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TempliPhi-PDF

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Materials

  • illustra TempliPhi™ 100/500 Amplification Kit
  • 0.6mL PCR tubes

Procedure

  1. Thaw sample buffer (red cap) and reaction buffer (blue cap) on ice.
  2. Transfer 5 μL of sample buffer to a small PCR tube for each template to be amplified.
  3. Add template to sample buffer
    • Dilute 1μL saturated overnight culture in 10-100 μL water. Use 0.2-0.5 μL.
    • Use a small portion of a colony. Avoid transferring any agar.
    • Dilute 1μL glycerol stock in 50μL water. Use 0.2-0.5 μL.
    • Use 1pg-10ng of purified plasmid DNA (volume < 0.5μL)
  4. Heat sample to 95°C for 3 mins to denature and cool to 4°C.
  5. Mix 5 μL reaction buffer with 0.2μL enzyme mix for each reaction. (Make up a master mix for multiple reactions.)
  6. Transfer 5 μL TempliPhi premix to the cooled sample.
  7. Incubate at 30°C for 4-18 hours.
    • Incubate overnight for optimal results but 4 hours should be sufficient if there is not too much inhibitory material like agar or rich medium.
  8. Heat to 65°C for 10 mins.
  9. Cool to 4 °C.
  10. Send DNA for sequencing.
    • Use 2μL in 12μL for sequencing at the biopolymers facility
    • Previously, for sequencing on the Knight lab sequencer, we used 1μL

In Vitro Kinase AssaGriffiny-PDF

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Assay of Tyrosine Kinases Using Synthetic Peptides

Todd Miller (State University of New York at Stony Brook)

Small synthetic peptide substrates are especially well suited for applications such as assays of tyrosine kinases in permeabilized cells or for enzyme kinetic studies. Although a number of different techniques are available to separate the phosphorylated peptides from other assay components, the most commonly used method is to use peptides containing basic residues. These peptides bind to phosphocellulose paper at low pH, while labelled ATP does not and is washed away.

Starting Materials

  • Protein Kinase (stock solutions of 1-10 mg/ml pure kinases): for these assays, we have used both purified kinases as well as crude cell extracts. For cell extracts, we recommend the addition of a phosphatase inhibitor (e.g. 0.1-1 mM sodium orthovanadate). For each enzyme, it is important to determine the optimal buffer, ionic strength, and pH for activity. If these conditions have not been established, the protocol listed below can be used as a starting point.
  • Peptide Substrate (stock solution of 10 mM): peptide substrates typically contain one tyrosine in a phosphorylation site motif. A recent compilation of phosphorylation sites can be found in Biochimica et Biophysica Acta 1314 (1996) 191-225. In addition, the peptide substrates should have a net positive charge to facilitate binding to phosphocellulose filters used in the assay. For quantitative binding to the phosphocellulose paper, we recommend at least 2 basic residues and a free amino terminus. If a phosphorylation site motif is not known, a general tyrosine kinase substrate can be used. For example, “Src-related peptide” (RRLIEDAEYAARG; Sigma # A7433) is a substrate for many receptor and nonreceptor tyrosine kinases). For initial reactions, a peptide concentration of 0.7-1.5 mM should be used. To determine the kinetic parameters for phosphorylation of the synthetic peptide, a range of peptide concentrations is required (see below).
  • 5X Kinase Buffer: contains 5 mg/mL BSA (to prevent kinase adsorption to the assay tube), 150 mM Tris-Cl (pH 7.5), 100 mM MgCl2. Divalent cations are required for most tyrosine kinases, although some tyrosine kinases (for example, insulin, IGF-1, and PDGF receptor kinases) require MnCl2 in place of MgCl2 (or in addition to MgCl2). The optimal concentrations of divalent cations must be determined empirically.
  • ATP: a stock solution of 1-5 mM is convenient. Note that most tyrosine kinases have Km values for ATP in the range 10-150 µM, so for kinetic experiments it is important to use saturating concentrations of ATP to arrive at values of Km and Vmax for the peptides.
  • [gamma-32P]ATP:10 mCi/mL.

Tyrosine Kinase Assays

A standard tyrosine kinase assay is carried out in a volume of 25 µl:

  • 5 µl of 5X kinase buffer
  • 5 µl of 1.0 mM ATP (0.2 mM final concentration)
  • [gamma-32P]ATP (100-500 cpm/pmol)
  • 3 µl of 10 mM peptide substrate (1.2 mM final concentration)
  • tyrosine kinase
  • H2O to 25 µl

I) Before the experiments, prepare a cocktail containing enough buffer, ATP, and [gamma-32P]ATP to complete the assays. For experiments with the same peptide substrate concentrations, the peptide should be incorporated into the cocktail. For assays at different peptide concentrations, the substrate should be diluted and added separately to each tube. After dispensing the cocktail into 1.5 ml microcentrifuge tubes, place the tubes in a water bath at 30 degrees C. Reactions should be initiated by the addition of kinase and allowed to proceed at 30 degrees C.

II) After the desired time, terminate the reactions by adding 45 µl ice-cold 10% trichloroacetic acid (TCA) to each reaction. Vortex the reactions.

III) Spin for 2 minutes in microcentrifuge (10K rpm).

IV) Spot 35 µl of the supernatants onto 2.1-cm diameter Whatman P81 cellulose phosphate filter circles.

VI) Wash the P81 filter circles three times with 500 ml cold 0.5% phosphoric acid (5-10 minutes per wash). The progress of the washing steps can be followed by removing the P81 filter circle for a blank reaction and checking it with a Geiger counter.

VII) Wash once with 200 ml acetone at room temperature for 5 minutes.

VIII) Allow the filter circles to dry at room temperature.

IX) Put filter circles into scintillation vials and measure 32P incorporation by counting the pads dry in a scintillation counter. The specific activity of ATP in a kinase reaction (e.g., in cpm/pmol) can be determined by spotting a small sample (2-5 µl) of the reaction onto a P81 filter circle and counting directly (no washing). Counts per minute obtained in the kinase reaction (minus blank) are then divided by the specific activity to determine the moles of phosphate transferred in the reaction.

Kinetics of Peptide Phosphorylation

The kinetic parameters for phosphorylation of a synthetic peptide by a tyrosine kinase can be determined using a variation on the protocol above.

I) Carry out a reaction at a high concentration of peptide (see above) to establish that the peptide is a substrate.

II) Vary the enzyme concentration in the assay. The rate of peptide phosphorylation should be proportional to the enzyme concentration under the conditions of the assay. This experiment is also used to determine the amount of enzyme needed for the kinetic studies. To determine rates, a time course of peptide phosphorylation should be carried out. In this case, prepare a larger enzyme reaction (we use 150 µl). At the desired time points, withdraw 25 µl aliquots and transfer them to microcentrifuge tubes containing 45 µl of ice-cold 10% TCA, and analyze the reactions as described above. Phosphorylation of the peptide should be linear with time, and for measurement of kinetic constants the initial rates of reaction (5%) should be used.

III) Vary the peptide concentration in the assay. Use a plot of velocity vs. peptide concentration to get an initial estimate of the value of Km. A wide range of substrate concentrations (e.g., 20 µM to 2 mM) should be used in this initial measurement.

IV) To determine Km (peptide) and Vmax , vary the peptide concentration and measure the rate of phosphate transfer. A good range of substrate concentrations are the following multiples of Km: 0.125 x Km, 0.25 x Km, 0.5 x Km, 1.0 x Km, 2.0 x Km, 4.0 x Km, 8.0 x Km. The reactions should be carried out in triplicate for best results.

V) Kinetic constants are determined by weighted non-linear least-squares fit to the hyperbolic velocity vs. [substrate] plots using iterative programs such as NFIT (Island Products, Galveston, TX).

Agarose Gel Electrophoresis-PDF

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Agarose Gel Electrophoresis-PDF

Procedure

  1. Place the gel tray perpendicular in the electrophoresis chamber to create a casting bay.
  2. Wipe the tray clean with a Kim-wipe and level the tray using the bubble level.
  3. Weigh out the desired amount of agarose and add it to 100ml of 1X TBE in a 300ml flask ( Mike usually uses 1.3 – 1.4g).
  4. Microwave the flask on high for 80 seconds. Be sure that it does not boil over.
  5. Swirl the microwaved agarose in the flask until the solution becomes clear.
  6. Pour the melted solution into the casting bay and insert the comb.
  7. Prepare the DNA ladder by combining the following:
    1. 10ul DNA ladder
    2. 1ul SYBR green (100X)
    3. 1ul Bromophenol Blue
  8. Prepare the DNA Samples by combining the following:
    1. 40ul DNA preparation
    2. 4ul SYBR Green (100X)
    3. 4ul Bromophenol Blue
  9. Remove the comb from the cured gel and realign the gel in the chamber.
  10. Adjust the buffer level by adding 1X TBE to the chamber until the buffer just covers the gel. The buffer can be reused a few times.
  11. Pipette the samples into the wells
  12. Apply 150 volts and run for approximately 60 minutes.
  13. Photograph the gel using the UVP transilluminator system.

Notes

  • Our “1X TBE” is technically 0.5X TBE (5.3g/L Tris Base, 2.75g/L Boric Acid, and 2.9g/L EDTA) but for our purposes we call it 1X TBE.
    • Correspondingly, the 10X TBE is officially 5X.
  • Using TBE allows us to have such a high (150V) voltage. If you use TAE you need to use a significantly lower voltage and your run time will be longer. Despite that, TAE is advantagous in some cases, but Mike feels that TBE is better suited for his applications. This is discussed in detail in the consensus protocol.
  • Use the following table to determine the amount of agarose you want to use.

Choosing an Agarose Concentration

Agarose Concentration (g/100mL) Optimal DNA Resolution (kb)
0.5 1 – 30
0.7 0.8 – 12
1.0 0.5 – 10
1.2 0.4 – 7
1.5 0.2 – 3

 

DNA ligation using T4 DNA ligase-PDF

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Materials

  • We use the T4 DNA ligase from NEB
  • Deionized, sterile H2O
  • Purified, linearized vector (in H2O)
  • Purified, linearized insert (in H2O)

Ligation Mix

  • X μL vector (equivalent to ~50 ng, can use less)
  • Y μL insert 1
  • Z μL insert 2
  • 1 μL 10X Ligase Buffer
  • (9.5 – X – Y – Z) μL deionized H2O
  • 0.5 μL T4 DNA ligase
  • Reshma 19:12, 13 December 2007 (CST): I frequently digest 500 ng of each part and the destination vector. Then I use 2 μL vector, 3 μL insert 1 and 3 μL insert 2 where all three linearized fragments have been purified with a Qiagen minelute PCR purification.

Calculating Insert Amount

[math]\displaystyle{ \rm{Insert\ Mass\ in\ ng} = 3\times\left[\frac{\rm{Insert\ Length\ in\ bp}}{\rm{Vector\ Length\ in\ bp}}\right]\times \rm{Vector\ Mass\ in\ ng} }[/math]

(Equimolar ratios may be preferable.)

Procedure

  1. Add appropriate amount of deionized H2O to sterile 0.6 mL tube
  2. Add 1 μL ligation buffer to the tube.
    Vortex buffer before pipetting to ensure that it is well-mixed.
    Remember that the buffer contains ATP so repeated freeze, thaw cycles can degrade the ATP thereby decreasing the efficiency of ligation. It is recommended that you aliquot the Ligation Buffer into smaller quantities.
  3. Add appropriate amount of insert to the tube.
  4. Add appropriate amount of vector to the tube.
  5. Add 0.5 μL ligase.
    Vortex ligase before pipetting to ensure that it is well-mixed.
    Also, the ligase, like most enzymes, is in some percentage of glycerol which tends to stick to the sides of your tip. To ensure you add only 0.5 μL, just touch your tip to the surface of the liquid when pipetting.
  6. Incubate 20 mins on the benchtop.
  7. Place on ice until transformation.
  8. Generally 1 μL of ligation mix is sufficient for either chemical transformation or electroporation. The amount of salt in 1 μL ligation mix should not cause arcing.
  9. Optional Heat-inactivate by incubating at 65°C for 20 mins. Then do a purification step to remove PEG. (See notes on DNA ligation.

ChIP francais-PDF

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Overview

L’immunoprecipitation de la chromatine ou ChIP est une technique qui permet d’identifier les sites de liaison d’un facteur de transcription proteique sur l’ADN in situ.

Materials

  • Chromatin Immunoprecipitation (ChIP) Assay Kit d’Upstate, Catalog # 17-295
  • Préparation d’inhibiteurs de protéases Roche version MINI cat 04 693 159 : 25x = 0,42 ml eau par tablette (Version « complete EDTA-free » cat 04 693 132, c’est 2 ml par tablette)
  • tubes a microcentrifuge 1,5 ml
  • tubes a centrifuge of 15 ml
  • sonicateur style Branson
  • potter ou mortier en agate et azote liquide
  • anticorps polyclonaux valides pour le ChIP y compris anti-histone H3 d’Abcam pour *temoin positif et anti-IgG pour temoin negatif
  • equipement d’electrophorese et agarose 1-2%
  • pipettes
  • grants
  • glace

Procedure

  1. Se procurer les tissus de souris. Prévoir 1 ml de fixateur par 100 mg de tissus dans un Falcon pré-pesé de 15 ml. Transférer les tissus avec PBS, centrifugation 1’ à 500g et enlever le surnageant avant de peser. Garder au froid et fixer aussitôt.
    1. 10 ml fixateur :
        • 9,33 ml PBS
        • 270 ul formaldéhyde 37%
        • (20 ul 0,5M EDTA)
  2. Fixer dans quelques ml pendant 10 ou 15 minutes à température ambiante avec rotation.
  3. Pendant ce temps, sortir le tampon « SDS Lysis Buffer » à température ambiante si on poursuit dans la foulée.
  4. Centrifugation 4ºC pour 30 secondes à 500g et/ou décanter fixateur immédiatement.
  5. Reprise dans 5-10 ml pour une concentration finale de 0,125M glycine :
      • Pour 10 ml de solution arrêt :
        • 8,25 ml PBS
        • 100 ul Triton X100 à 10% (conserver stock au frais)
        • (PBS + Triton X100 = « PBT »)
        • 1,25 ml glycine 1M
        • 400 ul inhibiteurs de protéases 25x
  6. Agiter doucement pendant 5 minutes pour arrêter la fixation.
  7. Centrifugation 4ºC pour 5 minutes à 500g. Décanter.
  8. Rincer avec PBT / 1x inhibiteurs de protéases sans glycine, toujours sur glace. Centrifugation et décantation.
  9. Enlever le surnageant autant que possible, reprendre les tissus dans 0,1 ml de SDS Lysis Buffer (tampon à faire à la main ainsi puisque le kit ne suffira pas en volume!).
      • Pour 50 ml SDS Lysis Buffer :
        • 2,5 ml de SDS 20% (1% final)
        • 1 ml de EDTA 0,5M pH 8 (10 mM final)
        • 2,5 ml de Tris 1 M pH 8.1 (50 mM final)
        • 40 ml H2O
        • 5 tablettes d’inhibiteurs de protéases Roche version MINI cat 04 693 159 (1 tablette si non version Mini)
        • Vérifier pH à la fin pour pH 8.1 puis qsp 50 ml.
  10. Broyer en mortier agate avec de l’azote ou avec potter sur glace. Repartir le broyat pour l’équivalent de 5 mg tissu par tube Falcon de 15 ml, compléter chaque tube Falcon à 400 ul avec du SDS Lysis Buffer.
  11. Garder 4 tubes pour une manipe puis congeler les autres à -30ºC ou -80ºC. Garder lysat sur glace a tout moment.

—

  1. Prendre avec un embout biseauté 300 ul de la bouteille resuspendue de Protein A-agarose/ssDNA (60 ul par tube prévu), centrifuger doucement et décanter surnageant.
  2. Reprendre les billes dans 1 ml ChIP Dilution Buffer plus 2% bovine serum albumin (BSA). Incuber 10 minutes dans la chambre froide sur agitateur.
  3. Centrifuger 700g à 4ºC pendant 1 minute. Décanter surnageant et jeter.
  4. Reprendre les billes dans 1 ml ChIP Dilution Buffer et repartir 200 l dans 5 tubes.
  5. Aux billes (ajouter 0,8ml ChIP Dilution Buffer dans les 4 premiers tubes) :
    1. Ajouter anti-S (6 ug) pour Tube Expérimental (ES) S
    2. Ajouter anti-P (4 ug) Tube Expérimental (EP) P
    3. Ajouter anti-histone H3 (4 ug) au Tube témoin Positif (P)
    4. Ajouter anti-IgG (4 ug) au Tube témoin Négatif avec anticorps (N)
    5. Ajouter 1 ml ChIP Dilution buffer + 2% BSA pour Tube Blanc sans anticorps (B)
  6. Incuber avec agitation au frais pendant 2h.
  7. Centrifuger 700g à 4ºC pendant 1 minute. Décanter surnageants et (garder la prochaine fois !).
  8. Rincer avec 1 ml de ChIP dilution buffer + 2% BSA. Laisser 10-30 minutes à 4ºC en agitant.
  9. Centrifuger 700g à 4ºC pendant 1 minute. Décanter surnageants et jeter. Ajouter 1 ml de ChIP Dilution buffer.
  10. Centrifuger 700g à 4ºC pendant 1 minute. Décanter surnageants et jeter. Ajouter 60 l de ChIP Dilution buffer à chaque tube et réserver ces billes preparées au frigo.
  11. Sonication de 5 aliquots de 400 ul de chromatine dans l’eau glacée en récipient confiture :
    1. 6 x 20 secondes, espacées de 20 secondes, sur sonicateur Branson ou equivalent
    2. 1er sonication 30W, 2e et ensuite à 40W
    3. ne pas mousser ! sinon prendre un autre aliquot.
  12. Transvaser dans Eppendorfs 1,5ml et microcentrifuger 10 minutes 4ºC 13K g.
  13. Prendre surnageant et repartir par 200ul en 1 tube (B) et 400ul en 4 autres tubes (ES, EP, P, N) de 1,5 ml peu adhérents.
  14. [Prélever 80 ul de chaque tube et les pooler (400 ul) = Total Input DNA, 5% du chromatine soniqué au départ.]
  15. Ajouter à chaque tube de chromatine :
    1. 1100 ul (tube B) ou 900 ul (tube ES, EP, P, N) de tampon ChIP Dilution Buffer
    2. 50 ul inhibiteurs de proteases Roche 25x repris dans de l’eau
    3. 100 ul Protein A – Agarose / Salmon Sperm DNA (ou protein A-agarose maison)
    4. Incuber 1h à 4ºC avec agitation. Ceci débarrasse des protéines en gros excès « pre-clear ».
  16. Spin 700g 1 minute pour récupérer l’agarose au fond, reprendre surnageant dans de nouveaux tubes étiquettés et toujours sur glace.
  17. Le résidu de chromatine brute (200 ul) est utilisé pour vérification de la sonication sur gel agarose 1%.
    1. Ajouter 30 ul NaCL 5M et incuber en bloc chauffant à 99ºC 15’
    2. Refroidir et ajouter 2 ul RNase (DNase-free) et 3 ul protéinase K (stock 10-20mg/ml), incuber 37ºC 30’ puis 45ºC pendant 15’. (Sinon on voit un « smear » à cause de l’ARN et n’en parlons pas des protéines)
    3. Microcentrifugation vitesse maxi.
    4. Extraction phénol-chloroforme du surnageant
    5. Pour 250 ul de chromatine, ajouter 12 ul NaCl 5M et 0,5 ml EtOH 100% froid. Précipiter 30’ -30ºC ou 10’ sur carboglace.
    6. Centrifugation à 4ºC 30’ vitesse maxi.
    7. Rincer culot avec 500 ul EtOH 70% froid, décanter aussitôt (ou spin) et sécher culot.
    8. Reprendre dans 20 ul. Doser 1 ul sur spectrophotomètre ou Nanodrop.
    9. Déposer 5-10 ul (2-3 ug chromatine) sur gel en face d’un marqueur 100bp+. Prendre une jolie photo que tu publierais.
  18. Si la sonication s’est bien passée, ajouter aux surnageants appropriés, les billes Protein A-agarose-anticorps.
  19. Mettre dans la chambre froide sur agitateur la nuit.

—

  1. Préparer tampon d’élution frais à conserver pour la fin :
    1. 1% SDS final (pour 2 ml = 0,2 ml de 10% SDS)
    2. 0,1 M NaHCO3 final (pour 2 ml = 0,2 ml de 1M)
  2. Centrifuger 700g à 4ºC pendant 1 minute.
  3. Décanter surnageant – conserver de coté
  4. Sur les billes, ajouter :
    1. 1 ml de Low Salt Immune Complex Wash Buffer
    2. 10 minutes sous agitation 4ºC
    3. Centrifuger 700g à 4ºC pendant 1 minute. Décanter surnageant – conserver de coté
  5. Sur les billes, ajouter :
    1. 1 ml de High Salt Immune Complex Wash Buffer
    2. 10 minutes sous agitation 4ºC
    3. Centrifuger 700g à 4ºC pendant 1 minute. Décanter surnageant – conserver de coté
  6. Sur les billes, ajouter :
    1. 1 ml de LiCl Immune Complex Wash Buffer
    2. 10 minutes sous agitation 4ºC
    3. Centrifuger 700g à 4ºC pendant 1 minute. Décanter surnageant – conserver de coté
  7. Sur les billes, ajouter :
    1. 1 ml de TE Buffer
    2. 5 minutes sous agitation température ambiante
    3. Centrifuger 700g à 4ºC pendant 1 minute. Décanter surnageant – conserver de coté
  8. Sur les billes, ajouter encore une fois:
    1. 1 ml de TE Buffer
    2. 5 minutes sous agitation température ambiante
    3. Centrifuger 700g à 4ºC pendant 1 minute. Décanter surnageant – conserver de coté
  9. Sur les 4 tubes de billes, ajouter maintenant 250 ul du tampon d’élution.
    1. Vortexer brièvement, puis agiter 15 minutes température ambiante.
    2. Centrifuger 700g à 4ºC pendant 1 minute. Décanter surnageant – C’EST LA CHROMATINE – et mettre dans un tube 2 ml.
  10. Sur les billes, ajouter encore 250 ul tampon d’élution.
    1. Vortexer brièvement, puis agiter 15 minutes température ambiante.
    2. Centrifuger 700g à 4ºC pendant 1 minute.
  11. Décanter surnageant et ajouter aux surnageants précédents (500 ul total).
  12. Ajouter à chaque tube ainsi qu’au tube de 400 ul Total Input DNA (qsp 500 ul avec de l’eau): 20 ul NaCl 5M
    1. Incuber 4-6 h à 65-68ºC dans le bain sec (faire l’étape d’après en même temps)
  13. (Possibilité de les conserver à -20ºC)
  14. Ajouter aux 6 tubes (ES, EP, P, N, B et TID) :
    1. 10 ul EDTA 0,5M
    2. 20 ul Tris-HCl pH 6,5 1M
    3. 2 ul Protéinase K à 10 mg/ml
    4. Incubation 1h à 45ºC
  15. Y ajouter 1 ul RNAse pour 30 minutes à 37ºC.
  16. Protocol Qiaquick :
    1. Ajouter tampon PBI du kit qsp 2 ml et mélanger. Vérifier couleur jaune, sinon rectifier pH avec 10 ul acétate de sodium 3M, pH 5,0.
    2. Placer les colonnes Qiaquick marquées sur leurs tubes de collection. Y ajouter les echantillons jaunes E, P, N, B, TID par volume de 0,67 ml chaque.
    3. Passer ce volume en centrifugeant à la vitesse maxi pour 30 secondes. Vider le liquide passé et répéter 2x pour mettre tout l’ADN sur membrane.
    4. Laver la colonne : 750 ul tampon PE (avec son EtOH ajouté)
    5. Centrifuger 1 minute, vider le liquide passé.
    6. Remettre colonnes sur les mêmes tubes de collection, centrifuger à sec 1 minute. Placer sur des tubes propres de 1,5 ml.
    7. Eluer dans 30 ul de tampon EB [ou de l’eau ultrapure pH 7-8 si des ligations ensuivent].
  17. Garder à -30ºC ou passer tout de suite à la PCR.

 

Notes

  1. Depend enormement sur la sonication – aucune mousse ni rechauffement toleree !
  2. Depend aussi de la fixation – a varier entre 10 et 15 minutes total (y compris centrifugation) en faisant moins pour les histones et plus pour les petits facteurs de transcription
  3. Le labo de Peggy Farnham a UC Davis a quelques indications
  4. Les sites de Upstate et de Chemicon a quelques astuces aussi

References

Relevant papers and books

  1. O’Geen H, Nicolet CM, Blahnik K, Green R, and Farnham PJ. Comparison of sample preparation methods for ChIP-chip assays. Biotechniques. 2006 Nov;41(5):577-80. DOI:10.2144/000112268 | PubMed ID:17140114 | HubMed [OGeen2006]
  2. Elnitski L, Jin VX, Farnham PJ, and Jones SJ. Locating mammalian transcription factor binding sites: a survey of computational and experimental techniques. Genome Res. 2006 Dec;16(12):1455-64. DOI:10.1101/gr.4140006 | PubMed ID:17053094 | HubMed [Elnitski2006]

All Medline abstracts: PubMed | HubMed

 

Site-directed mutagenesis/Multi site-PDF

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Materials

  • Template plasmid purified from a dam+ E. coli strain (not JM110 or SCS110)
  • Mutagenesis primers (one permutation, each on the same strand)
  • PfuTurbo DNA polymerase (non strand-displacing) and associated reaction buffer
  • Taq ligase
  • dNTPs
  • ATP
  • PCR Thermocycler
  • Dpn I
  • Competent cells
  • unphosphorylated primers (1 for each mutation)

Mutagenesis PCR mix

25μL total reaction volume:

  • 2.5 μL of 10X Taq ligase buffer (need the NAD for Taq ligase)
  • 0.5 μL 100mM ATP
  • 1 μL 25mM each dNTP
  • X μL (50-100 ng) of dsDNA template
  • X μL of each oligonucleotide primer
    • For 1-3 primers, add 100 ng each primer. For 4-5 primers, add 50 ng each primer.
    • If primers are greater than 20% different in length, scale the amount of primer added so that primer is added in approximately equimolar amounts. See Stratagene QuikChange Multi Site-Directed Mutagenesis manual for details.
    • Austin uses 0.1μL of 40μM primer (each one).
  • 1 μL of dNTP mix (100mM total dNTP mix with 25 mM each individual dNTP)
  • ddH2O to a final volume of 22 μL

Then add

  • 1 μL of PfuTurbo DNA polymerase (2.5 U/μL)
  • 1 μL of Taq Ligase
  • 1 μL of T4 PNK

Procedure

This procedure is primarily derived from the Stratagene QuikChange Multi Site-Directed Mutagenesis manual with some modifications based on past experience.

  1. Design mutagenesis primers.
    • The primer should be designed so that the desired mutation occurs at the exact center of the primer with 10-15 bp of matching sequence on each side.
    • Primers should be 25-45bp in length with a melting temp of >=75°C. Stratagene recommends not using primers greater than 45 bp in order to avoid formation of secondary structure. Primers should have comparable melting temperatures.
    • See the Stratagene manual for more detailed information. In particular, adhere to their formula for calculating the melting temperature of your primers and design your primers to have a melting temperature >=75°C.
    • Primers should have at least 40% GC content and terminate in one or more C or G bases at the 3′ end.
    • PAGE purification of primers may improve mutagenesis efficiency. See here for links to information on oligo purification.
    • See designing primers for general advice on primer design.
  2. Purify template plasmid from a dam+ E. coli strain via miniprep.
  3. Set up mutagenesis PCR mix as described above.
  4. Run Reaction
    1. 37°C for 30 min (T4 PNK step)
    2. 95°C for 3 min
    3. 95°C for 1 min
    4. 55°C for 1 min
    5. 65°C for 2 min/kb of plasmid length minimum (is optimal temperature for Taq ligase)
    6. Run reaction for 30 cycles.
    • Stratagene recommends using a PCR machine with heated lid or overlaying the reaction with mineral oil.
  5. Cool the reaction to <=37°C
  6. Add 1μL DpnI restriction enzyme to the PCR tube directly. (Purification is not necessary at this stage).
  7. Incubate 6 hours at 37°C (even though the Stratagene manual only recommends 1 hr).
  8. Purify PCR product (not necessary, Austin transforms 3 μl directly).
    • I typically do this step using a QIAgen PCR Purification kit but any purification which removes the salts, dNTPs, oligos and proteins from the PCR should be fine.
  9. Transform purified DNA into highly competent cells.
  10. Screen the transformants for the desired mutation using colony PCR, restriction digest or sequencing as appropriate.

Notes

  • Stratagene does not recommend this protocol for insertions or deletions.
  • Apparently there are no primer spacing-dependent effects on mutagenesis efficiency (primers can be adjacent or far apart).
  • You can also phosphorylate the primers separately from the rest of the mutagenesis reaction

DNA Hybridization-PDF

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Overview

Hybridization of complimentary ssDNA oligonucleotides to create dsDNA linker.

Materials

  • 5 μL 2mM sense oligonucleotide
  • 5 μL 2mM antisense oligonucleotide

NOTE: If X = nmol of lyophilized oligonucleotide, add X/2 μL water to obtain 2mM final concentration.

Procedure

  1. Combine 5 μL sense oligonucleotide and 5 μL antisense oligonucleotide. Vortex and spin.
  2. Double seal the tube airtight with parafilm and press firmly in a weighted holder.
  3. Boil 800 mL water in a 1L beaker.
  4. Wait 1 min.
  5. Place the weighted holder with sealed oligonucleotides into the beaker.
  6. Incubate on a bench overnight or until water is about room temperature (~4 hr). At the end of this process, the complimentary ssDNAs are hybridized into a single dsDNA linker with sticky ends at 1mM final concentration.
  7. Perform two 1/100 dilutions (label all tubes clearly):
    • 2 μL 1mM linker + 198 μL H2O = 200 μL 10μM linker
    • 2 μL 10μM linker + 198 μL H2O = 200 μL 100nM linker
  8. Store 1mM, 10μM, and 100nM stocks at -40 °C in latest “Primer” box. Keep an aliquot of the 100nM linker in your own box for use in constructions.

 

DNA labeling-PDF

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Materials

  • Sodium Bicarbonate (pH 8.3, but acceptable range is 8.0 and 8.8): Na(CO3)2
  • Sodium Borate (pH 8.3)
  • Acetonitrile: ACN (HPLC grade)
  • Sodium Acetate (pH 5.2)
  • TE: 10 mM TRIS (pH 8.0) + 1 mM EDTA

Procedure

  1. Suspend DNA from IDT in 100 uL of 50 mM TEAA + 5% ACN
    1. To help dissolve, raise the temperature for 10 mins if necessary
  2. Spin down, the pellet will form powdery-crystal residue which is the particulate from the sample that is not DNA. Keep it just in case, since there might be some DNA in that pellet.
  3. HPLC to separate lengths (use Reverse Phase/Size Exclusion column)
    1. There will be one large fraction which is the fraction of interest
  4. Dry the elution/fraction in SpeedVac. After drying, suspend the dried fraction in MilliQ water. Check absorbance to determine the concentration in UV/Vis Spectrometer.
  5. For labeling the complement strand with ATTO 647, resuspend 100 mM (pH 8.3) Sodium Bicarbonate. For labeling template strand with Cy3B, use 100 mM (pH 8.3) Sodium Borate. Determine the concentration of dyes in DMSO using absorbance.

Add dye in excess (10x recommended). Dissolve the dye in anhydrous DMSO/DMF, prepared immediately before labeling reaction. Leave for 24-48 hrs at room temperature.

  1. Ethanol Precipitate. Add 150 mM Sodium Acetate and 2x volume of ethanol
    1. For a 100 uL solution, add 5 uL of Sodium Acetate and 250 uL of ethanol

Place in (-80 or -20) freezer at least 1 hr, overnight is fine. Remove supernatant and make sure all ethanol has evaporated off. #Resuspend in 50 mM TEAA + 5% ACN

  1. HPLC and collect unlabeled peak, labeled peak
  2. Dry in SpeedVacFor long-term storage, leave in TE. Add BSA to prevent sticking if you expect very long-term storage.

Stitching Genes by PCR-PDF

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Stitching Genes by PCR-PDF

Introduction

This method allows you to “stitch” genes or coding sequences together when there are no convenient restriction sites at the junction point. It is especially useful when the target gene is flanked by other genes that can’t be disrupted. The technique is by no means new, but this should keep people from asking me how to do it.

Protocol

Refer to this figure:

  • Step 1:

Scan your sequence and find restriction sites anywhere to the left and right of the gene (the left one can be in your gene). Make sure they are unique and they are not contained in the fragment you want to append to your gene. In the figure, they are marked “A” and “B”.

  • Step 2:

Design 4 primers. “A forward” primes toward your gene and anneals to the left of (or on) the restriction site “A”; “B reverse” primes back toward your gene and anneals past (or on) restriction site “B”; “stitch forward” anneals to amplify your appendage and has a tail that matches (anneals to) the left side of the fusion point (green in this case); ‘stitch reverse” reverse primes your appendage and has a tail that encodes the region just to the right of the fusion point. The dashed lines show where the primers match.

  • Step 3:

PCR your appendage. If it is small (like a His tag), you can just have the two primers form a cassette when annealed. In this case, the appendage is a bit bigger, so PCR is used to make the product from another template that contains the appendage. Only do about 20 cycles, you don’t need a lot and this will reduce errors.

  • Step 4:

Set up a PCR with a microliter of the “appendage” PCR reaction in step 3 and a microliter of a miniprep containing your plasmid with your gene of interest. Use primers “stitch forward” and “B reverse”. You will end up with the product shown. I usually gel purify this product for the next step. Again, you don’t need a lot, use about 20 cycles.

  • Step 5:

Set up a primer extention reaction with the gel purified product from step 4 and your plasmid. Use about half of the gel-purified product from step 4. Set the extension time long enough so the polymerase will extend to the left restriction stite “A”. Both strands of the PCR product will act as primers and you will end up with a heterogeneous population of single-stranded products. The products that you need are the “bottom strand” extensions that primed back toward your site “A”. About 20 cycles will do.

AFTER THIS STEP, add the restriction enzyme Dpn I to the PCR reaction and incubate for 1-2 hours at 37 degrees. You are eliminating the added plasmid. This is important.

  • Step 6:

Without cleaning the reaction from step 5, set up a new PCR that contains half of the PCR/degradation reaction from step 5. This is your template. Remember, you haven’t cleaned the reaction, so you only need to add PCR buffer for 50% of the volume. Use the normal primer and dNTP concentrations and add fresh polymerase. Use primers “A forward” and “B reverse”. 20 cycles (you have a lot of template). Remember to increase the extention time so the polymerase can read the whole desired product.

  • Step 7:

Clean the reaction, digest with restriction enzymes “A” and “B”. Also cut your plasmid with them. Ligate the fragment and you’re done.

 

Example gel:

An example of 3 different stitching reactions at steps 4, 5, and 6. I usually only check these by gel. Notice after step 5, the product seems to be greatly reduced, but there is a hetergeneous population of fragments that don’t resolve as sharp bands that are larger than the starting primers (PCR product of step 4). The final PCR amplifies the desired product so it can be purified.

Notes

You can switch the first stitching reaction so that the left side is added first. I set mine up so the shorter of the two arms is added first. The primer extension that adds the longer arm is reading off of the plasmid and will have very few errors. In doing this, you keep the larger amplifications to short segments and reduce your chanced of getting a mutation in the final product. Keeping the cycles reduced also helps because you don’t deplete the dNTPs and starve your enzyme.

Since your going through the trouble of building the fusion, incorporate some unique, user-friendly restriction sites in your stitched product so modifying the gene in the future is a lot easier.

PhoenIX Maxiprep Kit-PDF

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When using a new kit

  1. Assemble the cardboard column rack.
  2. Note that the resuspension, lysis, and neutralization buffer bottles are VERY hard to open.

Procedure

The steps below have been changed to accommodate our tubes and centrifuge.

  1. Place the PhoenIX™ Maxi column in the assembled column rack.
  2. Place a beaker underneath to collect flow through.
  3. Add 30 ml of equilibration buffer (gray cap label) to the surface of the column and allow the liquid to drain by gravity flow.
    • Note: It will take 15-20 minutes for the column to drain completely.
  4. Pellet 200 ml of bacterial culture by centrifugation at 3,000 x g for 20 minutes at 4°C.
  5. Remove all traces of liquid medium from the bacterial cell pellet by pouring.
    • Trace media can affect subsequent steps.
  6. Add 10 ml of RNase A-containing cell resuspension buffer (yellow cap label) to the cell pellet and vortex until completely resuspended.
    • There is some resuspension buffer with RNase A added stored at 4°C in 32-306. To make more, you’ll need to add RNase A to more resuspension buffer. The resuspension buffer (without RNase A) is in the box and the RNase A is stored at -20°C on the door.
  7. Transfer resuspended cells to 50 mL conical tube.
  8. Add 10 ml of Lysis Buffer (blue cap label) and securely cap the tube.
  9. Mix thoroughly by inverting until the lysate appears to be homogeneous (5-6 inversions). DO NOT VORTEX.
  10. Incubate 5 minutes at room temperature.
    • Note: Do not incubate for longer than 5 minutes or plasmid DNA might become irreversibly denatured.
  11. Add 10 ml of neutralization buffer (green cap label).
  12. Securely cap the tube and mix immediately by multiple inversions until a homogeneous suspension containing no viscous matter is obtained. DO NOT VORTEX.
    • Note: If preparing several samples at once, thoroughly mix each sample immediately after the addition of the neutralization buffer before adding the buffer to the next tube.
  13. Centrifuge at 9,000 x g for 20 mins at room temperature.
    • Note: The supernatant must at room temperature (18 – 25°C) prior to loading on the column.
  14. Verify that the qquilibration buffer has been collected in the beaker.
  15. Discard the flow-through and replace the container.
  16. Use a pipette to remove the cleared lysate supernatant from the centrifuged sample and add to the top of the equilibrated column.
    • Note: Do not pour lysate directly onto the column. Use a pipette to ensure that precipitate particles do not enter the column and cause clogging.
  17. Allow the lysate to drain by gravity flow (10-15 minutes).
  18. Discard the flowthrough and replace the empty container.
  19. Add 30 ml of column wash buffer (orange cap label) to the top of the column and allow the liquid to drain by gravity flow (10 minutes).
  20. Add 30 ml of column wash buffer (orange cap label) to the top of the column and allow the liquid to drain by gravity flow (10 minutes).
  21. Discard the flow-through.
  22. Replace the waste collection container with a 50 mL conical tube.
  23. Add 15 ml of elution buffer (pink cap label) to the top of the column.
  24. Allow the eluate to drain by gravity flow (5-10 minutes) into the centrifuge tube.
  25. Add 10.5 ml of room temperature isopropanol to the eluted plasmid DNA in the centrifuge tube.
  26. Mix and centrifuge at 9,000 x g for 40 minutes at 4°C.
  27. Pour out the supernatant taking care not to disturb the DNA pellet.
  28. Add 5 ml of room temperature 70% ethanol and wash the pellet.
  29. Centrifuge at 9,000 x g for 10 minutes at 4°C.
  30. Completely remove ALL of the supernatant from the pellet with a pipette.
  31. Air-dry the pellet for 10 minutes.
    • Note: Drying with a vacuum chamber is not recommended because over-dried DNA may be difficult to completely resuspend.
  32. Dissolve the plasmid DNA in 500 μl of water.
  33. Move to smaller tube.
  34. Take a spectrophotometer reading to assess concentration.

Notes

  • These spin steps may not be hard enough. Most of the purification steps are supposed to be at 12,000 x g.
  • You’re not supposed to let the column stand between steps.
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