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Chattopadhyay A, editor. Serotonin Receptors in Neurobiology. Boca Raton (FL): CRC Press; 2007.

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Chapter 5Identification of Novel Transcriptional Regulators in the Nervous System

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Increasingly, it is becoming recognized that transcriptional modulation of genes by sequence variations, DNA methylation, and alterations in transcription factor expression can play a major role in the behavioral phenotype throughout its development. Hence, the understanding of the transcriptional mechanisms that regulate key candidate genes for mental illness has taken on new significance. In a step-by-step approach we describe the crucial experiments to identify and characterize mechanisms of transcription regulation of the 5-HT1A receptor gene, which has been implicated in major depression, anxiety disorders, and suicide. These steps include: identification of transcription start sites by RT-PCR, primer extension, 5′-RACE, and RNase protection assay; characterization of upstream promoter, enhancer, and repressor regions using 5′-deletion luciferase reporter constructs and transfection assays in cell lines and primary cultures; identification of protein-DNA interactions using EMSA, super-shift and CHIP assays; cloning of novel transcription factors using yeast one-hybrid screening and functional characterization using Gal4-DBD system and by siRNA approaches. Using these approaches in combination with bioinformatics searches it is possible to identify and characterize the basic transcriptional regulatory mechanisms of virtually any candidate gene of interest and to identify specific proximal DNA elements and transcriptional factors that mediate its regulation.

Transcriptional Regulators and Behavior

The roles of transcriptional regulation in determining behavioral phenotype are beginning to be defined by exploiting new knowledge of the DNA elements and transcription factors that control brain-specific gene expression. For example, the brain-derived neurotrophin (BDNF) promoter has long been known as a target for the transcription factor CREB, which mediates calcium- and cAMP-dependent induction [1]. More detailed characterization has identified at least seven different promoters in the BDNF gene and differential regulation for these promoters [2, 3], and a novel calcium-regulated transcription factor (CaRF) [4]. In addition, novel roles for DNA methylation and methyl binding proteins at specific BDNF promoter sites are associated with distinct behavioral (social defeat stress) or pharmacological (depolarization, cocaine) stimuli leading to BDNF-induced alterations in synaptic plasticity [5–9]. Similarly, enhanced maternal care of rat pups increases serotonergic activation of the NGFIA transcription factor, which binds to its DNA element in exon 17 GR promoter, leading to altered acetylation and methylation, increased glucocorticoid receptor expression, and correlating with a reduced stress response in adult rats [10]. Thus, early lifetime changes in transcription factor activity due to environment can lead to lifelong changes in promoter methylation status, gene expression, and consequent behavioral alterations. The challenge is to understand how these changes take place, and the first step is the identification of promoter/enhancer regions of behavior-modifying genes.

The Serotonin System

The serotonin system of the brain is a major determinant of mood and emotional status [11, 12], and altered regulation of this system has been strongly implicated in mental illness. Serotonin is synthesized in a small population of neurons from the raphe nuclei that express the enzyme tryptophan hydroxylase (TPH) [13]. Another important site of regulation of the 5-HT system is the 5-HT reuptake transporter (5-HTT), which eliminates 5-HT from the synapse and is the target of serotonin-specific reuptake inhibitor antidepressants [14]. The release of 5-HT triggers activation of a wide variety of postsynaptic receptors. The 5-HT1A receptor is one of the most abundant 5-HT receptors in the brain [12] and functions as the predominant somatodendritic autoreceptor to inhibit the firing activity of serotonin neurons [15]. Desensitization of presynaptic 5-HT1A receptors is thought to accelerate antidepressant action [16], whereas recent studies in animal models indicate a predominant role for postsynaptic 5-HT1A receptors in anxiety and antidepressant action [17, 18].

Because of the potential importance of 5-HT1A receptors in major depression and anxiety, we have focused on identifying the transcriptional regulators of the 5′-HT1A receptor gene [19]. The 5-HT1A gene has the important advantage of being an intronless gene [20, 21]; hence, the contribution of intron sequences to its regulation is not a complication. The protocols described focus mainly on the 5′-regulatory region of the 5-HT1A gene but can be applied to regulatory regions of other genes, including those identified in introns or 3′-flanking sequence.


The first step in characterizing a promoter of interest is to identify the transcription start site or sites (TSS). The use of different TSS is an important source of genetic diversity for differential tissue expression, with different promoters driving expression of RNA variants. Thus, it is important to identify the number and location of the TSS of a gene in order to identify promoters and transcriptional regulatory mechanisms of interest in a particular tissue.

General Strategies

Initially, a DNA sequence spanning the candidate promoter site can be examined through one of several Web-based programs that detect eukaryotic promoters [22, 23]. As programs are algorithms based on previously described promoter features, a negative result (i.e., the program does not recognize the sequence as a promoter) does not mean absence of a promoter, whereas a positive result does not guarantee its presence.

In order to experimentally verify a putative TSS identified by bioinformatics, several methods can be used including primer extension, 5′-RACE, and RNase protection assays. These methods require some knowledge of the candidate region; in particular, a stretch of the 5′ end of the coding sequence long enough to design a primer. One way to verify or narrow down the TSS region is by RT-PCR of isolated RNA using an antisense primer within the coding region and amplification using progressively 5′ upstream primers [20]. As shown in Figure 5.1, primers that are upstream of the TSS will fail to amplify, hence localizing the approximate TSS region. Once the TSS region has been localized to a short sequence, a primer extension experiment can be performed to pinpoint the exact start site [20]. In this protocol, a gene-specific labeled oligonucleotide primes reverse transcription of the 5′ end of the target mRNA; the length of the cDNA is measured by urea-polyacrylamide gel electrophoresis. The reverse transcription will terminate right at the TSS. The labeled product is run on a polyacrylamide gel to determine its precise size and therefore the TSS location.

FIGURE 5.1. Progressive 5′ RT-PCR strategy to identify the rat 5-HT1A receptor transcriptional initiation region.


Progressive 5′ RT-PCR strategy to identify the rat 5-HT1A receptor transcriptional initiation region. Shown above is the location on the rat 5-HT1A receptor 5 region of the predicted PCR products from the common 3′-primer adjacent to the (more...)


The method of choice for identification of the 5′-TSS is rapid amplification of cDNA 5′-ends (5′-RACE) [24, 25]. Following ligation of a linker primer to the 5′-end of the cDNA, PCR amplification of the 5′ end of a cDNA molecule is initiated between two nested gene-specific antisense primers and nested linker primers, followed by purification and DNA sequencing of the resulting products. The advantage of this procedure over primer extension is that the precise DNA sequences of the 5′ end of the RNA transcripts are determined, which can verify the predicted TSS or identify novel TSS that would not be predicted. A major caveat with 5′-RACE (and primer extension) is that the cDNA may not be full-length, particularly in CG-rich regions due to secondary structure that stalls the reverse transcriptase. Although commercial additives (e.g., DMSO, etc.) can reduce this problem, the best way to confirm TSS identification is by using RNase protection, which does not depend on reverse transcription (see the following).

We have performed 5′-RACE making use of BD Biosciences Marathon-Ready cDNA library [26] (Remes-Lenicov et al., unpublished). Alternately, tagged cDNA libraries can be homemade, depending on the characteristics of the target RNA. For example, to investigate a transcript that is expressed only at low levels in a subset of neurons, it might be better to isolate RNA from enriched tissue rather than use a commercial whole-brain cDNA library. In any event, when purchasing or constructing a library, it should be remembered that active TSS might change, depending on tissue or stage of development.

When designing the specific primer for the target gene, the user-designed primer may be located in a region that is spliced out in a particular transcript (false negative PCR). On the other hand, choosing a primer too far from the 5′ end (or with an unexpectedly long 5′-UTR) may also cause a false negative result. The latter problem may be resolved by longer reverse transcription times, by using high processivity DNA polymerases, or setting a longer extension period in the PCR program. Another problem is that the cDNA may not be full-length due to stalling of reverse transcriptase, particularly in CG-rich sequences. As the forward primer is the common tag and the user-designed antisense primer is specific for the target gene, the specificity of amplification is dependent on the specificity of the antisense primers. By using two sets of primers, specificity is enhanced but even if specific, it should be considered that a single band can represent multiple distinct products of similar size coming from distinct RNA variants. It is also possible that multiple bands arise from the same TSS as byproducts of RNA degradation before the 5′-tag was added. In any 5′-RACE protocol, the final answer should come after purification and sequencing of each product. If the PCR results in too many products (a smear), a Southern blot using the relevant labeled genomic fragment can be done to identify products from the gene of interest, which can be verified by DNA sequencing. The identification of the TSS is an essential component in the study of the control of gene expression and opens the way to further investigate the promoter region(s).

RNase Protection Assay

An important and complementary method to identify the TSS is the RNase protection assay [27, 28]. In this assay, a synthetic RNA probe antisense to the target DNA and extending beyond the putative TSS is synthesized and labeled with [32P]-UTP by in vitro transcription from a plasmid- containing genomic sequence including the target TSS and 5-flanking sequence [20]. Typically, plasmids such as KS+ (Stratagene) or pcDNA3 (Invitrogen) can be used as they contain initiation sites for T7, T3, or SP6 RNA polymerase. The plasmid is linearized to terminate the probe, and the reaction is terminated using RNase-free DNase to remove plasmid DNA and the probe purified on urea-polyacrylamide gel. The labeled RNA probe is hybridized overnight to denatured poly-A+ RNA or total RNA from tissue that expresses the target gene and is digested with RNase at conditions that are optimized. RNase preferentially removes the single-stranded nonhybridized portion of the RNA–RNA hybrid. The remaining portion is then electophoresed on urea-polyacrylamide gel to determine the molecular size. For accurate size determination, the hybridized portion should be less that 150 bp. Because this method relies on RNase digestion rather than reverse transcription from an antisense primer as is the case for primer extension or 5′-RACE, it is not subject to the problem of incomplete cDNA synthesis. Hence, the RNase protection assay provides a complementary approach to identify the TSS.


Characteristics of Promoters

Once the transcriptional start site of a gene has been identified, the promoter region upstream of that start site can be characterized to elucidate its transcriptional regulation. There are two general types of promoters: TATA box and TATA-less promoters. TATA-containing promoters typically initiate transcription 25–35 nucleotides downstream of the TATA box and often have a CCAAT box sequence located with 50–100 nucleotides upstream [29, 30]. TATA-less promoters are often CG-rich with multiple GC boxes (sites for Sp1/MAZ) and initiate at multiple sites or at an initiator (Inr) motif [31–34]. Examination of potential promoter regions of over 1000 genes by Suzuki and collaborators (2001) revealed that almost all putative promoter regions contained GC boxes (97%) and initiator motifs (85%), whereas substantially fewer contained CAAT or TATA boxes (64% and 34%, respectively), and only a fraction of promoters are composite promoters (TATA and Initiator dependent). However, the presence or absence of a TATA box is not necessary for correct expression. The human and mouse 5-HT1A receptor promoters, for example, have a TATA-less promoter that instead responds to MAZ and Sp1, and have multiple initiation sites [21]. By contrast, the rat 5-HT1A receptor promoter has CCAAT and TATA boxes, in addition to MAZ/Sp1 sites, and initiates at a single major TSS located 58 bp downstream of the TATA box [20]. Nevertheless, both mouse and rat 5-HT1A receptors are expressed with a similar tissue and brain regional distribution [17, 35]. Beyond these well-characterized essential promoter elements, there are multiple enhancer and repressor elements that coordinately regulate developmental and tissue-specific gene expression [36]. It is also important to note that, whereas promoters are typically located immediately upstream of the transcription start site, there are many examples of distally located, intron or downstream enhancer or repressor elements [37–39].

Bioinformatic Analysis of Promoters

For determining where known or user-defined transcriptional regulatory elements might reside in putative promoters, programs like the TESS (Transcription Element Search System, can be indispensable. Many other programs exist to facilitate identification of transcriptional regulatory elements, including TRANSFAC and JASPAR [22, 23], and the methodology for determining consensus-binding elements has been well described [23]. However, identification of a consensus element in a gene does not mean that it is a truly functional element, and this must be tested by functional assays as described below. Oppositely, many DNA elements have degenerate or poorly defined consensus sequences that may not be identified by search programs or may appear numerous times in the sequence as false hits.

Gene Reporter Assays

Once a putative promoter or regulatory sequence has been identified, gene reporter assays are widely used to functionally characterize putative promoter, enhancer, and repressor regions. Reporter assays provide a sensitive readout of the transcriptional activity of a DNA segment and can be used to test the effect of specific treatments on the transcriptional activity of a promoter.

Reporter constructs are generated by subcloning putative promoter sequences upstream of the transcriptional start site into vectors carrying an assayable nonmammalian reporter gene such as luciferase, chloramphenicol acetyltransferase (CAT), enhanced green fluorescent protein (EGFP), or β-galactosidase. The rationale for using a nonmammalian reporter is that it removes background due to endogenous mammalian transcripts and protein that are present in transfected mammalian cells. Also, the reporter construct removes mRNA sequences of the gene that could be regulated by posttranscriptional mechanisms, allowing for a discrete and sensitive assay of transcriptional activity. Typically, a series of 5′-deletion reporter constructs are generated that terminate at a common transcription start site so that the activity of equivalently initiated transcripts can be assessed [20].

Reporter assays involve transfection of the reporter plasmid into cells and preparation of cell extracts for assay of the reporter protein activity. The luciferase enzyme, for example, catalyses a reaction that emits light upon addition of its substrate luciferin (D-luciferin for firefly luciferase or coelenterazine for Renilla luciferase), which can be monitored by a luminometer such as the Lmax II (Molecular Devices) that permits rapid measurement of reporter activity in 96-well plates. Such sensitive monitoring systems also permits the use of smaller numbers of cells, less transfection reagent and less DNA for reporter assays. Assuming that all constructs have the same start sites and posttranscriptional processing, the activity of the reporter protein should be directly proportional to transcriptional activity of the transfected plasmid. To compare different plasmids and transfections, a second reporter plasmid, such as β-galactosidase or Renilla luciferase under control of a strong viral promoter (e.g., SV40), is cotransfected in order to control for transfection efficiency. The activity of this second reporter can also be measured and used to normalize the respective luciferase values. The fast decay kinetics of luminescence can be a problem when reading a large number of samples. As reactions are read in serial order, the reaction times of the first and last samples can differ considerably. Using the luciferase encoded in Promega plasmids pGL3 or pGL4, we suggest setting the integration time such that all reactions are read within 2 min. Finally, although buffer, substrate, and cell lysates are best kept on ice, the reaction should proceed at room temperature. Typical controls include transfection of the vector (without insert), and a positive control using a strong promoter to drive reporter expression. To determine whether the insert is a promoter or enhancer, the insert is cloned in the antisense orientation relative to the reporter as promoters lack activity in the antisense orientation, whereas enhancers or repressors display orientation-independent activity. In addition to measuring basal activity of the constructs, induced activity can be measured upon treatment with pharmacological regulators (second messenger activators such as forskolin, phorbol esters, growth factors, etc.) or cotransfected regulators to identify regulatory elements in the promoter.

Once a region of interest has been narrowed down by the deletion analysis described above, the specific element can be defined by excising the region of interest and placing it adjacent to minimal promoter–reporter construct [40]. If possible, the element of interest can be further narrowed down using bioinformatics to identify potential consensus sequences or palindrome (inverted repeat) sites. Alternately, DNase protection analysis of the region of interest can be used to experimentally identify sites of protein-DNA interaction [40]. Once the element of interest is identified, its function can be verified by deletion or site-directed mutagenesis of conserved nucleotides to disrupt the element, and reporter assays done. In addition, the activity of the element subcloned upstream of a minimal promoter should be examined. Upon identification of a known DNA element, the role of cognate transcription factors can be examined by cotransfection of their expression plasmids with reporter constructs. Binding of these factors to the DNA element can be verified by EMSA or supershift (see the following). If the element is novel, can be used yeast one-hybrid screening to identify transcription factors that bind to the element (see the next section).

Transfection of Cell Lines

Transfection of cells in culture is a basic technique of molecular research; however, optimal methods for different systems can vary widely. One of the more commonly used and efficient transfection methods is liposome-mediated transfections [41], which package DNA into lipophilic vesicles that can fuse to the cell membrane and release the DNA they carry. Many companies market transfection reagents for different purposes, with optimal working protocols inevitably being determined by the end user. It is important to optimize the cell density, lowest effective concentration and ratio of DNA-lipid reagent, the time of transfection, and the length of time the cells continue to grow after the transfection has been stopped. Common lipid transfection protocols from Invitrogen (i.e., Lipofectamine Plus) and Qiagen (i.e., TransMessenger) call for 3–6 h transfections, followed by 24–72 h recovery posttransfection. One of the least expensive transfection protocols is the calcium phosphate [42] method, which was one of the first transfection methods used but, has the downfall of using substantially more DNA and at times being unreliable. Other methods for DNA delivery include DEAE-dextran, dendrimers, “gene guns,” viral infection, electroporation, and direct microinjection.

Transfection of Primary Neurons

The mechanism for transfecting primary neurons in a culture with any lipid-based reagent differs from that acting on other cells. The lipid and the plasmid form a complex that allows the latter to be taken up by the cell. Once inside, in actively dividing cells, the DNA becomes trapped in the nucleus, where it can be expressed [43, 44]. On the other hand, the DNA enters the neuronal nucleus by other means that remain to be determined. In practical terms, this means that neurons and nonneuronal cells should be plated at different densities. Although proliferating cells should be given space to divide following transfection, it is better to maximize the number of plated neurons. Confluence around 85 to 90% is optimal, as overcrowding can negatively affect transfection efficiency [45].

Interpretation of Reporter Assays

Although reporter assays represent an essential tool for examining regulatory elements, it is important to bear in mind that reporter constructs transfected in cell lines do not replicate the full regulation of genes that occurs in vivo. For example, the action of distal enhancers or regulators is lost, together with the effect of elements present in introns, the coding sequence or the 3′-UTR. Secondly, evidence is growing for the effects of a specific nuclear location or genomic environment on transcription, factors that do not apply to transfected plasmids [46]. In addition, the epigenetic history that might control a promoter in vivo cannot be reproduced unless purposefully replicated for the experiment. The activities of promoters driving effector plasmids will likely also lead to levels of these proteins that do not normally occur in vivo. Other features are lacking in transfected plasmids, such as histone packaging and chromatin condensation, which also may not reflect the in vivo situation. Thus, interpretation of reporter assays must be tentative until the role of specific DNA elements or transcription factors can be demonstrated in the endogenous gene as described below.

Transformed cell lines are often used for transfection experiments as their transfection efficiency is high and very reproducible; however, cell lines are transformed and may not accurately reflect regulation of genes in normal tissues, hence, the regulation should be verified in primary cultures or tissues if possible. However, some cell lines do retain characteristics of normal cells. For example, rat RN46A cells provide a model of serotonergic raphe cells [47], whereas human SK-N-AS neuroblastoma cells model presynaptic dopamine neurons [48, 49]. Nonetheless, even in cell lines that retain differentiated properties, the levels of expression may be different than that observed in normal cells, and hence the transcriptional activity of the promoter may be weak or subject to strong repression. For example, in RN46A cells the level of 5-HT1A receptors is much lower than in raphe neurons [50], in part due to strong repression by the novel repressor Freud-1 [40, 51]. To obtain optimal responses in reporter assays it is important to try to match the species of transfected promoter DNA with the cell species. It is also important to compare reporter activity in cells that express the gene of interest to activity in cell lines that do not express this gene (i.e., the use of the nonneuronal, serotonin-deficient rat L6 myoblasts for studying 5-HT1A promoter elements).


EMSA (Gel Retardation Assay)

The interaction of proteins with DNA is important to control DNA repair, recombination, and transcription processes. Electrophoresis mobility shift assay (EMSA) is useful in studying the gene regulation and determining the DNA-protein interaction. The assay is based on the observation that complexes of protein and DNA migrate through a nondenaturing polyacrylamide gel more slowly than the free DNA fragments. The gel shift assay is performed by incubating a purified recombinant protein, nuclear or cell extract with [32P]-end-labeled DNA fragment containing the putative protein binding site. The reaction products are then analyzed on a nondenaturing polyacrylamide gel. The specificity of this interaction is confirmed by competition experiment using unlabeled DNA fragment of interest.

Probe Design

The first step in EMSA is probe design. Double-stranded linear DNA fragments of the binding site of interest are used in the gel retardation assay and are typically 10–30 nucleotides in length. Synthesized target sequence oligonucleotides (either sense or antisense), including a surrounding sequence with 5′ single-stranded overhangs (for labeling with Klenow) are purified by polyacrylamide gel electrophoresis to remove incomplete products or HPLC- (high performance liquid chromatography) purified primers are used [40]. The primers are annealed and radiolabeled by incorporating [α-32P] dNTP during a 3′ fill-in reaction using Klenow fragment of E. coli DNA polymerase [52]. Alternately, 5′-end labeling is done using [γ−32P] ATP and T4 polynucleotide kinase, although we find that Klenow results in better and more stable labeling. For nonradioactive DNA labeling, a biotinylated or hapten-labeled dNTP is incorporated, then probed and detected by the sensitive or chemiluminescent substrate. Free radioactivity probe is removed using Sephadex G-50 column chromatography. For novel DNA elements it is important to identify the minimal sequence that is required for binding of a transcription factor to minimize nonspecific or irrelevant protein complexes. This can be done by DNase protection analysis [40] or by designing a series of unlabeled probes of shorter sequences and examining their ability to either compete for specific binding, or by using the shorter sequence as a probe to bind to the protein of interest.

Recombinant Protein

Secondly, a source of transcription factor is required: either cell extract or recombinant protein can be used. For cell extracts, a cell line or tissue that preferentially expresses the protein of interest is chosen: the presence of the transcription factor can be identified by EMSA or Western blot. For generation of recombinant proteins, several in vitro transcription/translation protein expression systems are presently available such as EcoPro (Novagen), TNT® Coupled Reticulocyte Lysate (Promega), and TNT® Coupled Wheat Germ (Promega). The major limitation of these systems is their low levels of protein expression, making it difficult to perform more than one experiment with the same preparation of protein as well as limiting detection abilities in EMSA. In addition, proteins in the lysate sometimes produce nonspecific background. Alternately, the most common method for recombinant protein expression is the use of bacterial expression systems. Multiple vectors (pTriEX4 and pET30, Novagen; and pGEX, GE Healthcare) allow for expression of large amounts of protein. The major limitation to this expression system is potentially inadequate posttranslational modification, and decreased solubility and stability of proteins in the lysis and elution buffers, sometimes limiting the quantity and quality of the resultant protein. In some cases these problems can be fixed with the use of denaturing conditions and dialysis for renaturing, but not all proteins refold properly and degradation/aggregation may occur following denaturation. Finally, another method for protein expression uses eukaryotic expression systems with transfection reagents and mammalian expressing vectors such as pcDNA3 (Invitrogen) and pTriEX4 (Novagen). A mammalian cell expression system is optimal because it provides protein cofactors and modifications that might be essential for protein–DNA interaction. However, the presence of endogenous proteins can often cause serious background problems or interfere with detection of the recombinant protein.

Binding Reaction

For EMSA, the labeled primer, protein extract, and competition primers are combined at room temperature. As a negative control, the protein extract is omitted. To initiate DNA binding, purified protein is premixed with Poly [d[I-C]] and incubated 20 min, then a 100- to 200-fold excess of unlabeled competitor sequence is added and incubated for 20 min. Finally, 50,000 cpm of labeled probe is added to the mixture. Any specific band should be eliminated by the presence of excess unlabeled specific competitor. The addition of a mutant or unrelated sequence will not compete with the labeled target, and the band will be preserved. The competition assay is a particularly important control for cell extracts, which may contain multiple non-specific or other protein–DNA complexes. At high excess (>50-fold), nonspecific competitor primers with weak sequence similarity to the labeled primer may partially compete for the band of interest, and competitors with no sequence similarity to probe and lacking repetitive sequences should be chosen. It is also essential to use Poly [d[I-C]] (Roche) or herring sperm DNA to reduce nonspecific binding. Poly [d[I-C]] is thought to be better at blocking nonspecific binding but may eliminate specific interactions, in which case use of herring sperm DNA is necessary [53].

Supershift EMSA

Electrophoretic separation is done using a nondenaturing gel to preserve the protein–DNA complexes and resolves based on the charge and hydrodynamic size, hence the migration of the complex does not necessarily correspond to its absolute molecular weight. In order to identify the proteins in the complex it is necessary to use antibody specific for the protein of interest to supershift the complex. If the protein of interest binds to the target DNA, the antibody will bind to that protein–DNA complex, decreasing furthermore its mobility and resulting in a “super-shift.” In some cases, the antibody may interfere with the protein–DNA interaction, resulting in a reduction or loss of the specific band, rather than a supershift [51]. An important control is to use preimmune serum to show that the effect of the antibody is specific; in addition, a control with antiserum in the absence of cell extract should be run to rule out nonspecific DNA binding in the antiserum. An alternative identification process would be the “shift Western blot.” This involves transferring the resolved protein–DNA complexes to stacked nitrocellulose and anion-exchange membrane. Labeled DNA probe is detected on the anionic membrane whereas protein captured on the nitrocellulose membrane can be probed with a specific antibody (Western blot) [54].

If for some reason EMSA does not result in a clear outcome, it is possible to demonstrate protein–DNA interaction with the use of a CHIP assay or yeast one-hybrid system [51].


Once promoter analysis has revealed a DNA element of interest, if the element identified does not match a known consensus sequence and it is clear from EMSA of relevant nuclear extracts that a specific protein–DNA interaction occurs, then a screen must be performed to identify the interacting proteins. There are several options for screening. We have tried screening lambda-GT11 bacterial expression cDNA library with labeled oligonucleotides for the DNA element but obtained weak and inconsistent signals (Ou and Albert, unpublished data). This may reflect the low DNA binding activity of transcription factors following lysis of bacteria. Another option is to use the DNA element as bait in an affinity purification protocol to isolate DNA binding proteins from relevant nuclear extracts. However, DNA binding proteins are often present in such low abundance that this may require large scale, multistep purification protocols. Finally, we have had success using the yeast one-hybrid approach, which provides an effective method for screening for DNA-binding proteins.

Yeast One-Hybrid Analysis

The yeast one-hybrid system is an in vivo genetic assay for isolating novel genes encoding proteins that bind to a target, cis-acting regulatory element. Yeast provides a useful system to screen for mammalian transcription factors as the background is low. However, enhancer or promoter elements may produce background problems if the yeast homologues are able to directly bind to target elements and activate transcription. In contrast, repressors reduce the expression of target genes and therefore do not contribute to the background signal.

To screen a library for a gene encoding a DNA binding protein (BP) of interest, a reporter construct is made using oligonucleotides containing at least three tandem repeats of the specific target DNA element placed upstream of the reporter (Figure 5.2). Multiple copies of the DNA element enhance sensitivity and specificity of the screen. We use a dual His3/LacZ reporter system to reduce background and maximize stringency. Once the target-reporter strain is isolated and selection conditions established, a cDNA-library fused to the Gal4 activation domain (GAL4-AD) is transformed into the strain for screening. Clones that encode a GAL4-AD fusion protein that are able to bind to the target DNA element recruit GAL4 to activate transcription of the selectable marker, allowing growth of the yeast clone (Figure 5.2). This clone is then subjected to β-galactosidase assay to finally examine the activity of the isolated clone. Here, it is important to isolate DNA and retransform naïve reporter yeast strain to verify that the activity is due to the GAL4-AD clone. The positive clones are sequenced to confirm the identity of the cDNA insert and that it is in-frame with the GAL4-AD. The positive clone must then be tested for its DNA binding activity by EMSA, and for its function using promoter analysis in mammalian cells as described above.

FIGURE 5.2. Yeast one-hybrid strategy for cloning DNA binding proteins.


Yeast one-hybrid strategy for cloning DNA binding proteins. Yeast one-hybrid cloning was done using a human brain cDNA library fused to the GAL4 activation domain (AD). The target sequence was composed of either 3 copies of the DRE (for cloning of Freud-1 (more...)

One-Hybrid Screening Protocol

The plasmids provided with the Matchmaker One-Hybrid System ( are: pHISi, pHISi-1, and pLacZi, which contain for selection the yeast HIS3 or URA3 genes, respectively. The target-reporter constructs are linearized, transformed into the yeast strain YM4271 (Clontech) (MAT a, strain for integration of the element) using PEG4000/Li acetate followed by DMSO shock, and plated on the appropriate selection plates (SD-his or SD-ura) at 30°C. For pLacZi, we use a qualitative β-galactosidase assay [55] directly on the plate and for pHISi and pHISi-1 to determine the optimum concentration of 3-AT (3-amino-1,2,4-triazole; a competitive inhibitor of the HIS3 gene product) and identify which of the two HIS reporter strains has the lower background of HIS3 expression. If the β-galactosidase assay indicates that the background lacZ expression is low, you can prepare a dual reporter strain for library screening by integrating the target-HIS3 with the lower background HIS3 strain into the target lacZi strain. If the β-galactosidase assay [55] indicates that the background is high, the library can be screened using the target-HIS3 reporter strain with the lowest background of HIS3 expression. Subsequently, the single or dual reporter strain is transformed with the Matchmaker cDNA library fused to GAL4-AD and containing a LEU2 selectable marker for transformants, and selected on SD-leu-his+3-AT (at the optimized concentration) plates. For the AD-library, pretransformed cDNA libraries are available or you can amplify a cDNA library (according to availability). Pretransformed libraries are more convenient. However, in our hands we obtain many copies of the same clones and fewer different clones. This could be due to amplification of the pretransformed libraries that reduced the diversity of clones present in the library. Another problem is that pretransformed libraries of tissues or cell lines of interest may not be available.

Chromatin Immunoprecipitation (CHIP) Assay

Chromatin immunoprecipitation (CHIP) is an excellent way to demonstrate that the transcription factor is associated with a gene of interest in vivo. In this approach, an antibody to the transcription factor of interest is used to immunoprecipitate the protein–DNA complex that has been cross-linked in whole cells. The immunoprecipitate is then subjected to PCR using primers that flank the DNA element of interest. The products are quantified using electrophoresis or Q-PCR. In order to visualize these complexes it is essential to have a tissue or cell line that expresses the protein of interest, as the transcription factors bound to a gene may vary depending on cell type and level of gene expression. Another limitation for this technique could be the sequence surrounding the DNA element, as it might be challenging at times to design an oligonucleotide sequence that will yield an adequate PCR product.

A critical step is to identify an antibody that is able to immunoprecipitate the protein (IP) of interest once it has been fixed to its cofactors and DNA target. Alternately, if no such antibody is available, a transfected cell line that stably expresses the transcription factor (preferably containing an epitope tag to allow the use of anti-epitope antibody) can be generated [56]. Many commercially available antibodies will now indicate if they are suitable for CHIP assays. The initial step to this technique is fixation of cells that should be kept at the same confluency from experiment to experiment to achieve reproducibility. Many fixation protocols have now been developed. Some use formaldehyde (FA) only, whereas others combine N-hydroxysuccinimide (NHS) ether chemistry with FA to crosslink protein complexes [57]. Optimizing the fixation time is important, as overfixation can lead to protein–protein or protein–DNA aggregation while insufficient fixation will fail to stabilize protein-DNA complexes [58]. Following fixation, it might be advantageous for some proteins to perform nuclear extraction in order to eliminate background from cytosolic proteins [57]. The most widely used kit and protocol is from Upstate with many modified versions, one of which includes addition of 212- to 300-μm diameter glass beads during the critical sonication step (Sigma) [59]. The most important modification to the Upstate protocol is an increase in the number of washes to reduce high background in the elution fractions [5]. It is also critical to use silanized tubes with low DNA and protein binding capacity, as well as to change tubes during the washes. The analysis of bound DNA can be done either by regular PCR or Q-PCR for quantitative results. If regular PCR is used, it is essential to identify the number of cycles required for amplification, so as not to reach a plateau level as this will prevent accurate data analyses. On the other hand, the data analysis with Q-PCR is complicated by a lack of internal standard. It is then essential to find a control that will allow for normalization between experiments, such as a no antibody control. For reproducible results it is important to use consistent amounts of tissue, sonication, wash, and solution conditions.

Although use of CHIP assay may present several challenges, particularly for characterizing novel transcription factors, it is a powerful approach to demonstrate protein-DNA interactions in vivo, and complements the DNase I protection analysis and EMSA approaches.


GAL4-DBD Hybrid System

Once a putative transcription factor has been identified, it is important to understand whether transcriptional regulation is an intrinsic property or whether it is indirect occurring through competition with other DNA binding proteins. Repressor-operator systems, such as the Gal4-DNA Binding Domain (DBD) heterologous hybrid system [60], provide useful methods to address the intrinsic transcriptional regulatory activity of a protein. The S. Cerevisiae transcription activator Gal4 is a protein of 881 amino acids [61] and is required for the proper expression of genes that encode galactose-metabolizing enzymes. The first 147 amino acids of Gal4 contains a cysteine-rich DBD, which binds to specific DNA sequences (UASG), whereas the C-terminal domain of Gal4 contains the acidic transactivation domain (AD) which recruits RNA polymerase [60]. The Gal4 DBD and AD can function independently as “cassettes” to confer their activities on fusion proteins (Figure 5.3). By fusion of the transcription factor to be examined with the Gal4 DBD, the activity of this transcription factor (enhancer or repressor) is conferred upon the fusion protein. Although mammalian cells lack endogenous Gal4, yeast Gal4 protein can be a functional transcription factor in these cells by using the firefly enzyme luciferase as a typical reporter gene. When transfected into mammalian cells, the Gal4-DBD fusion protein can bind to the UAS(G) sequence. By cotransfection of a luciferase reporter plasmid that has incorporated the UAS(G), the activity of the fusion protein is assayed.

FIGURE 5.3. Gal4-DBD-hybrid to assess intrinsic repressor activity of Freud-1.


Gal4-DBD-hybrid to assess intrinsic repressor activity of Freud-1. To assess the intrinsic repressor activity of Freud-1, the indicated plasmids were constructed including Gal4-DBD (Gal vector), Gal4-Freud-1, or LexA (positive control) and were cotransfected (more...)

For example, a Gal4-DBD fusion construct was generated by subcloning the protein of interest (Deaf-1, Freud-1) into a vector (EcoR1 site of the pBXG-1 vector, obtained from Reference 62), ensuring ORF is in frame with the Gal4-DBD [52, 63] (Figure 5.3). A reporter construct (X2G2P) containing two LexA and two Gal4 UAS sites upstream of the SV40 promoter-luciferase was cotransfected with the fusion construct to determine whether the protein of interest confers repressor or enhancer activity when it is recruited to the heterologous element and promoter. As the transfected reporter construct is at a 10- to 100-fold molar excess compared to endogenous promoters, this provides a reliable measure of the intrinsic activity of the fusion protein at the reporter construct. Using this assay we found that although Freud-1 displays repressor activity in all cell types examined, Deaf-1 displayed intrinsic repressor or enhancer activities, depending on the cell type [52, 63]. A bacterial repressor, LexA, has been shown to reduce gene expression in eukaryotes [64] and is used as a positive control for repressor activity. A protein that lacks intrinsic activity would not affect the activity of this construct. However, a negative result must be interpreted with caution as the structure and function of the protein may be compromised in the fusion. As a positive control, the fusion should be tested for activity at its cognate DNA element.

The mammalian two-hybrid system is another version of this technique that is useful for detecting protein–protein interactions in mammalian cells. In this system, both Gal4-DBD and Gal4-AD fusion proteins are cotransfected with the reporter construct containing Gal4-UAS and minimal promoter. If the two fusion proteins interact, this leads to recruitment to the reporter of the Gal4-AD fusion by the Gal4-DBD fusion protein. It is important to note that transcription activation domains other than Gal4, such as VP16 (viral protein 16) or B42, can also be used, each with its own associated advantages and disadvantages.

siRNA or Antisense Probes

Many gene targets are proposed for transcription factors but in vivo analysis of their activity is necessary to conclusively state their involvement in gene regulation. One useful approach is to use RNAi (RNA interference) [65] to knock down the transcription factor of interest and measure the effect on the gene of interest using QPCR to quantify RNA levels, Western blot to measure protein, or by cotransfecting reporter constructs to assay transcriptional activity. siRNA (small interfering RNA) has proven a useful approach for specific knockdown of gene expression. The RNAi-target RNA hybrid recruits an RNA-inducing silencing complex (RISC) which aids in cleavage of targeted RNA with the use of endo- and exonucleases [66]. The use of siRNA is advantageous due to fast production and lower interferon response compared to other methods such as shRNA (short hairpin RNA) [67, 68]. One disadvantage to the use of siRNA is the inability to make stably expressing cells lacking the protein of interest, which is possible with the use of antisense or shRNA technology. The knockdown time is also limited due to the lability of RNAi. This method is also quite costly, especially if the target protein is not knocked down with the first couple of siRNAs, in which case multiple siRNAs must be tested. If there is a need to target the same gene in multiple species, it can be challenging to find a common sequence that would specifically target the gene of interest in all species.

The most important step in siRNA technology is the design of siRNA molecules that specifically target the sequence of the gene of interest only, using BLAST search for validation. The targeting sequence is usually 21 ribonucleic acids in length. It is essential to make a negative control, ideally containing the same nucleotide content but lacking recognition, with similar %GC content. Several free Web-based design software programs (e.g., Invitrogen, as well as many predesigned siRNAs for known genes are available which guarantee “specific” knockdown. Importantly, at least one additional siRNA that targets your protein should be designed to verify the specificity of the effect. siRNA can be readily transfected into cultured cells using available reagents such as Lipofectamine 2000 (Invitrogen) and HiPerFect (Qiagen). A positive control such as BLOCK-iT Fluorescent Oligo (Invitrogen) for transfection efficiency is useful to optimize delivery of siRNA. The time period for efficient protein knockdown will vary, depending on the RNA and protein half-life of turnover, and it is critical to optimize this time for each target. In our hands 72-h exposure to siRNA leads to ~90% down-regulation of protein expression (unpublished data), but this will vary, depending on the protein. The siRNA approach can provide evidence that the transcription factor is functionally important for the regulation of the gene of interest. However, it cannot be concluded whether the effect is direct (via a specific DNA element on the gene) or indirect (via modulation of other genes such as transcription factor genes).


The methods described above provide a step-by-step approach to characterize the transcriptional regulation and regulators of a gene of interest. This approach provides mechanistic insight into the fundamental regulation of the gene, but other sites of regulation including enhancer or repressor elements located in introns or 3-flanking sequence. The identification of specific DNA elements and transcription factors provides a starting point for further studies of epigenetic changes such as alterations in histone modification (lysine acetylation, methylation, sumoylation, etc.) using CHIP assay or of DNA methylation (at CG dinucleotides) that occur at these regulatory sites [6, 8, 69]. Recently, modification of histone acetylation and DNA methylation have been used to produce alterations in learning and behavior [6, 7, 70, 71].

The finding of novel transcription regulators and DNA elements provides an important tool for understanding global gene regulation in the nervous system. As transcription factors generally regulate multiple gene targets that have the appropriate DNA element, these can be identified by bioinformatics searches for consensus DNA elements, or by gene array studies [72, 73]. Gene knockout studies of novel transcription factors have also revealed specific functions in systems regulation for specific transcription factors (Pet-1/DREAM) [74, 75]. Because these factors provide a global regulation of multiple genes within a system, they may become useful therapeutic targets for global modulation of gene expression.

The identification of transcriptional mechanisms provides a novel method for addressing the function of promoter polymorphisms. For example, the 5-HT1A receptor promoter polymorphism C(−1019)G was initially identified in the repressor region of the 5-HT1A gene. We have been able to show that the polymorphism is the target of at least two repressor proteins (Hes-5 and Deaf-1) and that the G-allele binds weakly or not at all to these factors, leading to derepression of the 5-HT1A autoreceptor [76]. We are currently using the novel DNA elements that we have identified to search for conserved elements and nearby functional polymorphisms in other candidate genes involved in mental illness.


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