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Project Documentation & Protocols: Maize Gene Discovery Project: Microarrays: Protocols

Contents: Index | Progress | Controls | Libraries | Arrays | Data | Protocols | Ordering | Links | FAQs

Instructions to Microarray Users
Before Handling Slides
Your Microarrays
Your Spiking Controls
Steps for Hybridization, Data Aquisition, and Data Analysis
Your Microarray Format
Navigating Your Microarray
Preliminary Analysis of your Microarray Data
1. Immobilization of DNA on Sigma Silane Slides
2. Preparation of Spiking Controls
3. Preparation of Total RNA (using TRIzol Reagent-GIBCOBRL)
4. Preparation of polyA+ mRNA (using Dynabeads Oligo (dT)25--Dynal)
5. Direct Labeling of polyA+ mRNA
6. Hybridization of Labeled Targets to Microarrays

Instructions to Microarray Users


Take care to handle slides only by the labels or edges to avoid smearing or leaving residues from your fingers on the slides. Your slides have been shipped to you in a polyfoam slide case. They should be stored dry (in the case, sealed within a plastic bag containing desiccant) at room temperature no longer than 12 months. Inside the case, slides may be shipped with 1 or 2 slides per slot depending on the number of slides you ordered. If there are 2 slides per slot, then the slides have been shipped back-to-back and must be separated and carefully reoriented before use so that the label side of both slides is up. The DNA is printed on the side with the label-If you can read the label, you have the correct side up and it is safe to set your slides down on a clean, dry surface.


Check the label of your slides for EST library and printing format information. The slide number provides the EST Library number /Microarray Format /Print Number / Slide number (see example next page). Click on the information button on the Microarray Elements Search page for information on the preparation and format of your slides.


We have enclosed several spiking controls on paper filter disks.


Please read the overview provided below before beginning your work. Note that you must have access to certain equipment before proceeding. Protocols for all steps are available on this webpage and on the University of Arizona webpage -see Maize Gene Discovery Protocols. We recommend that you start by following our DNA immobilization protocol that has been optimized for the slide coating on your slides. If you are a beginner, we encourage you to start with the protocols working in our lab. If you are not, we certainly encourage you to follow your own protocols for steps other than DNA immobilization and contribute back to us what works best for you.

1. Immobilization of DNA on slides. Your slides have been sent to you WITHOUT immobilization of the DNA. You will need to MARK THE BOUNDARIES OF YOUR ARRAY before immobilization and proceed with the protocol before you begin handling your slides to any extent-slides may remain in the slide box without DNA immobilization. You will need access to a UV cross-linking instrument. We suggest you try rehydration for 10 seconds on your first slide(s) and add time (10-20 seconds) if you feel you need larger spots-be careful or spots will run together.

2. Preparation of mRNA. For each hybridization, you will need ~4 µg of polyA+ mRNA for each treatment-if you plan to hybridize a single slide with mRNA from two different sources (e.g. a control and a treatment), you will need a total ~8 µg of mRNA per slide. We routinely isolate total RNA using TRIzol (GibcoBRL) and purify using DynaBeads Oligo (dT)25 (Dynal) according to manufacturers instructions.

3. Preparation of Spiking Control. If you plan to use one or more spiking controls, you will want to prepare them prior to direct labeling. A small aliquot of the spiking control can be added to the labeling reaction mix for each treatment.

4. Direct Labeling of polyA+ mRNA. We routinely incorporate label directly as Cy3- or Cy5-labeled dUTP (Cy3-, Cy5-dUTP from Amersham Pharmacia catalog numbers PA53022 and PA55022), using Sigma.s AMV-RT Kit (catalog number STR1-KT) according to manufacturers instructions. You will need access to the reagents above as well as to waterbaths at various temperatures.

5. Hybridization. We follow a modified version of the protocol recommended by the manufacturer for Sigma Screen Silane slides (Modified from Sigma Technical Bulletin MB-745 November 1999-Product No: A7718 ArrayHyb Buffer). You will need to purchase certain reagents (see Protocol) as well as have access to a reliable 60-65 C incubator.

6. Data Acquisition. You will need reliable access to a Microarray slide reader. If you are using a fluorescent dye to label your hybridization mix, your slides should be scanned within 8 hours after hybridization.

7. Data Analysis. Check the slide label for the Microarray format used to print your slide. Microarray Element Tables are provided on our webpage for each microarray format used for printing. A search table allows you to quickly identify each printed spot by location and provides EST and BlastX identification. A complete Microarray Element Table for each Microarray Format can downloaded in text format for import into spreadsheets and data analysis programs. You will need access to data analysis software-We use a program called ImaGene (BioDiscovery) that must be purchased. However data analysis software is often available as part of the slide reader software package and some programs are available as freeware on the internet.

Your Microarray Format

The label of your slides provides information on the EST library and printing format used to make your slides (note there may be more than one printing format for an EST Library). The slide number provides the EST Library number/Microarray Format /Print Number/Slide number (see example below). You can click on the information button below the Microarray Element Table on our webpage for specific information on the preparation and format of your slides.

Navigating your Microarray

Each slide is composed of eight subarrays. When a slide is oriented with the label at the bottom (see below), the subarrays are positioned in 4 Rows and 2 columns. The location of each spot in a subarray is given by 4 variables: Subarray Row, Subarray Column, Row within Subarray, Column within Subarray. Once you know the location of a printed spot, you can use the information in the Microarray Element Table to provide EST and BlastX identification. A complete Microarray Element Table for each microarray format can downloaded in text format for import into spreadsheets and data analysis programs.


Preliminary Analysis of your Microarray Data

Recommended protocols for immobilization of DNA on slides, preparation of spiking controls and RNA samples, labeling targets, and hybridization are given in the next section. After completion of your hybridization, you will need to scan your slide immediately and acquire an image of your hybridized microarray. Data analysis begins with the use of software to find spots and acquire signal intensity and background information for each spot. You will need access to data analysis software. Typically data analysis software comes with or is recommended for use with your scanner. Free data analysis software can also be found on various Microarray websites-we do not provide data analysis software.

A. Obtaining signal intensity data, obtaining a key to spot identity, and merging data and identity files

  1. Spot finding and obtaining signal intensity data with your data analysis software. Most data analysis programs need to know the number of rows and columns of subarrays on the slide and the number of rows and columns of spots within a subarray. Please familiarize yourself with a) the previous section, Navigating your Microarray and b) the Print Design information for your Microarray. Navigating your Microarray provides an illustration of a slide and a slide label. It tells how the Microarray format can be determined from the slide label. It also defines rows and columns and demonstrates how subarrays are organized in rows and columns and spots are organized into rows and columns within subarrays. Print Design information specific to your Microarray can be found by clicking the information button at the bottom of the Microarray Element Search Table. You must be sure to consult the Print Design Information that corresponds exactly to the format of your Microarray. Check the slide label for the Microarray format used to print your slide. Then page through the information provided on how your microarray probes were prepared. The Print Design section provides information on exactly how many rows and columns of subarrays are on your slide (Slide Format) and how many rows and columns of spots are within each subarray (Subarray Format). Keep in mind when spot finding that most of the last row of a subarray may be blank-we typically did not have enough EST elements to completely fill every subarray.
  2. Example: The 614 microarray has 8 subarrays printed in four rows and 2 columns (see Navigating Your Microarray for an illustration defining rows and columns). Within each subarray, the spots are printed in 45 rows and 45 columns. When spot finding, keep in mind there are several blank spaces in the last row of each subarray-this can be verified by consulting the Microarray Element Search Table. Select 614 as the Array type, 45 for Row # within Subarray, and blank for Stanford ID.

  3. Obtaining a key to identifying ESTs on your microarray. You will need to download a Microarray Element Search Table from our web pages--this provides a key to the identification of every printed spot on your Microarray. You must be sure to download the Microarray Element Search Table that corresponds exactly to the format of your Microarray. Check the slide label for the Microarray format used to print your slide. Then go to the Microarray Element Search Table and select the Array type that matches exactly the Microarray Format printed at the bottom of your slide. Most PC-based software will require that you import the search table as a text document-select TEXT under Format. Make sure the Show All Columns is selected and that no other information is entered in the table or you will not get a complete data table-if necessary, select RESET and start again. Next, select GO and the Search Table will appear. Simply use the Save As command to save this file on your personal computer as a text file. This will save your file as a tab-delimited text file under the name you gave it.
  4. Opening your Microarray Element Search Table within a PC-based spreadsheet. For example, we use EXCEL (Microsoft) to manipulate our data. Within EXCEL, we simply open the text file containing the Microarray Element Search Table we downloaded. We then select the delimited option and the tab option for the delimiter. When opening your Microarray Element Search Table within a PC-based spreadsheet, make certain that you select the columns titled StanfordID and GenBank Accession Number (and to be safe all the BlastX and Description text columns) and when given the option to specify column data format, select Text. If you do not do this, all ESTs with a Stanford ID such as 614053E07 may be put in scientific notation-and other unwanted results could occur depending on your program. You can now resave the Microarray Element Search Table in the appropriate format for your spreadsheet program.
  5. Merging the Microarray Element Search Table with your signal intensity data. Finally, you will want to merge our Microarray Element Search Table with your signal intensity data. Simply copy all of the information from the Microarray Element Search Table into your data analysis output table (we recommend you simply add our Microarray Element Search Table to the columns at the right of your output table-same worksheet). Here are some important steps to make certain that you have correctly identified each spot on your microarray:
    1. After making certain that the two data sets start on the same line, page to the end of the document and make certain that you have the same number of lines from your output data table as you do from our Microarray Element Search Table. If you do not, then you have either incorrectly analyzed your microarray with your data analysis software or you have imported the wrong Microarray Element Search Table.
    2. Most data analysis programs will identify each spot by the subarray row and column and the row and column within a subarray where the spot is located. Make certain you understand how the output from your data analysis software is organized (e.g. make certain that subarray rows and columns, and spot rows and columns are designated the same way we designate them-for example some programs may have row and column designations reversed relative to our output). Consult Navigating your Microarray if you need an illustration of how our slides are organized.
    3. Make certain that the two data sets are sorted exactly the same way. Our Microarray Element Search Table is sorted by Subarray Row, Subarray Column, Row, and Column. To verify that both data sets are sorted exactly the same way, page down through the merged file and make certain that spot location information from both data sets match exactly. If the spot location information does not correspond, you will have to select all of the columns from one of the data sets and sort them the same way the other is sorted.
    4. Finally, check signal intensity data to see if they make sense. For example, the 614 microarray is printed with 3 side-by-side replicate spots with blanks printed in several locations. Does your signal intensity data seem to indicate that replicates are correctly grouped and that spots identified as blanks have low signal values? If not, you may need to repeat the previous steps.

B. Computing Signal Intensity and Background, Data filtering, Calculation of Preliminary Statistics, Estimating Reproducibility, and Representing Data

Please keep in mind that there is currently no consensus on how best to analyze or represent microarray data. The following discussion proposes several considerations and data analysis steps-keep in mind that microarray data analysis is a constantly evolving field and our recommendations are likely to change. We recommend that you consult the literature for the protocols that are the most up-to-date and relevant to your experiments.

  1. Understanding How Signal Intensity Data are Calculated. You should familiarize yourself with your scanner and data analysis programs to insure you obtain the best raw data for your analyses. For example, many scanners allow the user to select laser intensity, photomultiplier intensity, and other variables. Since output files from scans become the primary data for the rest of the analyses, care should be taken to understand how best to optimize scanning with your instrument for your experiment and to record exactly how signal intensity data were acquired. Similarly, most data analysis software enables you to select how spots are defined and how signal intensity and background are calculated. Although primary data files can always be reanalyzed, similar care should be taken to insure that you understand how to optimize analyses with your software and how signal intensities for each spot were calculated.
  2. Correcting signal intensity for background. Some analysis programs may correct for background automatically-others provide background data. Data analysis software may compute a background correction for each spot, for an entire subarray, or the entire slide. To correct for background, the user would create a new column to hold the corrected signal intensity value (raw signal intensity minus background correction). For example, using ImaGene (BioDiscovery) software, we often calculate the corrected signal intensity by using the median signal intensity minus the median background for each spot.
  3. Low signal filtering. Low signal filtering enables very low and negative corrected signal intensity values to be set to a positive minimum that is essentially what we would call signal background. Low signal filtering helps minimize analysis problems such as negative or highly variable signal ratios for spots with essentially null signal intensity. To calculate this minimum, we typically use either the background information provided by the data analysis software or data for blanks. On every slide, we have included a series of controls including blanks or spots where the pins printed only buffer. We typically compute signal background either by taking a) the mean of the local background for every spot plus 2 standard deviations, or b) the mean of the blanks plus 2 standard deviations. Corrected signal intensity values less than the signal background, are then set equal to this minimum.
  4. Computing signal intensity ratios for each spot. Because printed elements differ in many characteristics important to hybridization (including length, sequence, GC content, and concentration), microarray experiments generally focus on the ratio of signal intensities from labeled targets derived from two different treatments. Typically, one of the treatments can be identified as the control or standard and the other as the experiment. Ratios are then computed by dividing the signal from the experimental population by the signal from the control population. For example, when we hybridize targets derived from ear and endosperm on our 605 Endosperm Microarray, we would designate the endosperm treatment as the standard. For each spot, we would then calculate a ratio by dividing the signal from the ear target by the signal from the endosperm target.
  5. Normalization of signal ratios. Factors other than treatment effects can produce ratios of signal intensities that differ from a value of 1-when such factors affect the ratio of all elements on the microarray, normalization of ratios is in order. Such factors include differences in the incorporation efficiency of fluorescent label, differences in the amount of RNA in the two samples, or other non-treatment effects. There are various methods to determine when and how to normalize signal ratios. The most common is to use correlation analysis to compute a correlation model. If only a small subset of the elements are likely to differ in response to treatment, data for all elements can be used in the correlation analysis. Otherwise a subset of elements (e.g. specific controls) is used for the correlation analysis. Corrected signal intensities in one channel are plotted against corrected signal intensities in the other channel. Correlation analysis provides the equation that describes the relationship between values. If the correlation model is significant and the slope of the trend line differs from 1-the equation can be used to normalize the data (e.g. compute normalized X values by multiplying the X values by the slope and adding the Y intercept value).
  6. Expressing Signal Ratios as positive/negative factors or log2, log10 values. In order to make our ratios a little easier to use, we frequently represent ratios less than 1 as negative numbers equivalent to the factor difference in expression between treatments. To do this, a) simply take the inverse of ratios less than 1 and b) multiply by -1. The result is that a ratio of 0.5 (two fold lower signal intensity by the experimental treatment relative to the control) is now expressed as -2. For statistical comparisons of ratios, we use a log transformation to linearize our ratio data. Both log10 and log2 transformations are frequently used.
  7. Calculating mean, standard deviation, and CV of replicate data. We often find it useful to start a new worksheet that summarizes raw data by the mean of replicate spots. If more than two replicates are printed, we also calculate the standard deviation and coefficient of variation (standard deviation/mean * 100) for replicate spots. We use mean data to compare signal intensities and ratios between slides. We use CV as measure of variability within replicates.
  8. Reproducibility of hybridization. We recommend at least two replicate microarrays be used for each experiment--many laboratories suggest that the two replicates be dye-reversal hybridizations (e.g. fluorescent dyes used to label treatments are reversed on the two microarrays). We typically use correlation analysis to estimate reproducibility of hybridization. We compare signal intensity values and signal ratios (log10 transformed) of replicates on the same slide. We also compare mean signal intensity values and mean ratios (log10 transformed) of replicates between slides. Pearson correlation coefficient can be used to describe the strength of the relationship between replicates. Examples of reproducibility data can be found on our Data pages. If reproducibility is low, more replicates are needed.
  9. Reporting Microarray Data. The focus of most expression studies is identifying microarray elements for which the level of signal intensity in one treatment differs from the signal intensity of the other. Frequently, experimenters highlight elements that differ in signal intensity by two-fold as having different expression profiles. Experimenters may represent their data several ways:
    1. Graphically. Typically mean signal intensities for the control population are graphed on the X axis and mean signal intensities for the experimental population are graphed on the Y axis. Elements that differ in signal intensity by two-fold or greater may be highlighted.
    2. Tabular. Genes with 2 fold differences in expression can be categorized by hybridization intensity (for example, 100-90, 89-75, 74-50, 49-25, 24-10, 9-0% maximum signal in the control channel). Categorizing data in this way has several benefits. First, it provides a method for visualizing gene expression profiles relative to hybridization intensity. Second, because ratios at the extremes have different problems, categorizing genes into high, medium, and low hybridization classes enables the experimenter to look at each separately. For example, at the high end, signal intensity in one channel may have saturated making differences in expression between treatments less apparent. At the low end small differences between treatments can produce large and often meaningless differences in ratios.
    3. Relative to Controls. Frequently levels of gene expression are described relative to various control genes. Typically, microarrays are printed with an assortment of housekeeping genes as well as genes with known patterns of gene expression. Also microarrays may include spiking controls or other genes thought to have low homology to the rest of the printed genes. These control genes may be used as standards for comparison. Please consult the Control Pages for details on the control elements included in our microarrays.

Once again, we recommend that you consult the literature for the analysis protocols that are the most up-to-date and relevant to your experiments.




(these slides need backing at 80 C immediately after printing, which is done at the microarray facility at U of A)

1. Mark the boundaries of the array on each slide using a diamond scriber (array will become invisible after post-processing). Be careful not to press so hard slides break.

2. Hold the slide (face down) over a 42 °C water bath for 5-10 seconds to the re-hydrate spots. Note with high density printing (center to center spacing 200 um or less), do not exceed 10 seconds re-hydration time (pre-warming the slides DNA-side up for 3-5 seconds over a 65 C heat block will reduce the chances for over re-hydration)

3. Snap-dry each slide (DNA side up) on a 65 °C heat block for 3-10 seconds until the humidity is gone. Repeat sets 2 & 3 three times.

4. Place arrays in a slide rack and UV crosslink DNA to glass with 65 mJ of 254 nm UV light. (We use a Sigma Humidity Chamber {Sigma H6644} to hold the slides. We use a Fisher Scientific FB UVXL-1000 UV Cross Linker set to 650 x 100 uJ/cm2 for crosslinking DNA).

5. Incubate slides for ~ 5 minutes in 1% SDS on orbital shaker. This step removes the unbound nucleic acids from the arrays and prevents them from binding elsewhere. (We use Sigma staining chambers and racks {Sigma S6141} for all rinses. Make sure the volume in chamber completely covers slides while shaking).

6. Gently plunge the slide rack in boiling water for 2 min to denature the DNA.

7. Rinse slides by plunging rapidly 10-20 times in a 100% ethanol bath (use a clean staining chamber and transfer slides to ethanol still in same staining rack).

8. Transfer slides to 50 ml disposable centrifuge tubes and spin slides at 50-100 x g for 2-5 min. This step must be done quickly and carefully to avoid streaking and smearing arrays.

9. Use arrays immediately or store in a slide box (room temperature, with desiccant).

2. Preparation of Spiking Controls (Modified from Hong Wang-University of Arizona 8/1/99)

we recommend to use National spiking controls (SP1-SP6) which is included in our arrays.

Spiking controls may be used, if desired, to compare the efficiencies of the labeling reactions in experiments where microarrays are probed with RNAs from two populations (e.g. control and treatment populations). We will supply different spiking controls on paper filter disks on request (see bottom of this protocol for a description of the controls). These same spiking controls have also been printed on the arrays you received. Instructions to prepare spiking controls appear below. If you choose to use a spiking control, you would introduce it into both of your RNA samples in equal concentration prior to labeling. Care must be taken to minimize pipetting errors so that hybridization efficiency of the spiking controls will accurately reflect efficiencies of the two labeling reactions. Frequently, several spiking controls are introduced into both labeling reactions, such that each control is at a different concentration (e.g. Control 1, 2, 3, 4, 5 at 2.0, 1.0, 0.2, 0.1, and 0.02 ng, respectively). Preparation of your controls will involve standard transformation and large-scale plasmid preparation, preparation of RNA by induction of the T7 promoter, and RNA purification. Use of a spiking control is not necessary for analysis of Microarray data.

1. Reconstitute plasmid DNA (~50 ng) on the filter paper disk by placing the disk in 25 µl TE and vortexing.

2. Aliquot ~10 ng of plasmid DNA for transformation. Store remaining plasmid DNA at -20 °C.

3. Transform E.coli using standard laboratory transformation protocols. Select for transformants LB plates with appropriate antibiotic (see control table).

4. Check clones by preparing a 500 µl miniprep, digesting with the restriction enzyme , and analyzing for single bands on a 1.0 % agarose gel.

5. Prepare a large-scale plasmid preparation (50 ml culture) of the confirmed clones with PEG purification. Measure the plasmid concentration.

6. Aliquot ~10 µg of plasmid DNA and repurify plasmid using a Qiagen PCR purification column. Elute the DNA in 30 µl Tris-HCl (10 mM, pH 8.0). Store remaining plasmid DNA at -20 °C.

7. Digest the total 30 µl with the restriction enzyme in a 100 µl reaction mix. After 2 hours, aliquot 1-2 µl and run a 1.0 % agarose gel to check the digestion.

8. Extract the linearized plasmid once with an equal volume of phenol:chloroform and then 3 times with an equal volume of chloroform.

9. Add 0.1 volume of 3 M sodium acetate (ph 5.2) and 2 volumes of ethanol. Mix thoroughly and precipitate at -20 °C for ~ 2hrs.

10. Spin at 12,000 x g at 4 °C for 15 min.

11. Wash pellet with 70% ethanol and dry with SpeedVac. Resuspend the linearized plasmid in 10 µl of TE (10 mM Tris-HCl and 1 mM EDTA, pH 8.0). Adjust final concentration to ~about 1.0 µg/µl.

12. Prepare transcription reaction mix. In a new Eppendorff 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), 1.5 µl RNase Block (40 units/µl), and 54 µl of DEPC treated water. Mix by tapping the tube gently.

13. Add 0.8 µl of T7 RNA polymerase (50 Units/µl). Mix by tapping the tube gently.

14. Incubate at 37 °C for 1-1.5 hours.

15. Add 2 µl RNase free DNase I and mix well. Incubate at 37 °C for 15 min.

16. Extract the reaction once with and equal volume of phenol:chloroform and 3 times with an equal volume of chloroform.

17. Add 0.1 volume of 3 M sodium acetate (pH 5.2) and 2 volumes of ethanol. Mix thoroughly and precipitate at -20 °C for ~ 2hrs.

18. Spin at 12,000 x g at 4 °C for 15 min. There should be a large white pellet at the bottom of the tube.

19. Wash the pellet with RNase free 70 % ethanol and dry by SpeedVac. Resuspend the pellet in 20 µl 1X TE (RNase free).

20. Take 1 µl to run a formaldehyde gel to check the transcript integrity. Measure the concentration by spectrophotometer.

21. Use transcripts immediately or store at -70 °C. Note, transcript quality diminishes significantly with each freeze thaw cycle.

If you would like other spiking controls (refer to Control Table on our web site) please contact us by email.

3. Preparation of Total RNA (using TRIzol Reagent-GIBCOBRL)

(Modified from Manufacturers instructions--see product insert for details or consult 'Protocols' at Different procedures can be adopted for making total RNA from different tissue types, like RNAwize from Ambion etc.

1. Homogenize 100-500 mg of tissue in liquid nitrogen. Add the frozen tissue to a prechilled mortar and pestle. Add enough liquid nitrogen to cover tissue. As liquid nitrogen evaporates (and for 1 minute thereafter) grind the tissue to a fine powder.

2. Add 1 ml TRIZOL per 100 mg of tissue and continue to homogenize. Sample may freeze at this point--continue grinding 1 minute after sample has thawed.

3. Transfer to RNase free centrifuge tubes.

4. Incubate 5 minutes at room temperature.

5. Add 0.2 ml Chloroform for 1 ml of TRIZOL and vortex 15 seconds.

6. Incubate 3 minutes at room temperature.

7. Centrifuge at 12,000 x g for 10 minutes.

8. Transfer the aqueous phase to a fresh RNase-free tube.

9. Precipitate by adding 0.5 ml isopropyl alcohol. Mix well by inverting, Incubate 10 minutes.

10. Centrifuge at 12,000 x g for 10 minutes. A very small pellet should be visible at the bottom of the tube

11. Discard supernatant by decanting. Quick spin. Remove remaining supernatant with pipetter.

12. Wash pellet by add 0.5 ml 75% ethanol (made with RNase-free water).

13. Centrifuge at 75,000 x g for 10 minutes.

14. Discard supernatant by decanting. Quick-spin in microfuge, if necessary. Invert the tube on a Kimwipe to draw off residual liquid.

15. Allow the pellet to air-dry for 5-10 minutes (e.g. until all of the ethanol has disappeared from the walls of the tube). DO NOT ALLOW THE PELLET TO FULLY DRY OUT.

16. Add 10 µl of RNase-free water and resuspend by pipetting 20-30 times. If necessary, heat at 65 °C for 10 minutes to dissolve RNA.

17. Quantify RNA using UV spectrophotometer and TE pH 8.0 as a buffer. Yield may also be quantified fluorometrically, e.g. using RiboGreen (Molecular Probes).

18. Check integrity of RNA sample and inspect for contamination with DNA by running 0.5-1.0 µg RNA with and without DNaseI treatment in a 2% agarose TBE gel (no need to run a denaturing gel).

19. Store at -20 °C for overnight or -80 °C for a longer term.


4. Preparation of polyA+ mRNA (using Dynabeads Oligo (dT)25-Dynal) (Modified from Manufacturers instructions--see product insert for details).

1. Transfer 75 µg of RNA to an RNase-free tube. Adjust volume to 100 µl with distilled DEPC treated water. If the total RNA has already been diluted more than 100 µl then add the same total volume of 2X Binding buffer (solution B) to the Dynabeads to obtain a 1X Binding buffer concentration in the resuspension step below.

2. Heat RNA to 65 °C for 2 minutes.

3. In the meantime, resuspend the Dynabeads by shaking the vial gently to obtain a homogeneous dispersion of beads. Transfer 200 µl of resuspended Dynabeads from the stock tube to an RNase-free tube. Put the tube in the Dynal MPC (Magnetic Particle Concentrator-a magnetic device). After 30 seconds (or when suspension is clear) remove the supernatant.

4. Remove vial from Dynal MPC and wash once by resuspending in 100 µl of 2X Binding Buffer. Again place vial in the Dynal MPC and wait 30 seconds before removing buffer.

5. Transfer the vial to a rack and resuspend the beads in 100 µl 2X Binding buffer. Add more 2X Binding buffer, if necessary, as described in step 1.

6. Add the total RNA to the Dynabeads suspension. Mix gently but thoroughly and anneal by gently inverting vial for 3-5 min at room temperature.

7. Place the vial in the Dynal MPC for 30 seconds and remove the supernatant.

8. Transfer the vial to a rack and wash twice with 200 µl Washing buffer (solution E) using the Dynal MPC. The Dynabeads must be mixed thoroughly in the washing buffer and the supernatant completely removed between each washing step.

9. Elute the mRNA from the Dynabeads by adding 10-20 µl of Elution solution and keep at 65 °C for 2 min. Immediately place the tube in the Dynal MPC and transfer the supernatant containing the mRNA to a new RNase-free tube. You can reuse the Dynabeads up to 4 times for the same RNA sample.

10. Quantify RNA using UV spectrophotometer and TE pH 8.0 as a buffer according to standard protocol (see insert). Check quality of RNA sample by running 0.5-1.0 µg RNA in a 2% agarose TBE gel (no need to run a denaturing gel).

11. If the eluted mRNA is not to be used immediately, store at -70 °C.

12. Regenerate the Dynabeads for reuse according to manufacturers recommendations up to 4 times.



1. Combine in a 0.2 ml tube:

Reagent Add
DNTP Mix(10 mM ea. dA, dC, dG, 2 mM dT) 2 µl
Cy3 or Cy5 dUTP(1 mM) 2 µl
primer oligo dT23(0.5µg /µl) 2 µl

mRNA template

-- µl
nuclease-free water -- µl  
Sub-Total Volume 25 µl  

2. Incubate at 65°C for 10min. Note dyes are light sensitive and so tubes should be covered with foil to shield from light.

3. Transfer to ice and add:

5X RT Buffer                                                  8 µl
Superscript II 0.1M DTT 2 µl 4 ul
RNase inhibitor (1U/µl) 1 µl

4. Incubate at 42 °C for 90min


EDTA (0.5M) 5 µl

NaOH (1M) 10 µl

7. Incubate at 65 °C for 10min.,

8. Cool to room temperature and add:

Tris-HCl (1M , pH 8.0) 12.5 µl

TE 400 µl

9. Apply to a Microcon-YM30 column and spin at 12,000 x g for 20 min, Wash column 4 times with 400 &181;l TE and spin 20 min.

10. Add 20 µl TE, mix well with pipette and elute by inverting the column and spinning at 12,000 x g for 5 min.

11. Use immediately or store in an opaque container at -20 °C for no longer than ~ 1 month.

12. You can test for label incorporation by diluting 1 ul of the product 10 100 times with TE and spotting 0.2 &181;l on a microscope slide. After drying, slide can be scanned at same laser power and PMT for both the channel. Quantify the spots and estimate the efficiency of incorporation for cy3 and cy5.

Reagents and Materials

DNTPs (Sigma DNTP100A-1KT)

Cy3~, Cy5~dUTP (Amersham Pharmacia PA53022, PA55022)

SuperscriptII (GIBCO-BRL)

Microcon-YM30 column (Millipore 42410)


1. Prepare a hybridization chamber by cutting a paper towel in half, then folding the paper towel to fit along the long axis of a of a 50mL screw-top tube. Wet the paper towel with 2X SSC, then drain the excess liquid and seal the tube. Pre-heat the tube to 60-65 °C (you can loosen lid and microwave 10 sec and then place at 65 °C until you are ready to use it).

2. Combine Cy3 and Cy5 labeled targets.

3. Prepare the hybridization mixture in a 1.5 ml microfuge tube as follows

(A=for 22x22 mm coverslip; B=for 22x40 mm coverslip):

1.5 µl 3 µl Liquid Block (Amersham RPN3601)
2.5 µl 5 µl20X SSC
1 µl 2 µl 2% SDS
20 µl 40 µl combined labeled targets, and ddH2O
25 µl 50 µl 

Final volume is determined by practicing with a glass slide, reaction mix without labeled targets, the pipetters to be used, and a hybrislip on a 65 °C heat block. The goal is to have complete coverage without having excess solution flow outside the coverslip (Sigma Hybrislip Z36 591-2). I typically use 25 and 50 µl for the small and large hybridizations, respectively.

4. Close the tube and apply a lid lock. Denature at 95 °C for 2min.

5. Transfer to ice, then spin down briefly in a microcentrifuge.

6. Preheat a blocked and denatured slide on a 65 °C heat block.

7. Apply the hybridization mixture to the array, and quickly cover the liquid with a hybrislip (coverslips for hybridization, Sigma Hybrislips Z36,591-2). Immediately, transfer the slide to the pre-warmed 50 mL hybridization chamber (step 1).

8. Incubate overnight (8-12 hrs) at 60-65°C. If background is consistently high you can reduce hybridization time to 8 hrs.

9. Wash the array with gentle shaking using the following three wash steps:

1. 2X SSC, 0.5%SDS 55 °C, 5 min.

(Coverslips should come off easily in this first wash)

2. 0.5X SSC room temp, 5min.

3. 0.05X SSC room temp, 5min.

use clean slide staining chambers (Sigma S6141) for each rinse. I put the slides in a slide rack and transfer them (in the same rack) from rinse to rinse. NOTE: Make sure the slides are completely covered by rinse solution while shaking and never let the slides dry-if you must delay between steps, leave the slides in the rinse step they are currently in until you can proceed. If the slides dry you will have high background.

10. Immediately spin dry at low speed in a centrifuge (50-100 x g for 2-5 min) to avoid streaking.

11. Scan immediately.

Last updated: Aug. 13, 2002

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