home information troubleshooting & help bench work questions
For frequently asked questions about a particular microarray platform, service, or experiment, please visit our service pages.
The Microarray Centre has compiled a list of bench work questions we are frequently asked.
Q: Are the microarrays ready to use?
Q: Is it necessary to block the microarrays before use?
A: No. However, some users have reported that they have used Bovine Serum Albumin (BSA; 0.5 mg/mL) as a blocking agent and this seemed to reduce background signals.
Q: Is it necessary to pre-hybridise the microarrays before use?
Back to Top of Pre-treatment and starting your experiment
Q: How do you isolate total RNA from cells and tissues?
A: There are many excellent kits available for the isolation of total RNA from cells and tissues. We currently use Qiagen's RNeasy kit and Gentra's VERSAGENE for total RNA isolation from cells and Trizol® reagent from Invitrogen for total RNA isolation from tissue samples.
Q: What is the best method for isolating RNA from Yeast?
A: It depends on the user. We have tried the phenol/chloroform method and additional purification with RNeasy kit. We found that additional purification with the RNeasy kit improves the quality of purified RNA.
Back to Top of Isolating RNA
Q: Your labelling protocol refers to AncT primer. What is AncT?
A: Anchored T (AncT) is an oligo dT primer (5' T20VN) to which two variable nucleotides have been added. The first variable nucleotide (signified V) represents any nucleotide except T, and the second variable nucleotide (signified N) represents any of the four nucleotides. This structure causes the primer to sit down on the RNA at the beginning of the polyA tail, thus it is "anchored".
Any company that understands standard nucleotide notation can produce mixed primers. Some companies may not use "V" and "N" or they may use some other notation. You should always talk to the company to ensure they know what you want.
Q: What is the difference between labelling with Direct/Indirect method?
A: Direct labelling is a fast one-step method while indirect labelling is two-step, incorporating an amine-modified nucleotide (amino-allyl dUTP) in the RT reaction and then coupling it with monofunctional forms of Cy3 and Cy5 dyes. In terms of the reagent costs, the indirect labelling method is about 30% less expensive but about 2 hours longer than the direct labelling method.
With our direct labelling protocol we recommend a minimum of 10 µg of total RNA. Our indirect protocol also calls for 10 µg of total RNA but we have found that we can reduce this requirement to 3-5 µg, if necessary, without much loss of signal.Certain tissues also seem to be somewhat problematic, and may require more RNA.
Q: Which labelling protocol is better, direct or indirect?
A: Neither is "better", but we've found that some RNA samples are better labelled by the indirect (aminoallyl) labelling method.
Q: Should we label each sample with both fluors?
A: Labelling each sample with both fluors is what is meant by Reciprocal Labelling or Fluor-Flips. Two separate labellings are performed where the two RNA samples are labelled opposite fluors in each case. Reciprocal labelling is necessary when using the direct labelling protocol. For indirect labelling, we recommend reciprocal labelling only when a sufficient number of replicates are being performed. When doing 1-3 replicates, we typically do not perform reciprocal labellings.
Q: Are there other labelling kits available?
A: Yes, commercially available kits that we have tried at the Microarray Centtr are CyScribe, ULS, Label IT, and Gibco Random Primer Labelling Kit. All work comparably well in comparison to our in house Direct and Indirect methods, however, cost per reaction is significantly more expensive for each of the kits.
Q: Can we store labelled probe at -20°C for over a week?
A: While we don't recommend freezing labelled-cDNA for longer than a few days, labelled-cDNA can be stored at –20°C for several weeks without significant degradation. Please refer to the Technical Note on the stability of labelled-cDNA stored at -20°C. We typically prefer to hybridise to an array the same day as labeling.
Q: Why incorporate "cold" dTTP/dCTP in the reverse transcriptase reaction of the indirect/direct labeling protocol?
A: Since the cyanine dye-dCTP (direct labeling protocol) and amino allyl-dUTP (indirect labeling protocol) conjugates are relatively large (and thus more difficult for the RT enzyme to handle), adding a small amount of dCTP (direct) or dTTP (indirect) to the reaction actually increases the amount of incorporation
Q: If one uses a chemical way to label RNA (as in the indirect labelling method), is reciprocal labelling necessary?
A: If only 1-3 replicates are being performed using the indirect labelling protocol, we typically do not perform reciprocal labelling. However, if more than 3 replicates are performed, we will do reciprocal labelling as the dyes may still incorporate slightly differently for the two samples (this also depends on how differences in the two samples being compared).
Q: Is there a way we can tell if our labelling reactions succeeded?
A: Yes, one can tell if their labelling reactions succeeded by hybridizing to the array and if appropriate controls light up then you know labelling worked. In addition, you could analyse the purified-labelled-cDNA on the NanoDrop to ensure a) you have DNA present (OD 260/280) and b) you have fluor incorporation (OD 550 for Cy3 and OD 650 for Cy5).
Q: Does the MAC have a protocol for labelling total RNA with Alexa™ dyes?
A: For using the Alexa dyes (Alexa Fluor® 555 and Alexa Fluor® 647 reactive dye decapacks for microarrays, set of 2 x 10 vials, from Invitrogen) we have found they can be substituted for Cy™ dyes using the standard indirect labelling protocol.
Q: In the indirect labeling protocol, why do you add hydroxylamine to the dye conjugation reaction after the 1 hour incubation (prior to purification)?
A: Hydroxylamine is used to neutralize the unreacted fluor. By adding the hydroxylamine prior to purification, samples that are going to be hybridised to the same array can also be purified together (without the risk of cross-labelling the two different samples).
Q: Are there ways to use less than 10 µg of total RNA for labelling?
A: There are amplified labelling methods that can be used to reduce the amount of starting material required. One method we have used with success is that published by Wang et al. (Nature Biotechnology, 2000. 18(4): p. 457-9), which uses a T7-mediated linear amplification of RNA. We are also working on exponential amplification procedures (manuscript submitted). We have successfully used the Ovation Aminoallyl RNA Amplification and Labeling System (NuGEN) for amplifying 5-100 nanograms of total RNA. We have also worked on exponential amplification procedures developed by Norman Iscove (Iscove et al. Representation is faithfully preserved in global cDNA amplified exponentially from sub-picogram quantities of mRNA. Nature Biotechnol. 2002, 20(9):940-943) that can amplifiy as little as 20 picograms of total RNA. An alternative to amplifying the RNA is to amplify the signal itself. Kits like PE’s TSA kit and 3DNA Dendrimer technology (Genisphere) are employed such amplification.
Q: What volume of labelled-cDNA do I need to put on a slide?
A: To calculate the volume needed for hybridisation (we have empirically determined that 20 pmoles of Cy-dCTP for each channel is optimal when hybridising arrays that cover the majority of the slide's surface):
- For Cy3-labeled cDNA: volume containing 20 pmoles = 3 / A550
- For Cy5-labeled cDNA: volume containing 20 pmoles = 5 / A650
Combine these volumes of each cDNA solution and adjust volume accordingly per array used.
The UHNMAC typically uses the entire amount of labelled-cDNA from each labelling reaction.
Back to Top of Labelling
Q: Does Cy3 incorporate better than Cy5?
A: It depends on the sample and the user; not all RNA labels with the same efficiency for Cy3 and Cy5 and that is why we recommend reciprocal labelling when using the direct-labelling protocol.
Q: How do I calculate Cy3/Cy5 frequency of incorporation?
A: Perform the absorbance measurements for each dye separately following purification of the labeled cDNA. Only three values are needed to perform these calculations:
- Absorbance at 260 nm (A260; measures DNA concentration)
- Absorbance at 550 nm (A550; measures Cy3 fluorescence)
- Absorbance at 650 nm (A650; measures Cy5 fluorescence)
To calculate FOI:
- For Cy3 incorporation: 58.5 x A550/A260
- For Cy5 incorporation: 35.1 x A650/A260
The results are expressed as the number of Cy-dCTP incorporated per 1,000 nucleotides of cDNA.
The numbers 58.5 and 35.1 are conversion factors calculated using:
average molecular weight of dNTPs = 324.5 g/mole
concentration of Cy-labelled ssDNA that absorbs 1 AU of 260-nm light = 37 ug/mL
- absorbtion coefficient of Cy3 = 0.15
- absorbtion coefficient of Cy5 = 0.25
- cDNA length reference point = 1000 nucleotides.
A260 relates to the concentration of DNA in the sample; A550 and A650 relate to the concentration of Cy3 and Cy5, respectively. Note that sample volume is not factored in; this is because volumes cancel out when cDNA and cyanine absorbance are measured simultaneously.
Back to Top of Cy5 and Cy3
Q: How much hybridisation solution should I use for each type of array?
A: To ensure proper coverage (thus preventing subsequent drying out) of the array, the following volumes of hybridization solution are recommended for each type of array. The volume of hybridisation solution used is actually determined by the size of the coverslip used. Generally, a 24 x 60 mm coverslip requires 80 µL of hybridisation solution and a 24 x 50 mm coverslip requires 60 µL of solution. Note that these volumes are prior to the addition of labelled probe.
- Mouse 15K array = 80 µL
- Human 19K array = 80 µL
- Yeast 6.4K array = 60 µL
- Human and Mouse CpG Island = 80 µL
- Face-to-face hybridisations = 80 µL
Q: Can I use different hybridisation/washing conditions with your protocol?
A: Users are welcome to try any protocols, however, we have found that "combining" parts of different protocols often leads to poor results. Our suggested protocols have been optimized for use on the arrays that we produce, which is not to say that other protocols will not work. Please have a look at our technical notes for more details.
Q: Can we use other hybridisation buffers besides DigEasy?
A: Yes but most other hybridisation buffers require hybridisations at 42°C-65°C. Hybridisations at higher temperatures can cause the slides to dry, which in turn will result in high background. Dig Easy Hyb enables us to hybridise at 37°C and we won't have to worry about drying out slides and high background.
Q: Are unhybridised arrays kept longer than 2 months still useable?
A: For best results, arrays should be used within 8 weeks of printing. Arrays older than 8 weeks can be used, however, experiments using "older" arrays may or may not work depending on individual environmental conditions and slide variability.
Q: Is it necessary to pre-hybridise the arrays with tRNA, PolydA-dT or human Cot-1 DNA to block repeats and non-specific hybridisations?
A: No. We don't pre-hybridise our cDNA arrays. We add calf Thymus DNA and Yeast tRNA in our hybridisation buffer to block any non-specific hybridisation.
Q: For the human array your protocol calls for the use calf thymus to block repetitive elements. Can the same blocking agents be used for the mouse or yeast arrays?
A: We use Calf Thymus DNA as a blocking agent but you could also use Sonicated Salmon Sperm or some other sonicated "junk" DNA. The same can be used for the Mouse or Yeast chips as for the Human chips. Think of it like a Northern Blot, you only use one blocking agent per blot even when samples of different origin are together. The calf thymus DNA is a blocking agent not specific to the sequences found on the chip, it is there to increase the hybridization kinetics.
Back to Top of Hybridisation
Q: How can I spin my slides dry if I don't have a plate-holder adapter for the centrifuge?
A: There are two alternatives. One is to use compressed air to "blow-dry" the slides. We advise users to place a 0.2 mm filter on the end of the tubing to prevent particles from being blown onto the slides during drying. The other method is to use a 50 mL Falcon tube with a few small Kim-Wipes tightly stuffed into the bottom. Place your slide inside the tube using forceps to handle the slide at the barcode-end. Spin in a 50 mL tube-sized bucket at 500 rpm for 5 minutes.
Q: How can I reduce background?
A: Ensure you have DIG Easy Hyb (or whatever hyb solution you are using) at the bottom of your hyb chamber. This prevents the drying of the hyb solution and labelled-cDNA to the array. Also be sure that your washes are stringent enough. Rinse your slides well after washing in SDS (or other detergents). Detergents can form micelles that can trap unincorporated fluor that could become dried on to the slide during the drying process. If the background is very weak, you could reduce it by simply lowering the PMT setting on the scanner.
Q: Can I rewash my slides? Can I reuse my slides?
A: Slides can be rewashed and sometimes "swirly" background from incomplete washing/rinsing can be removed. However, rewashing can also remove some of the signal. We strongly recommend that you save your image before rewashing, in case all signal is removed. The length of rewash, temperature, and washing buffer all vary between users. Read our Technical Note on rewashing for more details. We have found that rewashing is more likely to be successful if the slides are rewashed within hours of the initial wash. However, slides cannot be used for more than one hybridisation.
Q: How long will the fluorescent signal remain on the slide?
A: While it is ideal to scan your slides as soon after washing/drying as possible, you can scan your slides 2-3 days after washing. Be sure to store your hybridised slides in the dark, and preferably in a cool and dry place. Elevated ozone levels may lead to a decrease (or "bleaching") of the Cy5 signal. This is most often seen on humid days in the summer.
Back to Top of Slides
Q: When using the isopropanol precipitation method to cleanup my reactions, do I need a carrier such as glycogen or linear acrylamide? Will I be able to see a pellet after the precipitation?
A: You do not need to add anything to your labelled cDNA in order to get a pellet. Some protocols you may find do call for the addition of some carrier however we have compared with and without and have not found any appreciable difference. You may or may not see the pellet, usually it is barely visible, just seen as a coloured smudge on the wall of the tube. Frequently no pellet is seen at all and the results are still good. When the pellet is readily visible it could be due to incomplete hydrolysis of the RNA fragments, but this still looks fine in the end. Instead of using the isopropanol precipitation method, you may find the CyScribe™ GFX™ columns (GE Healthcare) provide excellent purification of the labelled-cDNA.
Back to Top of Miscellaneous