Chapter 1

Preparation of DNA printing plates

Task A: PCR Cocktail Preparation

Objective:

Preparation of mix to be added to yeast ORF specific primers for amplification of Saccharomyces cerevisiae genome ORFs (~65 96-well plates)

Procedure:

1. Prepare cocktail of PCR buffer, magnesium chloride, dNTPs, Taq, etc. for four plates (=40 mL) in a 50 mL conical tube as follows:

	One rxn	Four 96-well plates (400 rxns)
RESGEN ORF SPECIFIC PRIMERS:
Forward primer (20 uM)	5 uL	-
Reverse primer (20 uM)	5 uL	-
10X PCR buffer	10 uL	4000 uL
MgCl2 (1 M)	.05 uL	20 uL
100X dNTPs (25 mM each)	1 uL	400 uL
Yeast genomic DNA (0.2 ug/uL)	0.2 uL	80 uL
ddH2O	77.95 uL	31.36 ml
AmpliTaq (5 U/uL)	0.35 uL	140 uL
Total volume	100 uL	90 uL aliquots

2. Mix well by inverting tube
3. Store at –20 C if not to be used in less than 24 hrs (4 C if within 24 hrs)

Task B: PCR amplification of yeast ORFs

Objective:

Addition of premade PCR cocktail mix to yeast ORF specific primers, followed by thermal cycling for amplification of yeast genome ORFs (~65 96-well plates).

Procedure:

1. Pour cocktail into disposable trough. Using 12-channel pipetman dispense 88 uL of cocktail to the pre-aliquoted primers. Carefully mix by pipeting up and down 3 times
2. Place rubber mat over PCR plate. Spin plate at 500 rpm in table-top centrifuge, to remove air bubbles from mixture (very important)
3. Place plates in tetrad PCR machine. Maintain same orientation for all plates (eg, A1-12 always closest to back hinge) for tracking heat block geographic problems. Record plate/PCR block/time.
   Closing lid of tetrad PCR machine: Design of tetrad lids promotes evaporation during PCR reaction, increasing failure rate. To close lid properly, tighten until lid lifts up off the body of the PCR machine, then loosen lid JUST until it again touches the body of the PCR machine.
4. Run program with the following steps:

· pre-melting 92ºC (1’)

melting 92ºC (30")

annealing 56ºC (45")

synthesis 72ºC (3'30")

# cycles 36

· soak at 4-11ºC (hold forever)

5. PCR reaction takes approximately 3.5 hours
6. Remove from PCR machine
7. Store at 4 C if next step is less than 24 hours away (-20ºC if need more time)
8. Expect ~92-95% success rate after first pass
9. Following characterization of success rate by running gels (see below), re-amplify

failures.

Task C: Preparing agarose gels for electrophoresis

Objective:

To cast gels for running out products of each amplification reaction.

Procedure:

1. Prepare 800 ml of 1% agarose in 1X TAE (200 ml/gel tray times 4 trays, each for one 96-well plate)

2. Cook in microwave or on stirring heat block until boiling.

3. Allow to cool to 55-60ºC

4. Prepare 4 trays by taping ends well. Place four combs of 26 wells each into gel.

5. Adjust molten TAE agarose to 0.5 mg/mL ethidium bromide after having cooled to ~55-60ºC to avoid excess vaporization.

6. Pour 200 ml of cooled molten agarose into each prepared tray (above 65ºC may warp tray)

7. Allow to polymerize 20-30 minutes

8. Store in 1X TAE buffer containing 0.5 mg/mL ethidium bromide

Task D: Verification of PCR success by gel electrophoresis

Objective

To verify success of the PCR and also roughly size the products to verify identity. High resolution on the gels for exact sizing is not required, as this step is largely for quality control.

Procedure:

Agarose gel:	1% agarose, 1X TAE, 0.5 mg/mL ethidium bromide
Running Buffer:	1X TAE, 0.5 mg/mL ethidium bromide
Loading dye (6X):	15% Ficoll-400, 0.25% xylene cyanol FF, 0.25% bromophenol blue
DNA size ladder:	4 mL 1X TE buffer, 1 mL 1kb ladder, 1 mL 6X loading dye

1	Start with 3 uL of PCR reactions in PCR plates, after remainder is transferred to U-bottom plates (see next section). (can mix PCR + dye directly on parafilm or in a fresh plate)
2	Add 1 uL 6X loading dye to PCR reactions in PCR plates.
3	Load 6 uL DNA size ladder in lane #1 of each row.
4	Using a 12-channel pipettor, load samples A1-A12 into alternating lanes 2, 4,..., 24.
5	Load samples B1-B12 into alternating lanes 3, 5,..., 25.
6	Repeat this procedure for the remaining samples, such that two sequential rows of PCR reactions are loaded into a single row of wells in alternating lanes.
7	Run at 70-80V until the first dye band (XC FF) is halfway to the next row of wells.
8	Take a high (~1") and low (~6/30") exposure photographs. Compare to predicted ORF sizes and for the presence of significant doublets.
9	Repeat PCR rxns for failed ORFs. NOTE: For 2nd PCR attempt, sort failures by gene size, doublets, etc., and modify reaction conditions accordingly. For genes that still give PCR failures, design new primers, e.g. to amplify subregions of genes.

Notes:

1. Loading gels: easiest to mix and load 24 wells at a time. Aliquot 24 spots of 1uL gel dye, add 3uL of PCR reaction, mix by pipeting, and then directly load 12 wells of the gel. Repeat this for another 12 wells (i.e. two rows from the 96 well plate loaded into one row of wells on the gel). Apply low voltage while preparing other samples to prevent diffusion of the DNA during loading.
2. Running gels:

· Do not run short products (<300 bp) off the gel. To ensure this, run until blue dye is 2/3 down the gel.

· b. During gel running, be sure gel is centered in gel box and resting within the plastic frame: if the gel floats off the frame toward the cathode the DNA will run out the BOTTOM of the gel leading to false negatives.

Tasks E-L: DNA Cleanup

Task E: Transfer of PCR products to precipitation plates

Task F: Addition of isopropanol to PCR products

Task G: Precipitation of DNA by centrifugation
Task H: Aspirating isopropanol supernatants
Task I: Addition of ethanol to PCR products
Task J: Precipitation/wash of DNA by Centrifugation
Task K: Aspirating ethanol supernatants
Task L: Resuspension of dried ORF DNA

Objective:

To cleanup PCR products in preparing spottable DNA for arraying.

Procedure:

Task	Step
E	1.	[This step is optional and we will not be performing it in this course. Instead, we will precipitate the PCR products directly in the PCR plates.] Transfer PCR reactions to 96-well U-bottom tissue culture plates (Costar #3790).
		If gels have not been run, transfer 3 uL back to PCR plates for check gels (see above).
F	3.	Add 1/10 vol. 3M sodium acetate (pH 5.2) (10 ul to a 100 ul PCR reaction). Then add 1 volume (100 ul) isopropanol and pipet up and down 3 times. Change pipet tips between rows.
		Store at -20ºC for a few hours to overnight (this is optional.)
G	4.	Centrifuge the plates in Sorvall at 3500 rpm (RC3B rotor) for 1 hr. If precipitating in PCR plates, use plastic holders designed for PCR plates in order to support them during the spin.
H	5.	Invert plate over sink in order to remove supernatant. Alternatively, remove supernatant with 12-channel aspirator (Wheaton/PGC Scientific #851388)
I	6.	Add 100 uL of ice-cold 70% ethanol and centrifuge again for 30 min.
	7.	Dry the pellets in speed-vac for 10 min (if available).
J	8.	Resuspend DNA in 100 uL dH2O overnight.
	9.	Transfer in 10 uL aliquots to 384-well plates (Corning Costar #6502) to make 10 duplicate print plate sets.
K	10.	Dry down print plate sets in speed vac (if available).
		Tightly seal plates with aluminum foil (R.S. Hughes #425-3) for long-term storage at room temperature.
L	11.	Before use, resuspend one print set in 4-5 uL 3XSSC and allow to hydrate overnight.

Task M: Rearray into 384-well plates

Objective:

To reformat spottable PCR products from 96-well to 384-well plates, in preparation for arraying (converting from 9-mm to 4.5-mm well spacing).

Procedure:

This rearraying scheme ignores the most common 96 to 384 reformatting schemes, namely ones which use robots with 96 tips (eg, Beckman Multimek, Robbins Hydra, Tecan, Hamilton, etc.). These robots, fill a 384-well plate using four interlaced 96-well registers corresponding to the 4 constituent 96-well plates.

Most commonly, this is done in one of two ways:

1. "ZIGZAG", Beckman Multimek's Default: 4 registers anchored at the top left (96-well position A1) fill 384-well plate as follows: top left for register 1 is A1, register 2 is A2, register 3 is B1, and register 4 is B2.
2. "Clockwise", Robbins Hydra's default: again, 4 registers anchored at the top left (96-well position A1) fill 384-well plate as follows: top left for register 1 is A1, register 2 is A2, register 3 is B2, and register 4 is B1.

Other schemes also exist:

Chapter 2

Preparing to print

Poly-L-lysine coating of glass slides for microarraying

Glass slides can be treated with a variety of coatings for attaching DNA to the surface, but in our experience, poly-L-lysine coating is the most convenient and best performing method of attachment. Although it's possible to buy lysine coated slides commercially, most of them are not as good as those you can make on your own with adequate care. Many commercial slides we have seen have dirt on the surface or uneven coatings of lysine.

The coating procedure consists of first cleaning the slides thoroughly with a basic ethanol solution and rinsing, followed by immersing them in a buffered solution of lysine. The slides are then rinsed briefly and spun dry to achieve an even coating.

Materials	Quantity	Ordering information
Glass microscope slides	60	Gold Seal #3010
Slide rack with handle	2	Shandon Lipshaw #121 (800-245-6212) Each rack holds 30 slides
Slide chamber	6	Shandon Lipshaw #121 Each chamber holds 350 mL
Double distilled water	~5 L
NaOH	70 g
95% Ethanol	420 mL
Poly-L-lysine	70 mL	Sigma # P 8920
Tissue culture PBS	70 mL
Vacuum oven (45C)
Slide box (plastic only)	1	VWR #48443-806

Notes

Wear powder free gloves whenever you handle slides in this protocol.

Directions

1. Rinse the glass chambers with the metal slide racks thoroughly with distilled water and get rid of most of the water by shaking & air drying. Place 30 slides in each slide rack.

2. Prepare cleaning solution: Dissolve completely 70 g NaOH in 280 mL ddH2O. Add 420 mL 95% ethanol slowly and stir until completely mixed. The total volume is 700 mL (= 2 X 350 mL). If solution remains cloudy, add double distilled H2O until clear.

3. Pour cleaning solution into 2 empty slide chambers. Use the wire handle to dunk the rack of slides into the solution, plunge it up and down briefly and cover chambers with glass lids. Shake on an orbital shaker for 2 hr. Once slides are clean, they should be exposed to air as little as possible. Dust particles will interfere with coating and printing. You should also ensure that PBS solution is made up at this point. (1 litre PBS has 8 g Sodium chloride, 0.2 g potassium chloride, 1.44 g Sodium phosphate, dibasic, anhydrous and 0.24 g potassium phosphate, monobasic)

4. Quickly transfer racks to fresh chambers filled with double distilled H2O. While transferring, tilt the rack to drain as much of the cleaning solution as possible. Rinse vigorously by plunging racks up and down in the fresh water. Repeat rinses at least 4 times with fresh double distilled H2O each time. It is critical to remove all traces of NaOH-ethanol. The slides remain in the final rinse water till the next step, which is the lysine coating.

5. Prepare poly-L-lysine solution: 70 mL poly-L-lysine + 70 mL tissue culture PBS + 560 mL water. Use a plastic graduated cylinder and beaker. Pour this lysine solution into two clean slide chambers.

6. Transfer the racks of slides from the rinse water to the chamber with poly-L-lysine solution and shake for 1 hr on the orbital shaker.

7. Transfer rack to fresh chamber filled with double distilled H2O. Plunge up and down to rinse for about 1 minute.

8. Place slides on microtiter plate carriers (place paper towels below rack to absorb liquid). It is best to do this step (transfer from water to centrifuge carrier) right at the centrifuge to avoid exposing the wet slides to air for any length of time. Centrifuge for 5 min. at 500 rpm. This step ensures even coating and drying of the slides. Transfer the slide racks to dry empty chambers with covers for transport to vacuum oven.

9. Dry slides in the racks (with cover off chamber) in 45 C vacuum oven for 10 min. (Vacuum is optional.)

10. Transfer slides from the racks to a clean slide box. The slides should look perfectly clean when inspected against the light. Store slides in closed slide box (plastic only, without rubber mat bottom) till they are ready for use in the printing process.

10.

Normally, the surface of lysine coated slides is not very hydrophobic immediately after this process, but becomes increasingly hydrophobic with storage. A hydrophobic surface helps ensure that spots don't run together while printing at high densities. This may not be a problem with fine printing tips and adequate spot-to-spot spacing, so the coated slides may be ready for use in a day or two. After they age a few days, (up to a week) they are probably optimal. The hydrophobicity can be tested by observing how a drop of water beads off the surface (compare this to a just coated or un-coated slide). However, coated slides that have been sitting around for long periods of time (few weeks) are usually too old to be used. They often show opaque patches when held to the light and these result in high background hybridization from the fluorescent probe. It's useful to test print, hybridize and scan sample slides to determine slide batch quality. Inspect every slide before placing it on the arrayer to ensure you don't use one that has obvious problems like dirt, uneven coating or opaque patches.

Organization (Group of 4 students)

Common task-make PBS for everyone

Student 1 & 2	Wash chambers, place slides in rack	15 minutes
Student 3 & 4	Make cleaning solution
	Cleaning	2 hours
Students 1& 2	Make poly-l-lysine solution
Students 3 & 4	Rinse 4x, 1 rack each	15 minutes
All	Poly-L-lysine	1 hour
Students 2 & 3	Rinse, dry & store	15 minutes

Post-Processing of printed microarrays

After printing microarrays on poly-L-lysine coated glass slides, the slides need to be processed before they can be used for hybridization. Most importantly, the remainder of the lysine coated surface needs to be blocked to prevent non-specific attachment of the fluorescently labelled probe DNA to the free amine groups. Blocking is achieved by treating the surface with succinic anhydride (in an organic solvent), which forms an amide bond with the free amines. The processing step also rehydrates the array to make individual spots more even and probably also denatures the DNA to some extent.

Materials Needed

Materials for 30 arrays	Quantity	Order info
Humid chamber	1	Sigma #H 6644
Diamond scriber	1	VWR #52865-005
Slide rack with handle	1	Shandon Lipshaw #121
Slide chamber	2	Shandon Lipshaw #121
Succinic anhydride	6 g	Aldrich #23,969-0
1-Methyl-2-pyrrolidinone	325 mL	Aldrich #32,863-4

Inverted heat block (90-100 C)

Sodium borate (1M, pH 8), 15 mL Make this by dissolving boric acid in water and adjust pH with NaOH

double distilled H2O ~1 L

2 L glass beaker

95% ethanol 350 mL

Notes

Wear powder-free gloves for the entire procedure.

Directions

1. Mark the array boundaries: The spots on an array will become invisible after post-processing as the salt gets washed away. Since you need to know where to pipette the probe on, you have to mark boundaries of the array. The best way is to etch a couple of lines just outside the top and bottom of the array, on the back of the slide using a diamond scriber. If the arrays are not labelled, now is the time to do it. Labelling can be done with peel-off labels or with the diamond scribe (see array printing). You need to be able to orient the array properly using the labels.

2. Fill the bottom of humid chamber with 100 ml 1X SSC. Place arrays face down over the 1X SSC in the chamber and cover with lid. Spots will take up moisture and swell. Re-hydrate until array spots glisten, approximately 5-15 minutes. This is best monitored by looking at the spots with a magnifying glass. Allow the spots to swell slightly but not run into each other. Using a slightly warm solution (35-40 C) is one way to speed things up.

3. Quickly transfer slides with their array side up, one by one from the humid chamber to the smooth surface of an inverted heating block at 90-100 C. Snap-dry each array for a few (3-5) seconds. You can usually see a ‘wave’ of dryness sweeping across the array as the spots are snap-dried.

4. Place arrays in metal slide rack with wire handle. Keep it close to a clean, empty slide chamber on an orbital shaker. Be sure the rack is bent slightly inwards in the middle, or else the slides may run into each other while shaking.

5. Prepare blocking solution: Have three 350 ml glass chambers with lids available, as well as a 2 L glass beaker or large round pyrex dish with double distilled H2O to a height of about 3 in., just enough to cover the rack of slides. The water should be heated in the microwave or on a heating plate to close to boiling. At this time, prepare the 15 ml sodium borate in a 50 ml conical tube.

6. Dissolve 6 g succinic anhydride in approx. 335 mL 1-methyl-2-pyrrolidinone. The most efficient way to do this is to have a clean 500 ml graduated glass beaker with a stir bar on a stir plate. Weigh the succinic anhydride and drop it into the beaker. Pour 1-methyl-2-pyrrolidinone from the stock bottle into the beaker till it is approximately 335 ml and turn on the stirrer.

7. Immediately after the last flake of the succinic anhydride dissolves (about a minute), add the 15 mL sodium borate into the still stirring solution.

8. Immediately after sodium borate solution mixes in, pour the solution into empty slide chamber on the orbital shaker. Plunge slide rack rapidly and evenly in solution. Vigorously shake up and down for a few seconds, making sure slides never leave solution.

9. Shake on orbital shaker for 10-15 min. Ensure that the water in pyrex dish or glass beaker is maintained at a point just before boiling (95 C).

10. Transfer the rack of slides from the blocking solution to the 95C water. Gently plunge slide rack in hot water for 2 min.

11. Transfer the rack of slides to a slide chamber with 95% ethanol. Plunge slide rack a few times in the ethanol.

12. Take the chamber with slides in ethanol to the centrifuge and transfer the rack to the carriers. Load slides quickly and evenly onto the carriers to avoid streaking. Centrifuge slides and rack for 5 min. @ 500 rpm.

13. Transfer arrays to a clean slide box. They are ready for use immediately. Processed arrays are good for many weeks.

Organization (Group of 4 students)

All (in turn)	Etch slides	10 minutes
All (in turn)	Rehydrate and snap dry	20 minutes
Students 1& 2	Blocking	20 minutes
Students 3 & 4	Transfer to water, ethanol and dry, cleanup	10 minutes

PRINTING MICROARRAYS

A microarray printing run of more than a hundred microarrays with many thousands of elements each is always a fun and rewarding activity, but without the right attitude, it can become a chore that you dread. A typical print run takes about a day of non-stop printing depending on the number of printing