<?xml version="1.0" encoding="ISO-8859-1" ?>
<protocol_list>
<protocol xml:space="preserve"
  title="Frog Oocyte extraction"
  type="tissue_or_sample">
   1.0: What you need to do:

     Consult the frog database and determine which frog is to be used 
     Make sure to locate the following essential items BEFORE you start the surgery 
              Ca-free OR2 solution (OR2 solution is by default Ca-free), at least a couple of hundred ml 
              Collagenase, 80 mg 
              Two Corning 50 ml culture tubes 
              Surgical tools 
              A sterile needle 
              A packet of chromic gut sutures 
     1: Surgery

     Wear gloves. 
         Prepare MS222 solution 
          Locate the MS222 jar with its lid 
              0.9 to 1.5 grams of MS222 (3-aminobenzoic acid ethyl ester) per 300 ml 
               MS222 is stored in the freezer Box #5 
          Do not reuse the MS222 solution. This could contribute to disease problems. 
         Take the MS222 Jar and an ice bucket to the frog room. Place the correct frog into the jar. Put some ice in the ice
     bucket. Note that frog identify. This will be recorded in the surgery database 
         Wait for the frog to go under. This may take 10 to 20 minutes. While frog is in MS222 (20 minutes), prepare
     collagenase solution (see below) 
              Locate two Corning 50 ml tubes 
              Measure 40 mg (2 mg/ml) of Sigma Collagenase stored at -20 deg C into each Corning 50 ml tube 
              Add 20 ml of Ca-free OR2 solution into each tube 
         Fill a few Petri dishes with Ca-free OR2 solution 
          We will place the eggs initially to rinse out the excess blood etc. 
         Rinse surgical tools (forceps and scissors) in 95% ethanol. 
         Get out a 23 or 21 gauge needle and a packet of chromic gut sutures. 
         Put ice in a silver-color aluminum tray. 
         Remove frog and place on ice. Cover head with ice. Poke feet with needle to be sure frog is out (20 minutes
     should be enough time). 
         Insert needle under skin layer on either side (if frog has had a previous surgery, go to opposite side, if frog has
     had multiple surgeries choose side that is the best healed). Make an incision. Grab muscle layer with tissue forceps
     and make another incision. 
         Tease oocyte lobes carefully out. Cut and place in Petri dishes with OR2. Take many cells as it is better to be
     choosy about which oocytes to inject. 
     Post Op Care of the frog 
              Put two to three sutures in the muscle layer and two to three sutures in skin layer. 
              Put frog in Rubbermaid tub with lid (holes punched in) with a small amount of ddH2O and allow to recover
          2-3 hours in lab. 
              Using forceps, carefully tease apart lobes, releasing clumps of oocytes then place oocytes in the collagenase
          solution. 
         Place the cells in the two Corning tubes with collagenase and close the lids tightly. 
         Place on orbiter at almost 45 to 50 RPMs for 1 hour (or more) 
          It is best to check every 15 minutes and shake dish to see if cells are dispersing. 
         Wash cells with OR2. 
          Fill the tube with OR2 to the top and gently swirl, and then decant the solution. 
          Repeat this at least 4 or 5 times until the solution is very clear. It is very important to wash the cells very well
          to rinse away collagenase. 
         Wash cells with ND96 for a few times. 
         Fill a few petri dishes with ND96 and transfer the cells to the petri dishes using a fire-polished blunt, large
     diameter pasture pipette. 
          You want a single layer of cells. You may need to use more petri dishes if you have many cells. 
         Fill a 60 mm petri dish with ND96. 
          You will collect &quot;GOOD&quot; cells in this dish 
         Using a dissection microscope, select good oocytes using a fire-polished glass pasture pipette with an
     approximately 2 mm diameter. 
         Record the surgery and the egg appearance in the frog database. 
         PUT THE FROG BACK IN THE ORIGINAL TANK 
          This has been neglected a few times before </protocol>
<protocol xml:space="preserve"
  title="Establishment of a Primary Culture, Chick Embryo"
  type="growth"
>
Materials 

     Chick embryo (approximately 8 days old)
     70% (v/v) ethanol for swabbing
     Sterile scissors, forceps and probes
     Sterile petri plates
     Phosphate buffered saline (PBS)
     Trypsin, cold sterilized in a 125 ml sterile erlenmeyer containing a magnetic stirring bar
     Minimum Essential Medium
     Fetal Calf Serum
     Clinical centrifuge with sterile capped centrifuge tubes
     Culture flasks
     Inverted phase contrast microscope (Optional) 

Procedure 8 

   1.Candle an 8 day old egg to ensure that it is alive. This is easily accomplished by holding the egg in front of a bright light source; the embryo can
     be seen as a shadow. Circle the embryo with a pencil. 

   2.Place the egg in a beaker with the blunt end up, and wash the top with a mild detergent, followed by swabbing with ethanol. 

   3.Carefully puncture the top of the egg with the point of a pair of sterile scissors and cut away a circle of shell, thus exposing the underlying
     membrane (the chorioallantois). 

   4.With a second pair of sterile scissors, carefully cut away and remove the chorioallantoic membrane, exposing the embryo. 

   5.Identify and carefully remove the embryo by the neck, using a sterile metal hook or a bent glass rod, and place the embryo in a 100mm petri dish
     containing phosphate buffered saline (PBS). Wash several times with PBS by transferring the embryo to fresh petri plates. After removal of all
     yolk and/or blood, move the embryo to a clean dish with PBS. 

   6.Using two sterile forceps, remove the head, limbs, and viscera. Be sure to remove the entire limb by pulling at the proximal end. Move the
     remaining tissues of the embryo to yet another dish and wash with PBS. 

   7.Mince the embryo finely with scissors and transfer the minced tissue to a flask containing PBS. Allow the tissue pieces to settle. 

   8.Remove the PBS with a sterile pipette and add 25 ml of trypsin, a
proteolytic enzyme. Stir the solution gently at 37 deg. C for 15-20 minutes. 

   9.Allow the larger, undigested tissue pieces to settle and decant the supernatant into an equal volume of Minimal Essential Medium (MEM) + 10%
     Fetal Calf Serum (FCS). FCS contains protease inhibitors which will inactivate the trypsin. 

  10.Centrifuge the cells in MEM at 1000 rpm for 10 minutes in a standard clinical centrifuge. Remove the supernatant and resuspend the pellet in 25
     ml of fresh MEM + 10% FCS. 

  11.Remove 0.1 ml of the culture and determine cell concentration and viability as directed in the previous section. 

  12.Seed two 25 cm plastic culture flasks containing 25 ml of MEM + 10% FCS to a final concentration of 10 cells/ml. 

  13.Label and place your cultures in the tissue culture incubator at 37 deg.  C and examine daily for cell density and morphology. 

  14.Note any changes in the color of the media. Tissue Culture media has a pH indicator (Phenol Red) added in order to check on the growth of cells.
     The media initially is a cherry red (with slight blue haze) and turns orange and then yellow as the cells grow, thereby reducing the media. Should
     this color change occur within 24 hours, the culture is most likely contaminated and should be disposed of. 

  15.Examine the cultures using an inverted phase contrast microscope. This will allow observation of the cells without opening or disturbing the
     growth. 

  16.Make cell density determinations at 10 X magnification using a square ocular grid, as explained in Chapter One for the determination of area. 

  17.Plot the cell density on a log scale vs. time of culture. 

  18.Diagram the shape of the cells at each phase. 

Notes 

The cultures will develop differently than the suspension cultures. The viable cells will grow out of the trypsinized pieces of tissue and will remain in
contact with the bottom of the culture flask. They will continue to divide and migrate until the entire bottom of the flask is covered with a single layer of
cells (contact inhibition and the formation of a monolayer). 
</protocol>
<protocol xml:space="preserve"
  title="Protocol: Growth of PC12 cells"
  type="growth"
>

     Medium:    

          DME, high glucose (4500 mg/L) 
          5% fetal bovine serum (Gemini) 
          10% horse serum (Gemini) 
          1:100 dilution of Pen/strep (UCSD core) 
          stock solution:Pen G, 10,000 U/ml    
                      Strep sulfate, 10,000 µg/ml    
          Mix and filter sterilize  

     Grow at 37 deg. C, water jacketed incubator, 6% carbon dioxide    

     Cells double about every 48h    

     Split as necessary    

     After rinsing cells of medium, suspend them simply by pipetting (no trypsin needed).    

     Split at confluence usually, or whenever you wish.    

     From 1:4 to 1:10 as needed for your experiment.    

     6 cm plates are good for transfections and secretion studies.    

     Split onto poly-L-lysine-coated plates for secretion or transfection studies.    
     Change medium every 2 days, unless the cells are very dilute (high split).    </protocol>
<protocol xml:space="preserve"
  title="Stanford: Total Bacterial RNA Labeling with Random Hexamers."
  type="cdna_and_labeling"
>Arkady Khodursky, Ph.D.
Email: khodursk@cmgm.stanford.edu
Updated: 2/10/99

20-25ug of total RNA in 10-14 ul of diH20 should be purified using  RNA kit
or hot phenol extraction. After obtaining RNA sample it is important to get
rid of residual DNA with RNase-free DNAse followed by 1x PHOH,
1xPHOH/CHCl3, 2x CHCl3 extraction. Extent of RNA purity achieved by
PHOH-CHCl3 extraction is absolutely critical for successful labeling.

Above indicated amount of RNA should be mixed with 500ng of random pdN6
primer on ice in total volume of 15 ul.

Incubate at 65 C for 10&apos;.

Incubate on ice for 2&apos;.

Add 3 ul of  1 mM FluoroLink Cy3 (orCy5) dUTP (Amersham Cat# PA53022).

Add 11.6 ul an mix everything by pipeting 3-4 times on ice of reverse
transcription mix:
0.1 DTT - 3ul
5x 1st strand buffer - 6 ul
dNTPs (25 mM dATP, 25 mM dCTP, 25 mM dGTP, 10 mMdTTP) - 0.6 ul
SuperscriptII - 2 ul
(All from GibcoBRL Cat.# 18064-014)

Incubate the complete mixture for 10&apos; at RT.

Transfer to 42 C for 110&apos;.

Stop by adding 1.5 ul of 1n NaOH for 10&apos; at 65 C.

Neutralize with 1.5 
http://cmgm.stanford.edu/pbrown/protocols/index.html
</protocol>
<protocol xml:space="preserve"
  title="Stanford: Preparation of Fluorescent DNA Probe from mRNA: Human"
  type="cdna_and_labeling"
>Updated: August 9, 1999

   1.To anneal primer, mix 2ug of mRNA with 2ug of a regular or anchored oligo-dT primer in a total volume of 15ul: 
                 Cy3 
                       Cy5 
      mRNA 
                 2 ug 
                       2 ug 
      Oligo-dT 
                 2 ug 
                       2 ug 
                            (Anchored: 5&apos;-TTT TTT TTT TTT TTT TTT TTV N-3&apos;) 
      Total volume: 
                 15 uL 
                       15 uL 


   2.Heat to 70 deg. C for 10 min and cool on ice. 
   3.Add 15 uL of reaction mixture each to Cy3 and Cy5 reactions:

      Reaction mixture 
                                         . . . . . . . . . . 
                                                    Unlabeled dNTPs 
                                                                  Vol. 
                                                                        Final conc. 
      5X first-strand buffer* 
                                   6.0 
                                                    dATP (100 mM) 
                                                                  25 uL 
                                                                        25 mM 
      0.1M DTT 
                                   3.0 
                                                    dCTP (100 mM) 
                                                                  25 uL 
                                                                        25 mM 
      Unlabeled dNTPs 
                                   0.6 
                                                    dGTP (100 mM) 
                                                                  25 uL 
                                                                        25 mM 
      Cy3 or Cy5 (1 mM, Amersham) 
                                   3.0 
                                                    dTTP (100 mM) 
                                                                  15 uL 
                                                                        15 mM 
      Superscript II (200 U/uL, Gibco BRL) 
                                   2 uL 
                                                    ddH2O 
                                                                  10 uL 
                                   ----- 
                                                                  ----- 
      Total volume: 
                                   15 uL 
                                                    Total volume: 
                                                                  100 uL 


     * 5X first-strand buffer: 250 mM Tris-HCL (pH 8.3), 375mM KCl, 15mM MgCl2)

   4.Incubate at 42  deg.  C for 1.5-2hrs. 
   5.Degrade RNA by addition 15ml of 0.1N NaOH, and incubation at 70 deg. C for 10 min. 
   6.Neutralize by addition of 15ml of 0.1N HCL, and bring the volume to 500ml with TE (10mM Tris, 1mM EDTA). 
   7.Add 20 mg of Cot1 human DNA (Gibco-BRL). 
   8.Purify probe by centrifuging in a Centricon-30 micro-concentrator (Amicon). Look for nice concentration of the &quot;colored probe&quot; in the
     centricon. 
   9.Combine the separate concentrated probes (Cy3 and Cy5) into a fresh centricon, bring to a volume of 500ml with TE, and concentrate
     again to a volume of less than 7ml. 
  10.Add 1mL of 10m g/ml polyA RNA (Sigma, #P9403) and 1 ml of 10mg/ul tRNA (Gibco-BRL, #15401-011). 
  11.Adjust volume to 9.5ml with distilled water. 
  12.For final probe preparation add 2.1ml 20XSSC (1.5M NaCl, 150mM NaCitrate (pH8.0)) and 0.35ml 10%SDS. Final probe volume can
     be adjusted to between 12ml and 15ml according to preference (Larger volumes make manipulation of the covers slip easier). Be sure to
     adjust salt concentration proportionately. 
  13.Denature probe by heating for 2 min at 100 deg. C, and incubate at 37 deg. C for 20-30 min. 
  14.Place on the array under a 22mm x 22mm glass cover slip. 
  15.Hybridize at 65 deg. C for 14 to 18 hours in a custom slide chamber with humidity maintained by a small reservoir of 3XSSC. 
  16.Wash arrays by submersion and agitation for 2-5 min in 2X SSC with 0.1%SDS, followed by 1X SSC, and 0.1X SSC. 
  17.&quot;Spin dry&quot; by centrifugation for 2 min in a slide rack in a Beckman GS-6 tabletop centrifuge in Microplus carriers at 650 RPM for 2
     min. 
http://cmgm.stanford.edu/pbrown/protocols/index.html
</protocol>
<protocol xml:space="preserve"
  title="Stanford: Preparation of Fluorescent DNA Probe from mRNA: Yeast"
  type="cdna_and_labeling"
>
Last updated: 3/29/99 


 Materials for 2 reactions (Cy3 and Cy5) 
                               Qty 
                                          Order info 
 Heat blocks, 42C and 70C 
                               1 each 
 Oligo dT (2 ug/uL) 
                               5.0 uL 
 Superscript II RT kit 
                                          Gibco BRL #18064-014 
 50X dNTPs (25 mM; 10 mM dTTP)* 
                               1.25 uL 
 Cy3-dUTP (1 mM) 
                               3.0 uL 
                                          Amersham #53022 
 Cy5-dUTP (1 mM) 
                               3.0 uL 
                                          Amersham #55022 
 Microcon-30 filter 
                               3 
                                          Amicon #42410 
 TE (pH 7.4) 
                               ~5 mL 
 polyA DNA or RNA (10 mg/mL in TE) 
                               1.0 uL 
 20X SSC 
                               3.0 uL 
 Millipore filter, 0.45 um 
                               1 
                                          Millipore #UFC3OHVNB 
 . 
 *50X dNTPs stock 
                                          Final conc. 
 dATP (100 mM) 
                               25 uL 
                                          25 mM 
 dCTP (100 mM) 
                               25 uL 
                                          25 mM 
 dGTP (100 mM) 
                               25 uL 
                                          25 mM 
 dTTP (100 mM) 
                               10 uL 
                                          10 mM 
 ddH2O 
                               15 uL 
                               --------- 
                               100 uL 


 1. 
    PREPARE PRIMER-ANNEALED RNA: 
                                         Cy3 rxn 
                                                     Cy5 rxn 
    RNA (2 ug mRNA or 15 ug total) 
                                         12.9 (RNA #1) 
                                                     12.9 (RNA #2) 
    Oligo dT (2 ug/uL) 
                                         2.5 
                                                     2.5 
                                         --------- 
                                                     --------- 
                                         15.4 uL 
                                                     15.4 uL 
    Heat 10 min. @ 70C. Quick chill on ice. 
 2. 
    PREPARE RT COCKTAIL FOR BOTH RXNS: 
                                         X 1 
                                                     X 2.5 
                                                                  Final conc. 
    5X Superscript II buffer 
                                         6.0 
                                                     15.0 
                                                                  1X 
    DTT (0.1 M) 
                                         3.0 
                                                     7.5 
                                                                  10 mM 
    50X dNTPs 
                                         0.6 
                                                     1.25 
                                                                  500 uM; 200 uM dTTP 
    Cy3- or Cy5 dUTP 
                                         3.0 
                                                     --- 
                                                                  100 uM 
    Superscript II (200 U/uL) 
                                         2.0 
                                                     5.0 
                                                                  13 U/uL 
                                         --------- 
                                                     --------- 
                                         14.6 uL 
                                                     11.6 uL aliquots 

 3. 
     Add 3.0 uL Cy3 or Cy5 to respective primer-annealed RNAs. 
     Aliquot 11.6 uL of RT cocktail to each rxn for total volume of 30 uL. 
     Incubate 2 hr @ 42C. Place on ice. 
 4. 
     Place 500 uL TE (pH 7.4) each in two microcon-30 filters. 
     Add RT rxns to each microcon filter. Centrifuge 7 min. at top speed. 
     Optional: Repeat TE washes 1-2 times, or until all unincorporated dye is removed. 
 5. 
     Inspect filters. Centrifuge in 30 sec. intervals until volume is 10-20 uL. 
 6. 
     Invert filters into fresh tubes. Centrifuge 1 min. to harvest labeled cDNA. 
     Optional: For next day hybridization, store cDNA @ 4C. 
 7. 
     Mix together Cy3- and Cy5-labeled cDNAs. 
     Concentrate sample to ~10-12 uL using microcon filter or vacuum pump. 
 8. 
     Add 1 uL polyA DNA or RNA for non-specific hybridization. 
     Add 3 uL 20X SSC for total volume of 12-15 uL. 
 9. 
     Pre-wet millipore filter by adding 5 uL ddH2O. Centrifuge 1 min. at top speed. 
     Remove eluted water with pipet tip. 
 10. 
     Add probe to filter. (Pipet probe onto filter wall, not directly onto membrane.) 
     Centrifuge 1 min. at top speed. 
 =&gt; 
     PROBE IS NOW READY FOR HYBRIDIZATION. 
     REMEMBER TO ADD 0.3 uL SDS (10%) TO PROBE as stated in hybridization protocol.  
http://cmgm.stanford.edu/pbrown/protocols/index.html
</protocol>
<protocol xml:space="preserve"
  title="Stanford: Preparation of DNA Samples"
  type="tissue_or_sample">
Last updated: December 3, 1999 


YEAST ORF AMPLIFICATION USING RESGEN PRIMERS (new link)

                            One rxn 
                                    One 96-well plate 
                                    (100 rxns) 
 RESGEN SPECIFIC PRIMERS: 
                                                   Program: 92C (30&quot;), 56C (45&quot;), 72C (3&apos;30&quot;) / 36 cycles 
 Forward primer (20 uM) 
                            5 
                                    - 
 Reverse primer (20 uM) 
                            5 
                                    - 
 10X PCR buffer (Perkin Elmer) 
                            10 
                                    1000 
 MgCl2 (25 mM) 
                            8 
                                    800 
 100X dNTPs (25 mM each) 
                            1 
                                    100 
 Yeast genomic DNA (0.2 ug/uL) 
                            0.2 
                                    20 
 ddH2O 
                            70.45 
                                    7045 
 AmpliTaq (5 U/uL) 
                            0.35 
                                    35 
                            ---- 
                                    ---- 
                            100 uL 
                                    88 uL aliquots 
 . 
 RESGEN UNIVERSAL PRIMERS 
                                                   Program: 92C (30&quot;), 56C (45&quot;), 72C (3&apos;30&quot;) / 25 cycles 
 Forward primer (30 uM) 
                            3 
                                    300 
                                                   5&apos; gga att cca gct gac cac c 3&apos; 
 Reverse primer (30 uM) 
                            3 
                                    300 
                                                   5&apos; gat ccc cgg gaa ttg cca tg 3&apos; 
 10X PCR buffer (Perkin Elmer) 
                            10 
                                    1000 
 MgCl2 (25 mM) 
                            8 
                                    800 
 100X dNTPs (25 mM each) 
                            1 
                                    100 
 Yeast ORF DNA (0.5 ng/uL) 
                            4 
                                    - 
 ddH2O 
                            73.6 
                                    7360 
 AmpliTaq (5 U/uL) 
                            0.4 
                                    40 
                            ---- 
                                    ---- 
                            100 uL 
                                    88 uL aliquots 




GEL ELECTROPHORESIS OF PCR REACTIONS (quality control)

 Agarose gel: 
                1% agarose, 1X TAE, 0.5 mg/mL ethidium bromide 
 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). 
 2. 
     Pour gel with four combs of 26 wells each. 
 3. 
     Add 1 uL 6X loading dye to PCR reactions in PCR plates. 
 4. 
     Load 6 uL DNA size ladder in lane #1 of each row. 
 5. 
     Using a 12-channel pipettor, load samples A1-A12 into alternating lanes 2, 4,..., 24. 
 6. 
     Load samples B1-B12 into alternating lanes 3, 5,..., 25. 
 7. 
     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. 
 8. 
     Run at 70-80V until the first dye band (XC FF) is halfway to the next row of wells. 
 9. 
     Take a high (~1&quot;) and low (~6/30&quot;) exposure photographs. 
     Compare to predicted ORF sizes and for the presence of significant doublets. 
 10. 
     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. 




DNA PRECIPITATIONS =&gt; See Comparison of PCR Cleanup Protocols

 1. 
     Transfer PCR reactions to 96-well U-bottom tissue culture plates (Costar #3790). 
     Transfer 3 uL back to PCR plates for check gels (see above). 
 2. 
     Dry down volume in U-bottom plates to ~50 uL. (High temp. speec vacuum for 1 hr for 8 plates.) 
     The drying will be uneven, with wells around the edges experiencing more evaporation. 1 hr gets all the wells down to ~50 uL. 
 3. 
     Add 1/10 vol. 3M sodium acetate (pH 5.2) + 2.5 volumes ethanol. 
     Store at -20C for a few hours to overnight. 
 4. 
     Centrifuge in Sorvall RC-3B at 3500 rpm for 1 hr (H-6000A rotor, RCF = 3565 g). 
 5. 
     Remove supernatant with 12-channel aspirator (Wheaton/PGC Scientifics #851388). 
 6. 
     Add 100 uL of ice-cold 70% ethanol and centrifuge again for 30 min. 
 7. 
     Dry the pellets in speec-vac for 10 min. 
 8. 
     Resuspend DNA in 100 uL dH2O overnight. 
 9. 
     Transfer in 10 uL aliquots to 384-well plates (USA Scientific #2802-0384 or Corning Costar #6502) to make 10 duplicate print plate sets. 
 10. 
     Dry down print plate sets in speed vac. 
     Tightly seal plates with aluminum foil (R.S. Hughes #425-3) for long-term storage at room temperature. 
 11. 
     Before use, resuspend one print set in 4 uL 3XSSC overnight. 
 12. 
     Spot DNA onto polylysine slides with 16-tip or 32-tip arrayer. 
     Dry down used print plates for storage until next use. (One set of print plates can be used multiple times.) 
http://cmgm.stanford.edu/pbrown/protocols/index.html
</protocol>
<protocol xml:space="preserve"
  title="Stanford: Preparation of Fluorescent DNA Probe from mRNA: Human (Ash/Max protocol; 8/3/00)"
  type="hybridization"
> 
  1.To anneal primer, mix 2ug of mRNA with 4ug of a regular or anchored oligo-dT primer in a total volume of 10ul: 
       
                      Cy3 
                           Cy5 
      mRNA (1 g/l)
                      2 l 
                           2 l
      Oligo-dT (4 g/l)
                      1 l 
                           1 l 
                               (Anchored: 5&apos;-TTT TTT TTT TTT TTT TTT
                               TTV N-3&apos;) 
      ddH2O (DEPC)
                      7 l
                           7 l
                                
      Total volume: 
                      10 l 
                           10 l 
                                

   2.Heat to 65oC for 10 min and cool on ice. 
   3.Add 10 uL of reaction mixture each to Cy3 and Cy5 reactions: 
      Reaction mixture 
                           l 
                               . . . . . 
                                    Unlabeled dNTPs 
                                                  Vol. 
                                                         Final conc.
      5X first-strand buffer* 
                           6.0 
                                
                                    dATP (100 mM) 
                                                  25 uL 
                                                         25 mM 
      0.1M DTT 
                           3.0 
                                
                                    dCTP (100 mM) 
                                                  25 uL 
                                                         25 mM 
      Unlabeled dNTPs 
                           0.6 
                                
                                    dGTP (100 mM) 
                                                  25 uL 
                                                         25 mM 
      Cy3 or Cy5 (1 mM, Amersham) 
                           3.0 
                                
                                    dTTP (100 mM) 
                                                  10 uL 
                                                         10 mM 
      Superscript II (200 U/uL, Gibco
      BRL) 
                           2.0
                                
                                    ddH2O 
                                                  15 uL 
                                                          
      ddH2O (DEPC)
                           5.4
                                
                                     
                                                  ----- 
                                                          
      Total volume: 
                           20 l 
                                
                                    Total volume: 
                                                  100 uL 
                                                          


     * 5X first-strand buffer: 250 mM Tris-HCL (pH 8.3), 375mM KCl, 15mM MgCl2)

   4.Incubate at 42oC for 1 hr. 
   5.Add 1 l SSII (RT booster) to each sample. Incubate for an additional 0.5-1 hrs. 
   6.Degrade RNA by addition 15ml of 0.1N NaOH, and incubate at 70oC for 10 min. 
   7.Neutralize by addition of 15ml of 0.1N HCl, and bring the volume to 400ml with TE (10mM Tris, 1mM EDTA). Combine both probes into 1 centricon. [WASH 1] Spin
     7 min. at 14kRPM in a Centricon-30 micro-concentrator (Amicon). If repurification of cy dye flow-through is desired, do not combine probes until Wash 2. 
   8.WASH 2: Add 450 ul 1X TE and spin 7 min. at 14kRPM. 
   9.WASH 3: Add 450 ul 1X TE, 20 mg of Cot1 human DNA (Gibco-BRL), 2mL of 10m g/ml polyA RNA (Sigma, #P9403) and 2 ml of 10mg/ul tRNA (Gibco-BRL,
     #15401-011). Spin 7 min. at 14 kRPM. Look for nice concentration of the &quot;colored probe&quot; in the centricon. Try to concentrate to a volume of less than 10 ml. This
     volume is attained when the center of the membrane is dry and the probe forms a ring of liquid at the edges of the membrane. 
   10.Invert the centricon into a clean tube and spin briefly at 14kRPM to recover the probe. 

     For 22 mm x 40 mm arrays:....

   11.Adjust volume to 20ml with 1X TE. (12 ml for 22 mm x 22 mm arrays) 
   12.For final probe preparation add 4.25l 20XSSC (3.0 M NaCl, 300 mM NaCitrate (pH8.0)) and 0.75l 10%SDS (2.55 ml 20X SSC and 0.45 ml 10% SDS for 22 mm x 22 mm
     arrays). When adding the SDS, be sure to wipe the pipette tip with clean, gloved fingers to rid of excess SDS. 
   13.Denature probe by heating for 2 min at 100oC, and incubate at RT for 15-20 min. 
   14.Spin at 14kRPM for 10 min. 
   15.Place 24 l probe on the array under a 22mm x 40mm glass cover slip (12 ul for a 22 mm x 22 mm cover slip). 
   16.Hybridize at 65oC for 14 to 18 hours in a custom slide chamber with humidity maintained by a small reservoir of 3X SSC (spot around 3-6 l 3X SSC at each corner of
     the slide, as far away from the array as possible). 
   17.Ready washes in 250 ml chambers to 200 ml volume [2X SSC (Wash I) with 0.1%SDS (add this to the wash first), followed by 1X SSC (Wash II -- prepare two of
     these chambers), and 0.2X SSC (Wash III)]. Avoid adding excess SDS. The Wash I chamber and one of the Wash II chambers should each have a slide rack ready. 
   18.Blot dry chamber exterior with towels and aspirate any remaining liquid from the water bath. Be sure to aspirate in between the two chamber halves. 
   19.Unscrew chamber; aspirate the holes to remove last traces of water bath liquid. 
   20.Place arrays, singly, in rack, inside Wash I chamber (maximum 4 arrays at a time). Allow cover slip to fall, or carefully use forceps to aid cover slip removal if it
     remains stuck to the array. DO NOT AGITATE until cover slip is safely removed. Then agitate for ~15 sec. 
   21.Remove array by forceps, rinse in a Wash II chamber without a rack, and transfer to the Wash II chamber with the rack. This step minimizes transfer of SDS from
     Wash I to Wash II. 
   22.Wash arrays by submersion and agitation for 30 sec. in Wash II chamber, then Wash III (transfer the entire slide rack this time). 
   23.&quot;Spin dry&quot; by centrifugation in a slide rack in a Beckman GS-6 tabletop centrifuge at 600 RPM for 2 min 
   24.Scan arrays immediately.
 
http://cmgm.stanford.edu/pbrown/protocols/index.html
</protocol>
<protocol xml:space="preserve"
  title="Stanford: Hybridization of Arrays"
  type="hybridization"
>Last updated: August 10, 1999 


 Materials 
                      Qty 
                                 Order info 
 100C heat block 
                      1 
 65C water bath 
                      1 
 Hybridization chamber 
                      1 per array 
                                 Custom made by Die-Tech (see &quot;Drawings for custom parts&quot; on MGuide
                                 home page) or purchased at Corning Costar (#2551) 
 Cover slip, 22 mm X 22
 mm 
                      1 per array 
 3X SSC 
                      10 uL per
                      array 
 10% SDS 
                      ~1 mL 
 Fine-tip forceps 
                      1 
 Slide rack 
                      2 
                                 TBA 
                                                                                          &lt;= Each rack holds 8
                                                                                          slides 
 Slide chamber 
                      3 
                                 TBA 
                                                                                          &lt;= Each chamber holds
                                                                                          200 mL 
 ddH2O 
                      ~600 mL 
 20X SSC 
                      ~13 mL 


Directions for fluorescent DNA probes:

 1. 
    Final probe volume should be 12-15uL, at 4X SSC, containing competitor DNA etc. as required 
    (e.g., polydA, CoT1 DNA for a human cDNA array) 
 2. 
    Set up array in hybridization chamber. 
    Place 10 uL 3X SSC on edge of slide to provide humidity. 
 3. 
    Add 0.3 uL 10% SDS to probe. 
 4. 
    Boil probe for 2 min. Set aside several cover slips for next step. 
 5. 
    Pipet probe onto array, avoiding bubbles. 
    Using forceps to help, immediately lay down 22mm x 22mm cover slip over array, avoiding bubbles. 
    It is worth practicing this step several times with 10 uL water and a blank microscope slide. 
 6. 
    Close hybridization chamber and submerge in 65C water bath. 
 7. 
    Hybridize 4-24 hr. 
 8. 
    Prepare wash solutions #1 (1X SSC / 0.03% SDS), #2 (0.2X SSC), and #3 (0.05X SSC): 

         WASH 1 
                 WASH 2 
                         WASH 3 
 dH2O 
         190 
                 198 
                         200 
 20X SSC 
         10 
                 2 
                         0.5 
 10% SDS 
         0.6 
                 -- 
                         -- 
 Final vol.:
         200 mL 
                 200 mL 
                         200 mL 

 9. 
     Pour wash solutions in slide chambers, and place slide racks in washes #1 and #2. 
 10. 
     Disassemble hybridization chamber and quickly submerge array in wash #1. 
     (If the array is exposed to air while the cover slip starts to fall off, you may see high background fluorescent signal on the side of the array.) 
 11. 
     Let the array sit in wash #1 until the cover slip slides off. 
     Gently plunge slide rack up and down several times to wash the array; 
     be sure not to scratch the array with the loose cover slip. 
 9. 
     Manually transfer array to slide rack in wash #2 and rinse 2nd time. 
 10. 
     Move slide rack to wash #3 and rinse 3rd time. It is critical to remove all SDS. 
 11. 
     Centrifuge slides on microtiter plate carriers (place paper towels below rack to absorb liquid) for 5 min. @ 500 rpm. 
 12. 
     Scan array immediately. Notes: 
     - Be aware of Cy5 bleaching during scanning. 
     - We have found that arrays scanned immediately and two days after hybridization give nearly identical data (using GenePix 4000A
     scanner)  
http://cmgm.stanford.edu/pbrown/protocols/index.html
</protocol>
<protocol xml:space="preserve"
  title="Stanford: Preparation of Slides"
  type="other_protocol">
Last updated: October 6, 1999


 Materials 
                     Qty 
                                     Order info 
 Glass microscope slides 
                    60 
                          Gold Seal #3010 
 Slide rack 
                    2 
                          Shandon Lipshaw #121 (800-245-6212) 
                                                         &lt;= Each rack holds 30 slides 
 Slide chamber 
                    6 
                          Shandon Lipshaw #121 
                                                         &lt;= Each chamber holds 350 mL 
 ddH2O 
                    ~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 


 1. 
     Place slides in slide racks. Place racks in chambers. 
 2. 
     Prepare CLEANING SOLUTION: 
     Dissolve 70 g NaOH in 280 mL ddH2O. 
     Add 420 mL 95% ethanol. Total volume is 700 mL (= 2 X 350 mL); stir until completely mixed. 
     If solution remains cloudy, add ddH2O until clear. 
 3. 
     Pour solution into chambers with slides; cover chambers with glass lids. Mix on 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. 
 4. 
     Quickly transfer racks to fresh chambers filled with ddH2O. Rinse vigorously by plunging racks up and down. 
     Repeat rinses 4X with fresh ddH2O each time. It is critical to remove all traces of NaOH-ethanol. 
 5. 
     Prepare POLYLYSINE SOLUTION: 
     70 mL poly-L-lysine + 70 mL tissue culture PBS in 560 mL water. 
     Use plastic graduated cylinder and beaker. 
 6. 
     Transfer slides to polylysine solution and shake 15 min. - 1 hr. 
 7. 
     Transfer rack to fresh chambers filled with ddH2O. Plunge up and down 5X to rinse. 
 8. 
     Centrifuge slides on microtiter plate carriers (place paper towels below rack to absorb liquid) for 5 min. @ 500 rpm. 
     Transfer slide racks to empty chambers with covers for transport to vacuum oven. 
 9. 
     Dry slide racks in 45C vacuum oven for 10 min. (Vacuum is optional.) 
 10. 
     Store slides in closed slide box (plastic only, without rubber mat bottom) 
 11. 
     BEFORE PRINTING ARRAYS: 
     Check that polylysine coating is not opaque. 
     Test print, hyb and scan sample slides to determine slide batch quality. 
http://cmgm.stanford.edu/pbrown/protocols/index.html
</protocol>
<protocol xml:space="preserve"
  type="rna_prep"
  title="Stanford: Revised Protocol for FastTrack 2.0 mRNA Extraction Human Cells">
     Preparation of Cells

        1.Prepare or collect between 2x107 cells for each mRNA prep (will yield about 10-20µg of mRNA). If PBMCs from a whole blood sample
          are to be used, the sample should be prepared with Ficoll-Paque according to the manufacturer&apos;s protocol. The isolated PBMCs can then
          be used in this protocol.

        2.If cells are adherent, gently agitate and scrape cells from the surface of the flask with a cell scraper to suspend in the media. If cells are
          non-adherent, proceed to next step. 

        3.Place the suspension of cells in a 50ml conical tube, pellet at ~400xg for 5 minutes.

        4.Discard the supernatant, leave the pellet intact.

        5.Freeze the pellet in a dry ice/ethanol bath (or on liquid N2) until solid.

        6.Store at -70oC until ready for mRNA preparation.

     Modified FastTrack 2.0 Protocol

     Isolation of mRNA

        1.Add 15ml of Lysis Buffer (15ml Stock Buffer + 300 ul RNase/Protein Degrader) to the frozen pellet.

        2.The tissue homogenizer should be cleaned so that it is RNase-free. At a minimum will require immersion and homogenization in ddH2O
          and ethanol 3 times sequentially, may require boiling H2O, SDS, or 10% bleach in addition. An RNAse degrader such as RNAse Zap can
          also be used.

        3.Shear each sample with a 3-5 second run of the tissue homogenizer, essentially until the cell pellet is no longer visible. Clean
          homogenizer tip between runs.

        4.Incubate the sheared samples in Lysis Buffer at 45oC for 45 minutes.

        5.Add 950 µl 5M NaCl stock solution and mix.

        6.Shear DNA using a 21 gauge needle attached to a sterile 20cc syringe. This step involves drawing the lysate into the syringe through the
          needle and expelling it from the syringe, again through the needle. This step should be repeated 3 times.

        7.Add one tube of oligo(dT) cellulose to each sample.

        8.Incubate for 2 minutes at room temperature. Vortex the sample to resuspend the cellulose completely.

        9.Rock the tube gently in a horizontal position for 60-90 minutes at room temperature.

       10.Centrifuge the oligo(dT) slurry at 3,000 x g for 5 minutes at room temperature. Make sure the brake on the centrifuge is set at low for all
          centrifugation steps, as a high brake may disturb the pellet.

       11.Carefully aspirate and discard the supernatant, trying not to disturb the easily dispersed pellet.

     Washing Oligo(dT) Cellulose

        1.Resuspend oligo(dT) in 20ml Binding Buffer by vortexing.

        2.Centrifuge at 3,000 x g for 5 minutes at room temperature.

        3.Resuspend oligo(dT) in 10ml Binding Buffer by vortexing.

        4.Centrifuge at 3,000 x g for 5 minutes at room temperature.

        5.Resuspend oligo(dT) in 10ml Low Salt Wash Buffer by vortexing.

        6.Centrifuge at 3,000 x g for 5 minutes at room temperature.

        7.Repeat Low Salt Wash Buffer wash two more times.

        8.Resuspend oligo(dT) in 800µl Low Salt Wash Buffer using a 1-2ml serological pipette.

        9.Transfer about 800µl of the oligo(dT) slurry to a spin column seated in the included microcentrifuge tube.

       10.Centrifuge at 5,000 x g for 10 seconds at room temperature.

       11.Aspirate and discard the flow-through liquid in the microcentrifuge tube.

       12.Repeat the transfer and centrifugation steps until ALL of the oligo(dT) cellulose is in the spin column.

     Elution and Precipitation of the mRNA

        1.Place the spin column containing the oligo(dT) into one of the included clean microcentrifuge tubes.

        2.Heat elution buffer to 65oC in a bath (or about 10-15 seconds in a microwave).

        3.Resuspend the oligo(dT) in 200µl Elution Buffer (heated), using the pipette tip to gently swirl the cellulose, without puncturing the
          underlying spin column membrane.

        4.Centrifuge at 5,000 x g for 30 seconds at room temperature. Do not decant. Keep the eluent, it contains your mRNA.

        5.Again resuspend the oligo(dT) in 200µl Elution Buffer (heated).

        6.Centrifuge at 5,000 x g for 30 seconds at room temperature.

        7.Save the combined 400µl eluent in the microcentrifuge tube.

        8.Add 60µl of 2M sodium acetate and mix.

        9.Add 1ml of 95% ethanol (Do not use 100% ethanol, it contains fluorescent contaminants).

       10.Freeze at -70oC overnight.

       11.Thaw and centrifuge at 16,000 x g for 15 minutes at 4oC.

       12.Carefully aspirate all the ethanol from the mRNA pellet, taking care not to disturb the pellet.

       13.Resuspend the mRNA pellet in 20µl of Elution Buffer (heated as before).

       14.Determine the concentration of the mRNA by OD or &quot;eyeballing&quot; on an agarose minigel.

       15.Store mRNA at -70oC. 
http://cmgm.stanford.edu/pbrown/protocols/index.html
</protocol>
<protocol xml:space="preserve"
  title="Stanford: Sep-Pak and HPLC purification protocol for Cy dyes (un)incorporated with dNTP Ver. 4.2 (9/18/98)"
  type="other_protocol">Cyanine Dye Purification Protocol
Last updated 9-18-98 by Chih Long Liu 
                                

                
                      compiled by Chih Long Liu, with Kathleen Hayashibara and Linda McAllister; Brown Lab
                         [HPLC purification based on dNTP protocol by Suzanne Admiraal, Herschlag Lab]


                             Note: not all of the colored or underlined words in the protocol are links!
                                  Brown Lab Members: Click here for operating guidelines.


Preparation of salt buffers

     The buffers are used for both HPLC purification and Sep-Pak purification.

High salt buffer (HSB):

     1M triethylammonium acetate, pH 8.0 
     10% acetonitrile

Low salt buffer (LSB): 

     25mM triethylammonium acetate, pH 8.0 
     10% acetonitrile (this is 40-fold dilution of HSB + 10% acetonitrile)

HSB, 800 mL 

          To 500mL ddH2O, add approx. 25-30 mL glacial acetic acid 
          Slowly add 111.3 mL (80.95 g) HPLC grade triethylamine; the reaction is exothermic; the solution will become cloudy - wait until it
          clears before adding more) 
          Add dropwise an additional 10-15 mL acetic acid to adjust the pH to 8.0. 
          Add 80 mL acetonitrile (HPLC grade) 
          Adjust the final volume to 800 mL with ddH2O.

LSB, 1000 mL

          To 25 mL HSB, add 100 mL HPLC grade acetonitrile 
          Raise final volume to 1L with ddH2O

*Filter, and then sparge both buffers before use. Makes enough for approx. 9 runs and 2 purges.

Desalting the sample for Sep-Pack Purification

Materials:

     HPLC grade acetonitrile (a set of three runs consumes around 40 mL) 
     20 ml ddH2O, 0.22 mm filtered 
     15 mM TEAA (this can be substitutied with 0.750 mL HSB and 49.25 mL ddH2O, 0.22 mm filtered, to produce 50 mL) 
     Sep-Pak cartridges, C18 (Waters #20515); 1 column per cy dye. Can be reused twice. 
     12 mL syringe (1 per dye)

Pre-load protocol:

   1.Wash Sep-Pak with 10 mL of acetonitrile. 
   2.Wash with 5 mL of water. 
   3.Wash with 10 mL of 15 mM TEAA.

Sample Preparation, Loading, and Elution:

   1.The column may be loaded up to 150 A260 units of sample (based on the A260 reading for both labeled and unlabeled dNTPs in the filtrate),
     lyophilized down to 3-5 mL (volumes of 6.5-7.5 provided reasonable yields, and volumes of u p to 10 mL have been successful, but with lower
     yields). 
   2.Add 1 M TEAA to the sample, to a concentration of 15 mM. 
   3.Load the sample onto the column. The dye should be visible near the top of the column. Wash the column with 5 mL of 15 mM TEAA. 
   4.Elute with 3 mL of acetonitrile. Discard the initial colorless drops, collect the colored eluent into a 2 mL centrifuge tube, and discard the
     remaining solution. The collected volume will be approximately 1.0-2.0 mL. Some colored material will rema in on the column after elution.

Immediately wash (pre-load protocol) and process any subsequent lots, reusing the column only twice for each lot. If the column is allowed to sit for
more than 5 minutes, yields may decrease 5-10% or more. Avoid reusing the column more than two times (a total of three uses), as yields may begin to
diminish. Pool the lots together, check UV absorbance, and calculate yield. Expected yields for this stage are 70-80% for either Cy dye. 

HPLC Preparation

   1.Lyophilize the entire Sep-Pak purified material, keeping the dyes separate (see Lyophilization and Redilution section). Resuspend with a small
     volume (200-500 ml) of 0.2 M TEAA. 
   2.Centrifuge the sample through a 0.45 mm filter. 
   3.Check the absorbance at 260 nm and, if necessary, dilute the sample down to a concentration that will not exceed the capacity of the column
     (Dionex NucleoPac[tm] 9 ´ 250 PA-100; P/N 43011 (anion exchange) column listed bel ow). Typically a 1 ml injection loop is used to inject a
     10 mM sample of unlabeled dNTPs + Cy-dUTP.

HPLC Purification

Column used: Dionex NucleoPac[tm] 9 ´ 250 PA-100; P/N 43011 (anion exchange)
HPLC Equipment: Perkins Elmer Series 200 LC Pump, Perkins Elmer UV/Vis Detector LC 295

     1. Equilibrate column in buffer system, 3 mL/min flow rate (general purge method)

          1.1. 5 min low salt buffer (LSB) or A
          1.2. 2 min gradient to 100% high salt buffer (HSB) or B
          1.3. 5 min HSB or B
          1.4. 2 min gradient to 60% LSB or A
          1.5. 5 min 60% LSB or A

     2. Load sample into injection loop. See &quot;Final Considerations&quot; on the next page for sample lot size and other issues.

     3. Run gradient (3 mL/min flow, chart recorder 0.5 cm/min)

          3.1. 5 min 60% LSB, 40% HSB
          3.2. 10 min gradient to 80% HSB
          3.3. 1 min gradient to 100% HSB
          3.4. 10 min 100% HSB
          3.5. 1 min gradient to 60% LSB
          3.6. 5 min 60% LSB

Final Preparative Procedures

     For a preparative run of 150 nmoles Cy-dUTP (note &quot;Final Considerations&quot;), the recovered fractions of purified Cy-dUTP is eluted in 5 ml of
     volatile TEAA buffer that can be removed by lyophilization.

Lyophilization and Redilution

(Return to HPLC Preparation.) 

   1.Pool fractions of each Cy dye into a 50 mL conical tube. Punch holes into the cap using an 18-gauge needle. Avoid exceeding more than half the
     volume of the tube, as the samples may have a tendency to splatter during degassing or travel up the tube during lyophilization. 
   2.Freeze the samples on dry ice and transfer them to a lyophilization chamber. Wrap the chamber with foil to protect the samples from light. 
   3.Lyophilize the samples to completion (approx. 12-15 hrs for HPLC-purified samples, 6-10 hrs for Sep-Pak-purified material). HPLC-purified
     samples will have the appearance of a residue until most of the TEAA is removed. 

     Note: Steps 4 and 5 are for HPLC-purified samples only.

   4.Dilute the samples back up to 8-10 mL with 0.22 mm filtered ddH2O and lyophilize until the samples again resemble residue. This time, it may
     require up to 24 hours of lyophilization. 
   5.Repeat Step 4. Lyophilization is complete when the sample has the appearance of a film and/or specks of solid residue. Resuspend the sample in
     a small volume of 10 mM phosphate buffer, pH 7.0. Bear in mind that some of the Cy-dUTP will coat the upp er regions of the tube walls and
     may not be readily visible. Check absorbance of the sample. A560/660/A260 ratios should be around 17 for cy 3 and 25 for cy 5.

Expected overall yields: cy 3: 50-70%; cy 5: 45-65%.

Final Considerations

     Currently 150nmol of Cy-dUTP contained in a 10 mM mixture of labeled and unlabeled dNTPs is loaded onto the column listed above,
     depending on the contents and concentration of the filtrate; PCR flow-through samples tend to be cleaner and may allow for larger injection lots.
     Thus, the flow-through from PCR and RT reactions may be kept separate so that they can be processed separately. The Sep-Pak purification
     desalts the sample and is likely to remove other impurities present in the sample. HPLC purif ication can still be performed without Sep-Pak
     purification, but salt content prohibits injection of more than 50nmol. Furthermore, yields tend to be lower if the sample is not desalted prior to
     HPLC injection. 
http://cmgm.stanford.edu/pbrown/protocols/index.html
</protocol>
<!--
<protocol xml:space="preserve"
  title="TIGR: Data Collection, Normalization, and Analysis"
  type="other_protocol">Differential gene expression is assessed by scanning the hybridized arrays using a confocal laser
 scanner capable of interrogating both the Cy3- and Cy5-labeled probes and producing separate TIFF
 images for each. As is the case with arraying robots, there are a number of manufacturers that produce
 scanners capable of detecting Cy3 and Cy5 (see Table 4) and most are planning to release instruments
 capable of detecting additional dyes.
  

 Slide Scanning 

 We are currently using the ScanArray 3000 produced by GSI Lumonics. This scanner uses red and
 green Helium-Neon lasers operating at 633nm and 543nm to excite Cy5 and Cy3, respectively.
 Hybridized slides are scanned first in the Cy5 channel, and then the Cy3 channel, as Cy5 is more
 susceptible to photodegradation than Cy3. Data from each fluorescence channel is collected and stored
 as a separate 16-bit TIFF image. These images are analyzed to calculate the relative expression levels
 of each gene and to identify differentially expressed genes. The analysis process can be divided into
 two steps - image processing and data analysis. Figure 5 shows a typical hybridization image
 produced when things work well. The contrast in this image has been adjusted to allow faint spots to
 be easily visualized. Important aspects of the hybridization to note are the low level, uniform
 background and the good signal-to-noise

 Image Processing

 Image processing involves three stages. First, the spots representing the arrayed genes must be
 identified and distinguished from spurious signals that can arise due to precipitated probe or other
 hybridization artifacts or contaminants such as dust on the surface of the slide. This task is simplified
 to a certain extent because the robotic arraying systems used to construct the arrays produce a regular
 arrangement of the spotted DNA fragments. However, variable intensities and uneven slide
 backgrounds as well as some irregularities in the gridded arrays complicate the problem slightly.
 Generally, problem of grid spot location is coupled with estimation of the fluorescence background.
 For microarrays, it is important the background be calculated locally for each spot, rather than globally
 for the entire image as uneven background can often arise during the hybridization process. The second
 step in analysis of the array images is the estimation of background.
 Following spot identification and local background determination, the background-subtracted
 hybridization intensities for each spot must be calculated. There are currently two schools of thought
 regarding the calculation of intensities - the use of the median or the mean intensity for each spot. As
 array analysis generally uses ratios of measured Cy3 to Cy5 intensities to identify differentially
 expressed genes, the mean and the integrated intensities are operationally equivalent. In comparisons
 of intensities measured for normalization controls spiked into the labeling reactions, we have found
 mean intensities to give more consistent results and consequently we use these in subsequent
 calculations (V. Sharov and J. Quackenbush, in preparation).
 A number of image processing software packages are available and are listed in Table 5. We have
 developed a software package called TIGR_Spotfinder for image processing
 (&lt;http://www.tigr.org/softlab/&gt;). TIGR_Spotfinder uses a thresholding algorithm that separates spots
 from the background, allowing a grid to be laid across the spots. Having found a grid, spots are found
 within each grid element, local background is calculated, and background-subtracted, integrated
 intensities are calculated in both the Cy3 and Cy5 channels. Measured intensities are entered into the
 Molecular Analysis of Gene Expression (MAGE) database, a Sybase relational database specifically
 designed to capture gene expression data.

 Data Normalization and Analysis

 Following image processing, the data generated for the arrayed genes must be further analyzed before
 differentially expressed genes can be identified. The first step in this process is the normalization of
 the relative fluorescence intensities in each of the two scanned channels. Normalization is necessary to
 adjust for differences in labeling and detection efficiencies for the fluorescent labels and for
 differences in the quantity of starting RNA from the two samples examined in the assay. These
 problems can cause a shift in the average ratio of Cy5 to Cy3 and the intensities must be rescaled
 before an experiment can be properly analyzed. 
 The normalization strategies that can be used are based on some underlying assumptions regarding the
 data and the strategies used for each experiment should be adjusted to reflect both the system under
 study and the experimental design. The primary assumption is that for either the entire collection of
 arrayed genes or some subset of the genes such as housekeeping genes, or for some added set of
 controls, the ratio of measured expression averaged over the set should be one.
 Depending on the experimental design, there are three useful approaches for calculating normalization
 factors. The first simply uses total measured fluorescence intensity. The assumption underlying this
 approach is that the total mass of RNA labeled with either Cy3 or Cy5 is equal. While the intensity
 for any one spot may be higher in one channel than the other, when averaged over thousands of spots
 in the array, these fluctuations should average out. Consequently, the total integrated intensity across
 all the spots in the array should be equal for both channels. Alternatively, one could add a number of
 controls in increasing but equimolar concentrations to both the labeling reactions and the sum of the
 intensities for these spots should be equal. A second approach uses linear regression analysis. For
 closely related samples, one would expect many of the genes to be expressed at nearly constant levels.
 Consequently, a scatterplot of the measured Cy5 versus Cy3 intensities should have a slope of one.
 Measured intensities for added equimolar controls should behave similarly. Under this assumption,
 one can use regression analysis techniques to calculate the slope. This is then used to rescale the data
 and adjust the slope to one. A third approach has been described by Chen et al. (2). They assume that
 some subset of housekeeping genes exists and that for these, the distribution of transcription levels
 should have some mean value ? and standard deviation ? independent of the sample. In this case, the
 ratio of measured Cy5 to Cy3 ratios for these genes can be modeled and the mean of the ratio adjusted
 to 1. Chen and collaborators describe an iterative procedure to achieve this normalization and we have
 implemented their algorithm and a variation of it that uses the entire data set, as well the total intensity
 and linear regression normalization, into a data visualization and analysis tool called TIGR
 ArrayViewer. TIGR ArrayViewer is freely available and can be obtained
 through &lt;http://www.tigr.org/softlab/&gt;. In any normalization approach, care must be taken in handling
 genes expressed at low levels. Statistical fluccuations in the measured levels can cause a significant
 variation in the ratios that are calculated and inefficiencies in labeling for either of the two dyes can
 cause these low intensity genes to disappear from the arrays. Typically, we only use spots in the final
 analysis where the intensities in both channels are two standard deviations above background.
 Following normalization, data are typically analyzed to identify genes that are differentially expressed.
 Most published studies have used a post-normalization cutoff of two-fold up- or down-regulation to
 define differential expression; the approach defined by Chen et al. (2) provides confidence intervals
 that can be used to identify differentially expressed genes. In order to separate genes that are truly
 differentially expressed from stochastic changes, we typically conduct three independent microarray
 assays starting from independent mRNA isolations and define differential expression based on their
 consensus.  
 Conclusion 
 The examination of gene expression using microarrays holds tremendous promise for the identification
 of candidate genes involved in a variety of processes. Indeed, the experiments that have been described
 to date have confirmed known patterns of expression and provided information on genes of unknown
 function. However, most applications have to date only allowed the identification of genes
 differentially expressed at significant levels. The true challenge, and the promise of this technique,
 will be to use it to identify genes that are consistently up- or down-regulated by 10 or 20% yet play
 significant roles in the development and progression of disease. This will require the analysis of data
 from multiple experiments and the correlation of patterns of gene expression with additional
 experimental and clinical information. Recently a variety of techniques including hierarchical
 clustering (3) and self-organizing maps (11) have been applied to the analysis of microarray
 expression data across multiple experiments. However, each of these depends on having reliable and
 reproducible data from each microarray assay. The laboratory techniques outlined here have allowed
 reproducible hybridization results such as those shown in Figure 5. Although these protocols will
 likely continue to evolve, we believe that they represent a reliable starting point for those beginning
 microarray experimentation. 
http://www.tigr.org/tdb/microarray/conciseguide.html
</protocol>
-->
<protocol xml:space="preserve"
  title="TIGR: Hybridization"
  type="hybridization">The goal in any hybridization is to obtain high
 specificity while minimizing background. We have
 developed protocols that give reproducible,
 high-quality hybridization results while maximizing
 the measured fluorescence from the array. 
 Aminosilane coated slides bind DNA with high
 efficiency. Prior to hybridization, the free amine groups
 on the slide must be blocked or inactivated, otherwise
 nonspecific binding of labeled cDNA to the slide can
 deplete the probe and produce high background.
 Although the slides can be blocked chemically, we have
 found a simple prehybridization in a solution
 containing 1% bovine serum albumin to be extremely
 effective in eliminating nonspecific binding of the probe
 to the slide. 
 Prehybridization has the additional advantage of
 washing unbound DNA from the slide prior to the
 addition of the probe. Any DNA that washes from the
 surface during hybridization competes with DNA
 bound to the slide. As the kinetics of solution
 hybridization is much more favorable than surface
 hybridization, this can dramatically decrease the
 measured fluorescence signal from the microarray. All
 prehybridization and hybridization washes are carried
 out in microscope slide staining trays (VWR Cat#
 25461-003).

 Prehybridization

 1.Prepare prehybridization buffer containing 5´SSC,
   0.1% SDS and 1% bovine serum albumin (BSA;
   Sigma Cat# A-9418).
 2.Prepare 2´ hybridization buffer containing 50%
   formamide, 10´SSC, and 0.2% SDS.
 3.Place slides to be analyzed into a Coplin jar (VWR
   Cat# 25457-200), fill with prehybridization buffer,
   and incubate at 42oC for 45 minutes.
 4.Wash the slides by dipping five times in room
   temperature MilliQ water.
 5.Dip the slides in room temperature isopropanol and
   air dry.
 Slides should be used immediately following
 prehybridization. We have found that hybridization
 efficiency decreases rapidly if the slides are allowed to
 dry for more than one hour. 

 Hybridization

 1.Combine 10ml each of purified Cy3- and
   Cy5-labeled probes, mix well and add
       COT1-DNA
       (20mg/ml)
                             1ml
                                       (LifeTechnologies; Cat#
                                               25279-011).
       Poly(A)-DNA
       (20mg/ml)
                             1ml
                                  (Pharmacia; Cat# 27-7836-01).

   to block nonspecific hybridization.
 2.Heat the probe mixture at 95oC for 3 minutes to
   denature.
 3.Centrifuge the probe in a microfuge set at maximum
   angular velocity for 1 minute.
 4.Combine the probe with an equal volume of 2´
   hybridization buffer that has been heated to 42oC.
 5.Apply the labeled probe to a prehybridized
   microarray slide and cover with a 22mm´60mm
   polyethylene hydrophobic coverslip (PGC Scientific
   Cat# 62-6504-06). 
 6.Place the slide in a sealed hybridization chamber
   (Corning Costar Cat #2551), add 20ml of water to the
   chamber at the end of the slide.
 7.Place the sealed chamber in a 42oC water bath and
   incubate for 16-20 hours. 
 8.Remove the array from the hybridization chamber,
   taking care not to disturb the coverslip. 
 9.Place the slide in a staining dish containing
   low-stringency wash buffer containing 1´SSC and
   0.2% SDS at 42oC.  
 10.Gently remove the coverslip while the slide is in
   solution and agitate for 4 minutes. 
 11.Wash the slide at high-stringency in a staining dish
   containing 0.1´SSC and 0.2% SDS at room
   temperature, agitating for 4 minutes.  
 12.Wash the slide in 0.1´SSC, agitating for 4 minutes.  
 13.Allow the slides to air dry. 
http://www.tigr.org/tdb/microarray/conciseguide.html
</protocol>
<protocol xml:space="preserve"
  type="cdna_and_labeling"
  title="TIGR: RNA Labeling">The ability to label small quantities of starting material is an important consideration for the study of
 expression in rare patient samples and consequently, we have focused on decreasing the quantity of
 starting material required. Probes for microarray analysis are prepared from RNA templates by
 incorporation of fluorescently labeled deoxyribonucleotides during first strand cDNA synthesis. Either
 total or poly(A+) RNA can be used in the reverse transcription reaction. Oligo(dT) labeling of total
 RNA provides consistently high-quality probes from smaller quantities of starting RNA and without
 the expense of poly(A+) purification. Figure 4 shows the results of microarray hybridizations using
 labeled total or poly(A+) RNA prepared from the same cell lines. An analysis of the fluorescence
 intensities for the elements in arrays hybridized with probes prepared from 1.5mg of poly(A+) RNA
 (the equivalent 50-100mg of starting total RNA) and 4mg of total RNA indicate that total RNA
 labeling provides comparable probe activity without any increase in background fluorescence. 
   
   
 Typically, we prepare labeled probes using Cy3- and Cy5-dUTP (Amersham Cat#s PA53022,
 PA55022), although Cy-labeled dCTP (Amersham Cat#s PA53021, PA55021) can be used with an
 appropriate change in the concentrations of unlabeled dNTPs in the reaction. We have investigated a
 number of reverse transcriptases, including AMV and MMLV and have found that Superscriptä II RT
 (LifeTechnologies; Cat# 18064-014) generates probes with significantly greater activity (data not
 shown).  
   
   
 It should be noted that both Cy3 and Cy5 are photosensitive and care should be taken to minimize
 exposure to light during the labeling, hybridization, washing, and scanning processes. Upon receipt,
 Cy-labeled nucleotides should be aliquotted into single-use light- proof tubes and stored at -20 deg. C
 until needed. All reactions should be carried out in foil-wrapped tubes and all hybridizations and
 washes in foil-wrapped containers.  
   
   

 Probe Labeling and Purification

  
 1.Prepare a labeling reaction master mix containing 500mM dCTP, 500mM dATP, 500mM dGTP,
   100mM dTTP, Cy 3-dUTP/Cy 5-dUTP, 400U Superscriptä II RT, 1mM dithiothreitol (DTT) and
   1´ RT buffer. We typically prepare sufficient quantities for 20 labeling reactions and store the
   unused solution at -20 deg. C:
                       5´ RT buffer
                                       120ml
                       DTT (5mM)
                                        60ml
                       dATP (100mM)
                                         3ml
                       dCTP (100mM)
                                         3ml
                       dGTP (100mM)
                                         3ml
                       dTTP (100mM)
                                        0.6ml
                       DEPC?d
                       WATER
                                      110.4ml
                              Total:
                                       300ml

 2.To 10mg of total RNA (or 2mg poly(A+)) in a microfuge tube, add 2mg of oligo(dT) (18-20mer;
   Life Technologies Cat# Y012120) and DEPC-treated WATER to a total volume of 10ml.
 3.Incubate the reaction mixture at 70oC for 10 minutes, and chill on ice for one minute.
 4.To the RNA, add:
                   RT labeling mix
                                            15ml
                   Cy3- or Cy5-dUTP (1 mM)
                                             3ml
                   Superscriptä II RT
                   (200U/ml)
                                             2ml

  
 5.Mix thoroughly and incubate at 42oC for 2 hours.
 6.Briefly centrifuge the reaction and add 1.5ml of 20mM EDTA to stop the reaction.
 7.Add 1.5ml of 500mM NaOH and heat at 70oC for 10 minutes to degrade the RNA
 8.Neutralize the reaction by adding 1.5ml of 500mM HCl
 9.Unincorporated fluorescent nucleotides are removed by glass fiber filtration using GFX columns
   (Pharmacia Cat# 27-9602-01) and the instructions provided by the manufacturer. 
 10.Elute the purified products using 50ml of TE pH 8.0 and dry the probe to completion in a
   speedvac.
 11.Resuspend the probe in 10ml of DEPC treated WATER. 
http://www.tigr.org/tdb/microarray/conciseguide.html
</protocol>
<protocol xml:space="preserve"
  title="TIGR: RNA Extraction"
  type="rna_prep">Impurities in RNA preparations can have an adverse effect both on labeling efficiency and the stability
 of the fluorescent labels that are used for microarray expression analysis. We have found that Trizol
 ä (Life Technologies; Cat# 15596-014) gives consistently high quality RNA from cell culture and
 many tissue samples, although additional steps must be taken to remove polysaccharides when
 extracting RNA from some tissues. Trizol extraction is quick and produces a high yield of total RNA. 

 RNA Extraction

  
 1.Aspirate media from the cells and wash once with Phosphate Buffered Saline (PBS). 
 2.Add 5ml PBS and scrape cells from the plate.
 3.Transfer cell suspension to a 50ml polypropylene conical-bottom tube (Falcon Cat# 352070).
 4.Wash the plate with an additional 1ml PBS and add the suspension to the tube.
 5.Pellet the cells by centrifugation at 2300 rpm (900´g) for 3 minutes at 4oC and discard the
   supernatant.
 6.Add 2ml Trizolä per ~2´106 cells (approximately one 150mm plate of fibroblasts) to the pellet and
   pass the suspension through an 18 gauge syringe several times to disrupt the pellet.
 7.Incubate the sample at room temperature for 5 minutes. 
 8.Add 0.4ml of chloroform (0.2ml/1ml Trizol) and shake vigorously for 1 minute 
 9.Incubate at room temperature for 2 minutes 30 seconds. 
 10.Remove cellular debris by centrifugation at 4000rpm (2700´g) for 15min at 4oC. 
 11.Transfer the supernatant to 1.2 ml microfuge tubes (0.5ml/tube) and an equal volume of
   isopropanol to precipitate the RNA.  
 12.Incubate at room temperature for 15 minutes. 
 13.Centrifuge at 15,000rpm (21,000´g) for 15 minutes to pellet the RNA. 
 14.Discard the supernatant and resuspend the pellet in 70% ethanol. The RNA can be stored in 70%
   ethanol at -20 deg. C until use. 
 15.Prior to use, centrifuge at 15,000 rpm (21,000´g) for 15 minutes at 4oC and discard supernatant.  
 16.Resuspend the pellet in diethylpyrocarbonate (DEPC) treated water or RNase-free TE buffer for
   labeling. 
http://www.tigr.org/tdb/microarray/conciseguide.html
</protocol>
<!--
<protocol xml:space="preserve"
  title="TIGR: Array Fabrication"
  type="other_protocol">Microarrays are constructed by arraying PCR amplified cDNA clones or genes at high density on
 derivatized glass microscope slides. For the analysis of expression in most eukaryotes, expressed
 sequence tag (EST) data represent the most extensive data for gene identification. ESTs are
 single-pass, partial sequences of cDNA clones, and they have been used extensively for gene
 discovery and mapping in humans and other organisms. The EST approach has been widely adopted;
 more than 71% of all GenBank entries and 40% of the individual nucleotides in the database are
 derived EST sequences (9).
 Generally, cDNA clones are selected to represent as many unique transcripts as possible. There are a
 number of analyses of these data that attempt to identify unique human transcripts within the EST
 data, the two most widely used are UniGene (1) (&lt;http://www.ncbi.nlm.nih.gov/UniGene/&gt;) at the
 National Center for Biotechnology Information (NCBI) and the TIGR Human Gene Index (7)
 (HGI; &lt;http://www.tigr.org/tbd/hgi/hgi.html&gt;) at The Institute for Genomic Research (TIGR). While
 both UniGene and HGI are based on EST clustering, the TIGR protocol assembles the ESTs within
 the clusters, producing Tentative Human Consensus (THC) sequences. 

   
   
 We selected cDNA clones for array construction using the TIGR HGI as part of a program to
 assemble a 30,000 gene clone set. THCs were chosen for representation in the clone set with
 preference given to those containing known genes or those with mapped positions; additional THCs
 were selected to represent as yet uncharacterized transcripts. For each target THC, a single cDNA
 clone was identified based on the EST content of the THC assembly.  

I.A. PCR Amplification and Clone Preparation

  
 cDNA clone inserts can be amplified by PCR from plasmid miniprep DNA or directly from clones in
 culture. In high-throughput applications, amplification of clones from culture has the advantage of
 being both more cost efficient and less labor intensive with lower cross-contamination rates than
 amplification from plasmid DNA. Our amplification success rate from culture is equivalent to that we
 have achieved using plasmid templates. For more than 30,000 clones, our success rate for single-band
 amplification is approximately 87.5%; 6.3% of reactions yield multiple or weak bands and 6.2% fail to
 amplify. Results from a typical amplification using the protocol described below are shown in Figure
 1.
 PCR amplification

   
   
 The cDNA clones that are widely available through the IMAGE consortium distributors - The
 American Type Culture Collection (ATCC), Research Genetics, and Genome Systems - have been
 cloned into a variety of vectors. While the majority have both M13(-21) and M13REV priming sites.
 However, many have point mutations in either of these two ?universal? priming sites. We have
 designed alternative M13 primers that avoid these point mutations and that have allowed amplification
 of clone inserts from all of the vectors we have encountered to date. These new universal
 amplification primers are:  
   
   
     M13 FWD: 5&apos; GTT TTC CCA GTC ACG ACG TTG 3&apos;  
     M13 REV: 5&apos; TGA GCG GAT AAC AAT TTC ACA CAG 3&apos;  
   
   
 Clone inserts are amplified using the following protocol:  
   
   
 1.Selected clones are inoculated into 96 well deep-well blocks (Qiagen; Cat # 19573) containing1.2 ml
   of LB/Ampicillin (50 mg/ml) and incubated for 16 hours at 37ºC and 200 rpm in a shaking
   incubator. A 100 ml aliquot of each is archived for future use in microtiter plates containing 10%
   glycerol at -80 deg. C.  
 2.Following overnight growth, 5ml of culture suspension are transferred into a 96 well Falcon
   U-bottom plate (BD Biosciences; Cat # 353077) containing 95ml of MilliQ water.  
 3.Microtiter plates containing diluted culture are heated to 95oC for 10 minutes in a laboratory oven to
   lyse the cells and release the plasmid clones.  
 4.Prior to PCR, cellular debris is removed by centrifugation at 1200´g for 3 minutes in a centrifuge
   equipped with microtiter plate carriers.  
 5.Clone inserts are amplified in 50ml PCR reactions in 96 well reaction plates (Perkin-Elmer Applied
   Biosystems Cat # N801-0560). A reaction master mix is prepared for each reaction plate:  
   MilliQ water
                              3.2ml
   10´ PCR buffer
                              500ml
   MgCl2 (25mM)
                              500ml
   M13 Forward primer
   (10mM)
                              100ml
   M13 Reverse primer
   (10mM)
                              100ml
   dNTP mix (2mM per
   dNTP)*
                              500ml
                                    (Life Technologies Cat#
                                    10297-018)
   (* 10ml of each dNTP as 100 mM stock in 460ml of MilliQ water)
   Platinum Taq (5U/ml)
                              20ml
                                    (Life Technologies; Cat#
                                    10966-018)
                  Total:
                             4.92 ml

 6.For each clone, add 48ml of master mix to 2ml of culture supernatant in 96 well PCR plate.
 7.Reactions are amplified in an thermocycler (MJ Research; PTC-225 Tetrad) using the following
   cycling protocol:
               95oC ´ 2min
                                     Initial Denaturation


               4 deg. C forever

 Reaction clean-up
 For efficient binding of the amplified clone inserts to the slides, it is essential to remove
 unincorporated nucleotides and primers from the reaction products. While there are a variety of
 techniques that can be used, we have found filtration using 96 well multiscreen filter plates (Millipore;
 Cat # MANU 03050) to give excellent DNA product recovery without any significant contamination at
 relatively low cost. 
 PCR products are cleaned using the following filtration protocol: 
 1.Transfer PCR product (50ml) to the Millipore filter plate. 
 2.Place the filter plate on a vacuum manifold filtration system (Qiagen, Cat # 19504 or Millipore Cat
   # MAVM0960R) and filter at a pressure of 15in (380 mm) Hg for 10 minutes or until the plate is
   dry.  
 3.Add 30ml of MilliQ water to each well and filter at 15mm (380 mm)Hg for 5-10 minutes or until
   the plate is dry.  
 4.Repeat step 3. 
 5.Remove plate from the manifold filtration system. Add 60ml of MilliQ water to each well and place
   on a shaker. Shake vigorously for 10 minutes to resuspend the DNA. 
 6.Manually pipet the purified product to a new 96 well plate. 
 7.Plates containing the purified PCR products are then sealed using a cap mat (VWR; Cat #
   40002-002) and stored at 4oC for future arraying. 

 I.B. Array Printing

 Microarrays are prepared by printing PCR amplicons suspended in either a high salt or other
 denaturing buffer onto poly-L-lysine or aminosilane coated glass microscope slides using a
 high-speed robotic system. This process was originally described by Patrick Brown and collaborators
 (8) at Stanford University and they provide plans so that others can replicate their arraying robot
 (&lt;http://cmgm.stanford.edu/pbrown/mguide/index.html&gt;). However, there are a number of companies
 that are selling robotic systems for microarraying and these are listed in Table 2. We use a microarray
 robot built by Intelligent Automation Systems (IAS) of Cambridge, Massachusetts. Based on a high
 precision, four-axis Seiko robotic arm, the IAS arrayer uses a 12-tip print head to array DNA samples
 from either 96- or 384-well microtiter plates onto as many as 100 silanized glass microscope slides.
 With an average spot size of 130 µm and the capability to adjust the spot-to-spot spacing, the IAS
 arrayer can spot 19,200 elements (the contents of 200 microtiter plates) or more onto a single slide.
 Both the slide surface and the spotting buffer are critical components for reproducible, high-fidelity
 micorarray analysis. Most published reports have used high salt buffers such as 3´SSC to print DNA
 on poly-L-lysine coated slides (10). Our analysis suggests, however, that aminosilane offers a more
 consistent surface with lower background fluorescence. There are a number of commercial vendors for
 aminosilane coated slides (see Table 2), but Corning CMT-GAPS? aminosilane coated glass
 microscope slides have been the most consistent. Using our protocol in number of side-by-side
 comparisons, the CMT-GAPS? slides produce approximately half the background fluorescence of
 poly-L-lysine slides while yielding signal intensities that are consistently higher (data not shown). In
 addition, the spot morphology on CMT-GAPS? slides is much more uniform, with fewer ?doughnuts?
 than on any of the alternatives we have investigated.
 We also investigated a number of different spotting chemistries to determine which provides the best
 results in subsequent hybridization assays. Figure 2 shows the results of a comparison between 50%
 dimethyl-sulfoxide (DMSO) and 3´SSC as a spotting buffer. We have found that PCR products
 printed onto CMT-GAPS? aminosilane coated glass microscope slides (Corning, Cat# 2550) using
 50% DMSO as a printing buffer provides the best substrate for hybridization, giving the greatest
 hybridization intensities. Using 50% DMSO as a printing solution has a number of additional
 advantages. DMSO denatures the DNA allowing better binding to the slide and providing more
 single-stranded targets for hybridization. Further, DMSO is hygroscopic and has a low vapor
 pressure, allowing DNA prepared for arraying to be stored for long periods of time without significant
 evaporation.
 The print head on our arrayer and most others use ?quill? pens that use capillary action to draw fluid
 into the spotting pens and surface tension interactions to dispense solution onto the slide. The Arrayit
 ChipMaker3ä microspotting pins (TeleChem International Inc.) are very durable and can reproducibly
 generate high-quality spots with good precision; all array images shown were printed with the same
 set of ChipMakerä 3 over more than six months.A variety of parameters such as the robot arm
 acceleration, temperature, and humidity control both spot morphology and size. We have found
 printing to be optimal at approximately 45% relative humidity and a constant temperature of 72oF
 (22oC). Changes in humidity and temperature have a significant impact on the size and morphology of
 spots, as well as the efficiency of DNA binding to the slides and these must be carefully controlled to
 provide the consistent spotting. Figure 3 shows the effects of varying humidity and temperature on
 spot morphology and DNA retention. DNA samples were spotted onto the slides as described above
 while temperature and humidity levels were recorded on a chart recorder. During the printing,
 temperature and humidity levels were allowed to vary continuously from 72ºF (22.2ºC) and 45-50% to
 a low of 62ºF (16.7ºC) and 40-45%and a high of 80ºF (26.7ºC) and 80-85% respectively. Following
 hybridization with a vector specific probe, we were able to reconstruct the optimal printing conditions
 by using the chart recorder data to assign temperature and humidity values to the spots. 
 Arraying 
 1.Add equal volumes of purified PCR product to DMSO in a 96 well V-bottom plate (Corning; Cat #
   3897). Typically, 5ml of each are used to prepare spotting plates that can be used to print 100 or
   more slides.  
 2.Slides to be printed are marked with a diamond-tipped pen, dust is removed by blowing the slides
   with high-pressure nitrogen gas, and the slides are placed in the arrayer. Care must be taken not to
   touch the surface of the slides as oils adversely affect the ability of the slide surface to bind DNA.  
 3.Microtiter spotting plates are loaded into the arrayer and PCR products are spotted onto the slides at
   72oF and 45% relative humidity.  
 4.Following printing, the slides are allowed to dry and spotted DNA is bound to slide by
   UV-crosslinking at 90 mJ using a Stratalinkerä (Stratagene, Cat# 400071) and baking at 80oC for
   two hours.  
 5.Printed slides are stored in a light-tight box in a bench-top dessicator at room temperature until they
   are to be used for hybridization.  
http://www.tigr.org/tdb/microarray/conciseguide.html
</protocol>
-->
<protocol xml:space="preserve"
  title="Schena and Davis: Preparation of total RNA from cultured human cells"
  type="rna_prep">Reagents and Equipment

   ·         RPMI media (Sigma)

   ·         Phosphate Buffered Saline (Sigma) 

   ·         GTC solution

   ·         Low speed (Beckman)

   ·         Ultracentrifuge (Beckman)

   ·         Ultra-Clear centrifuge tubes, 13 x 51 mm (Beckman).

   ·         CsCl (United States Biochemical)

Method

       1.      Obtain a confluent flask containing 50 ml of cultured human cells (e.g. Jurkat line J25).

       2.      Set up 5 flasks (162 cm2) containing 100 ml per flask of RPMI media.a

       3.      Split confluent cells 1:10 and grow 48 hrs.

       4.      Add inducer (e.g. phorbol ester) and grow an additional 24 hrs.

       5.      Transfer cells (500 ml total) to a 500 ml centrifuge bottle.

       6.      Pellet cells by centrifugation in a JA-10 rotor for 5 min at 3,000 rpm.

       7.      Remove and discard media.

       8.      Resuspend cell pellet in 10 ml phosphate buffered saline (PBS).

       9.      Transfer to a 15 ml conical tube.

       10.  Pellet cells by centrifugation in a clinical centrifuge for 5 min at 1,000 rpm.

       11.  Remove and discard PBS.

       12.  Freeze cell pellet (~5 x 108 cells) in liquid nitrogen and store at -80 deg. C.

       13.  Obtain cell pellet (~5 x 108 cells) from -80 deg. C.

       14.  Add 12 ml of GTC solutionb and resuspend pellet by vortexing.

       15.  Draw the cell lysate through a 19 gauge needle 10 times using a 12 cc syringe.c

       16.  Set up six ultracentrifuge tubesd containing 3.0 ml each cesium chloride solution.e

       17.  Overlay 2 ml of the GTC cell lysate onto the CsCl layer of each tube. 

       18.  Pellet the RNA by centrifugation in an SW50.1 rotor at 35,000 rpm for 12 hrs at 25 deg. C. 

       19.  Pour off and discard the supernatant by inverting each tube.f

       20.  Dry tubes in an inverted position for 60 min on a layer of paper towels.

       21.  Add 100 µl of 1X TEg to the bottom of each tube.h

       22.  Heat for 1 min at 65 deg. C to aid in dissolving the pelleted RNA.

       23.  Pipet vigorously to dislodge and dissolve pelleted RNA.

       24.  Combine the six 100 µl RNA samples. 

       25.  Use 0.4 ml fresh 1X TEg to isolate residual RNA from each tube.

       26.  Extract the 1.0 ml RNA sample twice with 0.5 ml phenoli to remove residual protein.

       27.  Extract twice with ether to remove trace organics.

       28.  Divide the RNA sample into two 500 µl aliquots in eppendorf tubes. 

       29.  Add 50 µl 2.5 M sodium acetate (depc-treated) and 1.0 ml ethanol to each tube.

       30.  Pellet the RNA by centrifugation for 15 min at 25 deg. C in a microfuge.

       31.  Remove and discard the supernatant.

       32.  Dry the RNA pellet in a speedvac.

       33.  Resuspend the RNA pellet in 0.4 ml 1X TE.g 

       34.  Concentration should be 7 mg/ml for a total yield of 3 mg RNA per 500 ml cells.

       a.       RPMI media is supplemented with 10% fetal bovine serum, 100 µg/ml streptomycin and 500 U/ml penicillin.

       b.      Dissolve 60 g guanidine thiocyanate, 0.5 g sodium N-lauroylsarcosine, 5.0 ml 1M sodium citrate in 100 ml of H2O. Sterile
         filter. Add 0.5 ml ß-mercaptoethanol.

       c.       Shearing chromosomal DNA increases RNA yield.

       d.      Ultra-Clear centrifuge tubes.

       e.       Dissolve 95 g CsCl and 20 ml 0.5 M EDTA (pH=8.0) in 100 ml of H2O. Sterilize by    filtration. Add diethyl
         pyrocarbonate (depc) to 0.1%. Autoclave.

       f.        DO NOT allow the supernatant to drain back onto the RNA pellet.

       g.       1X TE [1 mM Tris, 0.1 mM EDTA (pH 8.0)] is treated with 0.1% depc to inactivate ribonucleases.

       h.       Avoid the inner sides of the tube which may contain trace ribonuclease.

       i.         Phenol solution contains phenol/chloroform/isoamyl alcohol (25:24:1 v/v).
http://www.wenet.net/~telechem/DNA-Microarray-Protocols/
</protocol>
<protocol xml:space="preserve"
  title="Schena and Davis: Preparation of polyA+ mRNA from total human RNA."
  type="rna_prep">Reagents and Equipment

   ·         Oligotex mRNA Midi Kit (Qiagen).

Method

   1.      Thaw total RNA (~7 mg/ml) stored at -80 deg. C.

   2.      Mix 150 µl RNA (~1 mg), 150 µl 2X binding buffer, and 55 µl Qiagen Oligotex-dT resin.

   3.      Heat 3 min at 65 deg. C to denature the RNA.

   4.      Cool to room temperature 10 min to allow annealing of mRNA to resin.

   5.      Pellet the resin containing bound mRNA by spinning for 2 min in a microfuge.

   6.      Resuspend the resin in 600 µl of wash buffer by vortexing vigorously.

   7.        Pellet the resin by spinning for 2 min in a microfuge.a

   8.      Resuspend the resin in 600 µl of wash buffer.

   9.      Transfer to a Qiagen spin column.

   10.  Spin out the wash buffer by centrifugation for 30 sec in a microfuge.

   11.     Resuspend the resin in 33 µl of 80 deg. C elution buffer.b

   12.  Spin for 30 sec and transfer the eluant containing the mRNA to a new tube.

   13.  Elute the remaining mRNA from the oligotex resin in the spin column with two additional 33 µl volumes of 80 deg. C elution buffer.

   14.  Combine the three 33 µl mRNA fractions.

   15.  Add 10 µl of 2.5 M sodium acetatec and 220 µl of 100% ethanol.

   16.  Pellet the mRNA by centrifugation for 15 min at 25 deg. C in a microfuge.

   17.  Dry the mRNA pellet in a speedvac.

   18.     Resuspend the pellet in 11 µl of 1X TE (pH=8.0).c

   19.     Use 1.0 µl for quantitation.d

   20.  Dilute to 1.0 µg/µl. 

   21.  Yield should be 20-25 µg mRNA from 1.0 mg of total RNA.

   a.       Carry out the first wash in an eppendorf tube (ie. by &quot;batch wash&quot;) to remove particles and debris that clog the spin column.

   b.      Heat the eppendorf tube containing the spin column to 80 deg. C for 30 sec to assist in mRNA elution. Failure to do so will decrease
     mRNA yield!

   c.       Solutions treated with 0.1% diethyl pyrocarbonate to inactivate ribonucleases.

   d.      Mix 1.0 µl in 0.5 ml TE and determine the absorbance at 260 nm (OD of 1.0 = 40 µg/ml mRNA).
http://www.wenet.net/~telechem/DNA-Microarray-Protocols/
</protocol>
<protocol xml:space="preserve"
  title="Schena and Davis: Amplification and purification of cDNAs for microarray manufacture"
  type="other_protocol">Materials and Equipment

     PCR primers modified with a 5?-amino-modifier C6 (Glen Research #10-1906-90) 
     96-well thermal cycler (PCR system 9600-Perkin Elmer) 
     96 well PCR plates (MicroAmp 96-well Perkin Elmer #N801-0560) 
     Taq DNA polymerase (Stratagene #600139) 
     PCR Purification Kit (Telechem #PCR-100) 
     Flat-bottom 384-well plates (Nunc #242765) 
     Micro-Spotting Solution (TeleChem #MSS-1). 

Method

       1.Add 1.0 µl of plasmid DNA (10 ng/µl) into each well of a 96-well plate. 
       2.Add 99.0 µl of PCR mix which contains 10 µl of 10X PCR buffer (500 mM KCl, 100 mM Tris-Cl pH 8.3, 15 mM Mg2+,
         0.1% gelatin), 10 µl of dNTP cocktail (2 mM each), 1.0 µl primer 1 (100 pmole/µl)a, 1.0 µl primer 2 (100 pmole/µl)a, 1.0 µl
         plasmid DNA (10 ng/µl)b, 76 µl H20, and 1.0 µl Taq Polymerase (5 units/µl). 
       3.Amplify cDNAs using 30 rounds of PCR (94 deg. C, 30 sec; 55 deg. C, 30 sec; 72 deg. C, 60 sec). 
       4.Purify the PCR products a 96-well PCR Purification Kit. 
       5.Elute products with 100 µl of 0.1 X TE (pH 8.0). 
       6.Dry products to completion in a speedvac. 
       7.Resuspend each PCR product in 7.5 µl Micro-Spotting solution. 
       8.Transfer to a flat bottom 384-well plate (Nunc) for arraying. 

                 a. Use of generic primer pairs (~21mers) to vector sequences allows
                 high-throughput processing. Amino-linked cDNAs are made during PCR using
                 21-mers that contain a C6 amino modifier (Glen Research) on the 5&apos; end of
                 each primer.

                 b. Plasmid DNA can be prepared by alkaline lysis and purified. The 96-well REAL prep (Qiagen #SQ811 and
                 #19504) facilitates rapid preparation.
http://www.wenet.net/~telechem/DNA-Microarray-Protocols/
</protocol>
<protocol xml:space="preserve"
  title="Schena and Davis: Microarray manufacture and processing"
  type="other_protocol">
http://www.wenet.net/~telechem/DNA-Microarray-Protocols/
</protocol>Reagents and Equipment

     Micro-spotting robot (Various) 
     Stealth Micro Spotting Device (TeleChem) 
     SuperAldehyde Substrates (TeleChem) 

Method

       1.Obtain silylated (active aldehyde) microscope slides (TeleChem). 
       2.Print amino-linked cDNAs using a micro-spotting device according the to manufacturer?s instructions. 
       3.Allow printed microarrays to dry overnight in a slide boxa. 
       4.Soak slides twice in 0.2% SDS for 2 min at room temperature with vigorous agitationb. 
       5.Soak slides twice in ddH2O for 2 min at room temperature with vigorous agitation. 
       6.Transfer slides into ddH2O at 95-100 deg. C for 2 min to allow DNA denaturation. 
       7.Allow slides to dry thoroughly at room temperature (~5 min). 
       8.Transfer slides into a sodium borohydride solutionc for 5 min at room temperature to reduce free aldehydes. 
       9.Rinse slides three times in 0.2% SDS for 1 min each at room temperature. 
      10.Rinse slides once in ddH2O for 1 min at room temperature. 
      11.Submerge slides in ddH2O at 95-100 deg. C for 2 secondsd. 
      12.Allow the slides to air dry and store in the dark at 25 deg. C (stable for &gt;1 year). 

             a. Drying increases crosslinking efficiency. Several days or more is acceptable.

             b. This step removes salt and unbound DNA.

             c. Dissolve 1.0 g NaBH4 in 300 ml phosphate buffered saline (PBS). Add 100 ml 100% ethanol to reduce bubbling.
             Prepare JUST PRIOR to use!

             d. Heating the slides greatly aids in the drying process.
<protocol xml:space="preserve"
  title="Schena and Davis: Generating control mRNAs by in vitro transcription"
  type="rna_prep">
http://www.wenet.net/~telechem/DNA-Microarray-Protocols/
</protocol>Equipment and Reagents

     RNA Transcription Kit (Stratagene) 

Method

       1.Obtain a heterologous cDNA clone inserted into a Bluescripta vector. 
       2.Digest 10 µg plasmid DNA in a 25 µl reaction with a restriction enzyme that cuts at a unique site downstream of the polyA
         tractb. 
       3.Extract the linearized plasmid once with phenol, once with phenol/chloroform/isoamyl alcohol (25:25:1 v/v), and three times
         with etherc. 
       4.Incubate the extraction for 5 min at 65 deg. C and vortex gently to drive off the residual ether. 
       5.To the 25 µl linear plasmid, add 0.1 vol of 2.5 M sodium acetate and 2 vol of ethanol. Mix by vortexing. 
       6.Pellet the linearized plasmid by centrifugation in a microfuge for 5 min at room temperature. 
       7.Remove and discard the supernatant and dry the DNA pellet in a speedvac. 
       8.Resuspend the linearized plasmid DNA in 10 µl of 1X TE (10 mM Tris-Cl and 1 mM EDTA) pH 8.0 for a final concentration
         of ~1.0 µg/µl. 
       9.To a microfuge tube, add 20 µl of 5X transcription buffer, 4 µl linearized plasmid (1.0 µg/µl), 16 µl rNTP mix (2.5 mM each),
         4 µl DTT (0.75 M), 3 µl RNase Block (20 units/µl), and 52.5 µl H2O. Mix by tapping the microfuge tube gently. 
      10.Add 0.8 µl of T3 or T7 RNA polymerase (50 units/µl)d. Mix by tapping the microfuge tube gently. 
      11.Incubate the reaction for 30 min at 37 deg. C. 
      12.Following the incubation, add 10 µl of 2.5 M sodium acetate. 
      13.Extract the 110 µl reaction once with phenol/chloroform/isoamyl alcohol (25:25:1 v/v) and three times with ether. 
      14.Incubate the extraction for 5 min at 65 deg. C and vortex gently to drive off the residual ether. 
      15.Add 220 µl of ethanol and pellet the in vitro transcripts by centrifugation for 5 min in a microfugee. 
      16.Remove and discard the supernatant and dry the DNA pellet in a speedvac. 
      17.Resuspend the in vitro transcript in 10 µl of 1X TE (10 mM Tris-Cl and 1 mM EDTA) pH 8.0 for a final concentration of
         ~4.0 µg/µlf. 
      18.Use in vitro mRNAs as controls for gene expression monitoring (see protocols 7, 9). 

       a.Clones should be as divergent as possible. Plant (e.g. Arabidopsis) clones work well for human and yeast experiments and
         vice versa. Clones should have a polyA tract of 20-40 nucleotides to allow for priming by oligo-dT in the reverse
         transcription reaction. 
       b.Linearize the plasmid (e.g. BamH I) such that the in vitro transcript contains the polyA+ site, but as little vector sequence as
         possible. 
       c.The extractions remove any trace ribonucleases and prevent degradation of the in vitro transcripts. 
       d.The choice of whether to use T3 or T7 is made by selecting the polymerase that will produce a sense transcript from the cloned
         cDNA. The sense strand will contain the polyA+ tract as the top strand at the 3? end immediately upstream of the restriction
         site used to linearize the plasmid. 
       e.The in vitro transcription reaction is highly efficient. A large RNA pellet should be visible upon centrifugation. 
       f.Transcript integrity can be assessed with formaldehyde gels or by alkaline agarose electrophoresis of cDNAs generated from
         the mRNAs by oligo-dT priming and reverse transcription (see protocol 7). 
<protocol xml:space="preserve"
  title="Schena and Davis: Generating fluorescent cDNA controls by linear PCR"
  type="cdna_and_labeling">Equipment and Reagents

     Perkin Elmer 9600 Thermal Cycler (or equivalent) 
     QIAquick PCR purification kit (Qiagen #28106) 

Method

       1.Obtain a heterologous cDNA cloned into a plasmid vector. 
       2.Amplify the cDNA insert by PCR using cDNA-specific primers. 
       3.Purify the amplified cDNA insert using a QIAquick column. 
       4.Evaporate the sample to dryness in a speedvac. 
       5.Resuspend the purified cDNA insert in 10 µl 1X TE (10 mM Tris-Cl and 1 mM EDTA) pH 8.0. 
       6.To a microfuge tube, add 71 µl H20, 10 µl 10X PCR buffer (500 mM KCl, 100 mM Tris-Cl pH 8.3, 15 mM MgCl2, 0.1%
         gelatin), 10 µl dNTPs (2 mM each), 5 µl Cy5-dCTP (1 mM)a, 2.0 µl 20-mer oligonucleotide (100 pmole/µl), 1 µl cDNA PCR
         product (0.5 µg/µl). Mix by tapping the microfuge tube gently. 
       7.Add 1.0 µl Taq DNA polymerase (5 units/µl). Mix by tapping the microfuge tube gently. 
       8.Generate fluorescent, single-stranded cDNAs by linear amplification of the template according the the following regime:
         [denature at 95 deg. C for 2 min, amplify for 30 cycles of (94 deg. C 30 sec, 55 deg. C 30 sec, 72 deg. C 30 sec), extend at 72 deg. C 3 min, hold at
         4 deg.  until ready to purify]. 
       9.Purify the fluorescent linear amplification products on a QIAquick column. 
      10.Evaporate the purified products to dryness on a speedvac. 
      11.Resuspend the pellet in 50 µl of 1X TE (10 mM Tris-Cl and 1 mM EDTA) pH 8.0. The concentration of the fluorescent,
         single-stranded cDNA should be ~40 ng/µl. 
      12.Add 1.0 µl of the 40 ng/µl fluorescent control per 20 µl hybridization buffer (Protocol 7, step 18) to provide a fluorescent,
         single-stranded cDNA control at ~2 ng/µlb. 

       a.Alternate fluors such as F12-dUTP, L5-dCTP and Cy5-dCTP can also be used. 
       b.Controls of this type provide a measure of hybridization and scanning, independent of an enzymatic labeling step such as
         reverse transcription (see Protocol 7). A 2 ng/µl single-stranded product should produce an intense fluorescent signal
         equivalent to an abundant cellular transcript. Multiple fluorescent cDNAs can be used generate a concentration series. 
Reagents and Equipment

     StrataScript RT-PCR kit (Stratagene #200420) 
     Oligo-dT 21 mer (treated with 0.1% DEPC to inactivate ribonucleases) 
     100 mM dATP, dCTP, dGTP, dTTP (Pharmacia #27-2050, -2060, -2070, -2080) 
     1 mM Cy3-dCTP (Amersham #PA53021) 
     1 mM Cy5-dCTP (Amersham #PA55021) 
     1 mM fluorescein-12-dCTP (DuPont #NEL-424) 
     SuperScript II RNase H- Reverse Transcriptase (Gibco BRL #18064-014) 

Method

       1.In a micro-centrifuge tube, mix 5.0 µl of total polyA+ mRNA (1.0 µg/µl)a, 1.0 µl of control mRNA cocktail (0.5 ng/µl)b, 4.0
         µl oligo-dT 21mer (1.0 µg/µl), and 27.0 µl of H2O (DEPC-treated). 
       2.Denature mRNA 3 min at 65 deg. C. 
       3.Add 10.0 µl 5X first strand buffer (250 mM Tris-HCL (pH 8.3), 375 mM KCl, 15mM KCl), 5.0 µl 10X DTT (0.1M), 1.5 µl
         Rnase Block (20 U/µl), 1.0 µl dATP, dGTP, dTTP cocktail (25 mM each), 2.0 µl dCTP (1 mM), 2.0 µl Cy3-dCTPc (1 mM)
         and mix by tapping the micro-centrifuge tube gently. 
       4.Add 1.5 µl SuperScript II reverse transcriptase (200 U/µl) for a total reaction volume of 50.0 µl. 
       5.Mix by tapping the micro-centrifuge tube gently. 
       6.Anneal Oligo-dT to mRNA 10 min at 25 deg. C. 
       7.Reverse transcribe the polyadenylated RNA for 2 hrs at 37 deg. C. 
       8.Add 5.0 µl of 2.5 M sodium acetate and 110 µl 100% ethanol at 25 deg. Cd. 
       9.Centrifuge for 15 min at 25 deg. C in a microfuge to pellet cDNA/mRNA hybridse. 
      10.Remove and discard supernatant and carefully wash pellet with 0.5 ml 80% ethanolf. 
      11.Dry pellet in a speedvac and resuspend in 10.0 µl 1X TE (pH 8.0)g. 
      12.Boil sample 3 minutes to denature cDNA/mRNA hybrids. Chill on ice immediately. 
      13.Add 2.5 µl 1 N NaOH and incubate 10 min at 37 deg. C to degrade the mRNA. 
      14.Neutralize the cDNA mixture by adding 2.5 µl 1 M Tris-HCl (pH 6.8) and 2.0 µl 1 M HCl. 
      15.Add 1.7 µl 2.5 M sodium acetate and 37 µl 100% ethanol. 
      16.Centrifuge for 15 min at full speed in a microfuge to pellet the cDNA. 
      17.Remove and discard supernatant and wash pellet with 0.5 ml 80% ethanolf. 
      18.Dry pellet in a speedvac and resuspend in 13.0 µl H2Og. 
      19.Add 5.0 µl 20 X SSC [3 M NaCl, 0.3 M sodium citrate (pH 7.0)] and 2.0 µl 2% SDS. 
      20.Heat at 65 deg. C for 30 sec to dissolve fluorescent mixture. 
      21.Centrifuge for 2 min. in a microfuge at high speed to pellet trace debrish. 
      22.Transfer supernatant to a new tube. 
      23.The final probe concentration will be ~0.25 µg/µl per fluor in 20 µl of 5 X SSC and 0.2% SDS. 

       a.Total mRNA purified from total RNA using Oligotex-dT (Qiagen). 
       b.Control mRNAs from in vitro transcription are doped in at molar ratios of 1:1,000, 1:10,000 and 1:100,000 assuming a 1.0 kb
         average length for mRNAs. 
       c.To label mRNA with other fluors, substitute Fl12-dCTP or Cy5-dCTP in the reaction. 
       d.Chilling or use of &gt;2 volumes of ethanol results in the unwanted precipitation of free label. 
       e.Pellet product on one side of the tube, then remove supernatant from the other side. 
       f.To prevent loss of pellet, centrifuge 1 min before removing 80% ethanol. 
       g.The product often smears up the side of the tube. Re-suspend thoroughly. 
       h.Insoluble material can lead to elevated background fluorescence. http://www.wenet.net/~telechem/DNA-Microarray-Protocols/
</protocol>
<protocol xml:space="preserve"
  title="Schena and Davis: cDNA microarray hybridization and washing"
  type="other_protocol">Reagents and Equipment

     Hybridization cassettes (TeleChem) 
     Microarray wash station (TeleChem) 
     ScanArray 3000, 4000, or 5000 (GSI Lumonics) 

Method

       1.Place the microarray in a hybridization cassette. 
       2.Add 5.0 µl of 5 X SSC + 0.2% SDS to the slot in the cassette for humidificationa. 
       3.Pipette 6.0 µl of fluorescent probe along the edge of a 22 x 22 mm cover slip. 
       4.Place the cover slip onto the microarray using forceps such that the sample forms a thin monolayer between the cover slip and
         the microarrayb. 
       5.Seal the hybridization cassette containing the microarray. 
       6.Submerge the hybridization cassette in a water bath set at 62 deg. Cc. 
       7.Hybridize for 6 hrs at 62 deg. C. 
       8.Following hybridization, remove the microarray from the hybridization cassette and place it immediately into the wash
         stationd. 
       9.Wash the microarray for 5 min at room temperature in 1X SSC + 0.1% SDSe. 
      10.Transfer the wash station and microarray to a second beaker containing 400 ml 0.1 X SSC and 0.1% SDS. 
      11.Wash the microarray for 5 min. at room temperature 0.1 X SSC and 0.1% SDS. 
      12.Rinse the microarray briefly in a third beaker containing 0.1 X SSC to remove the SDS. 
      13.Allow the microarrays to air dry. 
      14.Scan the microarrays for fluorescence emission in a ScanArray 3000. 

       a.Prevents drying of the sample under the cover slip. 
       b.Cover slips must be free of oils, dust and other contaminants. Lower the cover slip onto the microarray from left to right so
         that the sample pushes out air bubbles as it forms a monolayer against the microarray surface. Small air bubbles trapped under
         the cover slip exit after several minutes at 62 deg. C. 
       c.A temperature of 62 deg. C works well for cDNA-cDNA hybridizations. Lower temperatures should be used for hybridization to
         oligonucleotides. 
       d.Wash station should be placed in a 600 ml beaker containing 400 ml 1X SSC + 0.1% SDS. The microarray should be
         transferred quickly from the cassette to the wash station. Leaving the microarray at room temperature will lead to elevated
         background fluorescence. 
       e.The cover slip should slid off the microarray during the wash step. If the cover slip does not slid off within 30 sec, use
         forceps to gently remove it from the microarray surface. Failure to remove the cover slip will prevent efficient washing of the
         microarray. 
http://www.wenet.net/~telechem/DNA-Microarray-Protocols/
</protocol>
<protocol xml:space="preserve"
  title="Schena and Davis: Gene expression analysis with microarrays"
  type="other_protocol">Reagents and Equipment

     SuperAldehyde Substrates (TeleChem) 
     cDNA microarrays 
     Fluorescent sample derived from mRNA 
     Microarray gridding robot (Various) 
     ScanArray 3000, 4000 or 5000 

 

Method

       1.Manufacture a cDNA microarray containing array elements corresponding to genes of interesta. 
       2.Prepare a fluorescent sample from total mRNA by enzymatic labeling. 
       3.Hybridize the fluorescent sample to the microarray under conditions of high stringency. 
       4.Wash the microarray to remove unhybridized sample. 
       5.Scan the microarray with the ScanArray 3000 to collect fluorescent emission. 
       6.Quantitate the fluorescent emission at each position within the microarray. 
       7.Assign gene expression values by comparing the experimental data to the appropriate controls. 

       a.Microarray elements can be cDNAs of known or unknown sequence representing a restricted subset of genes or all of the
         genes in a given genome. Synthetic oligonucleotides made from expressed sequences (e.g. ESTs) have also been. 
http://www.wenet.net/~telechem/DNA-Microarray-Protocols/
</protocol>
<protocol xml:space="preserve"
  title="Schena and Davis: Mutation detection with oligonucleotide microarray"
  type="other_protocol">Reagents and Equipment

     Synthetic 15-mer oligonucleotides, amino-modified (Stanford PAN facility) 
     SuperAldehyde Substrates (TeleChem) 
     2X Micro-Spotting Solution (TeleChem) 
     Fluorescent sample derived from DNA 
     UniHyb Hybridization Solution (TeleChem) 
     Microarray gridding robot (Various) 
     ImaGene software (BioDiscovery) 
     ScanArray 3000 (GSI Lumonics) 

Method

       1.Prepare a 15-mera oligonucleotide microarray wherein the central (8th) position identifies the polymorphism or mutation in the
         fluorescent sampleb. Microarrays are made by spotting 10-100 pmole/µl oligonucleotide in 1X micro-spotting solution. 
       2.Process the microarray to remove unbound 15-mer. 
       3.Prepare a fluorescent sample by PCR amplification of the locus encompassing the polymorphism or mutationc. Use ~1/10 of a
         100 µl PCR reaction for hybidization of a sample that contains &lt;1,000 loci. Purify the sample prior to hybridization by ethanol
         precipitation or spin column purification. 
       4.Denature the sample by boiling for 2 min prior to hybridizationd. 
       5.Hybridize the fluorescent sample to the oligonucleotide microarray for 4 hrs at 42 deg. Ce. 
       6.Wash the microarray to remove unhybridized sample as follows: twice for 5 min each at room temperature in 2X SSC, once
         for 5 min at room temperature in 2X SSC. 
       7.Allow the microarray to air dry. 
       8.Scan the microarray at the highest PMT and laser settings that preserve linearity and minimize backgroundf. 
       9.Quantitate fluorescent intensities with ImaGene software. 

       a.Oligonucleotides must be coupled covalently to the solid support. We have used microscope slides with reactive aldehyde
         groups and primary amines on the oligonucleotides to mediate covalent end attachment. 
       b.The central or 8th position in a 15-mer is used to identify a single base polymorphism or mutation by hybridization. For a
         marker in which the wild type is a &quot;G&quot; and the mutant is a &quot;T&quot;, the two complementary 15-mers would be identical except at
         position 8 in which the wild type 15-mer would contain a &quot;C&quot; and the mutant oligonucleotide would contain an &quot;A&quot;. 
       c.Fluorescent primers spanning the allele of interest by ~60-bp will yield a product that hybridizes efficiently to the
         oligonucleotide microarray. 
       d.Double-stranded fluorescent products must be denatured prior to hybridization. Single-stranded fluorescent samples made by
         linear PCR are preferable. 
       e.Hybridization temperature should be ~10 deg. C below the Tm. 42 deg. C works well for 15-mers. The temperature should be adjusted
         for longer or shorter oligonucleotides. 
       f.On the ScanArray 3000, laser and PMT settings of 70% and 80%, respectively, work well for most genotyping applications. 
http://www.wenet.net/~telechem/DNA-Microarray-Protocols/
</protocol>
<!--
<protocol xml:space="preserve"
  title=""
  type=""></protocol>
-->
</protocol_list>

