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Home Protocols Microarrays Protocol for Microarray Post-Processing (short) – Medina Lab

Protocol for Microarray Post-Processing (short) – Medina Lab

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adopted from UCSF protocol: http://cat.ucsf.edu/pdfs/hybCourseProtocols_v2.7.pdf

             

 

1. Etching and Crosslinking

 

Materials

Diamond-tipped glass etching pen

UV cross linker

Metal slide rack (1)

 

1.      With the diamond-tipped etching pen, lightly mark the boundaries of the array on back side of the slide.

2.      Place the slides array-side up on a piece of filter paper or other similar solid support.

3.      Place the arrays in the UV crosslinker and crosslink with 60 mJ of energy.  One common confusion is that the crosslinker lists its energy in 100 µJ, so the display will say “600” (600 x 100 µJ = 60 mJ). Crosslink for about 2 min.

4.      Place etched, cross-linked slides in a clean metal rack.

 

2. Shampoo treatment

 

Materials

Glass slide dish (4)

95% EtOH made with filtered dH2O (400mL)

20X SSC, filtered (75mL)

10% SDS, filtered (10mL)

Filtered dH2O (~900mL)

Timer

Eppendorf 5430 table-top centrifuge

 

1.      Preheat Shampoo (3x SSC/0.2% SDS) to 65°C using water bath.

a.     500 ml solution:

                                               i.     75mL 20x SSC

                                             ii.     10mL 10% SDS

                                            iii.     H2O to 500mL

2.      Wash slide dishes with EtOH and allow them to completely dry.

3.      Pour heated shampoo into the first slide dish.

4.      Soak slides in shampoo for 5 minutes on the benchtop.

5.      Transfer the slides to a slide dish filled with room temperature water. Move the slide rack up and down a couple of times and transfer the slide rack to another slide dish filled with room temperature water. Incubate 1 minute.

6.      Transfer the rack to a slide dish filled with 95% EtOH. Incubate 30 seconds.

7.      Spin dry 3 minutes at ~50g (5430 Eppendorf). Placing a paper towel under the slide rack will help remove the EtOH. Use a second slide rack filled with slides as a balance.

8.      Leave slides in metal rack – proceed to blocking protocol.

 

 

3. Blocking

 

Materials

Succinic anhydride (5.5g) – stored in vacuum chamber

1-methyl-2-pyrrolidinone (335mL) – stored in flammable cabinet.

1M sodium borate pH 8.0 (15mL)

Glass slide dish (2)

Metal slide rack (1)

95% EtOH made with filtered dH2O (400mL)

Filtered dH2O (1L)

Timer

Eppendorf 5430 table-top centrifuge

4L beaker

500mL beaker

500mL flask

25mL serological pipet (2)

Stir bar

Pipet Aid

Shampoo’d slides

Clinical rotator – stored on shelf to the left of the hood.

Stirrer – stored on shelf to the left of the hood.

Magnet

 

1.      Clean the 4L and 500mL beakers with EtOH and wipe completely dry with a Kimwipe.

2.      Clear space in the hood and assemble all materials in the hood.

3.      Pour a small amount of 1-methyl-2-pyrrolidinone into the 500mL beaker. Verify that 1-methyl-2-pyrrolidinone is clear and colorless.  Do not use solvent that appears slightly yellow when you first pour it from the bottle.

4.      Pour 300mL 1-methyl-2-pyrrolidinone into the 500mL flask (use graduations on the flask).

5.      Add another 35mL 1-methyl-2-pyrrolidinone to the same flask using a 25mL serological pipet, so that the final volume is 335mL.

6.      Add 5.5g of succinic anhydride to 500mL beaker with stir bar.  Note that the stock bottle of solid succinic anhydride should be stored under desiccation and vacuum. Do not use if exposed to moisture! 

7.      Add the 335mL of 1-methyl-2-pyrrolidinone into the 500mL beaker.

8.      Turn on magnetic stirrer.   

9.      IMMEDIATELY after succinic anhydride dissolves, mix in 15 ml of 1M sodium borate pH 8.0 and allow to fully mix.

10.  Quickly pour the buffered blocking solution into a clean, dry glass slide dish (it is useful to have a magnet handy to prevent the stir bar from being poured into the slide dish).

11.  Plunge the slides rapidly into blocking solution and vigorously shake, keeping the tops of the slides under the level of solution, for 30 seconds.

12.  Put a lid on the slide dish, and let shake gently on a rotator for 15 minutes. 

13.  Drain excess blocking solution off slides for approximately 5 seconds and transfer slide rack to the 4L beaker filled with 1L of filtered water.

14.  Swish mix the rack gently by turning hand clockwise/counter-clockwise under the water for 60 seconds. 

15.  Transfer the rack to a glass dish of 95% EtOH and plunge to mix for 60 seconds. Make sure the EtOH is crystal-clear. Do not use if it appears to have particulates or if it appears cloudy.

16.  Spin dry 3 minutes at ~50g (5430 Eppendorf). Placing a paper towel under the slide rack will help remove the EtOH. Use a second slide rack filled with slides as a balance.

17.  After spinning, the slides should be clean and dry. Remove the slides from the rack and store in a plastic (not wood!) microscope slide box. Arrays may be used immediately. 

 

Protocol Variations and Tips

 

• If the methyl pyrrolidinone appears yellowish, DO NOT USE.

 

• Do not use succinic anhydride that has been exposed to moisture.

 

• If you observe streaks of DNA, or “comet tails,” on your hybridized array, the

initial plunge-mix of arrays into the succinic anhydride solution was too slow.

 

• UV crosslinking appears to enhance binding of long-oligo DNA to lysine slides.

However, crosslinking seems to have little or no effect on binding of PCR

product, as measured by hybridization intensity. If you choose to cross-link,

we suggest an energy of 60 mJ, applied after rehydration and before

shampooing or blocking.

 

• Try to minimize the time that the arrays are exposed to dust.  Keep them in a

box or submerged as much as possible.

 

• Not all lysine slides are created equal. Several people have noted that slides

can contain sub-regions where the lysine coating appears to be “weaker.” This

is manifested during the hybridization where the region once occupied by

spotted DNA is now able to bind fluorescent hybridization probe. This is

presumably due to spotted DNA leaving the surface during the boiling step,

since this step represents the harshest treatment. The resulting array scans

appear to have regions where all the spots are one bright uniform color. Two

protocol variations seem to mitigate this effect. The first is to simply omit the

boiling step which comes at the cost of some moderate sensitivity loss. The

second is to block the slides in the succinic anhydride solution for 5 minutes,

transfer to 95˚C water for 60 seconds, and then plunge back into the succinic

anhydride solution for another 10 minutes. After this, plunge into 95% EtOH

to remove organic solvent. The rationale being that “loose” DNA will leave the

slide during the boiling. The exposed amines will be blocked by the second

succinic anhydride incubation.