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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.

RAD-seq-PDF

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RAD-seq-PDF

Overview

This is a protocol for generating RAD libraries for Illumina sequencing. With this technique, 96 samples can be multiplexed into one sequencing library, and only tags adjacent to PstI sites are sequenced. This is a cheap way to both mine and genotype large numbers of SNPs.

Materials

Reagents

  • Quant-iT Picogreen kit (Invitrogen)
  • Qiagen gel purification kit
  • Qiagen PCR cleanup kit
  • From New England Biolabs:
    • PstI-HF, 20,000 U/mL
    • MspI, 20,000 U/mL
    • T4 DNA ligase, 2,000,000 U/mL
    • ATP
  • KAPA HiFi Library Amplification Kit, without primers. In the past we used Phusion High Fidelity PCR master mix from NEB, but KAPA is supposed to be better.
  • 100 bp DNA ladder
  • Gel loading dye that does NOT have bromophenol blue. Currently we use a home-made loading dye with Orange G, glycerol, and TE. NEB also makes an orange loading dye that works well. I have also used Promega GoTaq Green PCR buffer as a loading dye.
  • You will also need a black microtiter plate for the Picogreen assay.

Note: Although MspI and PstI are not completely inactivated by heat, the adapters are designed such that the restriction cut sites are not recreated by the ligation reaction. The final ligated products will therefore not be re-digested.

Note #2: To mine additional genomic positions, additional libraries can be made in which PstI-HF is replaced with NsiI-HF. These two enzymes have the same overhang, and therefore the same adapters can be used. The nucleotides flanking the overhang are different between these two enzymes, and therefore they cut at different sites. With NsiI, if using adapters designed for PstI, some of the adapters will recreate the restriction cut site, and so care must be taken to deactivate the enzyme with the heat inactivation step.

Note #3: To mine a much smaller number of genomic positions at much greater read depth, PstI-HF can be replaced with SbfI.

Oligonucleotides

PstI adapters

This is the most expensive part of the protocol other than the sequencing itself, since 192 oligonucleotides must be ordered.

Adapter 1 top: 5'GATCTACACTCTTTCCCTACACGACGCTCTTCCGATCTxxxxTGCA3'

Adapter 1 bottom: 5'yyyyAGATCGGAAGAGCGTCGTGTAGGGAAAGAGTGTAGATC3'

Where xxxx and yyyy are the barcode and its reverse complement, respectively.

Barcodes and oligo sequences are from Pat Brown’s lab (Thurber et al. 2013).

Media:PstI-barcodes.txt

More recently (April 2015) we designed new PstI adapters ranging from six to ten nucleotides long using Deena Bioinformatics.

Other oligos

MspI adapters:

  • A2top: 5'CGCTCAGGCATCACTCGATTCCTATCAGAACAA3'
  • A2bot: 5'CAAGCAGAAGACGGCATACGAGATAGGAATCGAGTGATGCCTGAG3'

Note that the MspI adapter sequences were changed in September 2017 to be compatible with the HiSeq 4000.

Illumina PCR primers:

  • PCR1: 5'AATGATACGGCGACCACCGAGATCTACACTCTTTCCCTACACGACGCTCTTCCGATCT3'
  • PCR2: 5'CAAGCAGAAGACGGCATACGA3'

Equipment

  • Nanodrop spectrophotometer
  • BioTek Synergy plate reader (for reading fluorescence)
  • Ordinary PCR machine
  • Agarose gel rig
  • UV transilluminator for gel excision
  • Bioanalyzer
  • real-time PCR machine (we just pay the core facility to do that part)

Procedure

Adapter prep

Top and bottom strands of adapters need to be annealed 1X Annealing Buffer, which is 10 mM Tris, 50 mM NaCl.

The annealing program is:

  • 95°C 5 minutes
  • Ramp down -0.1°C every 2 seconds (or -1°C every 20 seconds) to 25°C.

My protocol:

  • We have a stock plate of PstI adapters that are at 1 μM. I took a bottle of autoclaved 1X Annealing Buffer, added 45 μl to each well of a 96-well plate, then transferred 5 μl from the 1 μM plate to make a 0.1 μM working stock.
  • MspI adapters are ordered like normal oligos, and I have 100 μM concentrated stocks in TE. To make a 10 μM stock:
    • 20 μl A2top, 100 μM
    • 20 μl A2bot, 100 μM
    • 20 μl 500 mM NaCl
    • 2 μl 1M Tris
    • 138 μl nuclease-free water
    • Mix well, add 100 μl to each of two PCR tubes, and run them on the annealing program (“Adapt” on the PCR machine).

DNA quantification and dilution

Dilution to ≤200 ng/μL (usually, just a 10x dilution)

  1. Picogreen can accurately detect very small quantities of DNA, but is not accurate over 1 ng/μL. In the Picogreen assay, DNA is diluted 200X in solution, so DNA stock solution of up to 200 ng/μL can be quantified.
  2. Our DNA extraction protocol yields concentrations of up to 2 μg/μL (2000 ng/μL). Therefore, we need to dilute 10X to ensure that we are in the range that can be measured with Picogreen.
  3. Take a 96 well PCR plate, and add 18 μL 10 mM Tris or TE to 88 wells (11 columns).
  4. Using a spreadsheet that records which sample goes in which well, add 2 μL of DNA extraction to the 18 μL of buffer. You can quantify 88 samples on one plate.

Quantify your ≤200 ng/μL dilution plate using Picogreen:

  1. Take the tube of bright orange Picogreen reagent out ahead of time to thaw. Wrap it in aluminum foil to protect it from light. It is in DMSO instead of water, so it takes a long time to thaw and will immediately freeze solid if you put it on ice.
  2. The Quant-iT Picogreen kit comes with a lambda DNA standard at 100 μg/mL. Dilute some of the 20X TE that comes with the kit to 1X TE, and use it to make a 2 μg/mL dilution of the lambda DNA. (1:50 dilution.) (Alternatively, I have made a 8 μg/mL stock that can be diluted 4X at the time the standard column is set up.)
  3. For one plate (88 samples, 8 standards) make up 20 mL of 1X TE. (1 mL of the TE that comes with the kit, plus 19 mL sterilized filtered water.)
  4. The plate you need for the assay is a black, flat-well plastic plate. (Corning makes these.)
  5. Set up a standard curve in column 1 (or column 12, doesn’t matter). Pipette 100 ul of TE into wells B-H. Add 100 ul of your 2 ug/ml lambda standard each to well A and B. Pipette well B up and down to mix, then transfer 100 ul to well C. Pipette well C up and down to mix, then transfer 100 ul to well D. Continue through well G, and leave well H as a blank. (After mixing well G, you will simply throw out 100 ul.)
  6. Add 99 μL TE to the other 88 (or however many samples you are doing) wells . Add 1 ul of ≤200 ng/μL sample DNA (from the 10X dilution plate) to each well.
  7. Add 50 μL of Quant-iT reagent to 10 mL of 1X TE. This solution needs to be used within a few hours, even if it is protected from light. Add 100 μL of the solution to each well (both sample, standard, and blank).
  8. Picogreen bonded to dsDNA has an excitation maximum at 480 nm and emission maximum at 520 nm. The plate readers in IGB (BioTek Synergy HT) probably already have a picogreen program on them.

If you need to re-make the picogreen program, use the screenshots below:

  1. Read fluorescence intensity on the plate reader, and export it to Microsoft Excel.
  2. Make a scatterplot of fluorescence intensity of the standard vs. the standard concentration. Given that the samples were diluted 2000X, the standard concentration is multiplied by 200:
    1. Well A 2000 ng/μL
    2. Well B 1000
    3. Well C 500
    4. Well D 250
    5. Well E 125
    6. Well F 62.5
    7. Well G 31.25
    8. Well H 0
  3. In Excel, fit a trendline to the scatterplot and display the equation on the chart. Use this equation to estimate the concentration of the samples.

In most cases, the concentration estimate via Picogreen should be lower than the concentration estimate via Nanodrop. This is because Nanodrop measures DNA + RNA, whereas Picogreen only measures DNA.

Based on the Picogreen concentration estimates, dilute the DNA to 50 ng/μL in 10 mM Tris (and 0.1 mM EDTA, optional).

Notes for samples of concentration lower than 50 ng/μL:

  • If you have a lot of samples that are 30-50 ng/μL, you can dilute all samples for your library to 30 ng/μL or 40 ng/μL instead of 50. The amount of adapter that you add at the ligation step (see below) should be reduced proportionately.
  • For samples in the 10-50 ng/μL range, a cheap and efficient way to concentrate them is by isopropanol precipitation:
    • Combine 200 μL DNA sample, 20 μL 3M sodium acetate, and 200 μL isopropanol.
    • Mix well by inversion. Place in the freezer for at least an hour.
    • Spin down 10 minutes in the centrifuge.
    • Pour off the liquid, taking care to keep the pellet.
    • Add 200 μL 70% ethanol to rinse. Invert a few times.
    • Spin down 1 minute, then pour off the ethanol, again being careful not to lose the pellet.
    • Allow to dry on the lab bench.
    • Resuspend the DNA in 20 μL TE.
    • Requantify with Picogreen, then dilute to 50 ng/μL.

Restriction digestion and ligation

Restriction digestion master mix:

Ingredient For one sample For one plate
50 ng/ul DNA 5 ul –
10X NEBuffer 4 (or CutSmart) 1.5 ul 165 ul
PstI-HF, 20,000 U/mL 0.25 ul 27.5 ul
MspI, 20,000 U/mL 0.25 ul 27.5 ul
Nuclease-free water 8 ul 880 ul

(I have also used DNA at a concentration of 100 ng/ul because that was what Keck wanted for GoldenGate, so then I used 2.5 ul DNA and 10.5 ul water.)

Do this in a 96-well plate. Pipette the DNA into the wells and then add 10 ul of master mix to everything. Pick one well that will not have DNA in it. This will be an important control later on to demonstrate that this library was not contaminated with another library (which will have a different empty well).

Run the Digest program on the PCR machine: 3 hours at 37°C, then 20 minutes at 80°C.

Using a multichannel pipette, add 1.5 μL of 0.1 μM PstI adapters to their corresponding wells on the digestion plate. (Do add the adapter corresponding to the well that has no DNA in it.)

Ligation master mix, keep on ice until use:

Ingredient For one sample For one plate
10X Ligase buffer with ATP 1 ul 110 ul
10 μM MspI adapter 0.5 ul 55 ul
10 mM ATP 1.5 ul 165 ul
T4 Ligase, 2M U/mL 0.1 ul 11 ul
Nuclease-free water 5.4 ul 594 ul

Add 8.5 μL of ligation master mix to each well of the digestion plate.

Run on the “ligate” program on the PCR machine: 2 hours at 25°C, 20 minutes at 65°C.

Cleanup and amplification

  • Using a multichannel pipette and a PCR 8-well strip tube, pool all the columns together, adding 5 μL from each well of the plate to the wells on the strip tube.
  • Pipette the 60 μL out of each well on the strip tube into one 1.5 mL tube. Mix well so that all samples are combined evenly. Freeze or keep on ice.
  • Pour a 2% agarose gel with ethidium bromide. Make it nice and deep; my recipe is 4 g agarose, 200 mL 1X TAE, and 10 μL ethidium bromide solution. Use a wide-toothed comb.
  • Take 50 μL (or more depending on your well volume) of your pooled library and combine it with a loading dye that does not have bromophenol blue. I use 10 μL of a 6X loading dye containing 30% glycerol, 0.2% orange G, 10 mM Tris, and 1 mM EDTA.
  • I recommend cleaning out your gel rig and putting in fresh TAE, since you especially want to avoid any contamination from other Illumina libraries.
  • Run your ~60 μL of library plus loading dye on the gel. The lane with the library should have a lane of 100 bp ladder on either side of it. You can put multiple libraries on one gel, but leave several empty lanes between them.
  • The gel doesn’t need to be run very long. I would go 20 minutes at 100 V, or until the ladder bands below 500 bp are distinguishable.
  • The library should look like a smear. There may be some undigested DNA (a band in the 10’s of kb) but that is okay as long as most of the DNA is digested. There may also be a thick band of RNA and leftover adapter below 100 bp. (I have found that RNAse treatment removed most of that band but did not appear to improve DNA digestion.)
  • Using a clean razor blade for each library, cut out the smear between 200 bp and 500 bp (if using SbfI, instead cut from 200 bp to 1000 bp). There should definitely be DNA visible in this range.

Three pooled ligations ready for gel extraction, with GoTaq Green loading dye
Pooled ligations when NEB orange dye is used

  • Use the Qiagen gel extraction kit to purify the DNA out of this gel slice. Do include the optional steps of washing with QG after binding the DNA to the column, as well as letting the column sit in PE for 2-5 minutes before spinning (Phusion can handle contamination from agarose/salts, but KAPA HiFi cannot). Elute in the lower volume (30 μL EB).
  • Run the Illumina PCR:
    • 3 μL gel-extracted library
    • 2 μL 10 μM forward + reverse Illumina primers (PCR1 and PCR2)
    • 25 μL 2X Kapa Hi-Fi Master mix
    • 20 μL nuclease-free water
  • PCR program:
    • 98°C 30 seconds
    • 15 cycles of 98°C 10 seconds, 65°C 30 seconds, 72°C 30 seconds
    • 72°C 5 minutes
  • The first time you do this protocol, run 5 μL of the PCR product out on a 2% agarose gel. Look to see whether there is primer-dimer visible. If there is no primer-dimer visible, use the Qiagen PCR cleanup kit to purify the remaining 45 μL of PCR product.

Nine libraries post-PCR, with GoTaq Green loading dye. A second gel (with space in between libraries) will be needed for extraction of the libraries, to eliminate the primer-dimer.
Amplified libraries, run with NEB orange dye, ready for gel extraction.

  • If there is primer-dimer visible, run the remaining 45 μL of PCR product on a 2% agarose gel and extract the library (as was done pre-PCR). Follow the instructions in the Qiagen gel extraction kit as specified for sequencing. (After binding DNA to the column, do a wash with QG. When rinsing with PE, let sit for 2-5 minutes before spinning.) Typically I get primer-dimer, so I just do this extraction and skip the previous gel to test for primer-dimer.

Quality control

  • Quantify the purified PCR product using the Picogreen protocol as above. Expected concentrations are in the 10’s of ng/μL.
  • Make 4 μL of a 1 ng/μL dilution of the library, and submit it to the Functional Genomics center to run on a High Sensitivity DNA chip on the Bioanalyzer. There should be a smooth curve from around 200 to 500 bp. Any sharp peaks could indicate that the enzymes were cutting in a repetitive region of the genome, in which case it is best to choose different enzymes. Use the Bioanalyzer software to calculate the average fragment size.
    • If there is primer-dimer remaining in the library, it will be visible as a sharp peak at a lower molecular weight than the broad peak for the library. (The library pictured below does not have primer-dimer.)

  • Calculate the concentration of the PCR product in nM. Keck supplies a worksheet for this calculation. If [math]\displaystyle{ x }[/math] is the concentration in ng/μL, [math]\displaystyle{ y }[/math] is the average size in base pairs, and [math]\displaystyle{ z }[/math] is the concentration in nM, then [math]\displaystyle{ z = \frac{10^6*x}{649y} }[/math].
  • Dilute the purified PCR product to 10 nM in EB (10 mM Tris).
  • Give 20 μL of 10 nM library to the core facility (Keck). They will use real-time PCR to confirm a concentration of 10 nM. Using Illumina Hi-Seq, do one lane of 100 bp single-end reads.

Bioinformatics

Given the genome duplication present in Miscanthus, we have found that the UNEAK pipeline works well.

I have written an some R functions for importing the output of the UNEAK pipeline into adegenet or more generally into a numeric (0 and 2 for homozygote, 1 for heterozygote) matrix format in R.

I have also created TagDigger for cases where we already know what tag sequences we are looking for.

Notes

Please feel free to post comments, questions, or improvements to this protocol. Happy to have your input!

  1. List troubleshooting tips here.
  2. You can also link to FAQs/tips provided by other sources such as the manufacturer or other websites.
  3. Anecdotal observations that might be of use to others can also be posted here.

Please sign your name to your note by adding ”’*~~~~”’: to the beginning of your tip.

References and additional reading

This protocol was published in:

Lindsay V. Clark, Joe E. Brummer, Katarzyna Głowacka, Megan Hall, Kweon Heo, Junhua Peng, Toshihiko Yamada, Ji Hye Yoo, Chang Yeon Yu, Hua Zhao, Stephen P. Long, and Erik J. Sacks (2014) “A footprint of past climate change on the diversity and population structure of Miscanthus sinensis.” Annals of Botany. doi:10.1093/aob/mcu084. Free offprint

This protocol is based heavily upon that of:

Poland JA, Brown PJ, Sorrells ME, and Jannik J-L (2012) Development of high-density genetic maps for barley and wheat using a novel two-enzyme genotyping-by-sequencing approach. PLoS ONE 7(2):e32253. doi: 10.1371/journal.pone.0032253

Barcode sequences are published in:

Thurber CS, Ma JM, Higgins RH, and Brown PJ (2013) Retrospective genomic analysis of sorghum adaptation to temperate-zone grain production. Genome Biology 14:R68. doi: 10.1186/gb-2013-14-6-r68

Additional reading

  • Baird NA, Etter PD, Atwood TS, Currey MC, Shiver AL, et al. (2008) Rapid SNP Discovery and Genetic Mapping Using Sequenced RAD Markers. PLoS ONE 3(10): e3376. doi:10.1371/journal.pone.0003376
  • Catchen JM, Amores A, Hohenlohe P, Cresko W, and Postlethwait JH (2011) Stacks: building and genotyping loci de novo from short-read sequences. G3: Genes, Genomes, Genetics 1:171-182. doi: 10.1534/g3.111.000240
  • Davey JL and Blaxter MW (2010) RADSeq: next-generation population genetics. Briefings in Functional Genomics 9(5):416-423. doi:10.1093/bfgp/elq031
  • Davey, J. W., Cezard, T., Fuentes-Utrilla, P., Eland, C., Gharbi, K. and Blaxter, M. L. (2012), Special features of RAD Sequencing data: implications for genotyping. Molecular Ecology. doi: 10.1111/mec.12084
  • Elshire RJ, Glaubitz JC, Sun Q, Poland JA, Kawamoto K, Buckler ES, and Mitchell SE (2011) A robust, simple Genotyping-by-Sequencing (GBS) approach for high diversity species. PLoS One 6(5): e19379. doi:10.1371/journal.pone.0019379
  • Hohenlohe PA, Catchen J, Cresko WA (2012) Population Genomic Analysis of Model and Nonmodel Organisms Using Sequenced RAD Tags. In: Data Production and Analysis in Population Genomics, Pompanon F and Bonin A, eds. 235-260. doi:10.1007/978-1-61779-870-2_14
  • Peterson BK, Weber JN, Kay EH, Fisher HS, Hoekstra HE (2012) Double Digest RADseq: An Inexpensive Method for De Novo SNP Discovery and Genotyping in Model and Non-Model Species. PLoS ONE 7(5): e37135. doi:10.1371/journal.pone.0037135
  • Serang O, Mollinari M, Garcia AAF (2012) Efficient Exact Maximum a Posteriori Computation for Bayesian SNP Genotyping in Polyploids. PLoS ONE 7(2): e30906. doi:10.1371/journal.pone.0030906

 

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