Optimization of minuscule samples for use with cDNA microarrays

Available online at www.sciencedirect.com

J. Biochem. Biophys. Methods 70 (2008) 1048 – 1058 www.elsevier.com/locate/jbbm

Optimization of minuscule samples for use with cDNA microarrays Susan McLean Hunter a , Fiona C. Mansergh a,b,⁎, Martin J. Evans a a

b

School of Biosciences, Cardiff University, Museum Avenue, Cardiff CF10, 3US, Wales, UK Ocular Genetics Unit, Smurfit Institute of Genetics, University of Dublin, Trinity College, Dublin 2, Ireland

Abstract The recent advent of microarray technology and RNA amplification allows us to compare the expression profiles of thousands of genes from small amounts of tissue or cells. We have compared and contrasted various methods of RNA preparation, RNA amplification, target labelling and array analysis in order to achieve a streamlined protocol for microarraying small samples. We have concluded that usage of the NIA 15K cDNA array set, in combination with RNA extraction using the Mini RNA Isolation kit (Zymo), amplification with the RiboAmp kit (Arcturus), followed by indirect labelling via the Atlas™ PowerScript™ Fluorescent Labelling kit (using a modified protocol), is optimal with a material derived from either very early stage mouse embryos or individually picked embryonic stem cell colonies. Normalisation using the analysis package Limma (Bioconductor) with data normalisation by print tip Loess, using the “normexp” function with an offset of 50 for background adjustment, and incorporating A-quantile between array normalisation was best with our results. Furthermore, RT-PCR confirmation of array results is achievable without amplification, thereby controlling for amplification bias. These methods will be of great utility in mapping the transcriptome of embryonic and other small samples. © 2007 Elsevier B.V. All rights reserved. Keywords: Microarray; RNA amplification; RT-PCR; Embryonic

1. Introduction The advent of microarray technology [1] allows comparison of gene expression profiles of thousands of genes from cell and tissue samples. This technology can pinpoint differences between normal and diseased tissues, identify genes crucial to certain disease processes, and generate gene expression profiles unique to individual tissue samples and explore the effects of different developmental and physiological conditions [2]. Until recently, use of this technology has been limited by the amount of RNA that can be derived. cDNA printed microarray slides need a minimum of 5 μg/sample for useful signal (derived from approximately 107 cells), whereas tiny samples, such as those derived from early embryonic compartments, biopsy or from laser capture microdissection, yield far less than this [3,4]. The ability to achieve genome wide expression profiles from single

⁎ Corresponding author. Ocular Genetics Unit, Smurfit Institute of Genetics, University of Dublin, Trinity College, Dublin 2, Ireland. Tel.: +353 1 896 2484. E-mail address: [email protected] (F.C. Mansergh). 0165-022X/$ - see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.jprot.2007.11.011

cells (10–30 pg of RNA) would represent the ultimate goal of current technological improvements. By combining RNA amplification [5] with microarray technology, it is now possible to derive sufficient material from very small samples. The original amplification method, pioneered by Eberwine and colleagues [5], involves the use of a T7 promoter containing oligo-d(T) primer to prime first strand cDNA synthesis. RNase H is used to degrade the RNA in the resulting RNA/cDNA hybrid, leaving small residual RNA oligos which prime the formation of double stranded DNA. Antisense RNA can then be generated from this template using T7 RNA polymerase. Further rounds of amplification can be carried out using random hexamers as primers. The RNA template is once more degraded with RNase H and a second strand synthesised in the presence of the T7 oligo-(dT) primer which anneals to the polyA tail on the second strand. Hence, the T7 promoter is again added to the cDNA allowing further synthesis of antisense RNA. Improvements to the protocol have been made [4,6–8], furthermore, quality control using this method suggests that gene expression patterns are maintained through the amplification step [4,9]. Advances have also been made using signal

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amplification with avidin/biotin incorporation, followed by horseradish peroxidase based detection and with the use of dendrimers. These methods can reduce the input RNA required by 10–100 fold, but would still require approximately 105 cells, more than many small samples would provide [3]. PCR amplification of cDNA is also possible; yields are higher than with the Eberwine protocol, but there are concerns with regard to maintaining the relative transcript abundance of the starting material owing to amplification bias [3,4]. Microarrays are largely divided into 3 types. GeneChip® arrays (Affymetrix, Santa Clara, CA, USA), consist of quartz chips, printed with up to 500,000 oligonucleotide probes. These have the advantage of most complete genome coverage. Unfortunately, an issue with amplified material is the 3' bias introduced during multiple rounds of mRNA reverse transcription. Affymetrix have started to produce chips with oligos designed for 3' biased material, indeed, heart biopsies that yield an average of 50 ng of RNA have successfully been analysed using 2 rounds of amplification with an Affymetrix gene chip [10]. However, we would estimate the yield from 10 ES cell colonies (or 20 embryonic inner cell masses] at about 10 ng of RNA (see Table 1). Given the tiny amounts of RNA and the fact that some cDNA libraries contain an average insert size of N 1 kb, spotted arrays may represent a better source of array for material that has been amplified twice. Moreover, the amplified RNA products of some RNA amplification methods are incompatible with the orientation of the oligos most commonly printed on Affymetrix chips [4]. Spotted arrays consist of cDNAs from expression libraries which are most efficiently printed on glass slides of typical microscope size, although filters may also be used. Targets for filter arrays are normally labelled radioactively and hybridised to the filters one at a time. Phosphorous 33 is a very sensitive

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label and is well incorporated during reverse transcription, however, the tendency of radioactive label to “bleed” outwards from the spot reduces the allowable density of printing, moreover, the use of radiation is hazardous [2]. The analysis of microarrays printed on slides is usually carried out via labelling and hybridization of two differentially labelled targets to the same slide. Targets are most commonly labelled with the dyes Cyanine 3 and Cyanine 5 (Cy3 and Cy5), which are fluorescent dyes which are widely separated in excitation and emission spectra, have good photostability and yield, and show relatively good incorporation frequencies with reverse transcriptase [2]. Various dye specific labelling biases have been noted with these dyes however [3,11]. Direct incorporation of the dyes into reverse transcribed cDNA is the simplest method of labelling; the disadvantages of this are that Cy5 is larger than Cy3, is therefore less efficiently incorporated, (and is also less stable) [12]. Furthermore, reverse transcription tends to limit target to the 3' ends of RNA species owing to limited enzymatic processivity and RNA secondary structure [13]. A number of methods have been chosen to overcome this problem, including the development of modified reverse transcriptases that incorporate Cy5 more efficiently, indirect labelling of cDNA via direct incorporation of aminoallyl groups, then subsequent attachment of the cyanine dye to the aminoallyl group and labelling of the RNA itself. Labelling of the RNA itself avoids processivity and uneven incorporation problems related to reverse transcriptase. However, RNA target instability is a problem, furthermore, as the target is sense, it can only be used with cDNA or antisense oligonucleotide arrays. Indirect labelling is possibly the best way in which to overcome problems of uneven incorporation, however, this does not solve issues of 5' bias or improve the stability of Cy5. Moreover, even given the use of indirect labelling, gene specific dye bias is still

Table 1 RNA amplification Sample number

Amount of total RNA input to first round (source of RNA)

Amount of aRNA input to second round

Yield of aRNA after two rounds of amplification

1 (Message Amp kit) 2 (Message Amp kit) 3 (Message Amp kit) 4 (Message Amp kit) 5 (RiboAmp kit) 6 (RiboAmp kit) 7 (RiboAmp kit) 8 (RiboAmp kit) 9 (RiboAmp kit)

10 ng (ES RNA) 10 ng (ES RNA) 100 ng (ES RNA) 100 ng (ES RNA) 10 ng (ES RNA) 10 ng (kit control) 10 ES colonies 10 ES colonies 100 ng (kit control)

1.5 μg 1.1 μg 1.7 μg 2.4 μg 200 ng 600 ng Not tested Not tested 52.8 μg

10 (RiboAmp Kit, TRIzol) 11 (RiboAmp Kit) 12 (RiboAmp Kit) 13 (RiboAmp Kit, TRIzol)

19 inner cell masses 25 inner cell masses 27 inner cell masses 5 ES cell colonies

Not tested Not tested Not tested Not tested

3.3 μg 1.1 μg 2.9 μg 7.4 μg 42.6 μg 37.08 μg 26.9 μg 81.25 μg Not carried out, too much RNA from round 1. 5.22 μg 43.9 μg 60.24 μg 1.8 μg

Assessment of the relative efficiencies of RNA amplification using the Ambion Message Amp kit and the Arcturus RiboAmp kit. Yields are much lower and the amount of starting RNA does not have a direct linear relationship with the yield of aRNA from Round 2 of amplification, using the Ambion kit. Unmeasured input RNA from pools of 10 ES colonies each compares roughly with an input of 10 ng of total RNA from large scale RNA preps (5 preps of 10 ES colonies were carried out, the lowest and highest values are shown above). A negative control was run using the RiboAmp kit, with input of 8 μl of water. No RNA was obtained after 2 rounds of amplification. RNA preps were carried out using the Zymo Mini RNA isolation kit, unless otherwise stated (note lower amplification yields from TRIzol RNA preps). Methods for isolation of 3.5 day ICMs are given in GSE8881. The whole blastocyst at the 64 cell stage contains about 2 ng total RNA, inner cell masses at day 3.5 have been estimated as containing 48 cells approximately [20].

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the major source of inter-replicate experimental variation [11]. However, attempts to replace the cyanine dyes with Alexa Flour 555 and 647 resulted in lower signal intensities and higher background than was achieved with the original Cy dyes [12]. The National Institute of Ageing (NIA) 15 K set, comprised of 15,264 putative unique genes with an average length of 1.5 kb, was derived from various embryonic stages and was compiled to address the lack of coverage of genes crucial to early mammalian development [14,15]. Given the probe length, the optimal coverage of embryonically expressed genes and 3' bias, we deemed the NIA set optimal for our purposes. Using the NIA 15000 set array slides, we have been able to optimize the preparation of RNA from very small samples, RNA amplification, labelling of samples derived from 2 rounds of amplification, hybridization to slides, image and statistical analysis, and finally, confirmation of differentially expressed genes. These results should be of great utility to anyone trying to derive global gene expression data from minute amounts of starting material, especially when studying early embryonic development. 2. Materials and methods

pipette prior to lysis in the buffer supplied with the Zymo Mini RNA Isolation kit™ (Zymo Research, Orange, California, USA). TRIzol reagent (Invitrogen Ltd, Paisley, Renfrewshire, UK) and the RNeasy Mini kit (Qiagen, Ltd., Crawley, Sussex, UK) were also tested for use with both large and small quantities of RNA; RNA preps were carried out according to the manufacturer's protocol in both cases, although 0.1 μg of linear acrylamide was added to the first precipitation stage of the TRIzol protocol, in order to maximize RNA yield and aid pellet visualization. Given the tiny amounts of RNA we were working with, neither spectrophotometry nor analysis of RNA via Nanodrop gave meaningful results prior to amplification. This also wasted 1 μl out of 8 μl starting material. Another issue is also not just the starting RNA concentration, but also its amenity to amplification and subsequent labeling (impurities can impede subsequent enzymatic reactions). It has therefore been easier either to measure the concentration after amplification, or to assess the final quantity and quality of labeled cDNA both by spectrophotometry and slide gel analysis. It is possible to amplify meaningful amounts of RNA from single ES cell colonies, however, in practice, we found that pooling 5–10 single ES colonies improved reliability.

2.1. Culture of embryonic stem cells

2.3. RNA amplification

For the purposes of technical optimization, we used RNA prepared from embryonic stem (ES) cells (IMT-11 [16]). ES cells were maintained in their undifferentiated state in the medium MEM-Alpha (Gibco™, Invitrogen Ltd, Paisley, Renfrewshire, UK) supplemented with 10− 4 M β-Mercaptoethanol (Merck KGaA, 64293 Darmstadt, Germany), 2 mM glutamine and 10− 3 U/ml murine LIF (ESGRO™, Invitrogen, Ltd, Paisley, Renfrewshire, UK), 10% FBS and 10% NBS (selected batches, PAA Laboratories GmbH, Linz, A-4020 Austria). The cells were maintained at 37 °C in a humidified atmosphere with 5% CO2 on 0.1% gelatin (Stem Cell Technologies) coated tissue culture grade plastic ware (NUNC™, Fisher Scientific, Loughborough, Leics, UK).

Amplification using the Eberwine method and variations thereof [5–7] were attempted (see Appendix C for tabulated methods). Amplification was also carried out with the MessageAmp™ (Ambion, Huntingdon, UK) and RiboAmp® (Arcturus, Sunnyvale, CA, USA) kits, according to the manufacturer's protocols (see Table 1 for results). The RiboAmp protocol recommends the use of DNase treatment prior to amplification. We tested this recommendation by treating some samples with the DNA free kit™ (Zymo) prior to amplification. DNase and non-DNase treated samples, derived from the same RNA source, were then compared via array analysis as described below. Given that the amounts of RNA obtained from individual ES cell colonies were difficult to assess, for amplification method testing, concentration of input amounts of RNA were obtained via serial dilution of a larger scale RNA prep of known concentration.

2.2. RNA isolation 2.2.1. Large scale RNA preps ES cells were washed twice with DPBS-A and treated with trypsin-EDTA (Invitrogen Ltd., Paisley, UK). The cells were counted in a haemocytometer and appropriate cell numbers were pelleted by centrifugation at 200g. The QIAgen RNeasy™ Midi Kit (Qiagen Ltd., Crawley, Sussex, UK) was used according to the manufacturer's protocol for all large scale samples. These RNA samples were quantified and checked for degradation via OD 260/280 spectrophotometry (Camspec, Cambridge, UK) and gel electrophoresis using dissociating conditions (NorthernMax™ buffers; Ambion, Huntingdon, UK), used according to the manufacturer's protocol. 2.2.2. Small scale RNA preps For small amounts of ES cell RNA, plates were covered in PBS and individual ES colonies were picked using a sterile glass

2.4. Production of fluorescently labelled target RNA The same batch of cytoplasmic RNA from the ES line IMT11 was used for all experiments where different labelling methods were tested. All labelled targets were either used immediately or stored at − 20 °C protected from light. Cleanup of labelled cDNA was carried out as recommended by the kit manufacturers, or using the Qiaquick PCR purification kit (Qiagen), if a kit or recommendation was not supplied. 2.4.1. MICROMAX ASAP kit (Perkin Elmer, Shelton, CT, USA) Cytoplasmic RNA (10 μg) was labelled with Cy dye according to the manufacturer's protocol. Labelled mRNA was then purified using the Oligotex™ mRNA Mini Kit (Qiagen, Crawley, West Sussex, UK). Cy3™ and Cy5™ labelled RNAs

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were handled separately so that each could be analysed by spectrophotometry and slide gel electrophoresis later on. 2.4.2. CyScribe™ First Strand cDNA Labelling Kit (GE Healthcare, Little Chalfont, Buckinghamshire, UK) Target RNAs (10 ug) were labelled via the manufacturer's protocol and were subsequently purified using the Qiaquick PCR purification kit (Qiagen). 2.4.3. Clontech Atlas™ Glass Fluorescent Labelling Kit (Clontech, Saint-Germain-en-LayeFrance) The Clontech Atlas™ Glass Fluorescent Labelling kit was used according to the manufacturer's protocol. The NucleoSpin™ purification kit, as supplied with the Atlas™ Glass Fluorescent Labelling Kit, was used to purify labelled product. 2.4.4. Clontech Atlas™ PowerScript™ Fluorescent Labelling Kit (Clontech, Saint-Germain-en-LayeFrance) The Clontech PowerScript™ Fluorescent Labelling kit was used according to the manufacturer's protocol. For some samples, instead of using RNase H to degrade the RNA template, NaOH was used; 2 μl of 2.5 M NaOH was added, followed by incubation at 37 °C for 15 min, followed by neutralisation with 10 μl of 2 M HEPES. Precipitation and recovery of the aminoallyl conjugated cDNA was also enhanced by the addition of 0.5 μl of PelletPaintNF™ (Novagen, Merck Biosciences Ltd, Nottingham, U.K.). The QIAquick™ Cleanup Kit was used to purify labelled product. The manufacturers recommended a Pharmacia method for cleanup which worked by desalting. However, according to our slide gel analysis method, this left too much free dye. This method was later replaced with another, supplied by the manufacturer, than interfered with analysis of the labelled product. Therefore, the Qiagen kit was used in preference. 2.4.5. CyScribe™ cDNA Post-Labelling Kit (GE Healthcare, Little Chalfont, Buckinghamshire, UK) The CyScribe™ cDNA Post-Labelling kit was used with 5 μg of RNA according to the manufacturer's protocol (Table 2). Purification of the labelled targets was carried out using the QIAquick™ cleanup kit.

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2.5. Assessment of probe quality Analysis of the probes was carried out by spectrophotometric analysis and slide gel electrophoresis [17]. 1 μl of each labelled sample was run on a slide gel in order to assess the molecular weight spread and the level of residual free dye. Gels were then scanned at the appropriate wavelengths using a GeneTac™ LSIV scanner (Genomic Solutions, Cambridge, UK). The amount of cDNA synthesised and the incorporation of dyes were calculated by application of Beer–Lambert's law (A = ɛlc, A = absorbance, ɛ = extinction coefficient in cm− 1 M− 1, and c = concentration (M)). The FOI (frequency of incorporation) for each labelled sample, (number of Cy-dUTPs/1000 nucleotides), was calculated according to the formulae: FOI Cy3™ = OD550 × 58.5/OD260 and FOI Cy5 = OD650 × 35.1/OD260. 2.6. Using target labelled with Cy3/Cy5 The required amount of Cy3™ and Cy5™ labelled targets were combined in a sterile amber microcentrifuge tube and mixed thoroughly. The labelled cDNAs were reduced in volume to 18 μl using a rotary evaporator (Christ RVC 2-18, GMBH, Wertheim, Germany) at 30 °C. To the 18 μl of target, 1.0 μl of each Poly-dA30 (1 mg ml− 1) (MWG Biotech, Covent Garden, London, UK) and human COT-1 DNA(1 mg ml− 1) (Invitrogen) was added to the cDNA followed by denaturation at 95 °C for 3 min. Prior to hybridization, the volume of the labelled target was adjusted by the addition of 20 μl of hybridization solution (2× hybridization solution, 50% formamide, 10% SSC, 0.2% SDS) and water. The labelled target was added to the microarray under Hybri-slips (Sigma) and hybridised for 18 h at 42 °C. 2.7. Microarrays The printing and immobilisation of the slides was carried out by the Microarray Facility, School of Biosciences, Cardiff University (http://www.walesgenepark.co.uk/). If not used immediately, the microarrays were stored in a desiccator at room temperature, protected from light. MIAME compliant data is stored on the following site: ftp://watson-bios.grid.cf.ac.uk/ sequences/Susan/.

Table 2 Comparison of labelling kits Template removal

cDNA ng Cy3

cDNA ng Cy5

% cDNA/total Cy3

% cDNA/total Cy5

pmol dye Cy3

pmol dye Cy5

FOI Cy3

FOI Cy5

NaOH NaOH NaOH RNase H RNase H RNase H RNase H

547.60 440.30 616.05 4280.90 3537.20 4206.90 3888.70

573.50 458.80 455.10 4539.90 358.90 4221.70 3936.80

10.95 8.81 6.16 42.81 35.37 42.07 48.61

11.47 9.18 4.55 45.40 3.59 42.22 49.21

29.33 29.33 27.33 42.00 38.67 29.33 23.33

28.80 28.40 13.0 47.60 7.60 19.20 20.40

17.39 21.63 14.41 3.19 3.55 2.26 1.95

16.30 20.10 9.27 3.40 6.88 1.48 1.68

Kit, temperature, amount RNA labelled PowerScript kit 42 °C, 5 ug CyScribe Post-Labelling kit, 42 °C, 5 ug CyScribe Direct, 10 ug Atlas Glass kit 37 °C, 10 ug Atlas Glass kit 42 °C, 10 ug Atlas Glass kit 48 °C, 10 ug PowerScript kit 37 °C, 8 ug

Note that RNase H gives, ostensibly, far higher cDNA yield. Note; figures for the Oligotex kit are unavailable in this format; fines left in the labelled mRNA by the Oligotex mRNA purification interfered with OD readings, making dye incorporation impossible to assess. Labelling reactions were carried out twice (once for each dye).

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2.8. Blocking Of CMT-GAPS™ coated slides Immediately prior to use for hybridization, the slides were blocked to prevent non-specific binding of target DNA to reactive amine groups present in the Gamma Amino PropylSilane coating. Microarrays were placed in a 50 ml Coplin jar containing blocking solution (1% Bovine Serum Albumin (Fraction V) (Sigma-Aldrich), 5× SSC and 0.1% SDS) and blocked at 42 °C for 45 min. Slides were then dipped briefly (2 s each) five times in sterile, filtered Milli-Q™ water. This was followed by a single 5-second dip in room temperature isopropanol. The blocked slide was then air dried for approximately 10 min. After blocking, hybridization was carried out within 1 h. 2.9. Post hybridization Following hybridization, unbound label was removed by washing once for 10 min at 55 °C in 1× SSC, 0.2% SDS then twice for 10 min each at 55 °C in 0.1× SSC, 0.2% SDS. The slides were finally rinsed in filtered 0.1× SSC and dried with compressed air. The hybridised and washed slides were scanned in a ScanArray Express HT scanner (Perkin Elmer) at 65 pmt and 90% gain. The scan resolution was 10 μm and was run at full speed. 2.10. Analysis and normalisation Image analysis was carried out using the program ImaGene™ 5.5 (BioDiscovery). Normalisation and analysis was carried out using both GeneSight™ 3.5 (BioDiscovery) and Limma (BioConductor, http://www.bioconductor.org/), with data comparing use of DNAse versus no DNase digestion prior to amplification. These data comprise 4 array slides, two DNase+, Cy5, DNase-, Cy3 and the remaining two slides labelled DNase+, Cy3, DNase-, Cy5. 2.10.1. Normalisation and analysis using GeneSight™3.5 (BioDiscovery) The data was processed in the sequence: omit flag types (selected for individual analyses), floor values to 20, log2 transform the intensities, normalise with Loess then subtract the log2 green from the log2 red values and combine the data for each array. Histograms and scatter plots were generated. Differentially regulated genes were identified by first finding the spots more than 2-fold up or down regulated. A confidence analysis test was used to identify the statistically significant genes at the 99% level. 2.10.2. Normalisation and analysis using Limma The raw data text files were loaded into Limma using the commands in Appendix A. Diagnostic plots of the raw data were generated. First the data were normalised by print tip Loess with no background correction. The normalisation procedure was then repeated using the “normexp” function with an offset of 50 for background adjustment. The data were then normalised using no background correction and global

loess as a comparison with GeneSight. Diagnostic plots of the processed data were then generated. A linear model was fitted to the data and eBayes statistics produced. Statistically significant regulated genes were identified from the top table. 2.11. RT-PCR aRNA from DNase treated and untreated samples was analysed via reverse transcription followed by PCR (RT-PCR). Genes confirmed by PCR were used to identify the most accurate analysis methods. RT-PCR was also used to analyse tiny quantities of unamplified RNA in order to determine the best methods for gene confirmation. Reverse transcription was carried out using the SuperScript™ First-Strand Synthesis System for RT-PCR (Invitrogen) with random hexamers for amplified RNA, and oligo dT for unamplified RNA, via the manufacturer's protocol, with the following modification: 1 μl of a 1/10,000 dilution of 50 pmol 18SrRNA reverse primer was spiked into the oligo dT mix supplied with the kit, prior to RT. This was carried out in order to be able to use 18S rRNA as a housekeeping control; as it does not have a polyA tail, RT using a gene specific primer is necessary. PCRs were performed in a Biometra® UNOThermoblock™ (Biometra, Thistle Scientific, Glasgow, UK). 1–2 μl of the reverse transcription reaction was subjected to PCR in a mix consisting of 10× PCR buffer (Sigma-Aldrich), 200 μM dNTP, 02 μl Taq (Sigma-Aldrich), and 50 pmol of each primer. 2.11.1. PCR primers Primers were designed from NIA EST sequences using methods described previously [17]. Beta-actin and Gapdh primer sequences are also given in this reference. Other primer sequences are included in Appendix B. 3. Results 3.1. RNA isolation Phenol based methods of RNA isolation, such as TRIzol (Invitrogen), were shown to yield amplifiable RNA from 19 ICMs (Table 1). Yields were not as reliable as with the Zymo kit (results not shown). We were concerned that carryover traces of phenol might be occasionally inhibiting downstream enzymatic reactions, furthermore, loss of a tiny pellet at either precipitation step was also a possibility. The Qiagen RNeasy Mini kit eluted purified RNA in a larger volume than the Zymo Mini RNA isolation kit (30 μl versus 8 μl), requiring a further concentration step in one case but not the other. Use of the Zymo Mini RNA isolation kit was therefore judged optimal for our purposes. Given the difficulties of assessing RNA concentrations directly from unamplified RNA, the relative successes with which different RNA preparation methods worked could be assessed indirectly either via yield of cDNA from labelling after amplification, or via spectrophotometric readings after two rounds of amplification. Note that RNA from 10 ES cell colonies gives slightly higher aRNA yield than when a known

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input of 10 ng of RNA is used prior to amplification. We should note that, while the theoretical dynamic range of Nanodrop flourospectrophotometers starts at approximately 10 ng/μl, we have not found readings taken at these approximate concentrations to be at all reliable. 3.2. Amplification RNA amplification was initially attempted using the original method pioneered by Eberwine and colleagues [5]. Subsequent modifications to this protocol were included such as the addition of a few extra 5' bases and 3' d(T)s to the primer in an attempt to stabilize the binding of the T7 enzyme to its promoter. E. coli DNA ligase was added to the second strand reaction and the cDNA. Double stranded cDNA was cleaned with phenol:chloroform:isoamyl alcohol and Microcon-100 to reduce carryover of primers and deoxynucleotides into the in vitro transcription reaction [6]. A template switch strategy and second strand extension at 75 °C using Taq polymerase was also incorporated, as this had been demonstrated to improve problems with 3' bias [7]. A modified method, involving making sense RNA on magnetic beads (DynaBeads® Oligo(dT)25, Dynal) or polystyrene-latex (Oligotex ®, Qiagen) was also tested. Methods for these procedures are given in Appendix C, however, despite a step by step testing showing that each protocol step was working, the combined protocols failed to reliably yield enough aRNA for microarray experiments (maximum yield from 10 ng total RNA, 2 rounds of ampliõfication, Wang protocol: 362.4 ng, Luo protocol: 556.8 ng, Dynabeads, 0 ng). The MessageAmp kit produced by Ambion did not work well at the very low RNA input levels necessary for this project (after 2 rounds of amplification, 10 ng of total RNA was amplified to 3.3 μg). We should note that Ambion have since updated this kit. However, the Arcturus RiboAmp® kit gives higher levels of amplification (after 2 rounds of amplification, 10 ng of total RNA was amplified to 42.6 μg amplified RNA), therefore the decision was made to use this kit in all subsequent experiments. The Arcturus kit is routinely used for the amplification of ES cell colonies and, although colony size varies, 10 colonies picked from a plate has produced between 26.9 and 81.25 μg over 5 independent replicates. A ttest comparing the yields from 2 rounds of amplifications generated by the two kits gives the following significant result p = 0.0053 (n = 4 for each kit, mean = 3.225, SD = 2.89 for Message Amp, mean = 46.69 SD = 23.77 for RiboAmp).

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3.3. Fluorescent labelling and hybridization 3.3.1. Direct labelling Labelled RNA generated using the MICROMAX ASAP kit was difficult to run on slide gels, so the molecular weight spread and the amount of residual free dye could not be assessed. However, when hybridised to an array slide, the degree of hybridization was good with few background problems. Unfortunately, both Cy labelling reagents proved to have a very short shelf life and the Cy5 labelling reagent overall was considerably less stable than the Cy3. The CyScribe™ First Strand cDNA Labelling Kit Labelling is efficient and reliable, however, approximately double the amount of Cy5™ labelled target is required to achieve similar levels of fluorescence in both channels, which may result in loss of sensitivity, particularly with highly expressed genes (Table 2). 3.3.2. Indirect labelling Indirect labelling relies on the incorporation of aminoallyl modified nucleotides into cDNA derived from target RNA. The aminoallyl side chains are then used to link either Cy3 or Cy5 to this cDNA, thereby labelling it. The CyScribe Post-Labelling kit performs well with regard to incorporation of the dyes. Labelling in both channels was more even when compared to the direct incorporation of Cy dye modified dUTP. The Atlas™ glass and Atlas™ PowerScript™ labelling kits, both produced by Clontech, were compared. Labelling was carried out according to the manufacturer's protocols and the probes were analysed as before. The amount of cDNA obtained from the Cy3 and Cy5 reactions is reproducible and the picomoles of dye incorporated compares well with the best conditions described above (Table 2). All three kits performed comparably well. 3.4. RNase H versus NaOH degradation of RNA template after labelling We noticed that the amount of cDNA recovered with both Atlas kits seemed unreasonably high, given that the proportion of mRNA in cells is 1–5% of total RNA. These kits rely on RNase H to degrade the RNA template. The RNA fragments may be inefficiently removed by the purification protocol, giving artificially high cDNA recovery values. Furthermore, arrays hybridised with Atlas™ Glass and Powerscript™ labelled targets were not as bright as those labelled directly

Table 3 Labelled cDNA Experiment number and dye

Template removed by RNase cDNA synthesises(ng)

Dye incorporated (pmol)

FOI

cDNA synthesises(ng)

Template removed by NaOH Dye incorporated (pmol)

FOI

1 Cy3 2 Cy3 1 Cy5 2 Cy5

2002.0 1357.9 1491.0 1311.5

20.0 15.3 20.0 11.2

3.24 3.72 4.35 2.75

469.9 758.5 384.8 455.1

43.3 46.3 38.8 20.0

29.94 19.99 32.74 18.04

5 μg of cytoplasmic RNA was labelled per reaction, labelling was carried out using the Powerscript kit at 42 °C.

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Table 4 aRNA yield after 2 rounds of amplification Treatment of total RNA

DNase+ Dnase-

Yield of aRNA (μg) from 10 colonies of ES cells 57.3 94.0

Yield of cDNA (μg) from labelling 5 μg of aRNA

pmol of Cy dye incorporated

FOI

Cy3

Cy5

Cy3

Cy5

Cy3

Cy5

440.3 745.8

407 821.4

28 28

18 32.4

20.6 12.0

18.0 12.8

Yield of aRNA after 2 rounds of amplification and properties of labelled cDNA derived.

(personal communication Ngoc Nga Vinh), suggesting competition between unlabelled nucleic acids and labelled cDNA. To test whether the RNase H degradation of the mRNA template is responsible for the apparent excess of cDNA, the protocol for the Powerscript™ kit was modified. 5 μg of cytoplasmic IMT11 RNA was used to prepare targets using the Powerscript kit. NaOH processed PowerScript sample was compared with the CyScribe™ cDNA Post-Labelling kit, which uses NaOH degradation of the template as standard protocol. The amount of cDNA synthesised was now similar for both kits (Table 2). Further RNA template was destroyed by either RNase H or NaOH and the labelled target was analysed as before, but this time, targets were hybridised to array slides, scanned and subjected to image analysis. The amount of cDNA produced and the level of dye incorporation was consistent with the earlier results (Table 3). The FOI for both channels was much lower in the RNase H treated targets. The total number of good quality spots were counted on each slide (flagged 0 by ImaGene), and the mean was calculated. No significant difference in the number of positive spots was detected (mean = 3055, SD = 1571, n = 4 for RNase H, mean = 3168.5, SD = 963.5, n = 4 for NaOH. The data from the Cy3 and Cy5 RNase H and NaOH experiments were combined and filtered such that only spots flagged 0 on all four slides were analysed (760 spots). A scatter plot was generated and the trendline was calculated. The significance of the model was tested using linest and the Pearson coefficient functions in Microsoft Excel™ 2003. The

slope of the the trendline, 0.998 +/− 0.005, very close to 1, suggests that the method of destroying the RNA template has no consequences for the fluoresence intensity of the spots. The model produced is highly significant as indicated by both the Pearson Coefficient R (0.977) and the F-statistic (31,674.67, p b 0.0001). However, the NaOH treatment method can produce significantly more labelled target, enough for three arrays whereas the RNase treatment method generally only produces enough for one array (Tables 2 and 3). The NaOH method was therefore used in subsequent analyses, further modified by the addition of PelletPaintNF™ (Novagen, Merk Biosciences) to the ethanol precipitation of the aminoallyl modified cDNA, which reduces losses by improving pellet consistency and visibility. 3.5. DNase treatment of RNA samples prior to amplification DNA contamination of RNA samples is a potential source of systematic error. Such contamination could lead to gene specific variation in gene expression when present in samples to be amplified. On the other hand, as the tissue samples in this study are at the most only a few hundred cells, it is advantageous to carry out as few procedures as possible and thus avoid loss of material through multiple purification steps and degradation. Therefore, we compared the effects of DNase treated and untreated IMT-11 RNA derived from 10 ES cell colonies and prepared using the Zymo kit. Samples were then amplified,

Table 5 Table of top ten differentially expressed genes, DNase +/− experiment

Top ten differentially expressed genes. The data was processed by removing the background with normexp Offset = 50, normalisation within arrays by print tip Loess and normalisation between arrays by A-quantile or with none. Only the first 9 genes are statistically significant. p b 0.05. When processed by within array print tip Loess and without background removal or between array normalisation only the first six genes, highlighted in yellow are found to be significantly different at p b 0.05. Note actual p values of yellow list are different. Genes in red font were actually confirmed.

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quantified (Table 4) and labelled with the Powerscript kit (using NaOH). Equal amounts of cDNA were used to array 4 slides (incorporating fluor switching), slides were then scanned and analysed using ImaGene™ 5.5.3 (BioDiscovery). The data was inputted into both the GeneSight™ 3.5 (BioDiscovery) and Limma (BioConductor) for normalisation and selection of differentially regulated genes. 3.6. GeneSight™ (BioDiscovery) The analysis package GeneSight™ is designed for full integration with the image analysis program ImaGene™ and can be used to remove different categories of spot from the normalisation process by utilising the spot quality flags. Four

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genes were identified as differentially regulated, but on closer examination, two were flagged as empty on three out of four slides and another was very low intensity. However, H3056D06-3 was found by visual inspection to be a real result. 3.7. Analysis using the Package Limma (BioConductor) Limma (BioDiscovery) is an alternative statistical analysis package for microarrays. It is not specifically designed for use with ImaGene, however, so all spots must be included in analyses. When the data were only subjected to global Loess normalisation, no significant genes were found (Table 5). Print tip Loess with background adjustment and scale normalisation also gave no genes. Nine genes (identical both in rank and

Fig. 1. a: Gene confirmations, DNase +/− experiment. PCRs were carried out on two sets of samples using the following cycles; Gapdh; 94 °C, 5 min, (94 °C, 1 min, 62 °C, 1 min, 72 °C 1 min 20 s) × 27, 72 °C 10 min. H3073B10; conditions as above, but 45 cycles, H3050H04, as above, 27 cycles. Primer sequences are given in Appendix B. b: RT-PCR from unamplified RNA derived from 10 ES colonies. 18S = 18S rRNA, Gap = Gapdh, B-Act = beta-actin. All were run at 30 cycles, using the same conditions as described for Gapdh, Note bands in some of the DNase untreated No RT controls.

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identity) were found as being differentially detected using print tip Loess with background adjustment as described, alone and also with A-quantile normalisation (Table 5). The six genes found by within array print tip Loess without background correction and by print tip Loess with A-quantile normalisation are the first six genes in the list of nine albeit in a slightly altered rank order. The slides where no background correction was applied before normalisation appear to have compression of the signal intensity. Therefore it was decided to use normexp with an offset value of 50 with the command print tip Loess. Box plots and density plots of the arrays, before and after between slide normalisation, were generated (not shown), which showed that although datasets were very similar, some between array normalisation would be beneficial. The density plot shows very little variation between channels (after within slide normalisation) and some variation between slides. Scale normalisation and A-quantile normalisation were compared. A-quantile appears to give a better result. Intensity data for the nine spots were filtered further in MS Excel. All of the genes were above background + 2SD in at least one channel. Examination of the M-values for these genes reveals that all were at least 2-fold different (Table 5). The gene H3056D06, detected by GeneSight™ (BioDiscovery), is also on the list of 9 genes and has the highest M-value (− 1.857, Table 5). 3.8. Gene confirmations We used the gene lists from the DNase treated and untreated samples to optimize our procedure for confirming genes from amplified samples. DNase treated and untreated amplified RNA from 2 sample sets was reverse transcribed. Beta-actin and 18S rRNA were used as housekeeping controls. We were able to confirm differential expression of three clones, H3050H04-3, H3075D05-3 and H3073B10-3 (see Fig. 1). None of these were detected by GeneSight, implying that Limma (Bioconductor) is a better analysis tool. Print tip Loess with background adjustment as described, with A-quantile normalisation appears to be the optimal analysis method, as only the full list of 9 genes contained all 3 that were actually confirmed by PCR. We would conclude, however, that DNase treatment prior to amplification for array analysis makes very little difference. We also tested our ability to RT-PCR from small samples without any amplification. 2 sets of RNA from 10 ES cell colonies was prepared using the Zymo kit. The 8 μl eluates were divided in two; one half was treated with the TurboDNAfree kit (Ambion, DNase treatment), while the other halves were not. All four samples were then reverse transcribed using oligo dT and PCRed as above. 18S rRNA, beta-actin and Gapdh were strongly amplified at 30 cycles from both DNase treated and untreated samples. However, PCR amplification was also evident in noRT controls from the DNase untreated samples (no amplification occurred in noRT controls from the treated samples). Therefore, for the purposes of PCR, DNase treatment is essential (Fig. 1 a). Unfortunately, as a result of amplification in the noRT controls of untreated, unamplified samples, we cannot test whether changes in gene expression of the above 3

genes are due to DNase treatment, or the presence of small amounts of DNA contamination interfering with the subsequent amplification process. We should also note that, when using RTPCR to analyse aRNA from either 1 or 2 rounds of amplification using the Arcturus kit, we have found that the incidence of erroneous signals in the no RT controls is greatly reduced by treating the aRNA with DNase before RT-PCR. 4. Discussion The aim of this project was to develop techniques which would allow transcriptomic phenotyping of RNA from microscopic samples. We first considered optimal methods of RNA extraction. A prior study comparing TRIzol RNA purification with the RNeasy mini kit alone and with the sequential use of both concluded that use of the RNeasy Mini Kit alone gave the best results when microarray analyses of cDNA generated from RNA from the three methods were compared [18] We found that amplification of RNA prepared using TRIzol reagent (Invitrogen) could be highly variable, possibly owing to carryover of phenol traces or to loss of micro-pellets during isopropanol and ethanol precipitation steps. Degradation of RNA using TRIzol has been noted as more severe when compared with the silica based RNeasy Mini kit [18]. Silica columns also have the advantage that RNA fragments of less than 200 bases are washed through the column and eliminated from further processing. There are also safety and disposal issues associated with the use of phenol, which do not apply to the use of silica columns. Two silica column kits are available, the Zymo Mini RNA isolation kit and the Qiagen RNeasy Mini kit. The RNeasy Mini kit elutes in a volume of 30 μl, while the Mini RNA isolation kit elutes in a convenient volume of 8 μl. Elution volumes of 30 μl required concentration to a lower volume, a further step where material could be lost or degraded. The Zymo Mini RNA isolation kit gave more consistent results than TRIzol where low quantities of RNA were to be extracted, involved fewer processing steps than the Qiagen kit and was therefore used consistently thereafter. Amplification using “in house” methods was compared with the MessageAmp kit (Ambion) and the RiboAmp kit (Arcturus). Adequate and reproducible amplification required the RiboAmp™ kit (Arcturus). Best results were obtained from amplification of 10 or so pooled ES cell colonies, rather than single colonies (although this is possible). The development of labelling techniques was carried out using cytoplasmic RNA as aRNA takes several days to generate and is a valuable resource. However, our optimized labelling protocol has since been used with amplified samples very efficiently. The Micromax ASAP kit (PerkinElmer Life Sciences) conjugates the cyanine dyes directly to the N7 of the guanine and adenine residues of the RNA through a platinum universal linker [19]. Since RNA is labelled chemically, problems with sequence specific variation, dye bias and premature termination of reverse transcriptases should be eliminated. This chemical direct labelling produced excellent array data (not presented), however, the instability of the reagents reduced reliability and increased cost. Moreover, owing to

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carryover of reagents, OD and slide gel testing of labelled RNA was impossible. The CyScribe™ First Strand Labelling Kit (Amersham Pharmacia Biotech) uses a modified reverse transcriptase, CyScribe™ enzyme, to incorporate the cyanine dyes into cDNA reverse transcribed from target RNA. CyScribe enzyme was developed to overcome sparse and uneven incorporation due to the larger size of the cyanine dyes, especially Cy5™. The protocol required 10 μg of total RNA to produce barely enough Cy5™ labelled cDNA for a single microarray slide. Where RNA quantity is limiting, this is less efficient than other methods. Furthermore, despite the modified reverse transcriptase, unequal dye incorporation is still a problem. Integration of aminoallyl dUTP into the cDNA, followed by conjugation of the Cy dyes has a number of advantages. Firstly, the aminoallyl modified nucleotides are much less bulky than the dye modified molecules and thus are more efficiently incorporated into the cDNA. Secondly, the monoreactive dyes, Cy3™ and Cy5™, bind with similar avidity to the modified cDNA. Thirdly the dyes bound via a linker cause less interference with hybridization. There was no detectable difference in the performance of the 3 kits studied. However, the Atlas™ PowerScript™ Labelling Kit (Clontech) has been designed as a replacement for the Atlas Glass labelling kit (Clontech), with an improved reverse transcriptase that generates longer cDNAs. It also has a slightly simpler protocol than the CyScribe™ PostLabelling Kit (GE Healthcare) and has therefore been chosen as the labelling method of choice. However, we have modified the Powerscript protocol, by using NaOH instead of RNase H to degrade the template. RNase H removes the template as efficiently as NaOH. Degradation with NaOH allows better quality control: the amount of cDNA and the FOI of the labelled nucleotides can be estimated (cDNA with too high an FOI is inhibited from hybridization). Furthermore, the yield of labelled cDNA from the NaOH modified protocol was higher. PelletPaintNF™ (Novagen, Merk Biosciences) promotes pellet recovery during ethanol precipitation and increases yield. Treatment of the RNA samples with DNase prior to amplification has little effect on the yield of aRNA and its subsequent labelling. Furthermore, the detection profile of the genes is only very slightly altered. Under stringent analysis conditions, a maximum of nine genes are shown to be differentially detected. However, only three were confirmed as differentially regulated via PCR (one upregulated and two downregulated in the DNase treated samples as compared to untreated). We are not unduly concerned that, were we to relax the analysis conditions, which more differentially regulated genes would appear. We have used the same methods to compare various amplified embryonic samples with picked ES cell colonies; where real biological differences exist, the same analysis methods can throw up 1000 or more differences between two sample sets (experiments submitted to GEO; GSE8881). Why these 3 genes should appear changed when the same sample is treated with DNase is unknown. Formations of stable RNA/DNA duplexes at points in the amplification protocol may inhibit amplification of these particular sequences, when DNA

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contamination is present, owing to sequence specific stable secondary structure formation. For genes that appear upregulated, perhaps the contaminating DNA can be used for ectopic amplification, owing to sequence related affinities with the amplification primers. Primer sequence related hypotheses cannot be fully tested, owing to non-disclosure of primer sequence by the kit manufacturers. Regardless, given that this phenomenon applies to only 3 ESTs, the indications are that this is a sequence specific problem. It should be noted that if these 3 sequences are of particular interest to anyone carrying out an array experiment, the targets should be DNAse treated. None of these genes was detected by GeneSight, implying that Limma (Bioconductor) is a better analysis tool. Limma data normalisation by print tip Loess, using the “normexp” function with an offset of 50 for background adjustment, and the same analysis incorporating A-quantile between array normalisation both gave a list of 9 significant genes. Leaving out the background adjustment gave a list of 6 genes and missed one of the confirmed genes (see Table 5). It may be the case that in order to maximize detection of differentially regulated genes from similar experiments that a certain amount of noise is inevitable. It was possible to amplify all genes of interest from doubly amplified RNA. However, doing so does not eliminate the possibility of amplification bias affecting the results. We have found that it is possible to obtain abundant PCR signals from betaactin and Gapdh after 30 cycles following reverse transcription of unamplified RNA. Highly expressed genes should therefore be very easy to assay, while those expressed at lower levels may either need two rounds of PCR or a single round of amplification prior to RT. When checking expression levels of unamplified RNAvia PCR, however, it is essential to DNAse treat unamplified RNA, or signal will appear in the no RT controls. 5. Simplified method description We conclude that an optimized protocol for arraying minuscule samples on cDNA arrays involves the use of the Zymo Mini RNA amplification kit (Zymo) for RNA extraction, the RiboAmp kit (Arcturus) for RNA amplification, the PowerScript kit (Clontech) using a modified protocol for target labelling, followed by analysis using Limma (Bioconductor) as described. DNAse treatment prior to amplification does not greatly alter results. RT-PCR confirmation of the majority of differentially regulated genes should be possible from unamplified samples. This protocol will be of utility to anyone studying the transcription profiles of small samples derived from embryos, larvae of aquatic vertebrates, embryonic and adult invertebrates, human biopsies or samples obtained from laser capture microdissection, among others. Acknowledgements We would like to thank Dr. Mike Wride for a critical reading of this manuscript. We would also like to thank the staff of the Cardiff University Microarray Facility (Steven Turner and Victoria Workman) for all their help. This project has been funded by the BBSRC and the Wales Gene Park.

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