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Use this protocol with Chromatin Immunoprecipitation to detect the in vivo chromatin binding sites of a protein of interest.

There is also a special protocol for ChIP-Chip E. coli.


In progress... ~ cmc 12:55, 22 August 2006 (EDT)

Cy3-Cy5 Labeling kit

  • Invitrogen Bioprime Labeling Kit


Wash Buffer I

per 2 L (final conc.):

  • 600 ml of 20x SSPE (6x)
  • 1398 ml of ddH2O
  • 2 ml of 5% N-lauroylsarcosine (0.005%)
  • Filter and store at rm temp.

Wash Buffer II

per 2 L [Final]:

  • 6 ml of 20x SSPE [0.06x]
  • 1994 ml of ddH2O

Wash Buffer III

  • Anti-oxidant mixture in acetonitrile. (Purchase 1x from Agilent)


  • Schedules include the ChIP protocol.


  • Day 1:
    • x-link cells, bind Ab on beads, cell sonication, ChIP --> o/n incubation
  • Day 2:
    • wash, elute, reverse x-link --> freeze o/n
  • Day 3:
    • RNase, ProtK, T4 Fill-in, Blunt end ligation --> o/n incubation
  • Day 4:
    • LM-PCR, Cy3, Cy5 label --> o/n incubation
  • Day 5-6:
    • Hybridize arrays --> 40 hr. incubation
  • Day 7:
    • Wash and scan arrays, Analysis

I need results asap

  • Day 1:
    • x-link cells, bind Ab on beads, cell sonication, ChIP --> o/n incubation
  • Day 2:
    • wash, elute, reverse x-link, RNase, ProtK, T4 Fill-in, Blunt end ligation --> o/n incubation
  • Day 3-4:
    • LM-PCR, Cy3, Cy5 label (3 hr.), Hybridize arrays --> 40 hr. incubation
  • Day 5:
    • Wash and scan arrays, Analysis


X. Cy3-Cy5 Random Primer labeling

  1. From the normalized IP and wce PCR DNA above, 1 µg of DNA will be fluorescently labeled (=10µL of LM-PCR product).
  2. Add 24 µL water to above.
  3. Add 26.8 µL of 2.5 X random primer solution (Invitrogen Bioprime labeling kit).
  4. Boil 5 min in heatblock.
  5. Place tubes in icewater. Incubate 5 min.
  6. Add 8 µL 10X low T dNTP mix (1.2 mM dATP, dCTP, dGTP each and 0.6mM dTTP).
  7. Add 1 µL of cy5-dUTP to IP tube and 1 µL cy3-dUTP to input tube.
  8. Add 1 µL of high concentration Klenow (40U/µL, Bioprime kit).
  9. Incubate 20∞C 6hrs- O/T (keep samples in dark as much as possible from here on).
    • Alternative: incubate 37oC for 3 hrs.

XI. Clean-up reactions using CGH columns

  1. Column Protocol
    • Add 25 uL of H20 to samples to bring the volume up.
    • Add 400 uL of Buffer A to each tub, vortex for 30 sec.
    • Load sample onto column inside collection tube.
    • Spin 1 min at 11,000 x g, rm temp. Discard flow through
    • Add 600 uL of Purification Buffer B.
    • Same spin again, discard flow.
    • Add 200 uL of Buffer B to columns
    • Spin again and discard.
    • Put columns in new, clean tubes. Add 50 uL of dd H2O. Incubate at rm temp for 1 min.
    • Same spin again. Keep flow-through.
  2. Clean-up the Clean-up
    • Spin 10 min at 14K. Spin down your labeled material to remove fine white particulate. Carefully remove liquid. Leave a relatively large volume behind if necessary. Repeat as often as necessary (as many as 3 –5 times). This step is crucial. The particulate will bind to Agilent slides creating a lot of background.
  3. Precipitate DNA with 25uL of ammonium acetate and 300 uL of EtOH. Do not wash pellet; just dry sample preparation.
    • Redissolve in H2O and nanodrop. NOTE: The precipitation is not absolutely required, but IS highly recommended. Precipitation seems to affect reproducibility. Using precipitated material, there’s less difference in the results you get when comparing hybes on fresh slides and

XII. Array Hybridization

  • Note: keep samples in dark as much as possible (use foil as necessary)
  1. Thaw labeled DNA, combine Cy3 and Cy5 labeled samples (combine IP and wce).
  2. To each sample, add 23.5 uL of control DNA mix:
DNA Stock Concentration Final Concentration Volume (uL) per Sample
Labeled DNA n/a 10 µg n/a
Herring sperm DNA 10 ng/µl 750 ng 7.5
tRNA 8 µg/µl 40 µg 5.0
CoT 1 µg/µl 10 µg 10.0
Agilent oligo controls 1x 1.0

  1. Precipitate with 170 uL of amonium acetate and 750 uL of EtOH.
  2. Incubate precipitations at -20∞C or -80∞C for 20 min.
    • Labeled DNA precipitates less well than unlabeled.
  3. Spin 10 min, 14 K, 4∞C. Pipette off EtOH, do not wash, dry pellet.
  4. Resuspend in 100uL H20
  5. Add 400uL of Hyb buffer to each sample:
Hybridization Buffer Stock Concentration Final Concentration Volume (uL) per Sample
Na-MES (pH 6.9) 500 mM 50 mM 50.00
NaCl 5000 mM 500 mM 50.00
EDTA 500 mM 6mM 6.00
sarcosine 5% 0.50% 50.00
formamide 100% 30% 150.00
H2O 94.00

  1. Heat samples to 95°C for 3 min.
  2. Transfer tubes to 40°C heat block for 15 min. Prepare hybridization chambers during this step. #*It’s easiest to handle 4-6 hybes at a time.
    • Put a gasket slide in the metal base of hybe chamber: “Agilent” on gasket slide faces up. The barcode end of the gasket slide goes in oriented so that the barcode will not be exposed in the open areas of the metal base.
  3. Spin tubes 14K x g for 45 sec.
  4. Assemble hybridizations
    • Add 490.0 µl of hybridization mix to gasket slide. Try to lay hybe mix in one straight line, evenly distributed along slide without touching the gasket slide with the tip of the pipette.
    • Place array on hybridization mix. The “Agilent” on the array slide is always face down and should align with the “Agilent” on the gasket slide (the “Agilents” kiss in slide sandwich). Place the slide using a smooth, even motion with one end of the array slide touching the liquid first. A quick drop will create bubbles.
    • Add metal top. Leaving the chamber flat on the bench, slide screw assembly over the end of the chamber and tighten until the screw is as tight as you can make it using only your hands and a medium level of force.
    • Ensure free rotation of mixture around gasket periphery. You have to make sure the liquid can flow evenly throughout the chamber.
  5. Incubate at 40°C in rotating oven for 40 hours. Check for bubbles at ~24 hrs. Tap and rotate slowly to reduce bubbles.
  • Note: Gasket slides can be re-used. Check them carefully to make sure the gaskets are intact. Make sure they are dry and free of water stains or salt spots. (Ideally re-use gasket slides only 3-6 times.)
  • Tips for ensuring free rotation of mixture around gasket periphery. Lift assembled chamber slowly towards you, leaving one corner in contact with the bench. This will allow the hybe mix to flow to the bottom with a minimum of bubbles. Pick up the chamber and SLOWLY rotate it while gently hitting it with the palm of your other hand to prevent the liquid from catching on the gasket and creating bubbles. Keep watching the liquid. Try to make 3 – 4 full rotations without any bubbles.
  • If you find you’re creating a lot of bubbles as you turn the chamber, it’s probably not screwed tight enough – you’re letting air in as you turn and hit it. If you do get bubbles, they will tend to pop. For bubbles stuck on the edge of the chamber, rotate the chamber so the bubble is not touching liquid and let it dry out and pop. For bubbles in the liquid, rotate the chamber so the long axis of the chamber is parallel to the floor. The bubbles should eventually thin and pop. For bubbles stuck on the slide surface, hit the edge of the chamber closest to the bubble until it is free flowing, then treat them as above to let them pop. Small, free floating bubbles can be left alone. If the bubble is particularly stubborn, slightly loosen the screw which will increase the volume in the slide sandwich and encourage the bubble to pop.

XIII. Array Washing

  • Process one array at a time through step 3. Process groups of up to 10 arrays during steps 4 through 9.
  • Note: steps 1-3 should be done as rapidly as possible.
  1. Disassemble hybe chambers and transfer hybe sandwich to reservoir containing Wash Buffer I.
  2. While submerged, IMMEDIATELY use Agilent tweezers to separate gasket slide from array slide and gently swish array slide through liquid.
  3. Place array in second dish of Wash I containing a slide rack (if doing whole genome hybes, up to 10 arrays can accumulate in this rack).
  4. Place dish on oscillating shaker at 60rpm for 5 min (alt: gentle stirring with stir bar).
  5. Remove rack, quickly dip it into a new dish containing Wash II and transfer to a second dish containing Wash II and a stir bar. Wash for 5 min with stirring (gentle agitation at surface).
  6. Remove rack, quickly dip it into dish containing acetonitrile and transfer to a final dish containing Wash III and a stir bar. Wash for 30 sec with stirring like above.
  7. SLOWLY remove rack and slides from Wash III – it should take approx 10 sec to lift rack from solution (the slides should be dry at this point).
  8. Scan immediately or store in vacuum sealed bags or nitrogen environment.

XIV. Scanning Arrays

  1. Load Arrays into array holders. Place in Agilent Scanner add details
  2. Open Agilent Control Scanner program to warm up scanner.
  3. Go to Settings --> modify default. Select the scan size for the type of array you are using.
  4. Select Red/Green settings. Start with 60/80.
    • You may need to adjust this after evaluating your first scan.
  5. Highlight the first array, select "reset selection" to use the new default settings above.
  6. Scan first array. Check histogram for Red/Green balance.
  7. Adjust Red/Green if necessary and scan all other arrays.
    • Try bumping up the low color rather than bringing the high color down.
  • Visualize in Genepixpro.
  • Note: The red dye is more sensitive to oxidation/degradation.

XV. Analysis

Stripping Agilent Slides

  • Strip just prior to re-use, rather than immediately after use. (Better preservation of oligos and better removal of hybridized DNA.)
  1. Acetonitrile Wash
    • Add stir bar to glass dish for washing slides.
    • Add 250 – 300 ml of 100% acetonitrile (FPLC grade) to glass dish.
    • Arrange up to six slides in 10-slide capacity, glass slide rack.
    • Place slides and rack in glass dish.
    • Stir with medium low speed for 5 minutes. Longer than 10 minutes will likely damage the slide surface.
    • Slowly remove rack from acetonitrile. Slides should be dry. Go to Step II.
  2. Phosphate Buffer Wash
    • Put 600 ml of 100 mM potassium phosphate buffer (pH6.6, filtered) in a 2 liter beaker.
    • Microwave until buffer reaches 65°C (4 minutes using the microwave in Room 511).
    • With 2 minutes remaining, turn hotplate/stirrer to high heat to pre-heat the plate.
    • When buffer is warmed, transfer beaker to hotplate.
    • Add magnetic stir bar. Add slide rack containing slides. Add thermometer
    • Cover beaker with aluminum foil and insulating cover.
    • Start stirring at medium-low speed.
    • Heat until buffer reaches 100°C AND there is a steady stream of bubbles. This should take about 10 minutes.
    • Leave slides in hot buffer for 5 minutes. Buffer should reach a rolling boil during this time.
    • Remove slide rack and blot excess liquid on Kimwipes.
    • Quickly plunge slide rack and slides into glass dish containing 250 ml of 100 mM potassium phosphate buffer (pH6.6, filtered).
    • Let slides cool for 2 minutes.
    • Slowly remove rack from buffer. Slides should be dry. Usually, slides are used immediately after stripping. If you have to store them, store in vacuum sealed bags or high nitrogen environment.


  • please contribute user notes.