Methods to Study Transcription Factor Structure and Function

C H A P T E R

2 Methods to Study Transcription Factor Structure and Function Ivana L. Viola, Daniel H. Gonzalez Instituto de Agrobiotecnología del Litoral (CONICET-UNL), Cátedra de Biología Celular y Molecular, Facultad de Bioquímica y Ciencias Biológicas, Universidad Nacional del Litoral, Santa Fe, Argentina

O U T L I N E 2.1 Introduction

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2.2 In Vivo Functional Studies 2.2.1 Overexpression or Ectopic Expression of Transcription Factors 2.2.2 Transactivation Systems for Expression of Transcription Factors 2.2.3 Expression of Dominant Negative Forms of Transcription Factors 2.2.4 Expression of Dysregulated Forms of Transcription Factors 2.2.5 Gain-of-Function Mutants 2.2.6 Expression of Fusions to Activating or Repressor Domains 2.2.7 Inducible Systems

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2.3 Methods for the Analysis of In Vitro Protein–DNA Interactions 2.3.1 The EMSA Assay 2.3.2 Selex 2.3.3 Analysis of Transcription Factor–DNA Complexes by Footprinting Assays 2.3.4 Microarray-Based Identification of Transcription Factor Target Genes

2.4 Methods to Study Protein–DNA Interactions In Vivo 2.4.1 Chromatin Immunoprecipitation (ChIP) Assays 2.4.2 ChIP-chip and ChIP-Seq 2.4.3 DNA Adenine Methyltransferase Identification (DamID) 2.4.4 Identification of Transcription Factors Using the Yeast One-Hybrid Assay 2.4.5 Transient Assays to Analyze Protein–DNA Interactions In Vivo

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2.5 Analysis of Protein–Protein Interactions 2.5.1 Methods for Protein–Protein Complex Identification

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2.5.1.1 Yeast Two-Hybrid Assay (Y2H) 2.5.1.2 Tandem and One-step Tag-Based Affinity Purification

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2.5.2 Methods for Verification of Protein–Protein Interactions

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2.5.2.1 Coimmunoprecipitation 28 2.5.2.2 In Vivo Split Methods 28 2.5.2.3 Resonance Energy Transfer Methods 28

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References

2.1 INTRODUCTION

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present in the promoters of their target genes (Harbison et al., 2004). According to the basic combinatorial principles of gene regulation, a given DNA-binding protein contributes to the transcription of many genes with varying expression patterns by acting in conjunction

The regulation of gene expression is a complex process controlled by a network of interactions between different regulatory proteins and cis-regulatory sequences

Plant Transcription Factors. http://dx.doi.org/10.1016/B978-0-12-800854-6.00002-6 Copyright © 2016 Elsevier Inc. All rights reserved.

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2.  Methods to Study Transcription Factor Structure and Function

with several other transcription factors, each possessing its own unique expression pattern. In addition, transcription factors can be regulated by posttranscriptional mechanisms, allowing them to be present in an active or inactive state or in different subcellular compartments. Chromatin structure and modification states play key roles in determining the competence of transcription, and most DNA-binding proteins can recognize a broad spectrum of DNA sequences with a wide range of affinities. Furthermore, most DNA-binding proteins are members of protein families, with each cell type containing several family members that recognize similar DNA sequences. Based on these considerations, it is likely that multiple proteins will be capable of binding a defined control element in vitro, including several members of a particular family of proteins, and perhaps members of another family that recognize a similar or overlapping sequence. The challenge is to determine which of these proteins is capable of performing the protein–protein and protein–DNA interactions that allow it to regulate endogenous genes. In conclusion, despite intensive work, deciphering the transcriptional code is proving to be more difficult than the genetic code (Harbison et al., 2004). The development of methodologies that allow the characterization of transcription factor DNAbinding specificities has been crucial to understanding transcriptional regulation. Transcription profiling can be applied to stable loss- and gain-of-function transcription factor mutants to identify the global expression changes that are associated with the mutant phenotype, thereby facilitating placement of the transcription factor in a developmental pathway or process. Combining DNA motif discovery and information on the up- or downregulation of target genes can lead to a deeper understanding of molecular modes of action and the specificity of particular transcription factors. In this chapter, the different strategies that can be used to elucidate transcription factor functions as well as a number of techniques commonly used to determine DNA-binding specificities in vitro and to identify their primary targets in vivo are described.

2.2  IN VIVO FUNCTIONAL STUDIES To study transcription factor action in vivo, the same strategies usually used to study the function of genes, like the analysis of loss-of-function mutants and gene-silencing techniques, for example, can be used. Since many transcription factor families are composed of a large number of members with different degrees of functional overlap, sometimes the analysis of higher order mutants or alternative strategies must be used. Many of these strategies rely on the specific characteristics of transcription factor action and are discussed here.

2.2.1  Overexpression or Ectopic Expression of Transcription Factors The simplest way to modify the action of a transcription factor in plants is to increase its expression by fusing its coding region to a strong promoter (Figure 2.1A). The most popular promoter for this purpose is the 35SCaMV promoter, which brings about high expression levels in most plant tissues. Thus, the use of this promoter and of most promoters used for a similar purpose not only brings about an increase in expression but also expression in cells where the transcription factor is not usually expressed, thus giving rise to ectopic expression too. The phenotypic or gene expression changes brought about by the overexpression/ectopic expression of the transcription factor can be used to infer what the real function of the transcription factor may be. The utility of this approach is mainly based on its simplicity. However, the results obtained are subject to artifacts and must be regarded with care. First, because many of the effects observed are probably due to the presence of the transcription factor in cells where it is normally absent, thus it is difficult to extrapolate how this refers to the “normal” function of the transcription factor. In addition, expression in unusually high levels is likely to distort many of the interactions involved in transcription factor action. Since the establishment of specific interactions with other proteins and DNA are essential for correct transcription factor action, any distortion of these interactions is likely to be the cause of many of the changes observed. Expression at high levels is likely to bring about unspecific protein–protein or protein–DNA interactions. In the case of transcription factor families, for example, a member of a family may be forced to interact with transcriptional components or response elements that are usually bound by a different member of the same family. In families that form heterodimers, overexpression of one member may disrupt the entire set of active molecules formed by the different interacting members of the family. Nevertheless, overexpression approaches are useful in many cases, provided that they are combined with other functional data. Overexpression or the ectopic expression of transcription factors is also useful for technological purposes to obtain plants with desired characteristics.

2.2.2  Transactivation Systems for Expression of Transcription Factors Besides using expression over all parts of the plant to study transcription factor action, it may be useful to analyze the effect of expressing the transcription factor in defined groups of cells or developmental stages. For this purpose, the coding region of a transcription factor gene may be linked to promoter regions known to drive expression in defined places or stages (Figure 2.1A). In

A.  General aspects of plant transcription factors



2.2 IN VIVO FUNCTIONAL STUDIES

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FIGURE 2.1  Strategies to study transcription factor function. (A) The expression level/pattern of a transcription factor of interest (blue) can be modified by expressing it from either a strong, constitutively active promoter (like the 35SCaMV promoter) or another type of promoter that brings about its ectopic expression (gray). (B) A two-component transactivation system can be used to modify the expression of a transcription factor. In this case, the promoter of interest (gray) drives the expression of an artificial transcriptional activator (yellow and green); this artificial protein binds to control elements (violet) located upstream of the coding region of the transcription factor under study, thus promoting its expression. (C) The expression of truncated or modified forms of transcription factors can lead to dominant negative effects by competing with the “normal” functions of endogenous transcription factors or generating constitutively active forms. (D) Fusions of transcription factors to strong activating (green arrow) or repressor (red rectangle) domains can be used to modify/enhance the action of transcription factors. (E) Inducible two-component transactivation systems can also be used. In this case, the artificial transcription factor of a system like the one described in (B) is fused to a steroid hormone-binding domain (orange) that retains the transcription factor in the cytosol. Addition of the steroid hormone (black section) activates the transcription factor that is then able to induce the gene of interest. (F) Inducible one-component systems are also available. In this case, the steroid hormone-binding domain (orange) is directly fused to the transcription factor under study, which becomes inducible by addition of the hormone (black section).

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2.  Methods to Study Transcription Factor Structure and Function

addition to the obvious strategy of cloning the selected promoters in front of the transcription factor coding region, two-component systems were developed for this purpose (Moore et al., 2006). These systems are composed of an activation construct that expresses an artificial transcription factor from a desired promoter and an effector construct in which expression of the gene of interest is located under the control of responsive elements for the artificial transcription factor (Figure 2.1B). Two requisites for these systems are that the responsive promoter is inactive in the absence of the artificial transcription factor and that this factor does not activate endogenous plant genes. Thus, nonplant transcription factors and responsive elements are usually used. One of these systems is the pOp–LhG4 system (Moore et al., 1998). The effector construct of this system is a lac operator (from the Escherichia coli lac operon) fused to a plant minimal promoter. This construct, named pOp, does not direct gene expression when introduced in plants. The activation component encodes a fusion of the Lac repressor (LacI) to the activation domain of the GAL4 yeast transcription factor. When expressed in plants, this chimeric transcription factor, named LhG4, promotes expression of a gene linked to pOp. Thus, cloning the coding region of the transcription factor under study under the control of pOp allows its expression in different tissues/ stages just by crossing plants containing this construct with plants that express LhG4 under the promoter of choice. A different two-component system, based on the E. coli LexA repressor and the lexA operator, has also been developed (Zuo et al., 2000). Other two-component systems for use in plants are the GAL4/UASG systems (Elliott and Brand, 2008). In these systems, developed initially for use in Drosophila, the activating construct expresses the Saccharomyces cerevisiae GAL4 DNA-binding domain fused to a strong activation domain. The effector construct contains several copies of the upstream activating sequence bound by GAL4 (UASG) fused to a minimal promoter. An initial version of this system used the activation domain of the maize C1 transcription factor (Guyer et al., 1998). Later on, the system was adapted for use with the activation domain of the herpes simplex virus protein VP16.

2.2.3  Expression of Dominant Negative Forms of Transcription Factors Sometimes, overexpression of the modified forms of transcription factors can be used to alter the function of endogenous processes related to their action (Figure 2.1C). The logic underlying this strategy is that if a modified transcription factor that is able to perform only part of its normal functions is expressed in plants, it will disrupt processes in which it is normally involved, simply because it will compete with related endogenous

transcription factors for binding to other transcriptional components or to DNA. Typical versions of transcription factors used for this purpose include factors with active protein–protein interaction domains but inactive DNAbinding domains or vice versa. Since most of these strategies are based on successful competition of the modified transcription factor with endogenous components, overexpression is usually required to observe an effect. The drawbacks mentioned previously for the overexpression of native transcription factors are then also applicable in this case. The availability of modified transcription factor forms with considerably increased affinity for endogenous interacting partners may help to overcome this disadvantage.

2.2.4  Expression of Dysregulated Forms of Transcription Factors Since the activity of transcription factors is highly regulated, the expression of a modified form that has lost this regulation may give hints about normal transcription factor function. This type of strategy is useful when the transcription factor becomes constitutively active as a result of the loss of regulation. Changes that interfere with posttranscriptional modifications or stability sometimes originate constitutively active versions of transcription factors. An advantage of this strategy is that the modified transcription factor can be expressed under its own gene promoter and then in the amounts and places in which it is usually expressed. If a knockout mutant is available, then the modified form can be used to replace the endogenous factor and analyze the effect of its dysregulation. As such, this strategy is much more informative than overexpression techniques, but requires a deeper knowledge of the system and is not always applicable. For transcription factors whose expression is regulated by miRNAs, expression of a form that is resistant to the miRNA, obtained by introducing mutations that disrupt the mRNA–miRNA interaction but do not modify the coding capacity, can be used to analyze transcription factor function.

2.2.5  Gain-of-Function Mutants Natural or induced mutations sometimes bring about gain rather than loss of function of transcription factors. Apart from strategies like activation tagging, designed for this purpose and others that provide similar information than overexpression approaches, activating mutations may be due to changes in promoter regions that bring about misexpression of the corresponding gene, mutations that modify the stability or the regulation of the activity of the transcription factor or mutations that bring about dominant negative forms.

A.  General aspects of plant transcription factors



2.2 IN VIVO FUNCTIONAL STUDIES

2.2.6  Expression of Fusions to Activating or Repressor Domains Another strategy that can be employed to study transcription factor function is fusion to strong activating or repressor domains (Figure 2.1D). The logic of this strategy is that binding of the modified transcription factor to its target genes will bring about defined changes (either increase or decrease, respectively) in their expression, and consequent phenotypic changes, that can be used to infer function. This strategy is particularly useful when members of families with redundant functions are under study. Conversion of the transcription factor into a strong repressor or activator usually overrides or counteracts the action of endogenous proteins with similar functions. An advantage of this strategy is that it does not usually require overexpression of the modified transcription factor to observe an effect. In this way, the transcription factor can be expressed under its own gene promoter. A disadvantage is that the normal function of the transcription factor is disrupted in any case, since the repressor or activating domains fused to it most likely establish different interactions with the transcriptional machinery than its native form. It can be postulated that this strategy provides information about the genes with which the transcription factor under study is normally able to interact within the plant, but not about the nature or the magnitude of the changes in expression brought about by this interaction. A repressor motif that is usually used for this kind of strategy is derived from the EAR (ERF-associated amphiphilic repression) domain (Hiratsu et al., 2003). This is a plant-specific repression domain present in several transcription factors that brings about a decrease in gene expression by promoting changes in chromatin structure. This is due to interactions of the EAR domain with corepressors like TOPLESS and SAP18 (Kagale and Rozwadowski, 2011). A modified version of the EAR domain, called SRDX (LDL DLE LRL GFA), has been used to create chimeric repressor gene silencing technology (CRES-T; Mitsuda et al., 2011). SRDX has the advantage that it is a short amino acid sequence that can be fused to many different transcription factors to obtain a strong repressive form. The most widely used activation domain is derived from the herpes simplex virus protein VP16 (Sadowski et al., 1988). This is an acidic activation domain that is functional in many organisms, including plants (Wilde et al., 1994).

2.2.7  Inducible Systems Several inducible systems have been developed to express or activate transcription factors when desired. Induction or activation is usually attained by treatment

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of plants that contain the appropriate construct(s) with a chemical compound (e.g., a steroid hormone; Figure 2.1E). An advantage of this strategy is that it allows verification of the changes that take place soon after the transcription factor is expressed. In this way, changes not directly related with the action of the transcription factor are minimized. This is especially important when expression of the transcription factor brings about profound changes in plant development or growth or is even lethal to plants. Most inducible systems are two-component systems that can be used to express any gene of interest in response to chemical treatment (Figure 2.1E). However, inducible one-component systems can be used to study plant transcription factors. These systems are based on the fusion of the transcription factor of interest to the hormone-binding domain of a steroid hormone Type I nuclear receptor (Figure 2.1F). These receptors are animal transcription factors that are activated by binding the corresponding hormone to the hormone-binding domain. In the absence of the hormone, they are retained in the cytosol and are then unable to affect gene expression. It has been shown that fusion of the hormone-binding domain to many different transcription factors grants them the same regulatory properties, which can then be used to regulate their action (Schena et al., 1991; Lloyd et al., 1994). Fusions to the glucocorticoid and estrogen receptors can be used to regulate transcription factor action by dexamethasone and 17-b-estradiol, respectively. Fusions to the insect ecdysone receptor, which is activated by methoxyfenozide, have also been reported (Martinez et al., 1999; Padidam et al., 2003). An advantage of these systems is that they do not require protein synthesis for activation, since the transcription factor is already present in the cell, though inactive, at the time of induction. In this way, activation in the presence of a protein synthesis inhibitor (usually cycloheximide) can be used to analyze genes that are directly modulated by the transcription factor, thus avoiding secondary effects. Fusions to steroid hormone receptors have been used to assemble inducible two-component systems based on the two-component systems described earlier (Aoyama and Chua, 1997; Martinez et al., 1999; Zuo et al., 2000; Padidam et al., 2003; Craft et al., 2005). In these systems, the activation construct is made inducible by fusing the corresponding artificial transcription factor to a steroid hormone receptor (Moore et al., 2006). In addition, other two-component systems have been used for inducible expression. One of them uses the tetracycline (Tet) repressor encoded in the bacterial transposable element Tn10, for tetracycline-induced expression of a derivative 35SCaMV promoter that contains the Tet repressor binding site (Gatz et al., 1992). Another system uses the ethanol-inducible Aspergillus nidulans ALCR transcription factor (ALCR) to activate a synthetic promoter composed of ALCR target sites (from the

A.  General aspects of plant transcription factors

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2.  Methods to Study Transcription Factor Structure and Function

A. nidulans alcA gene) fused to the 35SCaMV promoter (Caddick et al., 1998). This system has been widely used for inducible expression in plants.

2.3  METHODS FOR THE ANALYSIS OF IN VITRO PROTEIN–DNA INTERACTIONS 2.3.1  The EMSA Assay To study DNA–protein interactions in vitro, a very simple, efficient, and widely used method, first described in 1981, is the electrophoretic mobility shift assay (EMSA, also known as gel mobility shift assay or gel retardation assay) (Fried and Crothers, 1981; Garner and Revzin, 1981). This assay is based on the fact that molecules of different size, shape or charge will have different electrophoretic mobilities in a nondenaturing gel. In the case of a DNA–protein complex, the interaction of the protein with DNA will generate a slower migrating species in relation to the free DNA (Figure 2.2A). EMSA can be carried out in simple steps that include labeling the DNA probe, preparation of the DNA–protein binding reaction and subsequent analysis on a native polyacrylamide gel. In general, oligonucleotides or DNA fragments ranking from 30 bp to 200 bp (base pairs) are used for optimal resolution of complexes versus free DNA, and the DNA can be labeled radioactively or with fluorescent dyes (Steiner and Pfannschmidt, 2009; Viola et al., 2012). In this way, proteins within a crude cell extract that specifically recognize a given cis-element can be analyzed by incubating a labeled DNA fragment with the extract to allow the formation of protein–DNA complexes. Extracts from different cell types or developmental stages can be analyzed to determine whether the binding activity is cell-specific or developmentally regulated. Through these gel retardation experiments it is possible to reveal a specific protein–DNA complex even when the protein is at low concentrations within the extract. To confirm that the detected protein– DNA complexes are the result of a specific interaction with a defined regulatory sequence present in the DNA fragment, a mutagenesis assay of this target sequence is necessary. In this case, the protein–DNA binding reactions will be achieved with the native and a mutant version of the labeled oligonucleotide (Figure 2.2A). Alternatively, competition assays can be established. In these assays, binding to a labeled DNA molecule is performed when there is an excess of unlabeled DNA carrying the same or different sequences. If the interaction is specific, then the binding to labeled DNA will be competed by the DNA carrying the same sequence but not by a different one. In any event, it is always advisable to include a “nonspecific” unlabeled DNA (like poly (dI/dC)) to avoid unspecific binding of proteins to labeled DNA. Besides being useful for qualitative purposes, EMSA has the added advantage of being suitable for quantitative

equilibrium and kinetic analyses (i.e., measurement of dissociation constants and association and dissociation rates; Gerstle and Fried, 1993). Furthermore, because of its very high sensitivity, EMSA makes it possible to resolve complexes of different protein or DNA stoichiometry (Fried and Daugherty, 1998) or even to detect conformational changes in proteins that might interact with the DNA of interest. A modification known as the electrophoretic supershift assay, in which inclusion of an antibody causes an additional retardment of the complex, can be used to identify the presence of a particular protein bound to DNA. A similar strategy can be used to analyze the formation of a complex between a given protein and another protein bound to DNA. Procedures that combine EMSA with western blotting or mass spectrometry have also been designed to identify DNA-binding proteins that recognize specific sequences (Hellman and Fried, 2007).

2.3.2 Selex Transcription factors modulate gene expression through sequence–specific interactions with their DNAbinding sites (Dervan, 1986). Accordingly, the identification of the DNA sequences to which transcription factors bind is a first step in determining their functions in biological processes. Advances in the determination of transcription factor binding sites using in vivo and in vitro techniques has contributed to deducing transcriptional regulatory codes (Harbison et al., 2004; Badis et al., 2009). One of these approaches, SELEX (systematic evolution of ligands by exponential enrichment), provides an excellent tool for deciphering protein DNA-binding sequence specificities in vitro. Initially described by Oliphant et al. (1989) and Blackwell and Weintraub (1990), the SELEX strategy involves the progressive selection of the specific target sequence of a transcription factor from a large combinatorial double-stranded oligonucleotide library through successive steps of binding and amplification. In addition to serving as a technique to establish the in vitro DNA-binding specificity of a protein, SELEX is also a powerful tool in determining whether a particular protein binds DNA in a sequence-specific fashion or not (Chai et al., 2011). In the SELEX assay, a very large oligonucleotide library containing all possible sequences of 20–30 bp flanked by nonrandom sequences of fixed length is used in a binding reaction with the transcription factor under study (Figure 2.2B). The fraction of DNA molecules bound to the protein is separated from the free DNA (i.e., by EMSA, nitrocellulose membrane filtration, using affinity surfaces and affinity tags, or crosslinking and antibody-based flow cytometry; Gopinath, 2007), and amplified by the polymerase chain reaction (PCR) using primers specific to the flanking sequences. Then, the population of amplified molecules is used in a new protein–DNA binding reaction.

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FIGURE 2.2  Methods to study protein–DNA interactions in vitro. (A) Electrophoretic mobility shift assay (EMSA). A protein extract or the recombinant protein of interest is incubated with a labeled DNA fragment and the protein–DNA complexes are resolved from free DNA by electrophoresis in a native polyacrylamide gel (lane 2 of gel image). To confirm the specificity of the interaction, a binding reaction with a mutated version of the DNA fragment is necessary (lane 4). Lanes 1 and 3 correspond to binding reactions without the addition of protein. (B) In the SELEX strategy, a labeled oligonucleotide library that contains a central random region between two constant arms is incubated with the purified recombinant transcription factor of interest (pink oval). Once the oligonucleotides selected by the protein are purified (e.g., by EMSA) the obtained population of DNA fragments is amplified by PCR and employed in a new round of selection. After the desired number of rounds of binding, selection, and PCR amplification, the DNA population is sequenced and the consensus sequence for the DNA-binding protein is obtained. (C) Footprinting assay. A DNA fragment labeled only in one strand is employed in a protein-binding reaction (tube B) and submitted to controlled digestion with DNAse I or attack with hydroxyl radical under conditions where only one cleavage per DNA molecule is produced. As a control, a reaction without protein is performed (tube F). The DNA regions protected by the bound protein can easily be identified by comparison of the patterns obtained after separating the bound (B) and free (F) DNA samples in a denaturing DNA sequencing gel. M, molecular marker. (D) Protein-binding microarray experiments use a high-density dsDNA microarray platform where every possible nucleotide sequence of a determinate length is represented at least once. The purified protein tagged with an epitope (e.g., GST) is allowed to bind those DNA probes containing their preferred sequences. After hybridization with a fluorophore-conjugated antibody specific to the epitope, the microarray is scanned and the spot intensity image is used to determine binding parameters.

A.  General aspects of plant transcription factors

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2.  Methods to Study Transcription Factor Structure and Function

By means of several rounds of selection and amplification, the population of oligonucleotides is then enriched in those that contain the DNA sequences specifically bound by the protein. To determine the consensus-binding sequence of the transcription factor these oligonucleotides are then sequenced. Analysis of the sequences of a high number of molecules from the population can be used to deduce the preferred DNA-binding sequence, which can then be verified by EMSA (Chai et al., 2011). Usually, the result is not a single DNA sequence but a “consensus” sequence in which defined nucleotides are more or less represented at defined positions of the target sequence. This may also give a hint about the importance of the different positions of the sequence for binding. In general, the initial rounds of SELEX are performed under nonstringent conditions, such as a high protein/ DNA ratio, whereas later rounds often include the presence of a nonspecific competitor (i.e., poly (dI/dC)) with the purpose of reducing nonspecific protein binding (Gopinath, 2007). Usually 4–18 rounds are required to complete the selection process (Tuerk and Gold, 1990; Huang et al., 1993; Grotewold et al., 1994; Nole-Wilson and Krizek, 2000) and competition assays with the different oligonucleotide populations are necessary to determine the number of rounds of SELEX sufficient for the significant enrichment of target sequences.

2.3.3  Analysis of Transcription Factor–DNA Complexes by Footprinting Assays The discovery more than 35 years ago that regulatory proteins protect the DNA sequences to which they are bound from nuclease attack has been exploited to identify cis-regulatory elements in diverse organisms (Galas and Schmitz, 1978). In footprinting or protection assays, DNA labeled at the end of one of its strands is subjected to a chemical or enzymatic modification that produces the hydrolysis of phosphodiester bonds at random positions (Tullius, 1989; Figure 2.2C). The conditions of the modification reaction are adjusted such that only one cleavage per DNA molecule is obtained on average. As a consequence, a population of labeled molecules whose size extends from the labeled end to each cleavage site is obtained. The rationale of the assay is that if a bound protein protects certain regions of the DNA, molecules from the corresponding sizes will be absent or less represented in the population if the assay is applied to a protein–DNA complex. Comparing the patterns obtained after resolving samples of protein-bound and free DNA by denaturing polyacrylamide gel electrophoresis, the regions protected by the DNA-binding protein can be deduced (Figure 2.2C). The use of a chemical sequencing reaction of the same fragment or another molecular weight marker allows determination of the sequence of the protected region with single-nucleotide resolution.

In the DNase I footprinting method, deoxyribonuclease I (DNase I) is used to carry out controlled digestion in which the DNA is digested randomly but only once per DNA molecule. Since DNase I is a large molecule, it cannot bind adjacent to a DNA-bound protein because of steric hindrance. Hence, the DNase I footprint gives a broad indication of the binding site, generally 8–10 base pairs larger than the site itself (Carey et al., 2013). When small molecules are used as cleavage reagents, more intimate contacts between nucleotides and proteins can be identified. One of the smallest reagents used in footprinting is the hydroxyl radical (Tullius, 1988; Viola and Gonzalez, 2011). Due to its high reactivity and lack of base specificity, the hydroxyl radical modifies nucleotides in a rather random fashion, irrespective of the DNA sequence, producing the cleavage of phosphodiester bonds and the elimination of nucleosides (Jain and Tullius, 2008). This lack of specificity allows the generation of cleavages at every position in the DNA molecule so that a pattern that represents molecules differing in one nucleotide from each other can be obtained, producing results of high resolution. This method then allows evaluation of the contacts established by a protein with each nucleotide of a DNA molecule in a single reaction (Tullius and Dombroski, 1986). Since only one strand of the DNA is labeled at a time, the contacts established with each nucleotide of a complementary pair can be distinguished. It should be kept in mind that, due to the nature of the modification (Balasubramanian et al., 1998), contacts not only with bases but also with the DNA backbone will be identified. Dimethyl sulfate is another reagent frequently used for footprinting (Shaw and Stewart, 1994, 2009; Tron et al., 2005). This reagent methylates G and A nucleotide bases. Cleavage of the DNA backbone at modified Gs can be achieved after treatment with piperidine, while treatment with NaOH causes cleavage at modified Gs and As. In addition, other DNA modifications can be used such as carbethoxylation and ethylation (Wissmann and Hillen, 1991; Manfield and Stockley, 2009). An alternative footprinting analysis involves modification of the target DNA before protein binding. In this case, DNA molecules with modifications that affect protein binding will not be incorporated in the protein–DNA complex. This will cause an enrichment of these molecules in the population of free DNA, while molecules with modifications that do not affect protein binding will be enriched in the population of bound DNA. Analysis of both populations in a denaturing polyacrylamide gel allows the identification of positions important for protein binding. This type of approach is called missing nucleoside analysis when the hydroxyl radical is used to modify DNA and methylation interference when dimethyl sulfate is used (Viola and Gonzalez, 2011). In the case of dimethyl sulfate, since specific positions of bases are modified (G N7, facing the major groove, and A N3,

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2.4 METHODS TO STUDY PROTEIN–DNA INTERACTIONS IN VIVO

facing the minor groove), specific protein–DNA interactions can be monitored.

2.3.4  Microarray-Based Identification of Transcription Factor Target Genes For many years, biochemical assays like those just described were used to characterize DNA–protein interactions. However, such approaches are generally laborious, not highly scalable and, although in vitro selections have permitted the sampling of a large number of potential DNA-binding sequences, the resulting sites provide only a partial view of the DNA-binding specificity of a transcription factor, as typically only the highest affinity binding sites are obtained. It is possible that lower affinity DNA sites that are functionally significant in transcriptional regulation of gene expression may not be detected (Walter et al., 1994; Amendt et al., 1999). New technologies have been developed that permit the analysis of DNAbinding sites at much higher resolution and in a more unbiased manner. Protein-binding microarrays (PBMs), based on DNA microarray technology, have been used for high-throughput characterization of the in vitro DNAbinding sequence specificity of transcription factors and other DNA-binding proteins. With this technology, it has been possible to identify sites that match the known DNA-binding motifs of transcription factors as well as new candidate regulatory sites (Bulyk et al., 1999, 2001; Mukherjee et al., 2004). Additionally, DNA-binding site data obtained from PBMs, combined with gene annotation data, comparative sequence analysis, and gene expression profiling, can be used to predict what genes are regulated by a given transcription factor. To date several groups have developed PBMs for high-throughput determination of the DNA-binding specificities of transcription factors (Mukherjee et al., 2004; Berger et al., 2006; Warren et al., 2006; Kim et al., 2009; Godoy et al., 2011). In PBM experiments, a DNA-binding protein of interest is expressed with an epitope tag that serves a dual purpose: to isolate the protein by affinity purification and to achieve its detection by means of an epitope-specific reporter, such as an antibody. Alternatively, directly labeled proteins can be used in the assay. The protein is then incubated on a double-stranded DNA microarray and, after a wash to avoid nonspecific binding, the signal intensities obtained at the different array positions are measured (Figure 2.2D). Three types of DNA molecules can be used to construct the dsDNA array: short double-stranded oligonucleotides created by primer extension, short double-stranded DNAs created using self-hairpinning oligonucleotides, or longer double-stranded DNAs resulting from PCR amplification of genomic regions (Doi et al., 2002; Bulyk et al., 2001; Mukherjee et al., 2004). Microarrays made of long PCR products have the advantage that they cover more sequence space with relatively few microarray features

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(Ren et al., 2000; Lee et al., 2002). However, the probability of calling a binding event correctly is less for a single binding site that is embedded in a long rather than a short sequence. Moreover, depending on the number and types of binding sites present within a single region, interaction may occur once or several times by one or several proteins at various degrees of affinity. For accurate information about strength and specificity, arrays that consist of short synthetic double-stranded oligonucleotides exhibit superior performance (Warren et al., 2006; Berger and Bulyk, 2009). PBM technology has several advantages over highthroughput in vitro selection methodologies. First, PBM data are more quantitative, since the signal within each spot on the microarray corresponds to numerous DNA– protein binding events. In addition, nonbinding sequences can be identified. Finally, PBMs can provide binding preference data for each DNA sequence variant present in the array (Bulyk, 2007).

2.4  METHODS TO STUDY PROTEIN–DNA INTERACTIONS IN VIVO Key to understanding transcriptional regulation by transcription factors is the identification of their direct target genes. In general, the analysis of loss- and gain-offunction mutants that present altered phenotypes often provides the first clues to transcription factor function. However, additional approaches are required to elucidate the signal transduction cascades modulated by the transcription factor. For example, induction of expression or nuclear localization of a transcription factor and measurement of gene expression, either at short timepoints after induction and/or in the presence of cycloheximide, have been used to identify putative direct target genes (Ueki et al., 2009; Abel and Theologis, 1994; Yoo et al., 2007 Menges and Murray, 2002). Several methods used to obtain information about the specific DNA fragments bound by transcription factors in vivo, including chromatin immunoprecipitation (ChIP), Dam methylase identification (DamID), and the yeast one-hybrid assay, are described in the chapter.

2.4.1  Chromatin Immunoprecipitation (ChIP) Assays Even though in vitro protein–DNA interaction assays, as EMSA or footprinting, permit the study of the DNA-binding specificities of transcription factors, these interactions may not reflect the situation in cells. Chromatin immunoprecipitation (ChIP) represents a valuable alternative to probing such interactions in vivo under physiological conditions and to estimating the density of protein at specific sites (Kuo and Allis, 1999). The

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first ChIP protocol was developed by Gilmour and Lis (1984, 1985) to monitor RNA polymerase/DNA association in E. coli and Drosophila. Basically, cells in the ChIP assay are treated with a crosslinking agent to covalently bind any DNA-binding protein to the chromatin. Then, the cells are lysed and the genomic DNA is isolated and sonicated to produce sheared chromatin. An antibody specific to the protein of interest is added to the sonicated material and used to isolate the protein with all attached DNA via immunoprecipitation (Figure 2.3A). The DNA is released by reversing the crosslinking and subsequently purified. Finally, to evaluate whether a suspected target gene is associated with a particular DNA-binding protein, a test for enrichment of the target fragment in the immuneprecipitate compared with controls is performed employing semiquantitative or quantitative PCR. Formaldehyde is most commonly used to crosslink DNA-associated proteins to DNA in ChIP experiments because it offers several advantages: it easily enters cells and, unlike UV irradiation, crosslinks proteins to DNA and proteins to proteins so that it is possible to isolate DNA fragments indirectly associated to the protein of interest. In addition, the crosslinks are reversible by heat. Because formaldehyde rapidly inactivates enzymes, it provides a “snapshot” of the interactions occurring within the cell. However, formaldehyde can disrupt the epitopes necessary for immunoprecipitation, either by modifying the antigen (Das et al., 2004) or simply by denaturing the protein. For the ChIP assay, a highly specific antibody is necessary and in most cases epitope tags are engineered into the protein. A key determinant of a successful ChIP assay is the quality of the antibody, as some antibodies work poorly or not at all for ChIP. In the plant research community, commercial antibodies against HA, GFP, and FLAG tags are commonly used (Yamaguchi et al., 2014). Depending on the approach, different controls may be used. If antibodies against an endogenous protein are employed, the use of a null mutant line as control is recommended (Wu et al., 2012), while a nontransformed parental line can be used in the case of an epitope-tagged protein (Yamaguchi et al., 2013). For genomic control, a locus unlikely to be bound by the factor of interest should be tested for each experimental and control ChIP reaction. Because of anatomical differences between animal and plant cells, such as rigid cell walls, high levels of cellulose and lignin, and large vacuoles in plant cells, several modifications are needed to establish efficient ChIP protocols for plant systems (Gendrel et al., 2002; Saleh et al., 2008; Kaufmann et al., 2010). Vacuum infiltration is applied to allow the formaldehyde to penetrate plant cells and nuclei. In addition, use of young plant tissues increases the yield of nuclei per gram of fresh weight (Yamaguchi et al., 2014). ChIP experiments allow the ability of candidate proteins to bind the control region of interest in vivo to be

examined. However, it is important to keep in mind that ChIP is not a functional assay (Wang et al., 2009). ChIP is further limited by its dependence on high-quality antibodies for the protein of interest and on protein–DNA interactions that are amenable to ChIP analysis. Some proteins are difficult to study using ChIP assays for a variety of reasons, including transient binding, inefficient crosslinking to DNA or epitope masking.

2.4.2  ChIP-chip and ChIP-Seq ChIP assays can also be used to globally map the target sites of a protein in an entire genome. In this case, the population of DNA fragments obtained after immunoprecipitation is analyzed using genome microarrays or high-throughput DNA sequencing (Figure 2.3A). These techniques are called ChIP-chip and ChIP-Seq, respectively (Wu et al., 2006). In ChIP-chip, the inmunoprecipitated DNA fragments are labeled in an amplification reaction and subsequently hybridized to DNA microarrays to identify the protein-bound fragments. These microarrays can contain PCR-amplified genomic regions or an array of oligonucleotides designed to tile a portion of the genome (“tiling arrays”), allowing the definition of the genomic binding sites of a given protein (Winter et al., 2011; Yamaguchi et al., 2013). ChIP-Seq is a more recent alternative that uses next-generation DNA sequencing for identification of genomic sites enriched in the population of bound DNA fragments (Mardis, 2007).

2.4.3  DNA Adenine Methyltransferase Identification (DamID) Another recently developed method, alternative to ChIP, utilizes the DNA adenine methyltransferase (Dam) from E. coli to allow the identification of in vivo binding sites of transcription factors (van Steensel and Henikoff, 2000). In the DamID assay, the target protein is expressed as a fusion molecule to the Dam that methylates adenine at the A N 6 position within the GATC DNA sequence (Brooks et al., 1983). Therefore, when the protein under study binds its native genomic binding sites, the Dam portion of the fusion will methylate GATC sites that are located in the vicinity of the bound region (Figure 2.3B). After genomic DNA extraction, the methylated sites are digested by means of the methyl-specific restriction enzyme DpnI, which cuts only at methylated GATC sites. The smaller DpnI digestion fragments are specifically amplified using a methylation-specific PCR protocol, labeled, and hybridized in a microarray in the same way as for a ChIP-chip assay (Vogel et al., 2007; Figure 2.3B). This method takes advantage of the absence of adenine methylation in eukaryotes and of a suitable GATC motif distribution in eukaryotic genomes (one site every 200–400 bp on average). This approach has been

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2.4 METHODS TO STUDY PROTEIN–DNA INTERACTIONS IN VIVO

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FIGURE 2.3  Methods to study protein–DNA interactions in vivo. (A) Chromatin immunoprecipitation (ChIP). Plant cells expressing the transcription factor of interest are treated with a crosslinking agent that covalently links DNA-binding proteins to DNA. Once purified, the genomic DNA is sonicated to produce fragments of 100–300 bp (1). The protein–DNA complexes corresponding to the protein of interest are specifically purified through immunoprecipitation using an antibody against the protein (2, 3). Finally, the crosslinks are reversed and the genomic fragments purified (4). Purified DNA can be analyzed by PCR or qPCR to confirm the hypothetical targets of the protein or by microarray (ChIP-chip) or sequencing (ChIP-Seq) for a genome-wide analysis (5). (B) DNA adenine methyltransferase identification (DamID). The DNA-binding protein of interest (TF, orange oval) is expressed and linked to the Dam methyltransferase from E. coli (violet oval); the Dam enzyme methylates the adenine of GATC sites present in the neighborhood of the target genes of the transcription factor (1). However, more distant sites are not modified. Upon isolation of genomic DNA (2), digestion using DpnI, which only cuts methylated GATCs, is performed (3). Then, adapters are ligated to the ends of cleaved GATCs for PCR amplification, labeling and hybridization in a microarray (4). (C) Yeast one-hybrid assay. The transcription factor under study or a protein from a cDNA library (pink oval) is expressed in yeast fused to the activation domain of GAL4 (AD, green oval) from plasmid pTF-AD. The yeast genome contains a fragment of a promoter of interest or specific DNA regulatory sequences (blue bar) fused to a minimal promoter (light blue bar). If the transcription factor binds to the promoter of interest, it activates transcription of a reporter gene (orange bar), which brings about a detectable signal or acts as a selectable marker. (D) Analysis of protein–DNA interactions in plant cells. A construct that overexpresses the transcription factor of interest (pink) is transiently introduced in plants along with a plasmid that contains the promoter of the putative target gene in front of a reporter gene. Moreover, a regulatory region of interest fused to a minimal promoter can be assayed. The expression levels of the reporter gene are indicative of the existence of protein–DNA interactions in vivo. A.  General aspects of plant transcription factors

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2.  Methods to Study Transcription Factor Structure and Function

used to identify in vivo binding sites in Drosophila (van Steensel et al., 2001) and Arabidopsis (Tompa et al., 2002). Unlike ChIP, DamID has the advantage that it does not require high-quality antibodies or the use of crosslinking reagents, eliminating the risk of artifacts (Lee et al., 2006). However, some proteins lose their genomic binding specificity when fused to Dam, and this method is less suitable for detecting rapid changes in protein binding, because the methylation patterns obtained in a typical DamID experiment represent the average of a time period of about 24 h or more (Helwa and Hoheisel, 2010). So, even if performed side-by-side, DamID and ChIP experiments suggest that both methods give overlapping data (Moorman et al., 2006; Tolhuis et al., 2006; Negre et al., 2006). The appropriate technique can be chosen according to specific advantages and constraints, the model system, and the protein of interest (Germann and Gaudin, 2011).

2.4.4  Identification of Transcription Factors Using the Yeast One-Hybrid Assay A simple and powerful method to identify and isolate transcription factors that can interact with the specific regulatory DNA sequence of interest is the one-hybrid screening of yeast (Y1H) (Li and Herskowitz, 1993). This system is a variant of the yeast two-hybrid system (Y2H) (Fields and Song, 1989), which will be described later. In the Y1H method, a DNA segment of interest (either a promoter or tandem copies of a putative transcription regulatory element) are cloned in a reporter vector upstream of a minimal yeast promoter and followed by the reporter/selection gene. The reporter vector is introduced into the yeast genome via homologous recombination, and the resulting reporter strain is then transformed with a cDNA library that expresses fusions to a constitutive yeast activation domain (Figure 2.3C). Proteins able to recognize the specific DNA elements present in the reporter construct will activate the expression of the reporter/ selection gene, thus allowing the identification of cDNA clones that encode the respective DNA-binding proteins to be identified (Sieweke, 2000). The potential to screen several million independent colonies simultaneously makes the Y1H system quick and extremely sensitive, allowing the cloning and identification of very low abundance transcription factors. In the Y1H system, known cis-elements as well as noncharacterized DNA fragments of promoters can be used to search for interacting DNA-binding proteins present in the cDNA library (Ouwerkerk and Meijer, 2001). In addition, there is a high level of conservation of ciselements across species, and cis-elements from wellcharacterized plants such as Arabidopsis and rice can often be used successfully to isolate transcription factors from other species for which neither genomic nor EST data are yet available (Reece-Hoyes and Walhout, 2012). Recently,

Ji et al. (2014) developed a protein–DNA method, termed TF-centered Y1H, to identify the motifs recognized by a defined transcription factor. Unlike classic Y1H, this system utilizes a random short DNA sequence insertion library as prey and a transcription factor as the bait. With this method, these authors revealed novel DNA motifs specifically bound by bZip proteins from Arabidopsis.

2.4.5  Transient Assays to Analyze Protein– DNA Interactions In Vivo As described earlier, identifying DNA-binding proteins that interact with a control region of interest is relatively simple. However, establishing definitively that a specific transcription factor directly regulates a target gene by binding to a defined element can be a difficult task. Hence, the hypothesis that an interaction is relevant can be tested by several different experiments of in vivo protein–DNA interaction, but it must be remembered that none of these experiments by themselves are conclusive. The information gained from each approach will contribute to determining whether and how the transcription factor in study regulates a subset of genes. To clarify whether a transcription factor activates a target gene, transient expression systems with reporter genes have been widely used (Figure 2.3D). There are many different transient expression methods that can be used in plants, like Agrobacterium infiltration, particle bombardment, and polyethylene glycol (PEG) mediated protoplast transformation (Yang et al., 2000; Ueki et al., 2009; Yoo et al., 2007). The common transient expression approach begins with cotransformation with a vector that drives overexpression of the DNA-binding protein and a reporter plasmid harboring a reporter gene regulated by the control region of interest (Figure 2.3D). A reporter activity assay is used to monitor the effect of the expressed protein. The reporter genes luciferase (Luc) and b-glucuronidase (GUS) are commonly used (Gallagher, 1992; Ow et al., 1986). In order that transformation efficiency (which would be expected to vary among independent transformations) can be normalized, a reference plasmid harboring another reporter gene driven by a constitutive promoter is cotransformed as well (Iwata et al., 2011). If overexpression results in activation of the reporter gene, a new experiment with a reporter plasmid containing a mutant-binding site can be used to demonstrate the requirement of the binding motif for the protein. A positive result with this type of experiment suggests that the DNA-binding protein can activate a gene under the control region of interest when both the DNA-binding protein and the control region are overproduced within the cell. However, it provides little evidence that the protein, when expressed at physiological concentrations, can regulate the endogenous target gene. Nevertheless, the hypothesis can be strengthened greatly by subjecting it to as many rigorous tests as possible, and

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2.5  Analysis of protein–protein interactions

after considering the data obtained from different assays it can be possible to establish whether a specific protein directly regulates a target gene or not. Another strategy for testing the relevance of a protein–DNA interaction is an altered specificity experiment (Carey et al., 2012). Although difficult to design and perform, this method has the potential to provide more compelling evidence that a protein–DNA interaction is relevant. In the altered specificity experiment, the DNAbinding domain of the protein of interest is mutated so that it recognizes a different DNA sequence. On the other hand, this new sequence recognized by the altered protein is then inserted into the control region of interest in place of the sequence element recognized by the wild-type protein. The altered specificity DNA-binding protein is then expressed in cells containing an endogenous gene or reporter gene regulated by the altered control region, and its capacity to regulate transcription is monitored (Shah et al., 1997). The altered specificity strategy can provide compelling evidence that a DNA-binding protein acts at a particular target site.

2.5  ANALYSIS OF PROTEIN–PROTEIN INTERACTIONS The vast majority of proteins do not operate alone but in complexes. Protein–protein interactions are important for coordinating cellular signaling events as well as metabolic functions in any cell. In the case of transcription factors, these proteins modulate transcription by interacting with other proteins, and their activities, DNA-binding capacities and subcellular localization are usually modulated by the action of other proteins or enzymes. Knowing which other proteins physically and functionally interact with a given transcription factor is essential to gaining insight into the molecular networks by which the transcription factor fulfills its function. Numerous techniques have been developed over the years to detect and study protein–protein interactions either in vitro or in vivo. In this section, the most common methods of protein–protein complex identification and validation are summarized.

2.5.1  Methods for Protein–Protein Complex Identification A typical protein–protein interaction study usually starts with an initial screen for novel binding partners. One of the best known methods of purifying protein complexes is the tandem affinity purification (TAP) method, which allows purifying the partners of a protein of interest from a protein extract. Among the in vivo methods, the yeast two-hybrid assay has become immensely popular because it enables interacting proteins from a cDNA library to be identified. A number of in vivo assays are variations on

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the theme of fragment complementation (Hu et al., 2002; Subramaniam et al., 2001; Spotts et al., 2002). 2.5.1.1 Yeast Two-Hybrid Assay (Y2H) For years, Y2H (Fields and Song, 1989) has been the predominant method to identify protein interactions. The Y2H assay relies on the modular nature of transcription activators, which consist of a DNA-binding domain (BD) and a transcriptional activation domain (AD) (Brent and Ptashne, 1985; Hope and Struhl, 1986; Keegan et al., 1986). The DNA-binding domain serves to target the activator behind the promoter of target genes and the activation domain contacts other proteins of the transcriptional machinery to enable transcription to occur. The two-hybrid system is based on the observation that the two domains of a yeast activator (e.g., GAL4) need not be covalently linked and can be brought together by the interaction of any two proteins. To analyze the interaction, two proteins X and Y, which are hybrid proteins where the GAL4 BD is fused to protein X and the GAL4 AD is fused to protein Y, must be constructed. In addition, these hybrids contain a yeast nuclear localization signal to carry the proteins into the nucleus. These two hybrids are expressed in yeast cells containing one or more reporter genes whose promoters contain elements recognized specifically by the BD of GAL4. If the X and Y proteins interact, they create a functional activator, by bringing the AD into close proximity with the BD, which is able to recognize and bring about expression of the reporter gene (Fields, 2009; Figure 2.4A). The two-hybrid system can be used to screen libraries of AD hybrids to identify proteins that bind to a protein of interest (de Folter and Immink, 2011). These screens result in the immediate availability of the cloned gene for any new protein identified. In addition, since multiple clones that encode overlapping regions of a protein are often identified, the minimal domain for interaction may be readily apparent from the initial screen (Iwabuchi et al., 1993; Vojtek, 1993). Several Y2H versions exist that differ in the respective protein fragments used as activation and DNA-binding domains or reporter genes. As reporter genes, E. coli lacZ (Fields and Song, 1989) and selectable yeast genes such as HIS3 (Durfee et al., 1993) and LEU2 (Zervos et al., 1993) are employed. However, even this elegant and powerful technique has its drawbacks. For example, the interaction is tested only in the nucleus of a yeast cell. Moreover, interactions that depend on cellular compartmentalization or on more than two interaction partners are not always detectable by this method. Proteins must be able to fold and exist stably in yeast cells and to retain activity as fusion proteins. The use of protein fusions also means that the site of interaction may be occluded by one of the transcription factor domains. Interactions dependent on a posttranslational modification that does not occur in yeast cells will not be

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2.5  Analysis of protein–protein interactions

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 FIGURE 2.4 

Methods to analyze protein–protein interactions. (A) Yeast two-hybrid assay (Y2H). A yeast strain containing a reporter gene with a promoter recognized specifically by the yeast transcription factor GAL4 is transformed with two plasmids each encoding the proteins under study, TFX and TFY, fused to the binding domain (BD; pink) or the activation domain (AD; green) of GAL4. If a physical interaction between TFX and TFY occurs, the functionality of GAL4 is restored and the reporter gene is expressed. (B) Tandem affinity purification (TAP) for the purification of protein complexes from plants. The bait protein is expressed fused to two tags (e.g., protein A, green hexagon, and calmodulin-binding peptide, blue bar), separated by a protease cleavage site for the tobacco etch virus (TEV) protease. In the first round of purification, complexes are isolated through binding of the protein–A tag to an immobilized immunoglobulin G (IgG) and eluted by addition of the TEV protease. In the second round of purification, the complexes are isolated through binding of the remaining tag to a second affinity column. After elution, the proteins are digested and the peptides obtained are analyzed by mass spectrometry for protein identification. (C) Bimolecular fluorescence complementation (BiFC). The protein of interest (X, pink oval) and the potential partner (Y, green rectangle) are expressed fused to inactive N-terminal and C-terminal fragments of the yellow fluorescent protein (YFP). The interaction between X and Y brings the fragments together and reconstitutes the native fluorescent protein. If the proteins do not interact, the fluorescent signal is not detected. (D, E) In the fluorescence resonance energy transfer (FRET) (D) and the bioluminiscence resonance energy transfer (BRET) (E) methods a donor/acceptor pair of proteins with overlapping emission/absorption spectra is employed. If the proteins under study (X and Y) interact, the donor molecule transmits energy to the acceptor molecule and the emission energy of the acceptor is detected. In FRET (D) one of the proteins is fused to a fluorescent donor molecule (CFP, cyan cylinder) while in BRET (E) it is fused to the luciferase RLUC (blue cylinder) which emits luminescence in the presence of the substrate coelenterazine. The other candidate protein is fused to the acceptor fluorophore (YFP, yellow cylinder).

detected. Many proteins will activate transcription when fused to a DNA-binding domain, and this activation prevents a library screen from being performed. However, it is often possible to delete a small region of a protein that activates transcription and hence remove the activation function while retaining other properties of the protein. Finally, the yeast two-hybrid assay is based on reporter gene expression as an indirect readout. Despite this, several systematic and stringently controlled experimental and bioinformatics studies have demonstrated that Y2H can produce data of excellent quality, often surpassing that of other assays (Braun et al., 2009; Huang and Bader, 2009; Venkatesan et al., 2009; Yu et al., 2008, 2011). Especially important is the elimination of both constitutive and spontaneous autoactivators – usually constructs containing the DNA-binding domain that activate the reporter in the absence of an interaction – by fusing the cDNA library to the AD. The Y2H approach in plants has been summarized in several excellent reviews (Uhrig, 2006; Lalonde et al., 2008; Zhang et al., 2010). 2.5.1.2 Tandem and One-step Tag-Based Affinity Purification While Y2H screening is the most utilized strategy to identify the partners of a protein of interest (Parrish et al., 2006; Tardif et al., 2007; Suter et al., 2008; Yu et al., 2008; Bonetta, 2010; Arabidopsis Interactome Mapping Consortium, 2011; Seo et al., 2011; Vernoux et al., 2011), this method is not applicable to the study of large multimeric protein complexes. One of the best-known methods for the isolation of protein complexes is TAP (Rohila et al., 2006; Van Leene et al., 2008; Rubio et al., 2005; Schoonheim et al., 2007). With this method, multicomponent protein complexes can be isolated from a cell lysate through affinity purification steps and then analyzed by mass spectrometry (MS) to identify purified proteins. The basic concept of TAP is the use of a so-called TAP tag, which is fused to a bait protein. The initial version of the TAP tag consisted

of two sequential affinity tags separated by a tobacco etch virus (TEV) protease cleavage site that enabled two consecutive purification steps and reduced the amount of nonspecific binding (Figure 2.4B). Once expressed in vivo (stable or transient) the TAP-tagged protein associates to its endogenous targets and, after lysis of the cells, the TAP-tagged protein is allowed to bind via the first part of the TAP tag (e.g., protein A) to a specific column. Then the TEV protease is added and the TAP-tagged protein is cleaved, leaving the first affinity tag on the column. The bait protein, still fused to the second part of the TAP tag (e.g., a calmodulin-binding peptide) is then bound to a second column (e.g., calmodulin-coated beads), which is rinsed and eluted (e.g., by a buffer-containing EDTA) (Figure 2.4B). A platform with the GS tag, which combines two IgG-binding domains of protein G with a streptavidin-binding peptide separated by two tobacco etch virus cleavage sites, was developed to analyze protein complexes in Arabidopsis thaliana cell suspension cultures. This GS tag outperforms the traditional TAP tag in plant cells, regarding both specificity and complex yield (Van Leene et al., 2008). In addition to the possibility of isolating multimeric complexes, TAP has the advantage that complex formation occurs in vivo (in planta) and that posttranslationally modified proteins (e.g., phosphorylated, glycosylated, or oxidized/reduced proteins) are present. Even if TAP is highly sensitive and selective (Rohila et al., 2006; Rubio et al., 2005; Chang et al., 2009; Braun et al., 2013), weak protein–protein interactions may be lost during the purification steps. Another problem is that a relatively large amount of starting material is required, which can make purification and identification of low-abundance binding partners of transcription factors a difficult task. Nowadays, many affinity tags are available for affinity purification followed by mass spectrometry methods (such as affinity purification and mass spectrometry, AP-MS) based on a single purification step. These systems

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employ the biotin peptide (Bio), the Flag epitope, c-Myc, His, HA, or the green fluorescent protein (GFP) as a tag (Fukao, 2012). In addition to GFP and its variants such as YFP and CFP (Cristea et al. 2005), magnetic beads conjugated to an anti-RFP antibody also work well for AP-MS analysis (Fukao, 2012). The interacting proteins detected by the fluorescent protein tag purification method can be confirmed by means of colocalization experiments in transgenic plants expressing fluorescent fusion proteins (see in later sections).

2.5.2  Methods for Verification of Protein– Protein Interactions One of the major challenges when working with protein–protein interactions is to distinguish specific from unspecific binding. For example, MS-based protein identification has become so sensitive that any protein–protein interaction screen will result in a large number of identified contaminant proteins. Therefore, the general consensus is to confirm protein–protein interaction data using one or more independent approaches to arrive at an accurate evaluation. In some cases, the original method of screening can be used. Ideally, in vivo confirmation of the interaction is necessary. To do this, coimmunoprecipitation, fluorescence resonance energy transfer (FRET) and bimolecular fluorescence complementation (BiFC) methodologies can be used. 2.5.2.1 Coimmunoprecipitation Coimmunoprecipitation is a technique in which a protein complex is purified from a cell lysate using an immobilized antibody against a known component of the complex and the interacting partners are analyzed by western blotting or MS (Ransone, 1995; Masters, 2004; Backstrom et al., 2007). Several strategies can be followed, using an antibody against an endogenous protein or a tagged fusion protein either transiently or stably expressed. Moreover, protein extracts can be incubated with recombinant bait proteins to trap interacting proteins (Swatek et al., 2011). As coimmunoprecipitation usually generates a significant background signal, it is important to perform appropriate negative controls (Ransone, 1995). 2.5.2.2  In Vivo Split Methods In split methods, also called protein fragment complementation assays (PCA), the two proteins to be tested for interaction are fused to two fragments of a reporter protein, neither of which by itself has reporter activity. If the two proteins interact, they may bring the two split components together such that they can fold and the functionality of the reporter protein is restored. Among these reporters, the yellow fluorescent protein (YFP) and its variants have been commonly used (Figure 2.4C). This

PCA method is referred to as bimolecular fluorescence complementation (BiFC; Kerppola, 2008). In BiFC, when two nonfluorescent fragments of an otherwise fluorescent protein are brought into close proximity by the interaction of two proteins fused to the fragments, complex formation can be visualized directly in a living cell by epifluorescence or confocal microscopy (Bhat et al., 2006; Ohad et al., 2007; Weinthal and Tzfira, 2009). Simultaneously, the subcellular location of the protein complex in the cell can be determined (Ohad et al., 2007; Weinthal and Tzfira, 2009). The BiFC assay can be performed by means of transient or stable expression of the fusion proteins (Waadt et al., 2014). Several binary plasmids for plant transformation have been developed for regular or gateway-based cloning (Gehl et al., 2009; Martin et al., 2009; Citovsky et al., 2008; Walter et al., 2004). Additionally, a series of multicolor BiFC (mcBiFC) vectors have been developed using spectral variants of the fluorescent proteins, which allows the interaction between a given “bait” protein and multiple “prey” proteins in living plant cells to be studied simultaneously (Lee et al., 2008; Waadt et al., 2008). Another split reporter that can be used is luciferase. The luciferase split assay was first described in mammalian cells (Luker et al., 2004) and later adapted to plants (Fujikawa and Kato, 2007; Chen et al., 2008). It is particularly useful in plants because the autofluorescence of photosynthetic pigments is usually a problem in fluorescencebased assays. Split luciferase was shown to be the most sensitive protein–protein detection method; it is also able to detect protein dissociation (Li et al., 2011). 2.5.2.3  Resonance Energy Transfer Methods Energy transfer techniques are additional strategies for assaying protein–protein interactions in living cells. The basic principle of these methods is that when the physical distance between two fluorescent proteins is small (<100 Å), energy can be passed from one to the other by a phenomenon called resonance energy transfer. In the case of the FRET method (Bhat et al., 2006), two fluorophores with overlapping emission/absorption spectra linked to proteins that might interact with each other are employed (Figure 2.4D). If the proteins interact and transition dipoles are appropriately oriented, the donor fluorophore is able to transfer its excited-state energy to the acceptor fluorophore. This energy transfer leads to a decrease in the donor’s fluorescence intensity and a decreased lifetime of the excited state. As the acceptor molecule is a fluorophore, an increase in the acceptor’s emission intensity is manifested. The efficiency of the resonance transfer depends upon the spectral overlap of the fluorophores, their relative orientation, as well as the distance between the donor and acceptor fluorophores (Pollok and Heim, 1999). Ideally, the acceptor should exhibit minimal excitation at the wavelength used to excite the donor fluorophore. Chromophore-mutated

A.  General aspects of plant transcription factors

REFERENCES

green fluorescent proteins (GFPs) with an excellent spectral overlap have been widely used in FRET studies as CFP/YFP and GFP/RFP variants (Pollok and Heim, 1999; Goedhart et al., 2007). The choice of fluorophores is a very important aspect in FRET. A comprehensive review article about this topic has been published (Shaner et al., 2005). On the other hand, the bioluminescence resonance energy transfer (BRET) method (Xu et al. 1999) uses a bioluminescent luciferase (RLUC), as a donor protein, which is genetically fused to one candidate protein, and a green fluorescent protein mutant (YFP) fused to the other protein of interest (Subramanian et al., 2004, Xu et al., 2007; Figure 2.4E). Interactions between the two fusion proteins can bring the luciferase and the fluorescent protein close enough for resonance energy transfer to occur, thus changing the color of the bioluminescent emission (Pfleger and Eidne, 2006). Both FRET and BRET are nondestructive in vivo assays that allow real time measurements of protein–protein interactions. Even if FRET can be used to detect the molecular interactions between two proteins, it has been suggested as a method to analyze conformational alterations within a single polypeptide (Van Roessel and Brand, 2002; Janetopoulos et al., 2001). However, FRET has the disadvantage that it requires an excitation light source that may cause photobleaching or phototoxicity, as well as autofluorescence of plant pigments and unintended biological effects owing to cellular photoreceptors. Second, not only the photon donor (often CFP), but even the photon acceptor (often YFP) may be partially activated by the excitation light source, contributing to background signal (Bhat et al., 2006). In this sense, BRET does not need an excitation light source and is measured against a background of complete darkness. Thus, all photons emitted by the YFP acceptor originate from the luciferase and are indicative of BRET. BRET, on the other hand, requires a substrate. In the case of RLUC, this substrate is coelenterazine, which is nontoxic and membrane permeable (Subramanian et al., 2004). A number of considerations must be taken into account before starting any fluorophore-based in planta protein– protein interaction assay (as FRET, BRET, or BiFC). First, these methods utilize tagged variants of the proteins of interest that can present alterations in their physiological parameters (such as subcellular localization, stability, and biological activity; Bhat et al., 2006). Thus, wherever possible, complementation of mutant phenotypes or analysis of protein activities in vitro must be carried out. Since proteins can be tagged either N- or C-terminally, all possible pairwise combinations should be tested when performing FRET or BiFC assays (Frank et al., 2005). Second, the type of promoter utilized to express the fusion proteins must be considered. Although expression is frequently driven by strong constitutive promoters, native gene promoters should be used to avoid possible artifacts that may either

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promote or inhibit the protein–protein interactions. Wherever possible, expression should take place in respective double-null mutants, since endogenous untagged copies of the proteins may interfere with the interaction assay. In summary, transgenic lines expressing both tagged proteins under control of their own promoters against a respective double-mutant genetic background must be used, whenever possible. It is important to note that these methods determine the existence of a close physical proximity between the two tagged fusion proteins, but do not constitute proof of a true protein–protein interaction. To state this, evidence of direct interaction by in vitro assays using recombinant proteins or by Y2H is required. An interesting way to validate in vivo assays is by using mutant versions of the proteins in residues critical for the interaction as a control. If the loss of the interaction is coincident with an altered plant phenotype and the native proteins colocalize, then the biological relevance of the protein complex is demonstrated.

Acknowledgments The authors acknowledge support from Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET, Argentina), Agencia Nacional de Promoción Científica y Tecnológica (ANPCyT, Argentina), and Universidad Nacional del Litoral.

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