di questa influenza notevolmente la digestione restrizione quanto sia forte il vostro sfondo con plasmide immutata sarà. 1,5 Trasformazione: Dopo la digestione DpnI il plasmide è pronto per la trasformazione in autorità competenti E. coli cellule. Basta aggiungere 5 microlitri dalla reazione digestione DpnI nelle cellule competenti ed eseguire la trasformazione come raccomandato nel vostro laboratorio. Nel nostro caso usiamo la procedura di shock termico incubando i batteri-plasmide miscela in ghiaccio per 30 minuti e poi lo shock termico a 42 ° C per 90 sec. Dopo aver aggiunto 200 ul SOC-soluzione, i batteri vengono incubati, agitando vigorosamente, a 37 ° C per 1 h. Dopo 1 ora i batteri sono placcati sulla scelta dei supporti di agar. 2,0 secondo giorno 2,1 screening di cloni parte 1: Scelta dei cloni Poiché l&#39;efficienza mutazione non è al 100% è necessario per il vostro schermo mutanti. Facciamo questo digestione di restrizione in cui verificare la presenza o l&#39;assenza del sito di restrizione che abbiamo aggiunto o eliminato attraverso l&#39;integrazione primer. A questo scopo siamo soliti prendere otto colonie e farle crescere per tutta la notte per la preparazione plasmide il giorno successivo. 3,0 Terzo giorno 3,1 screening di cloni parte 2: digestione con enzimi di restrizione marcatore Il terzo giorno si esegue un Prep Mini e la digestione con la conseguente restrizione degli enzimi marker. Sul gel poi si può vedere che uno dei tuoi cloni portano la mutazione desiderata e non quelli. Il metodo di screening con digestione restrizione funziona molto bene. Tuttavia, si dovrebbe confermare la mutazione di successo mediante sequenziamento.

<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.

Mutagenesi sito-è un metodo di mutagenesi, che fornisce un modo veloce per mutare un gene portato da un plasmide. La reazione tutto può essere fatto in un solo giorno. Con questo metodo, sarà necessario solo un paio di primer omaggio portando le mutazioni desiderato e una polimerasi correzione di bozze, come Pfu-polimerasi. Il plasmide di nuova sintesi può essere separato dal plasmide genitoriale digerire la reazione con il DpnI enzima di restrizione. Questo enzima digerisce solo il DNA metilato. Quindi solo il plas...

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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>
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<td>R0191</td>
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<td>DpnI or DpnI Fast digest</td>
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<td>fermentas</td>
<td>ER1701</td>
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<td>primers</td>
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homemade site directed mutagenesis of whole plasmids

Mutagenesi sito-specifica di plasmidi tutto è un modo semplice per creare varianti leggermente diverse di un plasmide originale. Con questo metodo il gene bersaglio clonato può essere modificato attraverso la sostituzione, cancellazione o l&#39;inserimento di alcune basi direttamente in un plasmide. Funziona semplicemente amplificando il plasmide intero, in modo non basati sulla PCR reazione termici. Durante la reazione di primer mutagenico, portando la mutazione desiderata, sono integrati nel plasmide di nuova sintesi. In questo video tutorial che mostrano un modo semplice ed economico per introdurre sostituzioni di basi in un plasmide. Il protocollo lavora con reagenti standard ed è indipendente dal kit commerciali, che spesso sono molto costosi. L&#39;applicazione di questo protocollo in grado di ridurre il costo totale di una reazione ad un ottavo di ciò che costa utilizzando alcuni dei kit commerciali. In questo video abbiamo anche commentare le fasi critiche durante il processo e dare istruzioni dettagliate su come progettare il primer mutagenico.

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

Homemade Site Directed Mutagenesis of Whole Plasmids

Mutagenesi sito-specifica di plasmidi tutto è un modo semplice per creare varianti leggermente diverse di un plasmide originale. Qui mostriamo un modo semplice ed economico per introdurre sostituzioni di basi in un plasmide standard utilizzando reagenti.

Mutagenesi sito fatto in casa Regia di Plasmidi Whole

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 Views.  Johannes Gutenberg-University Mainz, Germany. Mutagenesi sito-specifica di plasmidi tutto è un modo semplice per creare varianti leggermente diverse di un plasmide originale. Qui mostriamo un modo semplice ed economico per introdurre sostituzioni di basi in un plasmide standard utilizzando reagenti.

Video: 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.

Principi di Site-Specific ricombinasi (SSR) Tecnologia

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

Ricerca

Biologia

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.

Mutagenesi e Analisi funzionale dei canali ionici Heterologously Espresso in cellule di mammifero

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.

Introduzione al Laboratorio Worm: da Worms coltura a mutagenesi

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.

Protocolli di selezione mutagenesi e funzionale per l&#39;evoluzione Regia di proteine ​​in 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.

Esaminando la dinamica conformazionale delle proteine ​​di membrana In situ Con il sito-diretta Etichettatura fluorescenza

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.

Analizzando la funzione di GTPasi Small Microiniezione di plasmidi in cellule epiteliali polarizzate

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.

Singolo ovociti Mutagenesi bisolfito

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.

High-throughput screening funzionale utilizzando una casalinga 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 ...

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.

Generazione di<em&gt; Enterobacter</em&gt; Sp. YSU auxotrofi Utilizzando Transposon Mutagenesi

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 protocollo per la valutazione funzionale del Whole-Protein Saturazione mutagenesi biblioteche Utilizzando 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 ...