法の原理： このビデオで説明されている全プラスミドの変異導入を指揮サイトでは、直接プラスミドに数個の塩基の置換、欠失または挿入によってクローニングされた標的遺伝子を変化させることができる突然変異誘発法です。それ以外のPCRベースの熱サイクル反応で、全体のプラスミドを増幅することによって動作します。反応中に、元のプラスミドにミスマッチの形で、目的の変異を有する変異誘発プライマーは、、、新たに合成されたプラスミドに組み込まれています。反応から元のプラスミドを除去した後に変異プラスミドは、E.に変換されるこのメソッドの変異の効率が100％ではないので、 大腸菌。次の手順では、唯一のスクリーニングを目的としています。全体の手順は、3日かかりますが、主な部分は、1日以内に行うことができます。 1.0 1日目： 1.1熱サイクル反応： オリジナルのプラスミド：変異誘発プロトコルを指示このサイトは、最大10キロバイトまでのプラスミドで最高の作品。大きなプラスミドは、この方法で変異するのが少し困難であり、いくつかの忍耐と熱サイクル条件及び/またはコンピテントセルの調整がかかることがあります。さらに、作業対象とするプラスミドは、ダム+細菌の菌株から分離されていなければなりません。熱サイクル反応のためにあなたが変異するプラスミドの10から60 NGSが必要になります。 プライマー：あなたの熱サイクル反応をセットアップする前に手元にプライマーを持たなければならない。あなたはすべての変異プライマーの約150 ngを必要。あなたが10倍希釈100 PMの株式の1.5μlを取ればそれはOKです。あなたの突然変異誘発プライマーを設計する際に考慮すべきいくつかの簡単なガイドラインがあります： <ul><li>プライマーは、互いに相補的である必要があります</li><li>プライマーは、長さ25から45ヌクレオチドの間でなければなりません</li><li>オリジナルのプラスミドへのミスマッチの形の変異は、両方のプライマーに含まれている必要があります</li><li>不一致は、プライマーの中央と両側に少なくとも8ヌクレオチドが隣接する必要がある</li><li>プライマーは、少なくとも40％のGC含量を持つ必要があります</li><li>プライマーは、1つまたは複数のGSまたはCsで5&#39;と3素数を止めるべきである</li><li>プライマーは、リン酸化されることが必要ではない、また彼らは単に彼らがあるべき脱FPLCまたはPAGE精製、しなければならないのか。 </li><li> Tmの計算では、出荷証明書上の任意の式がTmが60℃より高くすべき必要はありません。 </li><li>スクリーニングの目的のためにそれはあなたの突然変異で制限部位を挿入または削除することが現実的です。ために遺伝子コードの変性のご所望の変異で制限部位を挿入するために多くの可能性があります。ニューイングランドバイオラボのウェブサイトは、適切な制限部位を検索するためのツールが用意されています。単にDNAの曖昧さのコードを使用して、あなたが望むのアミノ酸配列をコードするあなたのプライマー配列を入力してください。酵素のファインダーツールでは、制限部位が、ご希望の突然変異に平行に導入することができることを教えてくれます。この方法によってあらゆる可能性や実用的制限部位を見つけることができない場合には、突然変異の部位で遺伝コードを操作せずに、新しい制限部位を挿入することができます。その後、この制限部位が新たな突然変異誘発プライマーの挿入によって削除されるの変異のシリーズ用のテンプレートとしてプラスミドを使用することができます。あなたがプラスミドの同じ部位での突然変異の一連の操作を行うことを計画している場合は、このアプローチは非常に便利であるかもしれません。 </li></ul> DNA -ポリメラーゼ：この方法を使うには3&#39; - 5&#39;エキソヌクレアーゼ活性を示し、平滑末端を生成する耐熱性DNA -ポリメラーゼが必要です。我々は常にFermentasから組換えPFU -ポリメラーゼを使用してください。あなたが増幅ステップで問題を持っている場合は、より品質の高いポリメラーゼを試す可能性がありますが、ほとんどの目的のために標準PFUは十分だろう。のdNTP、ポリメラーゼ緩衝液と水との反応を完了します。 1.2熱サイクル条件サーモサイクラーは以下の方法で設定する必要があります。初期および定期的な変性段階を95℃で30秒に設定されていますプライマーのアニーリング温度は、複雑な数式で計算してはいけません。我々は、通常55℃、1分アニール時間にアニーリング温度を設定します。あなたが主に使用する25から30ヌクレオチドのプライマーの場合は、これは私達のための時間の95％に動作します。と60℃ - これが動作しないかどうかいずれにせよ、単に50の間にアニーリング温度を変えてみ延伸温度は、使用するポリメラーゼによって異なります。このデモでは、72℃の伸長温度を呼びかけFermentas、からPFU -ポリメラーゼを使用して伸長時間は、プラスミドのサイズによって異なります。我々は常にkbあたり1分を計算し、そのための1分を追加時間。例えば、9キロバイトプラスミドのために我々は、伸長時間として10分を選択することになります。 18サイクルは十分にさらに使用するためのプラスミド変異作成するだけで十分です。サイクル数を低く保つこと、また、時間を節約できます。 <img src="/files/ftp_upload/1135/figure1.png" alt="図1" width="600" height="287" />熱サイクル反応後の1.3 Gelcheck 成功した増幅、電気泳動を行うことによりチェックする必要があります。サイクリングが終了した直後に、1％TAEアガロースゲルに反応液5μlをロードする。増幅が成功した場合には、明瞭なバンドが表示されるはずです。新たに合成されたDNAは、ゲル上で明確に表示されていない場合ただし、全体の反応を沈殿させるために試みると、次のステップで変換のためにそれを使用することができます。私たちにとってこのことはほとんど働いていないが、それは試してみる価値はある。反応が動作しない場合は最善の方法は、アニーリング温度を調整することです。 1.4 DpnI消化： 変換する前にテンプレートを務めた元のプラスミドは、強力なバックグラウンドを防ぐために、反応から削除する必要があります。これは、DpnIで制限酵素消化によって行われます。この制限エンドヌクレアーゼは、メチル化プラスミドカット。メチル化される必要があるのに対し、その認識と制限部位は、配列GATCです。 DpnIで切断するときにダムから隔離された唯一のオリジナルunmutatedプラスミドを+株はカットさ、メチル化されていない新たに合成された変異プラスミドは、DpnIの影響を受けません。単に反応するDpnIの1〜2μLを追加し、37℃で少なくとも1時間振とうする。高速ダイジェストDpnIを使用する場合はインキュベーション時間は約15分に短縮することができます。あなたのDpnIとこの制限酵素消化のインキュベーション時間の質が大幅unmutatedプラスミドを持つあなたの背景がどのくらい強いかを決定します。 1.5変換： DpnIの消化後にプラスミドは、コンピテント大腸菌への形質転換のための準備ができています大腸菌細胞。単純にコンピテント細胞にDpnIの消化反応から5μlを追加し、ラボで推奨されている変換を実行します。我々のケースでは、氷上で30分間細菌プラスミド混合物をインキュベートし、42℃90秒で、熱衝撃、それによって熱ショックプロシージャを使用します。 200μlのSOC -ソリューションを追加した後、細菌が培養される、精力的に揺れて、37℃で1時間C 1時間後に細菌を寒天培地を選択することでメッキされています。 2.0 2日目クローンのパート1の2.1スクリーニング：クローンの選定突然変異の効率が100％ではないため、あなたの突然変異体をスクリーニングする必要があります。我々は我々が追加またはプライマーの統合により削除さ制限部位の有無をチェックする制限酵素で消化することによって行います。この目的のために我々は、通常8コロニーをピックアップし、翌日のプラスミド調製のために一晩、それらを成長させる。 3.0 3日目クローンのパート2の3.1スクリーニング：マーカー酵素と制限酵素処理三日目に、あなたのマーカー酵素でミニプレップし、その後の制限酵素消化を行う。ゲル上では、その後、所望の変異を運ぶとなるものではない自分のクローンのどれを見ることができます。 制限酵素消化を用いたスクリーニング方法は、非常にうまく機能します。それにもかかわらず、あなたは配列決定によってあなたの成功の変異を確認する必要があります。

<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.

部位特異的突然変異誘発は、プラスミドによって運ばれる遺伝子を変異させる高速な方法を提供する変異導入法です。全体の反応は、一日で行うことができます。この方法では、それだけで所望の変異とそのようなPFU -ポリメラーゼのようなプルーフリーディングポリメラーゼを運んで無料のプライマーのペアが必要になります。新たに合成されたプラスミドを制限酵素DpnIとの反応を消化し?...

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

全体プラスミドの部位特異的突然変異誘発は、元のプラスミドのわずかに異なるバリエーションを作成する簡単な方法です。この方法ではクローン化された標的遺伝子は、直接プラスミドに数個の塩基の置換、欠失または挿入することで変更できます。これは単に非PCRベースの熱サイクル反応で、全体のプラスミドを増幅することによって動作します。反応中に所望の変異を運ぶ突然変異誘発性プライマーは、、、新たに合成されたプラスミドに組み込まれています。このビデオチュートリアルでは、プラスミドに塩基置換を導入しやすいと費用対効果の高い方法を示しています。プロトコルは、標準試薬と連携し、多くの場合非常に高価な市販のキット、から独立しています。このプロトコルを適用すると、それは市販のキットのいくつかを使用して、かかるコストの8位に反応のトータルコストを削減することができます。このビデオでは、また、プロセス中の重要な手順についてはコメントおよび突然変異誘発プライマーを設計する方法に関する詳細な指示を与える。

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

Homemade Site Directed Mutagenesis of Whole Plasmids

全体プラスミドの部位特異的突然変異誘発は、元のプラスミドのわずかに異なるバリエーションを作成する簡単な方法です。ここでは、プラスミドを使用して、標準試薬に塩基置換を導入しやすいと費用対効果の高い方法を示しています。

全プラスミドの自家製のサイトが特異的突然変異誘発

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 ビュー.  Johannes Gutenberg-University Mainz, Germany. 全体プラスミドの部位特異的突然変異誘発は、元のプラスミドのわずかに異なるバリエーションを作成する簡単な方法です。ここでは、プラスミドを使用して、標準試薬に塩基置換を導入しやすいと費用対効果の高い方法を示しています。

ビデオ: 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.

部位特異的リコンビナーゼ（SSR）技術の原理

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

生物学

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.

ワームの研究室の紹介：培養ワームから変異へ

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.

内タンパク質の定方向進化のための変異導入と機能選択プロトコル E.大腸菌</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.

膜タンパク質の構造動力学を調べるその場で部位特異的蛍光標識を持つ

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.

偏上皮細胞へのプラスミドのマイクロインジェクションによる低分子量GTPaseの機能を解析する

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.

単一の卵重亜硫酸塩の突然変異誘発

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.

自家製デュアルグロールシフェラーゼアッセイを用いて、ハイスループット機能スクリーニング

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.

の世代<em&gt;エンテロバクター</em&gt;エスピー。 YSU要求株にはトランスポゾン突然変異誘発を用いて

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.

ハイスループットシーケンシングを利用し全体プロテイン飽和突然変異誘発ライブラリの機能評価のためのプロトコル

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

Simple Homemade Tools to Handle Fruit Flies&#8212;Drosophila melanogaster

The fruit fly, Drosophila melanogaster, is widely used both in biological research and biology education. Handling adult flies is common but difficult in practice, as adult flies fly. Demonstrated here is how to make some simple and cost-effective tools to address difficult issues in the handling of Drosophila. Holes in foam stoppers are made and pipette tips or funnels are inserted into the holes. Flies then move only in one direction into the pipette tip/funnel assemblage, allowing efficient control of the transfer of adult Drosophila into or out of a vial. Existing protocols have been modified for cool-anesthetizing flies by chilling in crushed ice and transferring them onto a cold, hard icepack surface. The icepack is covered with a piece of medical gauze that keeps immobilized flies from the condensed water when examined under a stereomicroscope. The flies are finally euthanized for counting and sorting or discarded by microwaving. A bottle-shaped cage has also been developed for collecting eggs, as well as a labor-saving device and accompanying protocol for cleaning glass culture vials.

Described here is the use of several homemade tools to transfer, chill, and kill adult Drosophila, as well as to clean glass culture vials and collect eggs. These tools are easy to make and are rather efficient in handling Drosophila.

フルーツハエを扱うシンプルな自家製ツール -ショウジョウバエメラノガスター

The fruit fly, Drosophila melanogaster, is widely used both in biological research and biology education. Handling adult flies is common but difficult in ...