Chapter 1

Preparation of DNA printing plates


Task A: PCR Cocktail Preparation



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



  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)




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


77.95 uL

31.36 ml

AmpliTaq (5 U/uL)

0.35 uL

140 uL

Total volume

100 uL

90 uL aliquots


  1. Mix well by inverting tube
  2. 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



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). 



  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)


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




Task C: Preparing agarose gels for electrophoresis



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



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



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.



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



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)


Add 1 uL 6X loading dye to PCR reactions in PCR plates.


Load 6 uL DNA size ladder in lane #1 of each row.


Using a 12-channel pipettor, load samples A1-A12 into alternating lanes 2, 4,..., 24.


Load samples B1-B12 into alternating lanes 3, 5,..., 25.


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.


Run at 70-80V until the first dye band (XC FF) is halfway to the next row of wells.


Take a high (~1") and low (~6/30") exposure photographs.

Compare to predicted ORF sizes and for the presence of significant doublets.


Repeat PCR rxns for failed ORFs.


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. 



  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



To cleanup PCR products in preparing spottable DNA for arraying.








[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).



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.)



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.



Invert plate over sink in order to remove supernatant.  Alternatively, remove supernatant with 12-channel aspirator (Wheaton/PGC Scientific #851388)



Add 100 uL of ice-cold 70% ethanol and centrifuge again for 30 min.



Dry the pellets in speed-vac for 10 min (if available).



Resuspend DNA in 100 uL dH2O overnight.



Transfer in 10 uL aliquots to 384-well plates (Corning Costar #6502) to make 10 duplicate print plate sets.



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.



Before use, resuspend one print set in 4-5 uL 3XSSC and allow to hydrate overnight.









Task M: Rearray into 384-well plates



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).




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.




Ordering information

Glass microscope slides


Gold Seal #3010

Slide rack with handle


Shandon Lipshaw #121 (800-245-6212) Each rack holds 30 slides

Slide chamber


Shandon Lipshaw #121
Each chamber holds 350 mL

Double distilled water

~5 L



70 g


95% Ethanol

420 mL



70 mL

Sigma # P 8920

Tissue culture PBS

70 mL


Vacuum oven (45C)



Slide box (plastic only)


VWR #48443-806



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



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.


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



2 hours

Students 1& 2

Make poly-l-lysine solution

Students 3 & 4

Rinse 4x, 1 rack each

15 minutes



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


Order info

Humid chamber


Sigma #H 6644

Diamond scriber


VWR #52865-005

Slide rack with handle


Shandon Lipshaw #121

Slide chamber


Shandon Lipshaw #121

Succinic anhydride

6 g

Aldrich #23,969-0


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



Wear powder-free gloves for the entire procedure.



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


20 minutes

Students 3 & 4

Transfer to water, ethanol and dry, cleanup

10 minutes




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 tips and elements to be printed.  Someone knowledgeable needs to be near the machine constantly to make sure everything is going smoothly.  Before you begin the run, you need the following:



1.      DNA to be printed, aliquoted in printing plates:  It’s best to have this in 384-well plates.  The best printing plates are Genetix 384-well plates (see DNA prep and precipitation section for details).  For printing, the DNA needs to be in about 4 ul of 3X SSC.  Obviously, you should know how the DNA was aliquoted.  The best method of storing print plates is to dry them down.  If DNA in water was dried down, you need to add 3X SSC and if it was dried down in 3X SSC, you will be adding water.  This might seem like an obvious point, but when there’s a production pipeline going with many people doing different steps, it’s easy to lose track!  If dried down DNA needs to be resuspended, add the water or SSC a few hours before beginning the print run.  After it is resuspended, keep the plate tightly sealed to minimize evaporative losses.

2.      Familiarity with the ArrayMaker software:  You should know how to do alignments and what the different parameters in the test print and print window mean.

3.      Glass slides: You need plenty (~150) of coated glass slides from batches that have been tested with test prints and hybridizations.  See poly-L-lysine coating and test print sections.

4.      Printing tips: You should have many spares in addition to the 16 or 32 that you are planning to print with.  Obviously, these should all be in good condition.

5.      Other materials: 3X SSC, double distilled water, test print DNA (100 ng/ul sheared salmon sperm DNA in 3X SSC), ¼’ wide labelling tape, magnifying glass.  Also useful is a dissecting microscope to inspect printing tips, and a camera attached to a monitor that will let you observe the arrays being printed.



Details for some of the following steps are covered in the software section:


Turn on the main power and computer, run ArrayMaker and home the stages

See the Mguide and software section for details.


Align all positions

It is necessary to make sure that all the positions are correctly aligned at the start of every print run.  Although none of the positions might have changed, there’s always the possibility that someone had last printed with a weird configuration of tips in a 32-tip holder and aligned that to the dry station and print plate, or used a slightly different print plate.  If you are not absolutely sure that all of the positions are perfect, align everything.  It’s a good idea to make sure anyway. Clicking on the Align button on the main window takes you to the align window.  In the Align window, clicking on the “Reset Z” button resets all the vertical positions to the ready position, which is safely above everything. You can then click on the buttons for the various positions and carry out alignments.  See the software section for details on alignment.  The tips in the rinse station (sonicator) should be aligned with only the slot immersed in the liquid.  Remember to set and save before exiting this window.  If you are printing with less than 32-tips (and using a 32-tip holder and dry station), cover the unused holes in the dry station with a piece of tape.


Printing tips

The quality of microarrays you print depends a lot on the printing tips (the DNA in the printing plates is another major factor).  The current generation printing tips pick up ~ 0.5 ml of solution and deposit a few nl with every tap on the glass surface.  A good printing tip should be able to print 100-200 spots before it needs to be loaded again.  A good set of 16 or 32 tips should produce uniformly sized, distinct, well spaced spots across the entire array.  Before beginning, inspect the tips carefully for obvious imperfections under a dissecting microscope.  You can observe how well a tip picks up liquid by just touching the tip to the surface of a 3X SSC solution.  With a good tip, you should see the solution wicking up in the slot in the center of the tip.  Make sure the outside is free of any salt or corrosive build-up, especially at the retaining clip or inside the slot.  Once the tips are in the holder and the top is on, use forceps or needle nose pliers to make sure all the tips slide up smoothly and that the spring pushes them back down when released.  You might need to clean the tip-holder holes of any build up.  Gentle sanding of the outside of the tip (where it rubs against the tip-holder) might also be called for occasionally.  Use a fine grade sandpaper for this.  If the shaft of the printing tip is slightly bent, it affects the smooth up and down movement and also alters the register of the different sectors of the test print array.  If this is the case, replace the tip to get a set that is well matched and evenly spaced.  Tips should be cleaned after every print run by sonicating in warm water and rinsing with ethanol.  Handle tips with great care.  Their points should never hit anything hard except the slides while printing.  Putting them in 200 ul pipette tips is a convenient way to store them.


Test prints

It is important to do test prints with the tips that will be used for the print run, immediately before beginning the run.  Tips that have performed well in the past may gradually deteriorate with improper cleaning and other abuse.  Load a sample print plate that is identical to the real print plates with 4-5 ml test print DNA in the top left 16 or 32 wells (beginning at A1) and use this for test printing on slides from the same batch that will be printed on.  The test print parameters (spacing and number of spots across) should match what the real array is going to have.  This can be calculated depending on how many elements total you need to print (see software section for details). Tape down a glass slide in position 1 on the platter (closest to the plate holder) for test printing.  If printing with 16 or 32 tips, it’s very convenient to simply print 24 spots across in each quad.  This way, one row of spots on the array corresponds to 1 or 2 384-well plates and makes it easier to localize imperfections to specific print plates.  However, this will limit the total number of spots you can print.  You can set the tips to load multiple times, which allows you to print thousands of spots on the same test-print slide.  You can use the same wash and dry parameters as for the actual run or shorten them slightly.


The liquid in the sonicator for cleaning the tips can be either water or SSC (0.5 X or 1 X).  Using SSC leads to somewhat better ‘priming’ of the tips and they can then print more spots before running dry, but depending on the DNA in the print plates, it could also lead to spots that tend to spread.  You should do a test print run using a spare sample plate of the actual DNA that will be printed to see how different wash solutions affect the spot quality.  Do test prints from different load positions on the 384-well plate to check the plate alignment, and on different slide positions on the platter to check the slide height adjustment on the platter.  Use of the plate positioner is strongly recommended.


Printing the real microarray

Once you have a set of printing tips working perfectly on the test print, start the array print run immediately.  Set the print parameters in the Print Array window.  To prevent carryover on the tips, it is necessary to do at least 3 cycles of cleaning, with 5-6 sec sonication and 8 sec dry times.  A few other things to note:  You of course know how many printing plates total you have, but set the spacing so that you have room for more.  Printing a plate of controls is useful. The control plate can include l DNA, salmon sperm DNA, poly dA, Cot1 or other low complexity repeat containing DNA, genomic DNA, 3X SSC, clones corresponding to doped in controls, dilutions of fluorescently labelled cDNAs, etc. as positive and negative controls. You should also try to leave room for re-printing a few plates should there be a problem with some of them.


When all the parameters are properly set, you are ready to begin.  Spray off the dust and glass shards on the platter and wipe the surface with a moist kimwipe.  Place your slides (wearing powder-free gloves) on the platter, making sure they are aligned straight and lying flat.  If printing less than 137 slides (a no-no!) you should fill the platter in the order they will get printed.  Use the ¼’ tape to tape down all slides along the edge.  Don’t put the tape on top of the dowel pins on the platter.  Use pieces of tape to tape across columns.  Taping securely is important as you don’t want your slides moving around during the print run.  Place the dust cover on as soon as you are done taping the slides down.


Spin down every print plate just before putting it into the plate holder.  You want to ensure there’s no liquid on the sides of the wells.  Be careful when taking off the seal.  Make sure the plate is placed in the proper orientation in the holder. Again, the plate positioner helps.  You will have a spreadsheet listing the contents of each well in the printing plates.  The order and orientation on the slide platter determines where each DNA is going to end up on the array, and only you can keep track of this.  It’s very useful to keep a log at the arrayer where the orientation and plate number of every plate can be recorded for each print run. When everything is ready, drop the first plate in and click the start button!


Taping diagram

Monitoring the run

Close monitoring is essential throughout the run.  A live CCD camera with a macro lens or attached to a microscope, and hooked to a monitor, makes things a lot easier.  You basically need to ensure that all the tips are printing well on all the slides.  It’s sometimes necessary to use the reload feature if some tips are unable to print till the end of the platter.  Often, the first few slides have larger spots; this is normal.  If they begin running in to each other a lot in the beginning of the platter, you might want to turn on the blot pad feature.  Check the level of liquid in the sonicator every few plates.  In case of printing problems, you can stop the run after making note of the plate and load number.  Then remove the first slide, put in a fresh one, and do a few test prints (with test print DNA) and diagnose the problem.  Is a single tip printing large spots?  The tip could be blunted or the slot could be misshapen or something could be clogging the tip.  Look under the scope and try cleaning it gently.  Has a tip stopped printing? Make sure it continues to slide smoothly up and down in the holder and that it’s touching the slide on the test print.  Are all the tips misbehaving?  It could be due to salt or corrosive build up on the tips.  You can move the stage to the right to immerse the tips briefly in warm water, and then inspect them.  After this make sure the tip holder bottom holes are dry and free of crud.  Are some isolated slides on the platter missing spots?  Maybe the Z-height has not been corrected for those positions. 


Remember, many seemingly disastrous runs are salvageable.  If a bad tip problem crops up in the middle of a plate, and you lose a lot of spots on the array, you can print that plate again at the end of the run if you’ve managed to fix the problem. It’s usually not worth ‘filling in holes’ on the array as you have to keep careful track of which slides are missing which spots and so on.  It’s far easier to reprint an entire plate at the end, which is why it’s good to leave room for a few extra plates.  When you stop and start up again, make sure that the printing starts exactly where you stopped by entering in the appropriate values for the current plate, slide and load in the Print window.  Stopping a print run to catch some sleep at night is unseemly.  Tag team instead and carry on the through the night.  You’ll get the arrays faster that way.


Ending the run

Once all plates are printed, you can begin another run immediately with the same print plates.  If you are not up to that (or if you don’t have enough fresh slides), it's best to dry down the plates in the speedvac and seal them for the next run.  The first thing though, is to remove the slides and clean the tips.  Carefully peel the tape off the slides without touching the surface of the arrays.  It helps to have an extra person to hold down one edge of the slides while the other peels tape from the other end.  Alternatively you can use the edge of a long strip of metal like a ruler to hold down the slides when the tape is pulled off.  Some people like to label the slides at this point, especially if using stick on labels.  Put the printing tips in fresh warm water in the sonicator and clean for a few minutes.  Inspect tips for any corrosive or salt crud inside the slot.  Cleaning with ethanol may also help.  Store tips dry.

Chapter 3

Microarray Hybridizations

Harvesting Cells from Liquid Culture


This is a very simple protocol for collecting yeast cells from liquid culture, basically just chilling, spinning, and freezing.



Collection tubes

Pre-chilled centrifuge

Liquid nitrogen




(1)        Fill a collection tube with ice, so that there is as much ice in the tube as there will be sample.

(2)        Pour the sample into the tube.  Mix by shaking.

(3)        Pellet the cells by centrifugation.

(4)        Decant, and freeze pellet by submersion in liquid nitrogen.

(5)               Store at –80 C.



There are many different variations on this protocol.  The key elements are to stop gene expression quickly, by cooling or freezing.  The method also works well if a fast spin (<3 min) is used and the samples remain at their experimental temperature.  Generally longer spins with ice to chill the cells are the easiest way to go.

RNA Preparation


RNA preparation is one of the most important steps in expression studies.  RNA that is undegraded and free from contaminants will work well when labeled cDNA is generated later.  We have found that acid phenol methods are robust and easy.  The general goal is to quickly take frozen cells and add hot phenol which will break open the cell wall and separating nucleic acids from unwanted cellular components.  It is generally helpful to perform the extractions multiple times to ensure that high quality RNA is prepared. 



Water saturated phenol             Solid phenol, melted and saturated with water

Sodium acetate buffer                           50 mM sodium acetate, 10 mM EDTA pH 5.0

10% SDS


95% ethanol

70% ethanol


Water bath at 65 C


3M sodium acetate pH 5.3

de-ionized water



(1)        Fill phenol resistant tubes with 10ml phenol.

(2)        Add 9ml sodium acetate buffer and 1ml 10% SDS.

(3)        Immerse in 65 C bath, until heated.

(4)        Warm frozen cell pellets to ~ 0 C.

(5)        Add 1ml phenol/buffer mixture to each cell pellet, returning each mixture to the larger tube and then vortex for 10s.

(6)        Heat at 65 C for 7-15 min, vortexing for 10s every 1-2 min.

(7)        Centrifuge to separate phases (about 10 min, 3,000 g).

(8)        Remove aqueous to second tube containing 10ml phenol.

(9)        Vortex briefly, separate phases again.

(10)      Remove aqueous to third tube containing 5ml phenol, 5ml chloroform.

(11)      Vortex briefly, separate phases again (15 min, 3,000g).

(12)      Transfer aqueous to a fourth tube.

(13)      Add 1/10th volume sodium acetate, and either one volume of isopropanol, or two volumes of ethanol.

(14)      Centrifuge to precipitate RNA (~30 min, 3,000g).

(15)      Wash RNA pellet with 70% ethanol.

(16)      Dry pellet at room temperature.

(17)           Re-suspend RNA in water.  Store at –80 C.



Again there are many variations in protocols.  Increased yield can be obtained by lengthening the hot phenol step, or by pelleting the cell debris and reextracting it with more hot phenol.  Phase-lock tubes (5’ -> 3’ inc) are a useful way to make the separations easier (and reduce them).  It is relatively important to quickly get the frozen cells into the phenol/buffer mixture to prevent gene expression changes.

Amino-allyl Labeling of cDNA by Reverse Transcription      


The amino-allyl based labeling procedure is far more cost effective, and yields fluorescent probes that are much brighter than can be obtained by direct incorporation.  There protocol is somewhat longer, but the efort is worthwhile. 


I. RT Reaction


Oligo dT/Random Prime RNA:




Oligo dT:pdN6



Total Vol. of RNA








Incubate RNA and oligo dT at 70° C for 10 min.

Chill on ice 10 min.


1x dNTPS:


500uM each dA, dC, dG

200uM aadUTP

300uM dTTP


50x recipe: FOR 2:3

10uL each dA,dG,dC

4uL aa-dUTP


6uL dT


cDNA synthesis                                                                                 






5x buffer




50x aa dUTP/dNTPs








SuperScript II












42 degrees for 2 hours





II. Hydrolysis

Add:     10ul 1N NaOH

                        10ul .5M EDTA

            Incubate: 15 min. at 65°C.


Neutralize: 25ul 1M Tris pH 7.4


III. Cleanup

To continue with the amino-allyl dye coupling procedure all Tris must be removed from the reaction to prevent the monofunctional NHS-ester Cye dyes from coupling to free amine groups in solution. 


            Fill one Microcon 30 concentrator with 450 ul water. 

Add neutralized reaction.

Spin at 12K for 8 minutes.

Dump flo-thru.

Repeat process 2X, refilling original filter.


Dry eluate in speed vac.


IV. Coupling

Resuspend cDNA pellet in 4.5ul water.

            Resuspend monofunctional NHS-ester Cy3 or Cy5 dye:

If using aliquot, resuspend in 4.5ul .1M NaBicarbonate Buffer pH 9.0


If using fresh tube of Cy3/Cy5, resuspend entire tube in 72ul water.

Aliquot 4.5ul x 16 tubes and dry in speed vac.

Resuspend aliquot in 4.5ul .1M NaBicarbonate Buffer pH 9.0 as above.


Mix dye and cDNA.

Let incubate 1 hour at RT in dark.


V. Quenching and Cleanup

Before combining Cy3 and Cy5 samples for hybridizations, the reactions much be         quenched to prevent cross-coupling.


                   Add 4.5ul 4M hydroxylamine.

                   Let reaction incubate 15 min. at RT in dark.


To remove unicorporated/quenched cye dyes proceed with Qia-quick PCR purification kit.


                   Combine Cy3 and Cy5 reactions.

                   Add 70ul water.

Add 500ul Buffer PB.

                   Apply to Qia-quick column and spin at 13,000 rpm in for 30-60 sec.

                   Aspirate off flo-thru.

                   Add 750ul Buffer PE and spin 30-60 sec.

                   Aspirate off flo-thru and repeat.

Aspirate flo-thru and spin for 1 min. at high speed to dry column.

                   Transfer to fresh epp. tube

                   Add 30ul Buffer EB to center of filter and let sit 1 min. at RT.

                   Spin at 13,000 rpm for 1 min.

                   Repeat elution step again.


VI. Hyb. Prep.

Dry down Qia-quick eluate in speed vac.

Bring volume to 18 ul with water.


Add:      3.6ul 20X SSC

        1.8ul polyA(10mg/ml)

             Optional:  filter in Millipore 0.45micron spin column.

Add:      0.54ul 10% SDS.

             Incubate reaction at 100° C for 2 min.

             Apply to prepared microarray.


Probe preparation by direct Cy-dNTP incorporation


Preparing labeled cDNA from RNA for expression studies may seem complicated but it is one of the simplest molecular techniques.  The major necessary components, RNA, RT, and fluor-dNTP are relatively straightforward to use.  Generally, RNA and a primer that will be used to initiate reverse transcription are mixed, heated (to allow annealing), and chilled.  A cocktail of enzyme, buffer, etc… is made for each of the fluors, and added to the RNA mixture.  The RT reaction is allowed to proceed, until the two probes are prepared to be mixed, when EDTA (and NaOH) is added to prevent DNA production.  The next key step is that the unincorporated fluor needs to be removed by using a centricon filter (which retains molecules with a certain MW or greater, 30 kD works well).  This purified cDNA is ready to be used as a probe.




Olgio dT 1.5 mg/ml (~18mer)

65 C heat block

42 C heat block

100 C heat block

0.1 M DTT

First strand buffer                                

Reverse transcriptase (Superscript II)   

Labeling mix                                         25 mM dATP, 25 mM dCTP, 25 mM dGTP, 15 mM dTTP



Stop solution                                        1 N NaOH, 0.1 M EDTA

Neutralization solution                           1 N HCl






(1)        Aliquot 15 ug of total yeast RNA or up to 2 ug of polyA+ RNA into each reaction tube (0.5 ml or 1.5ml).

(2)        Add 6 ug (for total) or 4 ug (for polyA+) olgio-dT.

(3)        Heat 65 C, 1 min.

(4)        Chill on ice.

(5)        Make a master mixture of:

                        6 ul first strand buffer

                        3 ul 0.1 M DTT

                        0.6 ul labeling mix

                        3 ul Cy-dUTP

                        2 ul reverse transcriptase

            for each reaction. 

(6)        Add 14.5 ul of master mix to each RNA/oligo-dT sample.

(7)        Incubate at 42 C for two hours.

(8)        Add 1.5 ul stop solution.

(9)        Heat 10 min 65 C.

(10)      Add 1.5 ul neutralization solution.

(11)      Mix differentially labeled sample in a centricon-30.

(12)      Add 400 ul of TE.

(13)      Centrifuge 9 min at 14,000g.

(14)      Empty flow through and add 500 ul water.

(15)      Centrifuge until 5-10 ul remains above the membrane

(16)      Collect labeled cDNA.



Several different steps can be excluded or included from the protocol presented here.  Degradation of the RNA by adding the stop solution is probably unnecessary, as is neutralizing the stop solution directly.  A vast excess of TE will do the trick.  One step that seems important is to keep the reactions to 30 ul, changing the volume of the reaction can affect the heating/cooling steps resulting in changes in the representation of the cDNA produced or available for hybridization.  Probes can usually be stored overnight in the refrigerator.
Microarray hybridizations



Labeled cDNA


10% SDS

2.5% SDS

20 mg/ml yeast tRNA


Cover slips

Hybridization chambers




(1)        Add 2 ul of 20X SSC to labeled cDNA mixture

(2)        Add 1 ul of tRNA.

(3)        Add 1 ul of 2.5% SDS

(4)        Bring final mixture to 14 ul.

(5)        Insert microarray into chamber, placing 10 ul water at one end of the chamber.

(6)        Pipet cDNA mixture onto center of array.

(7)        Place a cover slip over the array, prevent bubble formation.

(8)        Close chamber, place in 65 C bath for 4-24 hrs.

(9)        Remove chamber from  bath and array from chamber.

(10)      Carefully place array into slide holder in 1X SSC, 0.2% SDS.

(11)      Shake slowly until cover slip falls off.

(12)      Plunge slide ~20 times, transfer to 0.4X SSC, plunge ~20 times, transfer to 0.2X SSC, plunge ~20 times.

(13)           Spin array dry in centrifuge at 600 rpm.



There are more tricks to this protocol than a dog could ever learn.  Everybody does it differently and it doesn’t seem to make a substantial difference.  Some of the interesting differences are…

(1)               cover slips- glass, plastic, beveled edges…

(2)               hybridization volume- 10, 12, 14, 20 ul or more.

(3)               Various blocking agents (repetitive DNA, oligo dA…)

(4)               Cover slip placement (dropping, tweezers…)