Molecular Cloning Fourth Edition, A Laboratory Manual, by Michael R. Green and Joseph Sambrook

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Chapter 10: Nucleic Acid Platform Technologies

Rando Oliver, Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts 01605

INTRODUCTION

EVERYMICROARRAY EXPERIMENT IS BASED ON A COMMON format (Fig. 1). First, a large number of nucleotide spots are arrayed onto a substrate, typically a glass slide, a silicon chip, or microbeads. Second, a complex population of nucleic acids (isolated from cells, selected from in vitrosynthesized libraries, or obtained from another source) is labeled, typically with fluorescent dyes. Third, the labeled nucleic acids are allowed to hybridize to their complementary spot(s) on the microarray. Fourth, the hybridized microarray is washed, allowing the amount of hybridized label to then be quantified. Analysis of the raw data generates a readout of the levels of each species of RNA in the original complex population.

Microarrays can be run in one-color or two-color formats. In a one-color microarray, a single sample (e.g., RNA from liver cells) is labeled, and the abundance of a species of RNA is inferred from the intensity of the spot(s) complementary to the relevant gene. Because, for each spot, hybridization is affected by a panoply of factors, interpretation of single-color experiments can be complicated. Practically, however, many of these complicating factors are resolvable by using the proper bioinformatics tools during data analysis. Two-color microarrays (Fig. 1) use a competitive hybridization in which one nucleic acid sample is labeled with one color (green), and a related sample is labeled with a second color (red). Following hybridization and removal of unbound nucleic acid, the microarray is scanned with lasers to detect where the red- and green-labeled molecules have bound. The intensity of each spot is determined, and the red/green ratio is measured for each spot. During data analysis, this ratio can be used to measure the ratio of the amount of related nucleic acid molecules in the two samples. For example, if RNA from normal liver is tagged green and RNA from a liver tumor is labeled red, then red spots would represent RNAs that are up-regulated in the tumor, and green spots would represent down-regulated RNAs.

Tiling microarrays consist of regularly spaced oligonucleotides that provide dense coverage of a genome or portion of a genome. For example, yeast chromosome III was tiled using oligonucleotides 50 nucleotides long spaced every 20 bases; that is, spot 1 comprised bases 150, spot 2 bases 2170, and so forth. Tiling microarrays therefore interrogate continuously along a chromosomal path, enabling applications, such as transcript structure and protein localization, to be performed with almost complete genomic coverage. One disadvantage to tiling microarrays is that not all genomic sequence is equally suited to microarray applications. For example, not all 50-mers in a genome are unique, leading to gaps in the tiling path where hybridization-based approaches cannot determine whether the hybridizing material comes from copy 1 or copy 2 (or copy N) in the genome. Furthermore, hybridization-related properties such as the percentage of A and T residues and oligonucleotide melting temperature vary from probe to probe, although many issues like these can be resolved by normalizing two-color arrays. Normalization, which compensates for systematic technical differences between spots and/or arrays, clarifies the systematic biological differences between samples.


MICROARRAY APPLICATIONS

Although the majority of early microarray studies focused on mRNA expression profiling, microarrays can be used for any purpose that involves comparison of two nucleotide populationsfor example, comparison of RNAs extracted from normal and tumor cells. In this section, we list several examples (by no means exhaustive) of published microarray analyses.

mRNA Expression Levels

Early gene expression microarrays consisted of arrayed spots each containing a cDNA corresponding to a single gene. For example, one of the first yeast gene expression microarrays consisted of about 6000 spots composed of cDNA corresponding to nearly every gene in the Saccharomyces cerevisiae genome (DeRisi et al. 1997). cDNA microarrays have fallen into disuse because of the advent of oligonucleotide microarrays. Advances in oligonucleotide synthesis make it possible to generate gene expression microarrays composed of long oligonucleotides that have been designed to be complementary to a given gene. Oligonucleotides can be synthesized and sold in soluble form, which are then arrayed by a spotting robot, as with cDNA arrays. Alternatively, several commercial microarray suppliers use in situ oligonucleotide synthesis, in which oligonucleotides are printed directly onto the microarray substrate. This process allows for much more flexible generation of oligonucleotide microarrays and, thus, is used more often than soluble oligonucleotide arrays because a laboratory need not purchase an excess of oligonucleotides in order to run microarray experiments.

Monitoring Changes in the Transcriptome

A typical mRNA expression experiment begins with the isolation of poly(A) RNA from two related populations of cells, such as yeast growing at 30C and yeast that have been shifted from 30C to 37C. Fluorophore-labeled cDNA is prepared from each mRNA population. Typically, the cyanine dyes used are Cy5 (pseudo-colored red in microarray images) and Cy3 (pseudo-colored green). The labeled cDNAs are mixed and hybridized to a microarray. After hybridization and washing, the microarray is scanned with lasers that excite the fluorophores, and the image is processed, generating a Cy5/Cy3 ratio for every spot on the array. The microarray data are normalized, and a list of the Cy5/Cy3 ratios per gene is generated, which can be analyzed for genome-wide changes in the transcriptome.

The microarray data are normalized so that the average log2(Cy5/Cy3) is 0; this is a necessary step because the absolute intensity of either channel (Cy5 or Cy3) is subject to a wide range of modifying influences, such as input RNA level and labeling efficiency, among others. This normalization should always be kept in mind in interpreting microarray studies. For example, microarray results for a comparison between liver RNA and sperm RNA populations might indicate that a number of RNAs are relatively enriched in sperm, but the absolute numbers of those RNAs per nucleus are still probably lower in sperm than in liver given the disparity in RNA abundance in the two types of cells.

mRNA Abundance

The experiment described above is capable of providing information about the relative abundances of every mRNA expressed in the two cell populations. What about absolute expression levels? These can be assayed in two ways. First, single-color experiments using the Affymetrix platform provide a reasonable measure of mRNA abundancethe intensity of a given spot is related to the absolute abundance of its complementary mRNA in the original labeled population. Second, for two-color experiments, a straightforward way to estimate transcript abundance is to compare labeled mRNA with labeled genomic DNA, which, in principle, should be present in a single copy per gene (unless the gene in question comes from a multicopy family). Thus, gDNA is labeled with Cy3, mRNA is labeled with Cy5, and Cy5/Cy3 ratios provide a relative measure of mRNA abundance. These ratios can be converted to absolute mRNA abundance if the absolute abundance of any RNA is already known. Normalization can be used to set the absolute level of these RNAs, and all remaining RNA abundances can be inferred from their Cy5/Cy3 ratios relative to the Cy5/Cy3 ratios for the known standards.

Comparative Genomic Hybridization

Comparative genomic hybridization (CGH) is used to detect changes in DNA copy number across a genome (Pollack et al. 1999). In a CGH experiment, the two populations of genomic DNA are fluorescently labeled using Klenow DNA polymerase. Labeled DNA is compared using the same experimental design as for analysis of mRNA expression. Changes in copy number are reflected in the Cy5/Cy3 ratios. This method has been used most commonly in the study of tumors, where there is a well-documented role both for gene loss and for amplification in the process of oncogenesis. By analyzing changes in gene copy number, regional amplification or even whole chromosome copy-number changes have been characterized. Variants of this approach have also been used to characterize replication timing, as DNA isolated during S phase shows variations in copy number, based on whether or not the segment of genomic DNA has replicated.

Determination of Transcript Structure

Although changes in mRNA levels can be analyzed easily using microarrays constructed with one probe (spot) per gene, different methods are required to detect variations in transcript structure. These variations include changes in splicing patterns and in the start and stop sites for transcription. These and other changes in transcript structure can be detected and mapped using tiling microarrays. For example, transcriptional start sites can be mapped to 1020-nucleotide resolution by a variant of the absolute abundance hybridization method described above. Briefly, mRNA is labeled with Cy5 and hybridized against Cy3-labeled genomic DNA. In principle, an expressed RNA should appear as a long square wave of high Cy5/Cy3 ratios: Cy5/Cy3 ratios should be low in spots corresponding to genomic sequences that flank those complementary to the RNA. High Cy5/Cy3 ratios should start at spots containing sequences corresponding to the 5 end of the RNA and should continue to the end of the RNA (Fig. 2). Indeed, this technique has been used to determine transcriptional start sites to 20-nucleotide resolution in yeast (Yuan et al. 2005) and to detect and map novel transcripts in human cells (Shoemaker et al. 2001).

Variations in splicing patterns can also be detected and mapped using microarrays. With tiling arrays, expressed exons should appear as peaks of Cy5/Cy3. Another way to determine exon inclusion is to use an exon microarray, in which each spot on the array corresponds to a single exon. When probing an exon microarray, for a given tissue all of a gene's expressed exons will show high Cy5/Cy3 (Shoemaker et al. 2001).

Tiling microarrays have also been used to discover noncoding RNAs and novel genes. The reason for this is obviousa classical, expression microarray only covers what is known, whereas a tiling microarray can be designed to contain an entire genome. For example, tiling microarrays of chromosomes 21 and 22 revealed the nearly pervasive low-level transcription of the human genome (Kapranov et al. 2002).

Identifying RNAProtein Interactions

RNA molecules associated with specific RNA-binding proteins have been identified using immunoprecipitation assays to isolate a protein of interest. The associated RNA molecules (or regions of RNA) are then compared with nonspecific RNAs that have been isolated in control immunoprecipitations lacking antibody (Hieronymus and Silver 2003; Gerber et al. 2004).

Subcellular Localization of RNA Populations

The functional properties of RNAs (e.g., translation) can be assayed by microarray using more sophisticated fractionation techniques. In an early study, polysomal RNAs were analyzed by microarray techniques (Arava et al. 2003). Subcellular localization of RNAs can also be studied when appropriate purification techniques exist; for example, isolation of dendrites from neurons has been used to identify the relevant RNA populations (Eberwine et al. 2002).

Protein Localization Studies

At present, one of the most popular applications of tiling microarrays is known as chromatin immunoprecipitation on chip (or ChIP-on-chip; see Chapter 20). By this method, the location of a specific protein, such as a transcription factor, is assayed through the use of chromatin immunoprecipitation. Briefly, proteins are covalently cross-linked to the genome using formaldehyde, and chromatin is sheared by sonication to small (500 bp) fragments. Using an antibody specific to a given protein (with either an antibody to an epitope tag, a protein-specific antibody, or even an antibody against a specific modification state), DNA associated with the protein in question is isolated by immunoprecipitation. After washing, the cross-links are dismantled and genomic DNA is isolated. The DNA is typically amplified, and the amplified material is labeled and hybridized competitively against labeled amplification reactions from no-antibody controls or from pre-IP material. Analysis of ChIP-on-chip microarray results provides a snapshot of a transcription factor's location on the genome and has proven to be a tremendously powerful method for studying transcriptional regulation by transcription factors and chromatin (Ren et al. 2000; Iyer et al. 2001).

Nuclease Accessibility as a Structural Probe

Tiling microarrays can also be hybridized with genomic DNA that has been treated diagnostically with a nuclease before labeling the DNA. This method takes advantage of the fact that chromatin packaging of the genome dramatically affects nuclease accessibility. Two nucleases have been used in genome-wide studies. First, DNase Ihypersensitive sites have long been known to occur at genomic regulatory elements such as promoters, enhancers, and insulators. Here, chromatin is lightly digested with DNase I, sites of cleavage are isolated, and DNA surrounding cleavage sites is interrogated by tiling microarray analysis (Sabo et al. 2006). Second, micrococcal nuclease (MNase) is a nonprocessive nuclease with preference for the linker DNA between nucleosomes. Analysis of the DNA protected from MNase cleavage including comparison to whole genomic DNA has proven to be an invaluable tool in genome-wide nucleosome mapping studies (Yuan et al. 2005).

Splicing Microarrays

Although exon microarrays (see above) can monitor expression of each exon represented on the array, the structure of the transcript cannot be discerned. Specifically, exon connectivity is not detected using exon microarrays. For example, expression of exons 1, 2, 3, and 5 could indicate a single species containing all four exons, or two or more distinct species (1-2-5 and 1-3-5, etc.). To determine connectivity of exons requires the use of splicing microarrays, which are comprised of oligonucleotides complementary to specific splice junctions.

Resequencing and SNP Detection

The sequence specificity of DNA hybridization has also been leveraged for genome analyses such as resequencing and polymorphism detection. In these applications, each genomic location is represented on the microarray by several oligonucleotides that differ by a single nucleotide. In other words, if the genomic region of interest has the sequence AATGCCA, oligonucleotides containing AATTCCA, AATCCCA, and AATACCA will also be printed. Hybridization of genomic DNA to these oligonucleotides can be used to determine mutations or sequence polymorphisms at the site interrogated by the set of oligonucleotides.

One way to use microarrays for resequencing is the so-called sequence capture protocol (Hodges et al. 2007). A microarray is printed with oligonucleotides corresponding to genomic regions of interest. DNA or RNA is hybridized to the microarray, and all complementary sequences are retained on the microarray through the washing protocols. After retention of desired genomic regions via hybridization, this hybridized material is eluted and sequenced using high-depth sequencing methods (Chapter 11).


PERFORMING MICROARRAY EXPERIMENTS

Although the specific bound oligonucleotides and labeled probes and details of the analysis of microarray hybridizations differ depending on the experimental questions being asked, most microarray experiments involve six steps.

  • 1. Design a microarray.
  • 2. Print or purchase a microarray.
  • 3. Isolate and amplify the DNA or RNA probe material.
  • 4. Label the DNA or RNA with fluorescent groups.
  • 5. Hybridize the labeled probes to the microarray.
  • 6. Analyze the microarray hybridization results.

For the remainder of the chapter, we discuss each of these steps in turn. Protocols are provided for printing a microarray in-house (Protocol 1) and for amplifying DNA and RNA following isolation (Protocols 24). Several techniques for adding fluorescent moieties to the nucleic acids are provided (Protocols 58). Protocol 9 explains how to block the positive charges of the polylysines bound to homemade microarrays. Finally, there is a detailed generalized protocol for hybridization of labeled probes to a microarray, as well as scanning, formatting, and storing the microarray hybridization data (Protocol 10). A brief guide to microarray analysis is contained in Chapter 8.

Designing a Microarray

A great variety of microarrays are commercially available, and for most researchers, an off-the-shelf product will suffice. If not, customized microarrays can be ordered from several manufacturers or designed (and printed) in-house. The appropriate design of a microarray will depend on the application for which it is to be used. There are, however, some basic design principles that are common to most applications. We consider two types of basic microarray: the gene expression microarray and the tiling microarray. Specialized formats, such as splicing arrays and resequencing arrays, have been designed for different organisms, and we point interested readers to the relevant literature (Hacia 1999; Mockler et al. 2005; Blencowe 2006; Hughes et al. 2006; Calarco et al. 2007; Cowell and Hawthorn 2007; Gresham et al. 2008).

Gene Expression Microarrays

Designing oligonucleotides for microarrays requires expertise in bioinformatics, but some basic design properties are easily understood. First, choose an appropriate oligonucleotide length. The majority of oligonucleotide microarrays today are printed with oligonucleotides 5070 nucleotides long. Designing oligonucleotides of optimal lengths requires consideration of several factors, including signal strength, specificity, cost, and efficiency of synthesis. Shorter oligonucleotides are more likely to cross-hybridize to many different regions of a given genome and will often have very low melting temperatures, making hybridization technically problematic. A detailed analysis of oligonucleotide length versus sensitivity and specificity can be found in Hughes et al. (2001). A reasonable rule of thumb is that for most applications, 60-nucleotide oligonucleotides provide the best balance between these competing constraints.

The second consideration in oligonucleotide design is the degree of specificity or complementarity between the oligonucleotides on the microarray and the RNA species in the organism of interest. In general, the oligonucleotides should be designed so that they are complementary to only one RNA species. Commonly, BLAST searches (see Chapter 8) are used to screen the sequence of each oligonucleotide against the relevant genomic sequence to identify the potential for multiple hits in the genome of interest. Ideally, rather than focusing on a cut-off BLAST value, the user should select for each gene the oligonucleotide having the lowest-scoring second match in the genome.

The third design consideration is optimization of the hybridization properties. Several features can be optimized, but the most important is the melting temperature (Tm) of the oligonucleotide. Specifically, the Tms of all of the oligonucleotides on the array should be within as narrow a window as is feasible. Other characteristics, such as entropy (i.e., the complexity of the sequence), GC, and self-complementarity should be optimized as well. Software tools are available to help with oligonucleotide design, ranging from publicly available tools like ArrayOligoSelector (see below) to commercial services provided by microarray companies (for further details of oligonucleotide design, see Chapters 7 and 8).

As an example, consider a case in which an investigator wishes to design gene expression microarrays for a non-model organism with a recently sequenced genome. The first step in designing gene expression microarrays is, of course, to identify the genes. As this is routinely performed in any genome sequencing effort, we will assume that genes have already been predicted using standard tools (Zhang 2002; Ashurst and Collins 2003; Brent 2005; Solovyev et al. 2006).

ArrayOligoSelector

Once coding regions are identified, oligonucleotides need to be designed for each gene. Many programs exist for oligonucleotide selection, including ArrayOligoSelector (Bozdech et al. 2003a,b), a commonly used program that is freely available at http://arrayoligosel.sourceforge.net/ (for further details, see Chapters 7 and 8).

ArrayOligoSelector is designed to analyze a complete genome and prepare oligonucleotides of a user-defined length. For every oligonucleotide, ArrayOligoSelector calculates scores for uniqueness, sequence complexity, self-annealing, and GC content. Uniqueness is a measure of the theoretical difference in binding energy between a given oligonucleotide and either its perfect match or the next most homologous genomic sequence. Sequence complexity allows the user to filter oligonucleotides with homopolymeric tracts, which otherwise may cause hybridization problems. The self-annealing score is a measure of the secondary structure generated by the self-annealing of an oligonucleotide. Self-annealing is another potential source of hybridization problems. Finally, it is important to minimize variation in GC percentage among the oligonucleotide sequences, both to minimize Tm variation and to minimize variability in the fluorescence intensity among the spots.

When running ArrayOligoSelector, the following features can be specified: oligonucleotide length, GC, number of oligonucleotides per gene, sequences to mask, and uniqueness cutoff. Common oligonucleotide lengths are 60-mers or 70-mers. GC will vary depending on the GC of the genome in question and will typically be chosen as the genomic coding region average. The number of oligonucleotides per gene will depend on whether the microarray is to be purchased or printed in-house. If oligonucleotides are purchased for in-house printing, then cost and the printable spot density will preclude using more than one or two oligonucleotides per gene. Alternatively, if commercial arrays are to be used, then the number of spots available will determine the number of oligonucleotides to be chosen per gene. Masking sequences are not commonly specified, but if a problematic short repeat element is present in the genome, then it is sometimes valuable to mask it out of the microarray oligonucleotides. The uniqueness cutoff is typically left blank, which will result in the default value being used (for additional information, see the ArrayOligoSelector manual).

The output of ArrayOligoSelector can be filtered by the experimenter. For example, it is often desirable to use oligonucleotides located toward the 3 ends of genes because reverse-transcriptase-based labeling is more efficient in this region.

Tiling Microarrays

Oligonucleotide design for tiling microarrays is more straightforward than for gene expression microarrays because tiling presumes that essentially all of the oligonucleotides within a region of interest will be included on the microarray. The simplest tiling design involves choosing nucleotides 150 (say) of some region to be tiled as spot 1, nucleotides 2170 as spot 2, and so forth. Once all of the oligonucleotides have been designed, BLAST can be used to find oligonucleotides with multiple identical matches in the genome of interest, and these oligonucleotides can be removed. More subtle tiling designs incorporate a small amount of wiggle in the oligonucleotide location; thus spot 2 might run from nucleotides 1665, spot 3 might run from 4594, and so forth. By doing this, the process of matching hybridization properties, such as Tm and GC, is better than with a simple tiling microarray.

Printing or Purchasing a Microarray

Once oligonucleotides have been designed, microarrays can be printed commercially. Alternatively, oligonucleotides can be synthesized commercially or by an in-house core facility, and then printed in-house using a spotting robot (see Protocol 1).

Isolating and Amplifying Nucleic Acid Samples for Hybridization to Microarrays

Most of the sample preparation procedures for microarrays follow standard protocols, many of which can be found elsewhere in this manual. For example, comparative genomic hybridization (CGH) analyses use genomic DNA isolated from samples of interest, as described in Chapter 1. Gene expression or splicing studies use either total RNA or mRNA, purified as described in Chapter 6. Protein localization analysis starts with material isolated via chromatin immunoprecipitation (ChIP), as described in Chapter 20. Typically, however, the intended assays for many of these protocols are based on blotting techniques or quantitative PCR readouts, which often require only nanograms of material. Microarray labeling, on the other hand, typically requires several micrograms of nucleic acid; therefore, an amplification step is necessary before labeling.

General Notes on Amplification

Because amplification of the nucleic acid that will be used to generate labeled microarray probes occurs before the hybridization step, it must not bias representation of any particular sequences in the genome. Thus, unbiased (or minimally biased) whole-genome amplification protocols are a key component of many microarray applications.

Three amplification protocols are included in this chapter: two for DNA (Protocols 2 and 3) and one for RNA (Protocol 4). There are several practical items to keep in mind while performing an amplification protocol. First, when trying an amplification method for the first time, start with a large and easily obtainable pool of material (e.g., liver RNA) and amplify an aliquot of the original pool. Microarray comparisons between the amplified material and the original bulk pool then provide a valuable readout of amplification biases. A perfect amplification would result in a yellow array with no spots showing any differences between the bulk and amplified material. Second, avoid contaminating your sample with anything that might contain DNA or RNA. Even tiny amounts of foreign nucleic acids will be amplified, contaminating your sample and corrupting your microarray experiments. Always wear gloves and use filter pipette tips when performing the isolation and amplification protocols. Finally, always include a control amplification with water only (no DNA or RNA) to ensure that the reagents are not contaminated with amplifiable material.

RNA Amplification for Expression Profiling

Less common than DNA amplification, RNA amplification is nonetheless required in experiments that use small populations of cells, such as may occur in neurobiological studies using laser-captured cell populations. RNA amplification kits are available from several vendors (e.g., Ambion), or amplification can be performed as described in Protocol 4.

Generating Fluorescently Labeled Nucleic Acid Probes

Labeling nucleic acids for use in microarrays is similar for most microarray platforms, except for Affymetrix microarrays. For most platforms (homemade or commercial), fluorescent molecules are attached to DNA by the Klenow fragment of DNA polymerase I. For labeling RNA, reverse transcriptase is used to prepare labeled cDNA. Labeling methods can include a fluorescent dNTP in the labeling reaction (Protocols 5 and 7). Because this approach is rapid but expensive, a cheaper but lengthier alternative first incorporates aminoallyl nucleotides into the nucleic acid molecules and then couples the aminoallyl group to the fluorophore (Protocols 6 and 8). In all of the labeling protocols, the source nucleic acids can be either unamplified or amplified before labeling.

Hybridizing to a Microarray

Methods for hybridization of labeled probe materials to a microarray differ significantly when using home-printed microarrays versus commercial microarrays. For home-printed microarrays, slides must first be blocked because polylysines remaining on the slide will cause significant background binding to labeled material unless they are neutralized by reaction with succinic anhydride. Protocols 9 and 10 provide methods, respectively, for blocking and hybridizing to homemade arrays. When using commercial microarrays, hybridization protocols are typically provided. Following hybridization, microarrays are scanned in a bench-top scanner. The data are collected, formatted, and stored digitally for subsequent analysis.

Analyzing Microarray Data

The tools used for analysis of microarray data will depend on the experimental question being asked: Localization studies require a different set of tools than do gene expression studies. Some of the available resources are described in Chapter 8. Here, we outline some of the basic steps in data analysis.

The first step in microarray data analysis is to remove bad data (i.e., data from spots that were flagged because they were obscured by fluorescent precipitate, etc.), and to normalize the remaining data. Working with a .gpr file, which is the standard output of GenePix software, we typically eliminate all flagged probes and then work exclusively with the Log2 ratio data. More advanced users may consider using specific features, such as Foreground and Background intensities.

Most two-color microarray studies are normalized to an average Log2 ratio value of 0. The assumption implicit in this normalization is that there was no overall change in whatever is being measured. Thus, it is important to remember when looking at such normalized data that relative values are being measured, not absolute values. Practically, normalization can be performed by averaging the unflagged log ratio value, then subtracting this value from every one of the entries in the column. This can be done by hand in spreadsheet programs or using common commands in languages such as MATLAB, Perl, or R.

Once normalization is completed, the list can be sorted by log ratio value to identify genes that are dramatically up-regulated or down-regulated (if gene expression) or identify loci with high levels of enrichment of the protein in question. At this point, data analysis paths will diverge depending on the questions being asked. However, because many microarray studies involve multiple microarrays, it is often useful to cluster data to identify genes or loci that share similar behavior.

For clustering and visualization, numerous programs are available online. We use the classic Cluster and TreeView programs (Eisen et al. 1998), available online at SourceForge (http://sourceforge.net/) or via the Eisen laboratory website (http://www.eisenlab.org/). Because sample files that guide the formatting of files for clustering are also available, formatting will not be described here. Briefly, however, data will be loaded into Cluster, various thresholds will be set (fraction of data missing for a given gene, number of genes changing over some threshold, etc.), and one of several clustering algorithms will be used. The output of the clustering can be visualized with TreeView, allowing users to generate the classic heatmap view of their microarray data.

ACKNOWLEDGMENTS

The protocols in this chapter were modified from protocols written and generously provided by Ash Alizadeh, Chih Long Liu, Audrey Gasch, Jason Lieb, Bing Ren, and L. Ryan Baugh.


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WWW RESOURCES
Source for Array Oligo Selector http://arrayoligosel.sourceforge.net/..
Source for Cluster and Treeview programs http://sourceforge.net/ and http://www.eisenlab.org/.
Protocol 1: Printing Microarrays doi10.1101/molclon.000179

Most laboratories will choose to order microarrays from a commercial vendor, such as NimbleGen, Agilent, Affymetrix, or Illumina. These vendors sell a number of products encompassing genomes from the three domains of life. In addition, custom microarrays can be made to order by most of these companies. Microarray printing, however, is sufficiently straightforward that even small laboratories with extensive microarray needs may find it cost-effective and worth the effort to produce their own microarrays. Printing microarrays requires a spotting robot. Several commercial spotting robots are available, such as the OmniGrid series (Digilab Genomic Solutions). These instruments are expensive and will most frequently be housed in institutional core facilities or in large laboratories. As an alternative, designs for building a spotting robot have been available from the Brown laboratory at Stanford for many years. Protocols for building a spotting robot are beyond the scope of this chapter, but interested parties can find a protocol at http://cmgm.stanford.edu/pbrown/mguide/.

For typical microarrays, oligonucleotides are prepared at 2040 M in aqueous 3 SSC and are distributed into 384-well plates. Oligonucleotides are printed to polylysine-coated microarray slides. It is best to print when the humidity is at least 50. When the humidity is 2030, there can be problems with pins drying. To increase or maintain proper humidity, consider building a humidity-controlled enclosure around the robot. For robots lacking an enclosure, it may be adequate to run a humidifier or two during the printing procedure.

The details of a printing protocol vary depending on the robot being used; thus, follow the manufacturer's instructions. However, a typical workflow using a robot equipped with contact steel quill pins is provided in this protocol.


MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Recipes for reagents specific to this protocol, marked <R>, are provided at the end of the protocol. See Appendix 1 for recipes for commonly used stock solutions, buffers, and reagents, marked <A>. Dilute stock solutions to the appropriate concentrations.

Reagents

  • Oligonucleotides of desired sequences
    • Oligonucleotides are typically prepared at 2040 m in aqueous 3 SSC.
  • Salmon sperm DNA <A>, sheared (150 ng/L resuspended in 3 SSC)
  • SSC (3) <A>

Equipment

  • Desiccator
  • Nitrogen gas, compressed
  • Plates (384 wells)
  • Polylysine-coated slides
    • Although polylysine coating can be done in-house, in our experience, the failure rate is significant and purchasing commercial polylysine-coated slides (e.g., Erie Scientific) has proven to be the most cost-effective solution.
  • Slide box, plastic
  • Spotting robot


METHOD
Preparing to Print
Day 0

  • 1. Array the oligonucleotides into 384-well plates with identical volumes per well (typically 1020 L per well). If the oligonucleotide solutions were frozen, thaw the master plates overnight at 4C. If the oligonucleotides were dried, resuspend them in 3 SSC, and let them incubate overnight at 4C.
  • 2. Test that the spotting pins are all printing and that the arrayer is properly calibrated by performing test prints (Fig. 1). To closely approximate the viscosity of the actual print plates, use a test print plate consisting of 150 ng/L sheared salmon sperm DNA resuspended in 3 SSC. All pins should be able to print several hundred consecutive spots. In addition, test print in several locations on the slide platter to determine whether the platter has warped significantly since its last use.
    • If test prints indicate that the spotting robot is not performing as it should, see Troubleshooting.

Day 1

  • 3. Perform one more test print to make sure that the pins are still printing properly.
  • 4. Place slides gently onto the arrayer platter. Make sure that all slides are sitting flat and are firmly attached (by vacuum, clips, or tape, depending on the arrayer).
    • Handle the slides carefully to avoid producing minute glass chips, which might settle onto the surface of the slide.
  • 5. Blow dust off the slides with compressed nitrogen.
  • 6. If the arrayer has a sonicator water bath for cleaning the pins, fill it with fresh water.
  • 7. Transfer four 384-well oligonucleotide-containing print plates from 4C, and let them come to room temperature for at least 1 h.
  • 8. Centrifuge the plates at 1000 rpm for 2 min to remove condensation from the plate covers. Carefully remove a plate lid and the adhesive plate cover from one of the print plates.
    • Be careful not to jolt the plate because cross-contamination between wells will cause serious problems in any downstream data analysis.
  • 9. Place a plate into the spotting robot's plate holder.

Printing the Microarrays

  • 10. Start the print run. Keep careful notes about plate order and orientation. Knowing the plate order is necessary for creating the .gal file that maps spot position to gene name when microarray data are analyzed. Any mistakes during printing such as printing plates out of order (e.g., plate 3 before plate 2) can be corrected later on as long as they are noted.
  • 11. When a plate is finished printing and the print head has come to a complete stop, either let the plate evaporate in a hood if the plates are stored dry, or cover the plate with foil and its lid and store the plate at 80C.
  • 12. Insert the next print plate into the plate holder.
  • 13. When the print run is done, let the slides dry overnight, unless you are performing back-to-back print runs.

Day 2

  • 14. Transfer all slides from the arrayer into a plastic slide box. Store them in a desiccator.
  • 15. Power down the arrayer.


TROUBLESHOOTING

Problem (Step 2): Printing pins are not printing properly.

Solution: If any pins are not printing, four steps may be taken to improve performance. First, if dirt or other material has clogged the quill tip, wash the pins extensively in a sonicator bath. Second, if examination of a pin under a microscope indicates that the pin is clogged, try clearing the quill tip carefully with a razor blade, which may dislodge the contaminant. Third, subtle variation in pin length may prevent some pins from printing well; swapping pins within the print head may improve printing. Finally, if a pin continues to fail after all of the above steps have been taken, replace it with a fresh pin.

Problem (Step 2): Test plate printing is not uniform everywhere on the arrayer.

Solution: If the arrayer fails to print at particular locations, adjust the print height for that section of the platter. These adjustments can be made using the software that accompanies the robot.


DISCUSSION

A successful print run requires that print plates be handled carefully to prevent cross-contamination, that pins be checked regularly (i.e., after every 384-well plate or two) to confirm that they are still printing, and that the level of water in the bath (or sonicator) for washing pins be always full enough to cover the pins. We typically add 510 mL of water to the sonicator every 6 h during a print run, but this will depend on the ambient humidity.

To check that pins are still printing, shine a flashlight onto the slide at an angle. The salt deposited during printing is white, and when a pin stops printing the pattern of one sector deviates from the others. For example, if five rows of 20 spots have been printed, a perfect 520 rectangle can be seen, but if a pin has dropped out, then the rectangle will be incomplete.

If a pin stops printing, it may be possible to remove the pin carefully and clean it under a microscope if any dust is observed in or on the pin. If no dust can be seen, sometimes a pin can be restored by extensive sonication. Finally, replacing the faulty pin with a spare pin, or moving pins around in the print head, can be used as a last resort. After a print run has been paused, replace the first slide with a clean slide and perform another test print to check on the new arrangement. When restarting the spotting robot, be sure that printing recommences at exactly the location where the print run was paused.

WWW RESOURCE
Protocol for building a spotting robot http://cmgm.stanford.edu/pbrown/mguide/.
Protocol 2: Round A/Round B Amplification of DNA doi10.1101/molclon.000180

The goal of this procedure is to randomly amplify a sample of DNA to achieve the best possible sequence representation. This protocol has been used successfully to amplify genomic representations starting with <10 ng of DNA. The protocol consists of three sets of enzymatic reactions. In Round A, Sequenase is used to extend randomly annealed primers to generate templates for subsequent PCR. During Round B, a specific primer is used to amplify the previously generated templates. Finally, amplified material can be labeled as in Protocol 7 or 8. Alternatively, Round C in this protocol can be used to incorporate either aminoallyl-dUTP or Cy-dye-coupled nucleotides during additional PCR cycles. This protocol may be unsuitable for amplifying material smaller than 250 bp because such material will not be amplified uniformly. In those cases, Protocol 3 is recommended. This protocol was adapted from Bohlander et al. (1992).


MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Recipes for reagents specific to this protocol, marked <R>, are provided at the end of the protocol. See Appendix 1 for recipes for commonly used stock solutions, buffers, and reagents, marked <A>. Dilute stock solutions to the appropriate concentrations.

Reagents

  • aa-dNTP/Cy-dNTP mixture (100) <R>
  • BSA (500 g/mL)
  • DNA, isolated from the sample under study (10100 ng)
    • For example, for CGH analysis, genomic DNA is isolated as described in Chapter 1; for protein localization studies, DNA is isolated by chromatin immunoprecipitation (ChIP) as in Chapter 20.
  • dNTP mixture (3 mM)
  • dNTPs (100; 20 mM each nucleotide)
  • DTT (0.1 M)
  • MgCl2 (25 mM)
  • PCR buffer (10) (500 mM KCl, 100 mM Tris at pH 8.3)
  • Primer A: GTTTCCCAGTCACGATCNNNNNNNNN (40 pmol/L)
  • Primer B: GTTTCCCAGTCACGATC (100 pmol/L)
  • Sequenase (13 units/L) (US Biochemical, catalog no. 70775)
  • Sequenase buffer (5)
  • Sequenase dilution buffer
  • Taq polymerase (5 units/L)

Equipment

  • Agarose gel (1)
  • Microcon 30 spin column (Millipore)
  • Thermal cycler


METHOD
Round A: Extending Randomly Annealed Primers

  • 1. Prepare Round A reactions as follows:
    • As little as 10 ng of DNA can be effectively amplified by this protocol. As a control, set up a reaction in which the DNA is replaced with H2O.
  • 2. Denature the template DNA and anneal the primer by heating for 2 min at 94C and then rapidly cooling to 10C. Keep the reaction for 5 min at 10C.
  • 3. Assemble the reaction mixture:
  • 4. Combine the reaction mixture and the templateprimer mix. Place the tube(s) into a thermal cycler, and extend the primers as follows.
    • i. Ramp from 10C to 37C over 8 min.
    • ii. Hold at 37C for 8 min.
    • iii. Rapidly ramp to 94C and hold for 2 min.
    • iv. Rapidly ramp to 10C, add 1.2 L of diluted Sequenase (1:4 dilution), and hold for 5 min at 10C.
    • v. Ramp from 10C to 37C over 8 min.
    • vi. Hold at 37C for 8 min.
  • 5. Dilute samples with water to a final volume of 60 L.

Round B: PCR Amplification

  • 6. Prepare Round B reactions as follows:
  • 7. Place the tube(s) into a thermal cycler, and amplify the templates as follows.
    Run 1535 cycles, depending on the amount of starting material.
    • To optimize the number of cycles, it may be necessary to remove an aliquot every two cycles to monitor the progress of the amplification. It is best to use the minimal number of cycles that generates a visible smear (see Step 8).
      Make sure that there is no DNA in the negative control lane!
  • 8. Run 5 L of the reaction on a 1 agarose gel. A smear of DNA should be present between 500 and 1000 bp.
  • 9. Label the DNA using the Round C procedure, Protocol 7, or Protocol 8.

Round C: Cyanine Labeling or Aminoallyl Activation of DNA

  • 10. Prepare the Round C reaction:
  • 11. Place the tube(s) into a thermal cycler, and label the templates as follows:
    Run 1025 cycles.
    • The number of cycles should be determined empirically. The objective is to minimize the number of cycles required to yield 23 g of material for hybridization.
  • 12. If aa-dNTPs were used in Round C, desalt the sample to remove Tris buffer, which interferes with the dye coupling. Add 400 L of water to the sample in a Microcon 30, and centrifuge for 8 min at 12,000 rpm. Repeat again with 500 L of water.
  • 13. Proceed to Cy-dye coupling as described in Protocol 8, Step 11.


DISCUSSION

The output of this amplification protocol will be double-stranded DNA. Quantify the amount of DNA by absorbance (see Chapter 2), and calculate the fold amplification based on the amount of input DNA used. First-time users of this protocol may want to run some amplified DNA on an agarose gel to observe the size distribution.


RECIPE

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

aa-dNTP/Cy-dNTP Mixture (100)
REFERENCE
Bohlander SK, Espinosa R III, Le Beau MM, Rowley JD, Diaz MO. . Year: 1992. A method for the rapid sequence-independent amplification of microdissected chromosomal material. Genomics13: 13221324.
Protocol 3: T7 Linear Amplification of DNA (TLAD) for Nucleosomal and Other DNA < 500 bp doi10.1101/molclon.000181

Protocol 2 has been used extensively in genomic localization analysis and appears to work quite well for typical applications in which DNA is sheared by sonication to 500 bp. However, when DNA is sheared to a population whose modal size is <500 bp, bias in the PCR step skews representation of some genomic loci (Liu et al. 2003). In addition, a subset of applications requires amplifying DNA populations that are smaller than 500 bp; a notable example is ChIP on mononucleosomal DNA, which is 150 bp long (Liu et al. 2005). In these circumstances, T7 linear amplification of DNA (TLAD) is preferred because it more accurately maintains uniform representation of short DNA fragments during amplification than does the amplification method described in Protocol 2.

Amplification of double-stranded DNA by TLAD begins with the addition of a 3 tail of poly-thymidine to DNA by TdT. Second, the Klenow fragment of E. coli DNA polymerase is used, along with a T7-poly(A) primer, to generate a complementary strand that carries a 5 T7 primer. Finally, extension of the original T-tailed DNA strand yields a template suitable for T7-based transcription, which generates amplified RNA (aRNA). This technique avoids the jackpotting issues observed with PCR, in which an early amplification event leads to disproportionate representation of a particular sequence, because PCR follows exponential kinetics, whereas transcription is a linear amplification method.

This protocol takes some time to complete. Table 1 provides estimates of the time required for each procedure.


MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Recipes for reagents specific to this protocol, marked <R>, are provided at the end of the protocol. See Appendix 1 for recipes for commonly used stock solutions, buffers, and reagents, marked <A>. Dilute stock solutions to the appropriate concentrations.

Reagents

  • -Mercaptoethanol
  • Calf intestinal phosphatase (CIP) (New England Biolabs, catalog nos. M0290S or M0290L)
  • CoCl2 (5 mM)
  • Dideoxynucleotide tailing solution (8) (92 mM dTTP, 8 mM ddCTP) (Life Technologies)
  • dNTP mixture (5 mM) <A>
  • EDTA (0.5 M, pH 8.0) <A>
  • Ethanol (95100)
  • Klenow fragment of DNA polymerase I (New England Biolabs)
  • Mineral oil
  • NEB buffer 2 (10) (New England Biolabs)
  • NEB buffer 3 (10) (New England Biolabs)
  • NTP mixture (75 mM)
  • Reaction buffer, provided with kit
  • RNase-free H2O
  • RNase inhibitor
  • T7-A18B primer
    • The primer sequence is 5-GCATTAGCGGCCGCGAAATTAATACGACTCACTATAGGGAG(A)18[B], where B refers to C, G, or T. The primer should be HPLC, PAGE, or equivalent purification grade.
  • T7 RNA polymerase
  • Template DNA (maximum 500 ng per 10 L)
  • Terminal transferase (TdT) (New England Biolabs, catalog nos. M0315S or M0315L)
  • Terminal transferase buffer (5) (Roche, catalog no. 11243276103)

Equipment

  • MinElute kit (QIAGEN, catalog no. 28204)
  • RNase/DNase-free tubes (1.5 mL)
  • RNase-free PCR tubes (0.2 mL)
  • RNeasy Mini Kit (QIAGEN)
  • Rotary evaporator (e.g., SpeedVac)
  • Thermal cycler
  • Vacuum manifold (optional; see Step 17)
  • Water bath or heat block set to 37C


METHOD
CIP Treatment of Samples with Terminal 3-Phosphate Groups

Treatment of DNA with calf intestinal phosphatase (CIP) is only necessary if the source DNA has been sheared or treated with MNase. Treatment of the DNA with CIP removes 3-phosphate groups, leaving free hydroxyl groups on the 3 ends, which are necessary for efficient tailing by TdT. Failure to perform this step will likely reduce the yield of amplification products by 50.

  • 1. Set up the following reaction:
    Incubate the reaction for 1 h at 37C.
    • Each reaction can be scaled up to 100 L per tube.
  • 2. Clean up the DNA with a MinElute column. Follow the protocol supplied by the manufacturer. Elute the DNA in 20 L.
    • When working with <100 ng of DNA, the 10-L elution volume in the manufacturer's protocol may yield less than the 80 claimed by QIAGEN. Increase the elution volume to 1520 L, and reduce the volume of the DNA by drying, if necessary.

Tailing Reaction with Terminal Transferase

  • 3. Set up the tailing reaction:
    • Do not use the NEB buffer 4 supplied with the NEB terminal transferase because DTT in the buffer will precipitate the CoCl2 and inhibit the reaction. Use the cacodylate buffer (1 M potassium cacodylate, 125 mM Tris-HCl, and 1.25 mg/mL BSA at pH 6.6), either supplied with the Roche enzyme or purchased separately. Take precautions in handling this arsenic-containing buffer and use waste-disposal practices appropriate for your institution.
    • Do not freeze and thaw the dNTP mixes more than three times. Additional freezethaw cycles will degrade the dNTPs and will reduce the efficiency of the reaction.
    • Aim for 1 pmol of template molecules. The tested range is 2.575 ng of DNA per 10 L of reaction volume. Scale up the reaction volume accordingly for greater starting amounts. For ChIP samples, use a sensitive UV-Vis spectrophotometer or a fluorometer to quantify the amount of sample precisely. If the amount of DNA is unknown, scale up to a 20-L volume to ensure that there is enough TdT enzyme present for an efficient tailing reaction. Note that if insufficient enzyme is used, the efficiency of subsequent steps in the protocol will be significantly affected and result in significantly reduced yields (as little as 510 of normal expected yields).
    • We strongly suggest that the NEB terminal transferase be used for this protocol; TdT enzyme from other sources may not perform optimally. If using the Roche recombinant TdT, double the volume of enzyme.
  • 4. Add 12 drops of mineral oil to the top of the mixture to prevent evaporation of reactants during incubation. Incubate the reaction for 20 min at 37C.
  • 5. Stop the reaction by adding 2 L (per 10 L of reaction volume) of EDTA (0.5 M, pH 8.0).
  • 6. Clean up the DNA with a MinElute column. Follow the protocol supplied by the manufacturer. Elute the DNA in 20 L.
    • If the starting volume is 10 L, then add 10 L of water to bring the volume to 20 L before adding the reaction to the spin column. When working with <100 ng of DNA, the 10-L elution volume in the manufacturer's protocol may yield less than the 80 claimed by QIAGEN. Increase the elution volume to 1520 L, and dry the volume down if necessary.

Second-Strand Synthesis with Klenow Fragment Polymerase

  • 7. Set up the second-strand synthesis reaction:
    • If production of template-independent product is a significant problem, scale down the reaction volume while keeping the reagent concentrations (except for the T-tailed DNA) constant. See the end of the protocol for an example.
    • Do not freeze and thaw the dNTP mixes more than three times. Additional freezethaw cycles will degrade the dNTPs and will reduce the efficiency of the reaction.
    • NEB (early 2004) switched the supplied buffer for Klenow enzyme from EcoPol buffer to NEB buffer 2. This buffer should provide at least comparable yields to the old buffer and may actually increase yields up to 14.
    • Do not use mineral oil. Trace amounts of mineral oil appear to interfere with cleanup and in vitro transcription.
  • 8. Use the following program in a thermal cycler:
    • i. 2 min at 94C.
    • ii. Ramp from 94C to 35C at 1C/sec, then hold for 2 min to anneal.
    • iii. Ramp from 35C to 25C at 0.5C/sec.
    • iv. Hold for 45 sec at 25C (or up to 6 min).
      • During this time, add 1 L (5 U) of Klenow DNA polymerase. If necessary, centrifuge the tube to remove condensation from the top and sides of the tube.
    • v. 37C, 90 min.
    • vi. (Optional) 4C to temporarily halt enzyme activity until user returns to take reaction tubes out of cycler.
  • 9. Stop the reaction by adding 2.5 L of EDTA (0.5 M, pH 8.0) (the final concentration will be 45 mM).
  • 10. Clean up the DNA with a MinElute column. Follow the protocol supplied by the manufacturer. Elute the DNA in 20 L.
    • When working with <100 ng of DNA, the 10-L elution volume in the manufacturer's protocol may yield less than the 80 claimed by QIAGEN. Increase the elution volume to 1520 L, and dry the volume down if necessary. An elution volume of 20 L at this step increased yields by 3040 for a 50-ng sample.

In Vitro Transcription (IVT)

  • 11. In vitro transcription (IVT) requires that the double-stranded DNA (dsDNA) be in an 8-L volume. Dry down the eluate from 20 to 8 L in a rotary evaporator at medium heat for 1012 min (the drying rate is 1 L/min).
  • 12. Set up the in vitro transcription reaction in 0.2-mL RNase-free PCR tubes as follows:
    • If the IVT kit is new, combine the NTPs in one tube, then realiquot into four tubes. In the first three freezethaw cycles, yields drop 1015 after each cycle. If the NTPs go through more than three freezethaw cycles, each subsequent freezethaw cycle may drop the yield by as much as 50.
    • The buffer should be at room temperature. Adding cold buffer and dsDNA may cause the DNA to precipitate. If there is a precipitate, warm the buffer to 37C until the precipitate dissolves.
  • 13. Incubate the reaction overnight at 37C in a thermal cycler with a heated lid or in an air incubator.
    • The incubation can range from 5 to 20 h; typical is overnight, roughly 16 h.

Amplified RNA (aRNA) Purification Using RNeasy Columns

  • 14. Prepare the buffer (433.5 L per IVT reaction):
  • 15. Aliquot the mix into 1.5-mL RNase/DNase-free tubes.
  • 16. Transfer the contents of the IVT mix (from Step 13) to the RNase/DNase-free tube, and vortex gently and briefly.
  • 17. Add 250 L of 95100 ethanol, and mix well by pipetting. (Do not centrifuge!) Purify the aRNA through an RNeasy column by either the centrifuge method or with a vacuum manifold. aRNA Purification Using a Centrifuge
    • i. Apply the entire sample to an RNeasy Mini spin column mounted on a collection tube. Centrifuge the column for 15 sec at 8000g. Discard the flowthrough.
    • ii. Transfer the RNeasy column to a new 2-mL collection tube. Add 500 L of Buffer RPE (which must have ethanol added before use) and centrifuge for 15 sec at 8000g. Discard the flowthrough, but reuse the collection tube.
    • iii. Add 500 L of Buffer RPE onto the RNeasy column and centrifuge for 2 min at maximum speed.
    • iv. Remove the flowthrough, and pipette another 500 L of Buffer RPE onto the column. Centrifuge for 2 min at maximum speed.
      • This additional wash, which is not in the QIAGEN protocol, is necessary because of guanidinium isothyocyanate contamination in the eluted RNA.
    aRNA Purification Using a Vacuum Manifold
    • i. Apply the sample (700 L) to an RNeasy Mini spin column attached to a vacuum manifold. Apply vacuum.
    • ii. Shut off the vacuum, and pipette 500 L of Buffer RPE onto the RNeasy column. Apply vacuum.
    • iii. Repeat Step ii. Transfer the columns to 2-mL collection tubes. Centrifuge for 1 min at full speed.
    • iv. Return the column to the vacuum manifold, and add 500 L of Buffer RPE. Apply vacuum.
    • v. Transfer the column back to a 2-mL tube. Centrifuge for 1 min at full speed to completely dry the column.
  • 18. Transfer the RNeasy column into a new 1.5-mL collection tube, and add 30 L of RNase-free water directly onto the membrane. Centrifuge for 1 min at 8000g to elute the RNA. Repeat if expected; the yield is >30 g.
  • 19. Check the RNA concentration and purity by measuring the A260 and A260/A280.
  • 20. Proceed to Protocol 5 or 6 to add fluorescent label to the RNA.


TROUBLESHOOTING

In addition to the items below, it may be valuable to consult the troubleshooting section in the manufacturer's manual that accompanies the IVT kit.

Problem (Step 19): Amplified RNA appears to have been damaged by RNase.

Solution: Perform an IVT control, using 250 ng of the pTRI-Xef-linearized plasmid provided with the Ambion IVT kit. If not using the kit, use an appropriate amount of a dsDNA template that contains the pT7 promoter. Be sure that the chosen template has been used successfully as a template for T7 RNA polymerase. Yields should typically range from 100 to 140 g, limited by the 100-g binding capacity of the QIAGEN RNeasy column. If the yield from the IVT control template is poor, contamination with RNases can be assessed by running a 2 nondenaturing agarose gel in Tris-acetateEDTA (TAE) and ethidium bromide. An RNase-contaminated IVT sample will yield a smear of low-molecular-weight material. If RNase contamination is the cause, ensure that aerosol-barrier, RNase-free pipette tips are used, and that working surfaces are treated with RNase-decontaminating agents (e.g., RNaseZap; Ambion catalog no. 9780). This is particularly important when working with ChIP samples.

Problem (Step 19): Yields of aRNA are poor, but RNase contamination is not the cause.

Solution: If there is no RNase contamination detected, it is likely there may be problems with the IVT reaction conditions. Consider the following.

  • The NTP mix has gone through too many freezethaw cycles. NTPs are very sensitive to freezethaw cycles, and each one decreases the yield. Use a fresh IVT kit, and aliquot the NTP mix before use.
  • There has been excessive evaporation of the reaction volume during the incubation. The described IVT conditions (Steps 1113) were designed to limit evaporation and vapor volume during the long incubation period. Using mineral oil is not recommended because it may interfere with either the IVT reaction, the aRNA cleanup, or both.


DISCUSSION

The output of this protocol is amplified RNA (aRNA) suitable for labeling for microarray studies as described in Protocols 5 and 6. It is important to determine the amount of aRNA produced and calculate the mass amplification obtained. Typical amplifications result in at least a 200-fold increased mass yield. For example, 20 g of aRNA is synthesized from an input of 75 ng of DNA. Use a 12 agarose gel to assess the composition and quality of the amplified RNA. Unless knowing the size distribution is crucial, it is usually not necessary to run a denaturing gel. Within the resolution limits of an agarose gel, the amplified product may migrate 2040 bp more slowly on the gel. This shift is to be expected because of the addition of poly(A) tails, the tight size distribution of the poly(A) tail, and the sequence added by the T7 promoter. The size distribution of the poly(A) tail becomes particularly evident in amplification products produced from a single-size template, such as PCR products or a restriction-digested plasmid.

Second-Strand Synthesis with Limiting Primer Amounts

Occasionally a low-molecular-weight band may also appear near the bottom of the gel, at 100 bp. It has been observed under certain amplification conditions, usually when the concentration of starting material is significantly less than that of the primer during second-strand synthesis. In these cases, a substantial amount of small-molecular-weight material may be generated, which likely represents amplification product produced from IVT-valid template synthesized through the formation of primer dimers during second-strand synthesis. These products represent nonproductive material for downstream analysis, and if substantial amounts of this material are observed, then it is necessary to limit the amount of primer.

Limiting the amount of primer is important when amplifying from very small amounts of starting material. Not only will it decrease the amount of primer-dimer product, it can also increase the yield of the desired amplification product. Table 2 describes the single reaction volumes to use for a suggested mass range of starting material.

REFERENCES
Liu CL, Schreiber SL, Bernstein BE. . Year: 2003. Development and validation of a T7 based linear amplification for genomic DNA. BMC Genomics4: 19. doi: 10.1186/1471-2164-4-19.
Liu CL, Kaplan T, Kim M, Buratowski S, Schreiber SL, Friedman N, Rando OJ. . Year: 2005. Single-nucleosome mapping of histone modifications in S. cerevisiae. PLoS Biol3: e328. doi: 10.1371/journal.pbio.0030328.
Protocol 4: Amplification of RNA doi10.1101/molclon.000182

Gene expression profiling typically requires microgram quantities of mRNA, which can be difficult to obtain. In such cases, RNA must be amplified in order to have enough material for microarray labeling and hybridization. Currently, the most popular choice for amplifying RNA is to use a commercial kit, such as MessageAmp II (Ambion), a product with which we have had good success. These kits are expensive, however, and thus this protocol provides an alternative RNA amplification procedure adapted from Baugh et al. (2001).

This protocol generates amplified antisense RNA (aRNA) from limited quantities of total RNA (see Fig. 1). It is designed around maximizing yield and product length while minimizing template-independent side reactions. Template-independent reactions compete with the desired template-dependent reaction, an undesirable situation that becomes more severe as less RNA template is used. Amplification products dominated by template-independent product result in greatly reduced sensitivity and compression of differences in microarray hybridization experiments. Most notably, the oligo(dT) primer used in reverse transcription (RT) yields a high-molecular-weight product in the in vitro transcription (IVT) reaction independent of any cDNA template (Baugh et al. 2001). This reaction occurs under all conditions tested; the protocol is therefore designed to limit the amount of primer used to start with. In addition, high-molecular-weight, template-independent product is generated in the presence of biotinylated NTPs and the absence of any polymer when excessive amounts of T7 RNA polymerase activity are used. Template-dependent product of questionable molecular weight and limited functionality in downstream reactions can also be produced with excessive T7 RNA polymerase activity. Essentially, more yield is not always better. The protocol limits the amount of primer used by employing small cDNA synthesis volumes.

The key consideration in any amplification protocol, as noted in Protocols 2 and 3 for DNA labeling, is preventing representation bias in the amplified material. As with DNA amplification, a valuable initial experiment for investigators who are new to RNA amplification is to compare unamplified RNA and amplified RNA by microarray hybridization. The readout should be designed to reveal which sequences are over- and underrepresented after amplification, and by how much.

At the end of the protocol, quantify the mass yield of amplified material. A single round of amplification typically results in a fivefold to 20-fold mass conversion of starting material. If the first-round aRNA is used as a template for a second round of amplification, 200- to 400-fold amplification is typical.


MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Recipes for reagents specific to this protocol, marked <R>, are provided at the end of the protocol. See Appendix 1 for recipes for commonly used stock solutions, buffers, and reagents, marked <A>. Dilute stock solutions to the appropriate concentrations.

Reagents

  • Carrier
    • Add either 5 g of linear polyacrylamide (LPA) or 20 g of glycogen. If a second round of amplification is to be used (or other downstream reverse transcription reactions), then LPA is recommended over glycogen. LPA will slow the Microcon washes (from 1214 min to 2832 min for Microcon 100s at 500g and room temperature), but it does not inhibit reverse transcriptase.
  • DEPC ddH2O
  • DEPC-treated TE (pH 8.0)
  • DNA ligase, from Escherichia coli
  • DNA polymerase I
  • dNTP (10 mM)
  • DTT (100 mM)
  • (dT)-T7 primer
    • The primer sequence is 5-GCATTAGCGGCCGCGAAATTAATACGACTCACTATAGGGAGA(T)21V-3 (where V stands for A, C, or G).
  • Ethanol (70 and 95)
  • First-strand buffer (5), provided with kit
  • High-yield T7 in vitro transcription kit (e.g., AmpliScribe from Epicentre; similar kits from Ambion, Promega, and others are available)
  • IVT buffer (10), provided with IVT kit
  • NaCl (5 M)
  • NTP mixture (100 mM)
    • The NTP mixture contains 100 mM each of ATP, CTP, GTP, and UTP.
  • Phenol:chloroform
  • Random primers
  • Reverse transcriptase (SuperScript II; Life Technologies)
  • RNase H, from E. coli
  • RNase inhibitor
  • Second-strand synthesis buffer (5) <R> (Lifetech or homemade)
  • T4gp32 (single-strand binding protein; 8 mg/mL)
  • T4 DNA polymerase
  • T7 RNA polymerase, high concentration (80 U/L)
  • Total RNA (100 ng dissolved in water or TE; the concentration does not matter as the RNA will be dried down in Step 1)

Equipment

  • Bio-Gel P-6 Micro-Spin Column (Bio-Rad)
  • Heat block set at 65C and 70C
  • Incubator set at 14C16C
  • Microcon 100 spin column (Millipore)
  • Phase Lock Gel Heavy tubes (0.5 mL) (Eppendorf)
  • Thermal cycler (with a heated lid) or air incubator set at 42C and 70C
  • Tubes (0.6 mL)
  • Rotary evaporator (e.g., SpeedVac)


METHOD
Reverse Transcription

  • 1. Combine 100 ng of the (dT)-T7 primer with 100 ng of total RNA in a 0.6-mL tube. Reduce the volume under vacuum to 5.0 L.
    • Do not allow the RNA to dry out completely.
  • 2. Prepare the RT premix and place it on ice:
  • 3. Denature the RNA/primer mix for 4 min at 70C in a thermal cycler with a heated lid.
  • 4. Snap-cool the mix on ice and keep it on ice.
    • The volume may drop following denaturation.
  • 5. Add 5.0 L of ice-cold RT premix to the RNA/primer tube, and mix by pipetting. The final volume should be 10 L.
    • If there was evaporative loss during denaturation (Step 3), then add dH2O to adjust the volume to 10.0 L. Before committing your RNA, run these initial steps using a control nucleic acid to determine the volume loss.
  • 6. Incubate the RT reaction for 1 h at 42C in either a thermal cycler with a heated lid or an air incubator, but not in a water bath to reduce the chance of contamination.
  • 7. Heat-inactivate the reaction for 15 min at 65C.
  • 8. Chill the tube on ice.

Second-Strand Synthesis

  • 9. Prepare second-strand synthesis (SSS) premix:
    Chill on ice.
  • 10. Add 65 L of ice-cold SSS premix to the RT reaction tube, and mix by pipetting. Incubate for 2 h at 14C16C.
  • 11. Add 2 U (10 U) of T4 DNA polymerase, and mix by flicking and gentle vortexing. Incubate for an additional 15 min at 14C16C.
  • 12. Heat-inactivate the reaction for 10 min at 70C. Go immediately from 15C to 70C without letting the tube sit at room temperature to avoid undesirable enzyme activities.
  • 13. Add 75 L of phenol:chloroform (1:1), and mix by pipetting vigorously. Transfer the mixture to prespun Phase Lock Gel Heavy 0.5-mL tubes, and centrifuge for 5 min at 13,000 rpm.
  • 14. Prepare a Bio-Gel P-6 Micro-Spin Column per the manufacturer's instructions.
  • 15. Transfer the aqueous phase from Step 13 to the prepared P-6 column and centrifuge at 1000g for 4 min, recovering the flowthrough (80 L) in a clean 1.5-mL tube.
    • The flowthrough can be kept in the 1.5-mL tube or transferred to a tube of a different size, depending on the details of the IVT reaction steps. For example, it may be desirable to run the reaction in a thermal cycler; thus, the products can be held at 4C rather than overincubate them.
  • 16. Add the appropriate carrier and 3.5 L of 5 M NaCl for precipitating the DNA, and mix by vortexing. Add 2.5 volumes of 95 ethanol (220 L) and mix well. Precipitate for at least 2 h at 20C.
  • 17. Centrifuge to pellet the DNA at 13,000 rpm for 20 min.
  • 18. Carefully remove the supernatant. Wash the pellet with 500 L of 70 ethanol, and centrifuge for 5 min at 13,000 rpm.
  • 19. Carefully remove the supernatant. Pulse-centrifuge (up to full speed) the tube to collect all residual ethanol at the bottom.
  • 20. Remove the remaining supernatant with a pipette. Allow the pellet to air-dry for 23 min.

In Vitro Transcription

  • 21. Prepare the IVT premix at room temperature to avoid forming a precipitate:
  • 22. Add 40 L of IVT premix to the DNA pellet (from Step 20). Resuspend the pellet in the premix by gently flicking and vortexing the tube. Incubate the reaction for 9 h at 42C.
    • If the starting amount of total DNA was in the microgram range, then IVT yields might improve by using a 60- or 80-L reaction volume.
  • 23. Proceed with the cleanup and additional amplification, or freeze the IVT reaction at 80C.

Cleanup

  • 24. Add 480 L of DEPC-treated TE to the IVT reaction tube.
  • 25. Transfer the 500 L to a Microcon 100, and centrifuge at 500g until the volume is <20 L (1115 min at room temperature without LPA, 2832 min with LPA).
    • Processing a sample through a Microcon spin column is slower with LPA in the sample, although the results are fine. Alternatively, RNeasy columns can be used for cleanup.
  • 26. Add another 500 L of DEPC-treated TE, and centrifuge as before. Repeat this step one more time (three washes total).
    • If you intend to proceed with a second amplification, the final wash should be with dH2O, and with the filtrate should be reduced to a small volume.
  • 27. Measure and, if necessary, adjust the volume for downstream applications.
    • It may be comforting or worthwhile to quantify the yield and analyze the products by electrophoresis. Alternatively, proceed to the second round of amplification.

Second Round of Amplification

  • 28. Add 0.5 g of random primers to the aRNA (from Step 26). Reduce the volume under vacuum to 5.0 L.
  • 29. Denature the RNA for 5 min at 70C in a thermal cycler with a heated lid.
  • 30. Snap-cool the mix on ice. Let the tube rest for 5 min at room temperature.
  • 31. Prepare the RT premix and place it on ice:
  • 32. Add 5 L of room-temperature RT premix to the RNA, and mix by pipetting.
    • The final volume should be 10.0 L. If there was evaporative loss during denaturation (Step 28), then add dH2O to adjust the volume to 10.0 L.
  • 33. Incubate the reaction tube in a thermal cycler with a heated lid, programmed as follows:
    • i. 20 min at 37C.
    • ii. 20 min at 42C.
    • iii. 10 min at 50C.
    • iv. 10 min at 55C.
    • v. 15 min at 65C.
    • vi. Hold at 37C.
  • 34. Add 1 U of RNase H, and mix by vortexing gently. Incubate for 30 min at 37C and then heat for 2 min at 95C.
  • 35. Chill the tube on ice, and then centrifuge it briefly to collect the condensation. Place the tube on ice.
  • 36. Add 1 L of 100 ng/L (dT)-T7 primer while the tube is on ice. Incubate the tube for 10 min at 42C to anneal the primer.
  • 37. Prepare the SSS premix (minus ligase):
  • 38. Snap-cool the sample (from Step 36) on ice.
  • 39. Add 65 L of ice-cold SSS premix to the chilled reaction tube. Incubate for 2 h at 14C16C.
  • 40. Add 10 U of T4 DNA polymerase, and mix by gentle flicking and vortexing. Incubate for an additional 15 min at 14C16C.
  • 41. Heat-inactivate the reaction for 10 min at 70C.
  • 42. Perform phenol:chloroform extraction, Bio-Gel P-6 chromatography, and nucleic acid precipitation as per Steps 1320.
  • 43. Prepare the IVT premix as in Step 21.
  • 44. Add 40 L of IVT premix to the DNA pellet (from Step 42). Resuspend the pellet in the premix by gently flicking and vortexing the tube. Incubate the reaction for 9 h at 42C.
  • 45. Proceed with the cleanup as in Steps 2427, or freeze the IVT reaction at 80C.


DISCUSSION

The output of this protocol will be amplified RNA suitable for direct or indirect labeling for microarray hybridization (Protocols 5 or 6). As with DNA amplification, always quantify the mass yield of amplified material, and include control amplifications without input RNA to ensure that there is no contamination of any of your reagents.

First time users of this protocol will find it helpful to analyze amplified product on denaturing or native agarose gels (see Fig. 1 in Baugh et al. 2001). Identification of high-molecular-weight product in the No Template control likely indicates excess primer, and this can be minimized by decreasing primer concentration in the initial in vitro transcription reactions.

Another useful first-time control is to amplify RNA from an abundant RNA source and to competitively hybridize amplified RNA against the original RNA pool, ideally using a microarray containing probes at the 5 and 3 ends of genes. If the protocol is working well, aRNA and original RNA samples should be highly correlated, and 5/3 ratios should be close to 1. Lower 5/3 ratios indicate poorly processive in vitro transcription, which can be corrected by increasing input RNA mass to the protocol. Alternatively, lower 5/3 ratios may indicate failure to include the single-strand DNA binding protein T4gp32 in the in vitro transcription reaction.


RECIPE

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Second-Strand Synthesis Buffer (5)
REFERENCE
Baugh LR, Hill AA, Brown EL, Hunter CP. . Year: 2001. Quantitative analysis of mRNA amplification by in vitro transcription. Nucleic Acids Res29: e29. doi: 10.1093/nar/29.5.e29.
Protocol 5: Direct Cyanine-dUTP Labeling of RNA doi10.1101/molclon.000183

This is the simplest method to label RNA for use in expression analysis. RNA is reverse-transcribed using both oligo(dT) and random hexamers as primers. The random hexamers improve overall efficiency of labeling, especially at the 5 end of the RNA. Fluorescently labeled dUTP is incorporated into the cDNA. After reverse transcription, the RNA is degraded, and the labeled cDNA is purified from unincorporated Cy dyes. Finally, samples labeled with Cy3 and Cy5 dyes are mixed and combined with blocking nucleotides and used for hybridization, as described in Protocol 10.


MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Recipes for reagents specific to this protocol, marked <R>, are provided at the end of the protocol. See Appendix 1 for recipes for commonly used stock solutions, buffers, and reagents, marked <A>. Dilute stock solutions to the appropriate concentrations.

Reagents

  • Cy3-dUTP or Cy5-dUTP (GE Healthcare Life Sciences)
  • DEPC-treated H2O
  • HCl (0.1 N)
  • NaOH (0.1 N)
  • Oligo(dT) (2 g/L)
  • Random hexamers (4 g/L)
    • N6 random hexamer can be ordered from any oligonucleotide company and made up at 5 mg/mL in RNase-free TE or water.
  • Reverse transcriptase, 200 U/L (SuperScript II; Life Technologies)
    • First strand buffer <R> and DTT, both required for preparation of the master reagent mix in Step 3, are provided with SuperScript II.
  • Total RNA (from Protocol 3 or 4)
  • Unlabeled dNTPs (low dTTP) stock <R>

Equipment

  • Heat block set at 65C
  • MinElute kit (QIAGEN, catalog no. 28004)
  • Thermal cycler


METHOD
RT Reaction

  • 1. Prepare the following RNA/oligo reaction mixtures in separate microcentrifuge tubes.
  • 2. Heat the tubes for 10 min to 65C, and cool on ice to anneal the primers to the RNA.
  • 3. While the RNA/oligo reaction mixtures are incubating, prepare two master reaction mixtures, one containing Cy3-dUTP and one with Cy5-dUTP, in a volume sufficient to transfer 14.6 L of each to an RNA/oligo reaction mixture.
  • 4. To the Cy3 RNA/oligo reaction mixture, add 14.6 L of the Cy3 master reaction mixture (Step 3) to each tube. To the Cy5 RNA/oligo reaction mixture, add 14.6 L of the Cy5 master reaction mixture (Step 3) to each tube. Incubate for 1 h at 42C.
  • 5. Add 1 L of SuperScript II enzyme (200 U/L) to each reaction and thoroughly mix the reaction components with a pipette. Incubate for an additional 1 h.
  • 6. Degrade the RNA by adding 15 L of 0.1 N NaOH to each reaction and incubating for 10 min at 65C70C.
  • 7. Neutralize each reaction by adding 15 L of 0.1 N HCl.

Cleanup

If you want to determine the amount of fluorophore incorporated, then clean up the Cy5- and Cy3-labeled cDNA samples on separate MinElute columns. However, if thin coverslips and a small probe volume will be used during the hybridization (Protocol 10), it may be necessary to purify the Cy5- and Cy3-labeled cDNA samples together or use vacuum centrifugation to reduce the volume after elution.

  • 8. Add 600 L of Buffer PB (binding buffer) to each sample.
  • 9. Assemble a MinElute column on a 2-mL collection tube.
  • 10. Add the entire 660 L or 720 L (volume depends on whether each color sample is being cleaned up separately or together) to a MinElute column. Centrifuge the column for 1 min at 10,000g. Discard the flowthrough, and reuse the 2-mL tube.
  • 11. Add 750 L of Wash buffer PE to the column. Centrifuge for 1 min at 10,000g. Discard the flowthrough, and reuse the 2-mL tube.
  • 12. Centrifuge again at maximum speed for 1 min to remove residual ethanol.
  • 13. Place the column in a fresh 1.5-mL tube. Add 10 L of H2O to elute. Allow the Elution buffer to stand for at least 2 min.
  • 14. Centrifuge at maximum speed for 1 min. Add 10 L of H2O to elute. Allow the Elution buffer to stand for at least 2 min before spinning.
  • 15. Centrifuge at maximum speed for 1 min.
  • 16. Measure the volume of the eluate for each sample. The volume should be 18 L for each column.


DISCUSSION

The output of this protocol will be Cy5-labeled cDNA and Cy3-labeled cDNA. The DNA can be used immediately to hybridize to a microarray, or it can be stored in foil, to prevent bleaching, at 4C for up to a week. A useful spot check for labeling success is the color of the labeled material after unincorporated nucleotides have been removed (see Step 16). A good labeling will result in blue Cy5-DNA and red Cy3-DNA.


RECIPES

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

First-Strand Buffer
Unlabeled dNTPs (Low dTTP) Stock
Protocol 6: Indirect Aminoallyl-dUTP Labeling of RNA doi10.1101/molclon.000184

This protocol is slightly longer than the simpler direct-labeling protocol, but it is significantly cheaper because of the high cost of Cy-dNTPs used in Protocol 5. This labeling procedure is called indirect because the fluorescent moiety is not incorporated during the reverse transcription reaction. Instead, a reactive nucleotide analog (aminoallyl-dUTP) is incorporated during reverse transcription, the cDNA is isolated, and then the cyanine dyes are incorporated by binding with the aminoallyl group to produce the desired fluorescent cDNA.


MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Recipes for reagents specific to this protocol, marked <R>, are provided at the end of the protocol. See Appendix 1 for recipes for commonly used stock solutions, buffers, and reagents, marked <A>. Dilute stock solutions to the appropriate concentrations.

Reagents

  • aa-dUTP mixture (50) <R>
  • Anchored oligo(dT) (Life Technologies)
    • Anchored oligo(dT) primer is a mix of 12 primers, each having a sequence of 20 dT residues followed by two nucleotides VN, where V is dA, dC, or dG and N is dA, dC, dG, or dT. Thus, the VN anchor restricts annealing of the primer to the 5 end of the poly(A) tail of mRNA.
  • Cyanine dyes (typically Cy5 and Cy3) (GE Healthcare Life Sciences)
    • Dye comes dried and should be resuspended in 11 L of DMSO. This amount of material is sufficient for three labeling reactions. If the entire amount will not be used, prepare 3-L aliquots, dry down in a rotary vacuum, and store them at 20C.
  • DTT (0.1 M)
  • EDTA (0.5 M)
  • HEPES (1 M, pH 7.5)
  • NaHCO3 (50 mM, pH 9.0)
  • NaOH (1 N)
  • Reverse transcriptase (SuperScript II; Life Technologies)
  • Reverse transcriptase buffer (5) (Life Technologies)
  • RNasin (ribonuclease inhibitor)
  • Total RNA (from Protocol 3 or 4)

Equipment

  • Heat block set at 67C, 70C, and 95C
  • Lightproof box
  • MinElute kit (QIAGEN, catalog no. 28204)
  • Thermal cycler (42C)
  • Zymo column (Zymo Research, catalog no. D3024)


METHOD
Reverse Transcription to Make aa-dUTP-Labeled cDNAs

  • 1. Combine 30 g of total RNA and water to a final volume of 14.5 L. Add 1 L of 5 g/L anchored oligo(dT). Mix by pipetting. Heat for 10 min at 70C.
  • 2. Cool the tube on ice for 10 min.
  • 3. Pulse-centrifuge to bring the condensate to the bottom of the tube.
  • 4. Prepare the Master mix just before use (be sure to add the enzymes last):
  • 5. Add 14.5 L of Master mix to the tube of RNA. Pipette to mix. Incubate the reaction for 2 h at 42C.
  • 6. Incubate the tube for 5 min at 95C. Transfer immediately to ice.
  • 7. Add 13 L of 1 N NaOH and 1 L of 0.5 M EDTA to hydrolyze the RNA. Mix the reagents and pulse-centrifuge. Incubate the tube for 15 min at 67C.
  • 8. Neutralize the reaction with 50 L of 1 M HEPES (pH 7.5). Vortex the tube and pulse-centrifuge.
  • 9. Purify the reaction over a Zymo column to remove unincorporated nucleotides.
    • i. Add 1 mL of Binding buffer to the reaction. Mix by pipetting. Load half of the material onto the column. Centrifuge for 10 sec at maximum speed. Discard the flowthrough.
    • ii. Add the remainder of the reaction. Centrifuge for 10 sec at maximum speed. Discard the flowthrough.
    • iii. Add 200 L of Wash buffer to the column. Centrifuge for 30 sec at maximum speed.
      • Typically, multiple individual samples will be labeled simultaneously, with at least two reactions for a given two-color microarray.
    • iv. Add another 200 L of Wash buffer to the column. Centrifuge for 1 min at maximum speed. Discard the flowthrough, and centrifuge for 1 min at maximum speed.
    • v. Add 10 L of 50 mM NaHCO3 (pH 9.0) to the filter. Incubate for 5 min at room temperature. Centrifuge for 30 sec at maximum speed to elute the cDNA.
  • 10. Couple cyanine dyes to the aa-dUTP incorporated cDNAs by adding the eluted material directly to 3 L of Cy5 or Cy3 dye resuspended in DMSO. Incubate for 1 h to overnight at room temperature in a lightproof box or drawer.

Purification of Labeled cDNA

  • 11. Purify the cDNA using a MinElute kit, following the manufacturer's instructions.


DISCUSSION

The products of this protocol will be Cy5-labeled and Cy3-labeled cDNA, which are ready to hybridize to microarrays. This material can be stored in foil, to prevent bleaching, at 4C for up to 1 wk. A useful check for labeling success is the color of the labeled cDNA after unincorporated nucleotides have been removed: Good labeling will result in visible blue (Cy5) or red (Cy3) color in the cleaned up material.


RECIPE

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

aa-dUTP Mixture (50)
Protocol 7: Cyanine-dCTP Labeling of DNA Using Klenow doi10.1101/molclon.000185

Similar to the direct RNA labeling protocol (Protocol 5), direct DNA labeling with Cy-dCTP is the simplest and fastest method for labeling DNA. This is a standard Klenow labeling protocol in which Cy-dCTP is incorporated during the labeling reaction. After stopping the reaction, labeled nucleotide is separated from unreacted Cy-dCTP, and Cy3- and Cy5-labeled materials are combined for hybridization (Protocol 10). This protocol is suitable for many applications, including detection of copy number variation, nucleosome mapping, and other location analysis (e.g., ChIP-chip).


MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Recipes for reagents specific to this protocol, marked <R>, are provided at the end of the protocol. See Appendix 1 for recipes for commonly used stock solutions, buffers, and reagents, marked <A>. Dilute stock solutions to the appropriate concentrations.

Reagents

  • Cy3-dCTP and Cy5-dCTP (GE Life Sciences)
  • dNTP mixture (10) <R>
  • EDTA (0.5 M, pH 8.0)
  • Genomic DNA (or from Protocol 2)
  • Human Cot-1 DNA (1 g/L) (GIBCO)
  • Klenow fragment of E. coli DNA polymerase I (4050 units/L)
  • Poly(dA-dT) (Sigma-Aldrich)
    • Make a 5 g/L stock in nuclease-free water. Store the stock at 20C.
  • Random primer/buffer solution <R>
    • This can be obtained from Life Technologies BioPrime labeling kit or can be made in the laboratory.
  • TE (pH 7.4)
  • Yeast tRNA (GIBCO)
    • Make a 5 g/L stock in nuclease-free water.

Equipment

  • Heat block set at 37C and 95C100C
  • Microcon 30 spin column (Millipore)


METHOD

  • 1. Add 23 g of DNA to a microcentrifuge tube.
    • For high-complexity DNA, such as mammalian genomic DNA, reducing the fragment size by sonication or restriction digestion improves labeling efficiency. Shearing by sonication to 1000-bp average fragment size is sufficient for this application.
  • 2. Add H2O to the DNA to a final volume of 21 L. Add 20 L of 2.5 random primer/buffer mixture. Boil the tube for 5 min in a heat block set to 95C100C.
  • 3. Place the tubes on ice for 5 min.
  • 4. Add reagents in the following order:
    • Cy-UTP also works well, but if using UTP, adjust the 10 dNTP mix accordingly.
  • 5. Incubate the reaction for 12 h at 37C.
    • For improved labeling efficiency, add 1 L of Klenow after the reaction has been going for 1 h. Incubate the reaction for another hour.
  • 6. Stop the reaction by adding 5 L of 0.5 M EDTA (pH 8.0).

Cleanup of Labeled DNA

  • 7. Combine the Cy3- and Cy5-labeled samples, and add 450 L of TE (pH 7.4).
  • 8. Add the combined samples to a Microcon 30 spin column. Centrifuge the spin column for 1011 min at 10,000g in a microcentrifuge.
  • 9. Discard the flowthrough.
  • 10. If hybridizing to homemade microarrays, add blocking nucleotides at this step:
    • If using a commercial microarray, refer to the manufacturer's protocols for specific hybridization buffers and blocking nucleotides.
  • 11. Add another 450 L of TE, and centrifuge as in Step 8.
  • 12. Measure the volume of the liquid remaining in the column. If necessary, centrifuge for 1 min. The desired probe volume is 20 L. The exact volume will depend on the size of the coverslip that contains the microarray. For small microarrays this volume may be as low as 12 L (see Protocol 10, Step 3).
  • 13. Discard the flowthrough. Recover the labeled DNA from the filter by inverting the column onto a fresh collection tube. Centrifuge for 1 min at 10,000g.
  • 14. Proceed to hybridization with the microarray (Protocols 9 and 10).


DISCUSSION

The products of this protocol will be Cy5-labeled and Cy3-labeled cDNA, which are ready to hybridize to microarrays. This material can be stored in foil, to prevent bleaching, at 4C for up to 1 wk. A useful check for labeling success is the color of the labeled cDNA after unincorporated nucleotides have been removed: Good labeling will result in visible blue (Cy5) or red (Cy3) color in the cleaned up material.


RECIPES

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

dNTP Mixture (10)
Random Primer/Buffer Solution
Protocol 8: Indirect Labeling of DNA doi10.1101/molclon.000186

As with RNA labeling protocols, the main difference between direct and indirect DNA labeling protocols is a trade-off of cost and time. Indirect labeling of DNA takes 2 h longer than direct labeling but is hundreds of dollars cheaper.


MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Recipes for reagents specific to this protocol, marked <R>, are provided at the end of the protocol. See Appendix 1 for recipes for commonly used stock solutions, buffers, and reagents, marked <A>. Dilute stock solutions to the appropriate concentrations.

Reagents

  • aa-dUTP/dNTP mixture (3 mM)
    • Combine 6 L of 50 stock <R> used for cDNA synthesis and 44 L of dH2O.
  • Cyanine dyes (typically Cy5 and Cy3) (GE Healthcare Life Sciences)
    • Dyes come dried and should be resuspended in 11 L of DMSO. This amount of material is sufficient for three labeling reactions. If the entire amount will not be used, prepare 3-L aliquots, dry down in a rotary vacuum, and store them at 20C.
  • EDTA (0.5 M, pH 8.0)
  • Genomic DNA
    • Prepare genomic DNA according to your favorite protocol. Because the DNA should be fairly pure, avoid using quick and dirty methods. Commercial gDNA isolation kits are suitable, as in Chapter 1, Protocols 12 and 13. As a rule of thumb, the OD260/280 ratio should be at least 1.8.
  • Klenow buffer
  • Klenow fragment of DNA polymerase I
  • NaHCO3 (50 mM, pH 9.0)
  • Random hexamer
    • N6 random hexamer can be ordered from any oligonucleotide company, made up at 5 g/L in RNase-free TE/H2O.

Equipment

  • Heat block set at 37C and 100C
  • Lightproof box (see Step 11)
  • MinElute kit (QIAGEN)
  • Zymo columns (Zymo Research, catalog no. D3024)


METHOD
Genomic DNA Labeling

  • 1. For each array, combine 4 g of genomic DNA, 10 g of random hexamer (N6), and dH2O to a final volume of 42 L. Incubate for 5 min at 100C.
  • 2. Cool the tube(s) quickly on ice for 10 min.
  • 3. Pulse-centrifuge to bring the condensate to the bottom of the tube.
  • 4. On ice, add:
    Pipette to mix. Incubate for 2 h at 37C.
  • 5. Add 5 L of 0.5 M EDTA (pH 8.0) to stop the reaction.

Purification of Random Prime-Labeled DNA

  • 6. Add 1 mL of Binding buffer to the reaction. Mix well, and load half (527 L) of the reaction mixture onto each of two Zymo columns. Centrifuge the columns for 10 sec at maximum speed.
    • The large volume of Binding buffer is necessary to precipitate single-stranded DNA.
  • 7. Discard the flowthrough. Wash the column with 200 L of Wash buffer. Centrifuge for 1 min at maximum speed.
  • 8. Repeat Step 7.
  • 9. Add 10 L of 50 mM NaHCO3 (pH 9.0) to each column filter. Incubate for 5 min at room temperature.
  • 10. Elute the DNA from the column by centrifuging for 1 min at maximum speed.
  • 11. Couple cyanine dyes to the aa-dUTP-incorporated cDNAs by adding the eluted material directly to 3 L of Cy5 dye or Cy3 dye resuspended in DMSO. Incubate for 1 h to overnight at room temperature in a lightproof box or a drawer.

Purification of Labeled Genomic DNA

  • 12. Purify the cDNA using a MinElute kit, following the manufacturer's instructions.


RECIPE

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

aa-dUTP/dNTP Mixture (50)

Protocol 9: Blocking Polylysines on Homemade Microarrays doi10.1101/molclon.000187

Homemade microarrays are printed on polylysine-coated slides. The lysines form a positively charged surface that can bind nonspecifically to the acidic nucleic acids during hybridization, resulting in significant background fluorescence. Thus, a key step in microarray processing is blocking all of the surface lysines not associated with the oligonucleotides in the microarray spots. The -amino group of lysine is succinylated by reacting with succinic anhydride (Fig. 1). Because anhydrides readily hydrolyze in water, use only fresh reagents that have not had the opportunity to absorb much water.

The procedure is straightforward. Microarrays are, if necessary, rehydrated; excess liquid is removed by drying at a moderate temperature; and the succinylation reaction is performed. After the reaction is complete, the slides are washed and dried with ethanol, at which point they are ready for hybridization or they can be stored in a desiccator. Rehydration is necessary if the microarrays were desiccated after printing (see Protocol 1). When microarrays are stored in a desiccator, the spots typically dry out to form rings. Rehydration is important to restore the full spots.


MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Recipes for reagents specific to this protocol, marked <R>, are provided at the end of the protocol. See Appendix 1 for recipes for commonly used stock solutions, buffers, and reagents, marked <A>. Dilute stock solutions to the appropriate concentrations.

Reagents

  • Ethanol (95) (see Step 11)
  • 1-Methyl-2-pyrrolidinone
    • Use only HPLC grade. If it has become slightly yellowed, then it is no longer usable, having absorbed excess water.
  • Microarrays printed onto glass slides (either homemade [Protocol 1] or purchased)
  • Sodium borate (1 M, pH 8.0)
  • SSC (0.5) <A>
  • Succinic anhydride
    • The stock bottle of solid succinic anhydride should be stored under desiccation and vacuum (or under nitrogen).
      Do not use if exposed to moisture!

Equipment

  • Beakers (500 mL and 4 L)
  • Centrifuge, fitted with microtiter plate carrier
  • Glass-etching pen
  • Glass slide racks and wash chambers (e.g., Thermo Scientific Fisher, catalog no. NC9516192)
  • Gloves, powder-free
  • Heat block set at 80C (see Step 5)
  • Heating plate set at 80C
  • Humid chamber, for standard-size glass slides (e.g., Sigma-Aldrich, catalog no. H6644)
  • Microarray slide box, plastic
  • Orbital shaker
  • Stir bar that fits a 500-mL beaker
  • Water bath set at 80C


METHOD

  • 1. Select 15 slides for postprocessing. Handle all slides with powder-free gloves. Determine the correct orientation of each slide. With a glass-etching pen, lightly mark the boundaries of the array on the backside of the slide.
    • Marking the array boundary is important because after processing, the arrays will not be visible.
  • 2. Add enough distilled water to a large 4-L beaker so that a slide rack will be completely submerged when placed inside. Place the beaker on a heating plate and heat to 80C.

Rehydrating the Microarrays

  • 3. Fill the bottom of a humidifying chamber with 0.5 SSC. Suspend the arrays face-up over the 0.5 SSC, and cover the chamber with a lid.
  • 4. Rehydrate until all of the microarray spots glisten (usually 15 min at room temperature).
    • Allow the spots to swell slightly, but do not let them run into each other. Insufficient hydration results in less DNA bound within a spot, and too much hydration will cause spots to run together.
  • 5. Dry each array by placing each slide, with the DNA side facing up, for 3 sec on an inverted heat block set at 80C.
  • 6. Place the arrays in a slide rack. Place the slide rack into an empty slide chamber that is sitting on an orbital shaker.

Blocking Free Lysines on the Microarray Slides

  • 7. Prepare the blocking solution by measuring 335 mL of 1-methyl-2-pyrrolidinone into a clean, dry 500-mL beaker. Dissolve 5.5 g of succinic anhydride using a stir bar.
  • 8. Immediately after the succinic anhydride dissolves, mix in 15 mL of 1 M sodium borate (pH 8.0). Quickly pour the buffered blocking solution into a clean, dry glass slide dish.
  • 9. Plunge the slides rapidly into the blocking solution, and vigorously shake the slide rack manually, keeping the slides submerged at all times. After 30 sec, place a lid on the glass box, and shake gently on the orbital shaker for 15 min.
  • 10. Drain excess blocking solution from the slides for 5 sec. Submerge the slide rack into an 80C water bath. Gently swish the rack back and forth under the water for a few seconds. Incubate for 60 sec.
  • 11. Quickly transfer the rack to a glass dish of 95 ethanol, and plunge to mix.
    • Make sure that the ethanol is crystal clear. Do not use if it contains particulates or is cloudy.
  • 12. Take the entire glass dish with slide rack still submerged to a bench-top centrifuge. (Be sure to have an equivalently loaded slide rack ready to serve as a balance in the centrifuge.) Drain excess ethanol off the slides for 5 sec. Quickly, place the slide rack onto a microtiter plate carrier, and centrifuge in a bench-top centrifuge for 3 min at 550 rpm.
  • 13. After centrifugation, the slides should be clean and dry. Remove the slides from the rack and store them in a plastic microscope slide box. Arrays may be used immediately, or can be stored for at least 34 mo in a desiccator at room temperature.

REFERENCE
Simpson RJ. . Year: 2003. Peptide mapping and sequence analysis of gel-resolved proteins. In Proteins and proteomics, pp. 343424. . Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
Protocol 10: Hybridization to Homemade Microarrays doi10.1101/molclon.000188

Competitive hybridization of labeled probes to a microarray is conceptually similar to other hybridization methods, such as Southern blotting. For massively multiplexed microarrays, the adoption of two-color hybridization schemes has been a significant advance. The use of two colorstypically Cy3- and Cy5-labeled nucleic acidsmakes it possible to control for factors that affect hybridization intensity, including the number of labeled nucleotides and the Tm of each oligonucleotide. Thus, the difference in intensity among spots on a microarray can be quantified and analyzed to assess biological phenomena, like changes in gene expression or details of transcript structure.

Hybridization is conceptually straightforwardcold (nonfluorescent) blocking nucleotide is added to the mixed probe material, Hybridization buffer is added, and the mixture is applied to the microarray surface. Hybridization occurs overnight, after which the microarray is washed and scanned. The only technically challenging aspects of this protocol are application of the probe solution to the microarray surface and placement of the coverslip (Steps 8 and 9 below). Practice these steps, applying Hybridization buffer containing salmon sperm DNA, before using your fluorescently labeled DNA in an actual experiment. Develop the ability to work rapidly, and avoid introducing bubbles or scratching the array surface.

Many hybridization chambers are available commercially; these typically consist of a two-part metal chamber housing an internal cavity large enough to hold a microarray slide. Often there are small depressions in the internal cavity, which can hold extra buffer that will humidify the chamber. There is also a rubber gasket that forms a watertight seal around the chamber. Other hybridization systems include the Maui mixer, which moves hybridization solution back and forth over the array. For the user of homemade microarrays, however, these more expensive hybridization systems are an unnecessary expense.


MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution's Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Recipes for reagents specific to this protocol, marked <R>, are provided at the end of the protocol. See Appendix 1 for recipes for commonly used stock solutions, buffers, and reagents, marked <A>. Dilute stock solutions to the appropriate concentrations.

Reagents

  • Cot-1 DNA (GIBCO)
    • Cot-1 DNA is available commercially at 1 g/L. Before hybridization, concentrate Cot-1 DNA from 1 g/L to 10 g/L in a rotary vacuum device.
  • Cy5- and Cy3-labeled nucleic acids (from Protocol 5, 6, 7, or 8)
  • HEPES (1 M, pH 7.0)
  • Microarrays printed onto glass slides (either homemade [Protocol 1] or purchased)
    • If homemade, slides must be blocked as per Protocol 9 before hybridization.
  • Poly(A) RNA (10 g/L) (Sigma-Aldrich, catalog no. P9403)
  • SDS (10)
  • SSC (20) <A>
  • Yeast tRNA (10 g/L) (GIBCO, catalog no. 15401-011)

Equipment

  • Coverslips
    • Use either 2260 regular thin coverslips or 2260 Erie M-series lifter slips.
  • Dishes for washing microarrays (see Step 13)
  • Gloves, powder-free
  • Heat block set at 95C100C
  • Hybridization chambers
  • Lightproof box
  • Microarray scanner
    • When using homemade microarrays, the standard scanner is the GenePix 4000B (Molecular Devices).
  • Microarray slide box
  • Paper towels
  • Water bath set at 65C


METHOD
Preparation of the Hybridization Solution

If you have already combined the Cy dyes and added blocking material and buffer, skip to Step 4.

  • 1. Combine the following blocking nucleic acids:
  • 2. Combine Cy5- and Cy3-labeled nucleic acids.
  • 3. Prepare the complete hybridization solution as shown in the table below. To avoid introducing bubbles into the solution, do not vortex after adding SDS.
  • 4. Denature the probe by heating the hybridization solution in a water-filled heat block for 2 min at 95C100C.
  • 5. Let the probe sit for 10 min in the dark at room temperature.
  • 6. While the probe is sitting at room temperature, set up the necessary number of hybridization chambers. Open each chamber on a clean, flat surface, and place a microarray slide into a chamber with the array side facing up.
  • 7. Centrifuge the solution at 14,000 rpm for 5 min at room temperature.

Application of the Hybridization Solution to the Microarrays

  • 8. Carefully pipette the probe solution as a single drop onto one end of the microarray surface. Avoid creating any bubbles during pipetting, and be careful not to touch the microarray surface with the pipette tip (Fig. 1).
    • Leave 2 L of probe behind in the tube. If the probe precipitates upon application to the slide, see Troubleshooting.
  • 9. Carefully apply a coverslip by placing one edge of the coverslip on the slide near the probe and slowly lowering the other edge, using another coverslip as a lever and wedge to lower it (Fig. 1).
  • 10. Close the chamber, and immediately submerge it in a 65C water bath.
    • Be careful not to tilt the chamber. Metal tongs may be used to place the hybridization chamber into the water bath.
  • 11. If hybridizing multiple arrays, repeat Steps 810.
    • Be quick and efficient because the probes should not sit at room temperature for widely varying times. Try to have all the probes onto slides within 1015 min.
  • 12. Incubate the chambers for 1620 h at 65C.

Washing the Microarrays and Setting Up the Scanner

  • 13. Prepare dishes with the following solutions:
  • 14. If there is one, turn on the ozone scrubber connected to the scanner at least 15 min before scanning the arrays.
    • This is particularly useful in summer months when ozone levels are high. Bleaching of Cy5 dye becomes a significant problem in urban environments in the summer. If you choose not to use an ozone scrubber, consider performing hybridizations on cool or rainy days.
  • 15. Turn on the scanner, letting it warm up for at least 15 min to allow the laser outputs to stabilize.
  • 16. Launch the GenePix Pro software, and let it connect to the scanner and the network hardware key.
  • 17. Remove a hybridization chamber from the water bath. Working quickly, dry the chamber with a paper towel, open the chamber, remove the slide, and using your gloved hand, tap the slide against the bottom of the Wash 1A dish until the coverslip gently slides off the slide.
  • 18. Transfer the slide to a presubmerged rack in Wash 1B. Keep the slide in Wash 1B until all arrays have been transferred.
  • 19. Quickly transfer the rack from Wash 1B to Wash 2 (tilting the rack back and forth a couple of times to remove excess wash solution). Plunge the rack up and down several times in Wash 2. Incubate for 5 min with occasional plunging up and down.
  • 20. Quickly transfer the rack from Wash 2 to Wash 3 (tilting the rack back and forth a couple of times to remove excess wash solution). Plunge the rack up and down several times in Wash 3. Incubate for 5 min with occasional plunging up and down.
  • 21. As quickly as possible, transfer the rack of slides from Wash 3 to a tabletop centrifuge, with paper towels under the rack, and centrifuge at 500600 rpm for 5 min.
  • 22. Place the microarrays in a lightproof box, and begin scanning immediately.
    • Typically, four to five arrays are washed at a time. If more than five were hybridized, then the next set of slides should be washed as the last array from the first batch is being scanned.

Scanning the Microarrays

  • 23. Open the scanner door, and insert a slide with the array features facing down and the barcode/label closest to the edge of the scanner.
  • 24. Run a preview scan using the double-arrow button on top of the right-hand toolbar to visualize the array.
    • This performs a low-resolution (40-m) scan so that the location of array features can be visualized.
  • 25. After the preview scan is completed, select the Scan Area button in the left-hand Tools group. Use the mouse to draw a rectangle around the region containing features.
    • This limits the data scan to just this area, which reduces the time needed for each scan.
  • 26. Click on the Hardware Settings button on the right-hand side of the window to bring up the Settings box.
  • 27. Use the Auto Scale Brightness/Contrast button on the left-hand side to adjust the brightness and contrast of the image (both colors will be affected). Switch between the red and green channels by clicking on the respective radio buttons on the top left-hand side of the image tab.
  • 28. Set the Pixel Resolution to 10 m, then start the data scan. Zoom into the top half of the scanned image, and by eyeballing the image, adjust the PMT gain settings in both the red and green channels so that the average signal intensities balance out.
    • This takes some practice, although Step 29 describes how to do this with the aid of the intensity histograms. By eye, the goal is to balance the array so that the color yellow dominates, with roughly equal numbers of green and red spots. An array with mostly green spots will need the red channel PMT increased, and vice versa.
    • Normally, the PMT gain for the green channel (532 nm) is around 100 less than the red channel (635 nm). Adjust the settings so that any landing lights (bright spots printed near the top of the array corresponding to highly expressed genes, etc.) and some positive control spots on the array are saturated, but not many of the other features. Saturated pixels are drawn as white on the image.
    • Overall, the goal is to minimize the length of time spent scanning the array, and because some color imbalance can be normalized out later, it is more important to be fast than to be absolutely precise with regard to color balance.
  • 29. Alternatively (to the previous step), view the Histogram tab to check on the red/green ratio. Set the Min and Max Intensity fields in the Image Balance area on the left-hand side to 500 and 65,530, respectively. The goal is for the Count Ratio field to be 1.0, although this is not as important as having the two histogram curves as close to overlapping as possible.
  • 30. Once the PMT Gains for both channels are balanced, stop the scan by hitting the red Stop Scan button on the right-hand side of the window. Change the pixel resolution in the Hardware Settings window to 5 m, and ensure that the Lines to Average field is at 1. (More will not be detrimental to the scanned image, but the scanning will take much longer.)
  • 31. Start the scan again with the new settings. Record the PMT gains for each array in an Excel spreadsheet, along with any special observations/notes for the slide (e.g., the red signal is very dim, etc.). Save the spreadsheet in the same folder as the image files.
  • 32. When the entire array is scanned, click the File button on the right-hand side of the window. Select Save Images, and from the Save As Type field, select Single-Image TIFF Files. Make sure that both wavelengths are selected. Name the arrays with the identifier printed on the label, then hit the Save button.
    • There should be two TIFF (.tif) files, one with the data from the green channel (532 nm) and one with the data from the red channel (635 nm).
  • 33. Scan the next array, repeating Steps 2332. Balancing the two PMTs must be done for every microarray, as different RNA or DNA samples label with different efficiency.
  • 34. After all of the slides have been scanned, quit the GenePix Pro program, and turn off the scanner.

Gridding the Microarrays

  • 35. Open the GenePix software, and open the image file in question.
  • 36. Open the .gal file, which describes what DNA is found in each spot. This file will either be provided by the microarray manufacturer or it can be generated using the ArrayList generator found in GenePix.
  • 37. Align each pattern block (corresponding to one print pin) over the image so that image spots are roughly aligned with circles in the .gal file.
  • 38. Find features for each block by pressing [F5]. GenePix will move .gal file circles to cover the closest spots and will also flag spots where the intensity is too low.
  • 39. For each block, scan the entire sector by eye, and manually flag spots that are scratched or appear to be fluorescent dust rather than DNA.
  • 40. After all blocks have been flagged, hit the Analyze button to extract the .gpr file.
  • 41. At this point, either save the .gpr output file or choose Flag features to automatically impose thresholds.
    • We typically use this to flag all spots with a signal-to-noise ratio in each channel (532, 635) of <3.
  • 42. Save the .gpr file.


TROUBLESHOOTING

Problem (Step 8): When adding the hybridization solution to the microarray slide, the probe precipitates.

Solution: With some home-printed microarrays, the labeled probe can precipitate when added to the slide at room temperature. This leads to a pin prick morphology in which the precipitated probe fills the depressions created during the printing process. To mitigate this phenomenon, place the hybridization chamber onto a 55C60C heat block before the addition of the hybridization solution, and add hybridization solution while the chamber sits on the heat block.

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