dna–protein interactions- methods for detection and analysis
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DNA–protein interactions: methods for detection and analysis
Bipasha Dey • Sameer Thukral • Shruti Krishnan •
Mainak Chakrobarty • Sahil Gupta •
Chanchal Manghani • Vibha Rani
Received: 24 September 2011 / Accepted: 16 February 2012 / Published online: 8 March 2012
� Springer Science+Business Media, LLC. 2012
Abstract DNA-binding proteins control various cellular
processes such as recombination, replication and transcrip-
tion. This review is aimed to summarize some of the most
commonly used techniques to determine DNA–protein
interactions. In vitro techniques such as footprinting assays,
electrophoretic mobility shift assay, southwestern blotting,
yeast one-hybrid assay, phage display and proximity ligation
assay have been discussed. The highly versatile in vivo
techniques such as chromatin immunoprecipitation and its
variants, DNA adenine methyl transferase identification as
well as 3C and chip-loop assay have also been summarized.
In addition, some in silico tools have been reviewed to pro-
vide computational basis for determining DNA–protein
interactions. Biophysical techniques like fluorescence reso-
nance energy transfer (FRET) techniques, FRET–FLIM,
circular dichroism, atomic force microscopy, nuclear mag-
netic resonance, surface plasmon resonance, etc. have also
been highlighted.
Keywords DNA–protein interactions � Footprinting �Electrophoretic mobility shift assay � Southwestern
blotting � Phage display � Yeast one-hybrid assay �Chromatin immunoprecipitation assay �Biophysical techniques
Introduction
Association of DNA with proteins is a phenomenon of utmost
importance. In effect, almost all aspects of cellular function,
such as transcriptional regulation, chromosome maintenance,
replication and DNA repair depend on the interaction of
proteins with DNA. Activation of genes by DNA-binding
proteins is a fundamental regulatory mechanism involving the
chromatin modifying and transcription complexes to initiate
the RNA synthesis [1]. Such DNA-binding proteins have
diverse roles and may function as structural proteins making
up the nucleosome, enzymes modulating chromatin structure
to control gene expression, transcription factors, and also as
cofactors. One of the most widely studied examples of DNA-
binding proteins is the transcription factor. TFs association
with DNA is considered to be extremely critical in develop-
ment processes and in response to environmental stresses.
Also, in humans their dysfunction can contribute to the pro-
gression of various diseases [2].
In view of such an important role played by DNA–
protein interactions, various techniques have evolved over
the years to elucidate them. Each technique, with its own
advantages and drawbacks, serves a very specific purpose.
In brief, the techniques cater either of the two parts of the
interaction: protein (molecular weight, identity, domains
etc.) or DNA (general sequence, specific sequence, alter-
native sequences etc.).
This review has been focused to aptly summarize some
of the most important in vitro, in vivo, in silico and bio-
physical techniques to study DNA–protein interactions,
owing to the pivotal role played by DNA-associating pro-
teins in various cellular processes. The review shall assist a
researcher to understand and evaluate various DNA–pro-
tein interaction techniques and use them appropriately for
their research.
All the authors have contributed equally.
B. Dey � S. Thukral � S. Krishnan � M. Chakrobarty �S. Gupta � C. Manghani � V. Rani (&)
Department of Biotechnology, Jaypee Institute of Information
Technology, A-10 Sector-62, Noida 201307,
Uttar Pradesh, India
e-mail: vibha.rani@jiit.ac.in
123
Mol Cell Biochem (2012) 365:279–299
DOI 10.1007/s11010-012-1269-z
In vitro techniques to study DNA–protein interactions
There are several techniques to determine the in vitro
DNA–protein interactions experimentally. Some of the
well known in vitro techniques are footprinting assay,
southwestern assay, electrophoretic mobility shift assay,
yeast one-hybrid assay, phage display and proximity liga-
tion assay.
Footprinting assay
Foot printing assays are based on the principle of protec-
tion of protein-bound DNA from degradation. The tech-
nique is used to decipher the specific sequence to which a
DNA-binding protein or molecule binds. The procedure
employs chemical or enzymatic digestion of naked- and
protein bound-DNA oligomers. Both the reactions are then
compared using gel electrophoresis. The segment of the
DNA bound by the protein appears as an empty stretch
‘footprint’ in the protein-bound reaction when compared to
the continuous fragments produced by naked DNA diges-
tion (Fig. 1a).
Foot printing has been a valuable tool for elucidating
sequence specificity and dissociation constants of a variety
of ligands binding to DNA. The agent used to cleave DNA
is called the probe. The smaller the probe, the higher is the
resolution provided, but its chances for cleaving DNA
under the bound protein also increases. The enzymatic
digestion methods include the use of DNAse I, MNase [3],
methidiumpropyl-EDTA�Fe(II) (MPE) [4, 5], copper phe-
nanthroline [6], uranyl photocleavage [7, 8], hydroxyl
radicals [9–13] and iron complexes [14]. Comparisons
between different probes used for foot printing provide
useful information on their relative merits and demerits
[15–18].
DNAase I footprinting is the most commonly used
footprinting assay. DNAase I is a double-strand-specific
endonuclease, which binds to the minor groove to break
phosphodiester bonds. The technique was developed by
Galas and Schmitz [19] for visualizing the binding of the
lac-repressor protein to the lac-operator sequence. The
footprinting technique employs use of a single end-radio-
labeled, synthetic or natural, DNA fragment. The fragment
is incubated with either crude or purified protein sample,
Fig. 1 In vitro techniques to study DNA–protein interactions
280 Mol Cell Biochem (2012) 365:279–299
123
under appropriate binding conditions, allowing the protein
to bind to its specific DNA sequence. The protein-bound
fragment and the control (i.e. naked fragment) are then
subjected to DNAse I treatment in an appropriate buffer,
with varying concentrations and time periods. Both the
samples are then run on a denaturating polyacrylamide gel,
processed and imaged [20].
There are various key points for this technique [21].
First, by altering experimental conditions, DNAse is
allowed to partially digest the fragment, assuming a single
nick per fragment. This creates a range of fragments which
differ from one another by single nucleotide, hence pro-
viding high resolution for the protein-binding sequence.
Upon examining variety of experimental footprinting gels,
it is evident that if all the sequences were cleaved without
any sequence-dependent specificity by DNAse I, all bands
would have been of similar intensity, but DNAse I has
partial sequence specificity, resulting in some sites
becoming hyper sensitive and thus showing a more intense
band. Second, end labeling of DNA is for a specific pur-
pose. In a single reaction, DNAse will cut both the strands
leading to mixture of ?ve and –ve strand fragments which
are further separated on the denaturating gel. The purpose
of radio labeling DNA is to provide a clue as to which of
the two anti-parallel strands does the protein under con-
sideration bind. Thus, in a 50 labelled reaction only the 50–30 strand sequence information is provided on the final
exposed film. Subsequently, it is common to digest both
?ve and –ve strand labeled fragments in separate tubes and
then run them along side each other. Lastly, there are a
variety of methods to analyze the final footprinting image.
These methods range from visual inspection to creating a
differential cleavage plot on the basis of densitometric
analysis. Further there are techniques available for quan-
titative analysis of binding affinity [22].
The purpose of a denaturing gel is to make sure that
fragments show up on the gel only as single-stranded DNA.
Often the naked DNA is chemically sequenced and run on
the same gel, as a marker for finding the exact sequence of
the footprint. However, some precautions must be consid-
ered. First, it is important not to titrate too much DNA with
the protein sample. This shall cause a large amount of
DNA to remain unbound from the protein and thus sus-
ceptible to DNAse I attack. This fragmented DNA shall
show up at the place of the usual footprint, making it harder
to detect. Second, incubation of crude samples with the
fragment requires the presence of competitor DNA to
exclude the non-specific binding proteins from showing a
footprint. The limitation of this technique is that it does not
provide identity of the protein [20]. Because of the large
molecular weight of DNAse I, its attack is easily sterically
hindered, by the bound protein. Other probes for nicking
DNA, like free radicals may not be hindered so easily and
hence have a chance of nicking a few bases, under the
bound protein itself.
Apart from proteins, footprinting technique is also used
to elucidate the binding of other small molecules like drugs
to DNA [22]. The modifications of this technique use
automation and capillary electrophoresis along with fluo-
rescent labeling [23, 24]. A technique using DNAse I
digestion followed by sequencing called DNAse-seq is
often used for genome-wide studies [25]. Protocols using
automated infrared sequencers, allowing long range and
highly sensitive DNAse I footprinting have been developed
[26]. Also using Streptavidin-bound oligonucleotides for
protein binding and subsequent DNAse I digestion makes
the technique more convenient [27]. The technique can
also be used to fish out a protein of interest, from a crude
sample. It is used for quantitatively measuring the disso-
ciation constant of protein–DNA interaction [28–30].
Though initially in vitro, the technique has been adopted to
be used as an in vivo foot printing assay, involving per-
meabilization of cells followed by DNAse I-mediated
cleavage and ligation-mediated PCR [31, 32]. Drug–RNA
footprinting methods have also been developed [33]. It is
now known that Gold (Au)-DNA conjugates change their
surface plasmon resonance (SPR) wavelength depending
on the length of the DNA oligo attached. Comparing the
SPR wavelength of a control protein-bound DNA–Au
conjugate, with the experimental DNAse I or any other
probe digested protein-bound DNA–Au conjugate provides
information on the exact number of nucleotides from end,
where the protein under consideration is bound. This recent
advancement provides a label-free, quantitative, real-time
measurement of nuclease activity and footprint of a bound
protein without running a gel [34].
Electrophoretic mobility shift assay (EMSA)
EMSA is a relatively simple in vitro technique to study
DNA–protein interactions. Its novelty lies in its application
to deduce the binding parameters and relative affinities of a
protein for one or more DNA sites or for detecting protein–
nucleic acid interactions with the aim of comparing the
affinities of different proteins for the same sites [35]. It is
based on the principle that DNA–protein complexes are
heavier and move slowly when subjected to non-denaturing
polyacrylamide or agarose gel electrophoresis as compared
to unbound free probe. Since, the rate of DNA migration is
shifted or retarded when bound to protein, the assay is also
referred to as a gel shift or gel retardation assay. The DNA
sequence is provided externally and incubated to crude cell
protein lysate. Following this, the DNA and extracted
proteins are incubated together in a binding reaction and
separated on a gel. The DNA probes used may be radio-
labeled or dyes specific to stain DNA and protein may be
Mol Cell Biochem (2012) 365:279–299 281
123
used to visualize the DNA–protein interaction. In general
poly (dI-dC) is added to abolish any non-specific binding.
A supershift assay can be performed to specifically assert
the DNA–protein interactions by using an antibody specific
to the protein of interest. By incubating antibody along
with DNA–protein sample followed by gel separation, the
DNA–protein–antibody complex can be visualized as a
supershifted band. Competition assays may also be per-
formed using unlabeled specific and non-specific oligo
duplexes (Fig. 1b).
EMSA can be used qualitatively to identify sequence-
specific DNA-binding proteins in crude lysates and, in
conjunction with mutagenesis, to identify the important
binding sequences within a given gene upstream regulatory
region. EMSA can also be utilized quantitatively to mea-
sure thermodynamic and kinetic parameters. This tech-
nique poses several advantages. The most significant
benefit of EMSA is its ability to resolve complexes of
different stoichiometry or conformation. Another major
advantage is that the source of the DNA-binding protein
may be crude nuclear or whole cell extract, in vitro tran-
scription product or a purified preparation. In addition, the
relatively low ionic strength of the electrophoresis buffer
helps to stabilize transient interactions, permitting even
labile complexes to be resolved and analyzed by this
method [36–39].
An additional variation to the conventionally used
EMSA is capillary electrophoretic mobility shift assay
(CEMSA) which allows the rapid separation and quan-
titation of DNA–protein interactions, in uncoated capil-
laries with no gel matrixes, using high-sensitivity laser-
induced-fluorescence detection of fluorescein-labeled
DNA. Capillary electrophoresis (CE) separates analytes
on the basis of their mass-to-charge ratio and elutes
complexes in the order of free protein, protein/DNA
complex, and lastly DNA [40]. A rapid and quantitative
procedure has also been developed that permit accurate
assessment of specific DNA–protein interactions on a
scale more than 100-fold, below the minimum signal
necessary for EMSA by using a laser-induced fluores-
cence detection system [41].
IDEMSA is another modification of EMSA that
combines immunodepletion with the traditional EMSA
and supershift assays. In this, the nuclear or cytoplasmic
extracts are depleted of the specific protein by incubation
with the relevant antibody and protein A-sepharose. The
depleted extracts are then analyzed for the presence of
protein by the EMSA and supershift assay. This tech-
nique poses the advantage of combining results of im-
munodepletion and supershift to determine the protein
composition of a particular protein–DNA complex and
also the localization of the dimer to a specific complex
[42].
Southwestern blotting
This technique combines the principles of southern and
western blotting and is primarily used for elucidating the
molecular weight of the protein in a protein–DNA com-
plex. Though a super shift assay, an extension of an EMSA
experiment, provides more information on the nature and
hence the molecular weight of the protein, often there are
no antibodies known for the bound protein. Thus, in cases
where no preliminary knowledge of the DNA-binding
protein is available, southwestern blotting provides at least
some minimal information like molecular weight.
The experimental procedure involves, a modified wes-
tern blot using labeled oligonucleotides instead of anti-
bodies as probes. In brief, the crude or purified
cytoplasmic/nucleic/whole cell extract containing the pro-
tein of interest, is resolved on an SDS-PAGE, followed by
electrophoretic transfer of the proteins from the gel to a
membrane under conditions favouring renaturation of the
proteins. The membrane-bound proteins are then incubated
with oligonucleotides to which the protein of interest
putatively binds. The membrane is developed, photo-
graphed and only the band corresponding to the bound
oligo appears in the final picture (Fig. 1c). Aligning the
band on the developed picture with the SDS-PAGE posi-
tion of the protein at that band, marks the protein bound to
the oligo and provides information about its molecular
weight [43–46]. The SDS-PAGE provides the information
of the molecular weight, while the blotting allows the
protein to bind to the sequence. The labelling is required to
mark the spot of the bound protein–DNA complex [47].
A 2-D gel electrophoresis, instead of SDS-PAGE and
on-blot digestion of the DNA-bound protein followed by
LC–MS/MS, analysis provides better information about the
molecular weight of protein [48, 49]. Non-radioactive
methods for southern blotting make the procedure less
cumbersome [50, 51]. Moreover, using differently labelled
oligos on the same blot would provide information on the
binding affinity of various mutants of the oligo. The same
blot is probed with different probes by using alkaline
phosphatase to strip the signal of the bound probe [52]. A
further modification uses the southwestern blot itself as a
substrate for nuclease footprinting or other types of foot-
printing like chemical nuclease and methylation protection,
thus identifying the exact DNA sequence where the protein
binds [53]. To differentiate the specific from the non-spe-
cific binding on the blot, a rapid dimethylsulphate (DMS)
protection assay has been developed, which distinguishes
between them on the basis of conditions that specific
binding creates, making the complex impervious to DMS
[54]. Though southwestern blotting is primarily a technique
for knowing the molecular weight of protein binding to a
known DNA sequence, it can also be used to find the
282 Mol Cell Biochem (2012) 365:279–299
123
sequence of DNA that a particular protein binds to [55].
While screening various oligos, caution is advised for
cDNA expression libraries screened by southwestern
methodologies [56]. Southwestern histochemistry is also an
important modification, allowing in situ identification and
localization of DNA-binding proteins. It uses oligonucle-
otides instead of antibodies to probe a specific protein in a
histological sample. Incubation of the labelled oligonu-
cleotide with the crude or purified cytoplasmic/nucleic/
whole cell extract, followed by cross-linking using UV
light and subsequent resolution of the extract by gel elec-
trophoresis, is an alternative to blotting [57–59].
Apart from these modifications, another modification
would be coupling the chromatographic separation of
proteins with SDS-PAGE for each fraction. This shall
provide better information on the characteristics, purifica-
tion properties and molecular weight of the protein. Fur-
ther, this technique is restricted to blotting because
oligonucleotides cannot be directly made to penetrate a
SDS-PAGE and bind to the cognate proteins. Hence, if in
future, some protein-resolving oligonucleotide-permeable
gels are developed, the blotting procedure can be avoided
and hybridization can happen on the gel itself.
The disadvantage of this technique is that DNA-binding
proteins involving multiple subunits may get dissociated
during the SDS-PAGE step and hence evade detection.
Even the proteins which are monomers may not renature
properly on the blot to recognize their binding sequence.
Proteins requiring co-factors for DNA binding are difficult
to detect on blot, unless those specific co-factors are added
[60].
Yeast one-hybrid assay (Y1H)
The Y1H, a modification of the yeast-two hybrid assay, is a
sensitive technique for identifying and analyzing proteins
that bind to a specific DNA fragment of our interest. In
1993, Wang and Reed [61] first used the Y1H to clone the
gene encoding the olfactory neuron-specific transcription
factor OLF-1.
The concept of this assay, like the yeast-two hybrid
assay, exploits the same basic finding that most eukaryotic
transcription factors have two physically separable
domains called the activation domain (AD) and the DNA-
binding domain (DB/DBD). If these domains are separated
from each other, it results in a functionally inactive tran-
scription factor that cannot recruit RNA polymerase at its
corresponding promoter to start transcription [62].
In the yeast-two hybrid assay, which is used to study
protein–protein interactions, a protein X is translationally
fused to AD, while the other protein Y is translationally
fused to DB and both are expressed in the same yeast cell.
The DB-Y fusion is often referred to as the ‘bait’ and the
AD-X fusion is called the ‘prey’. If X and Y interact within
the yeast cell, it brings the AD and DB in close physical
proximity to reconstitute the functionally active transcrip-
tion factor and allows the expression of a downstream
reporter gene. Thus, yeast cells expressing the reporter
gene show that the proteins X and Y interact with each
other.
On the other hand, in the one-hybrid system, the bait is
replaced by a DNA sequence of our interest and the
interaction of a protein X with the bait sequence is assayed.
If X interacts with the bait DNA sequence, it results in
bringing AD-X fusion close to the promoter, allowing AD
to activate the RNA polymerase and result in the expres-
sion of the downstream reporter gene. While AD recruits
RNA polymerase, X plays the role of BD since the BD
fusion protein is absent here. Since this assay contains only
one-hybrid encoded on a vector, it is called the ‘Y1H’. In
other words, the one-hybrid assay can be used to trap any
protein (X) having a binding-domain specific for any target
DNA sequence (Fig. 1d).
The one-hybrid assay offers maximal sensitivity because
detection of the DNA–protein interaction occurs while
proteins are in their native configurations. In addition, the
gene encoding the DNA-binding protein of interest is
immediately available after a library screening [63].
The procedure first involves the construction of a vector
carrying the bait-sequence upstream of a reporter gene
promoter. Transforming the yeast cells with this bait–
reporter construct generates a yeast reporter strain to be
used for the assay. The bait sequence and reporter gene
may remain on the vector or can be integrated into the
chromosome. The integration of the construct into the yeast
genome is preferred and is ensured using high-frequency
homologous recombination sites flanking bait–reporter
region on the vector. The transformants are screened by
marker selection and are then again transformed with the
vector encoding a DNA-binding protein fused to Gal4p
AD. The library of AD-DNA-binding protein is screened to
check for potential DNA–proteins specific to the bait
sequence, which is reported by the expression of the
reporter genes [63]. The bait sequence can either be an
artificial site having several tandem repeats of the
sequence, or it can be a partial site or a fully functional site
in situ [64].
The reporter gene most commonly used is HIS3, which
allows growth of yeast cells showing the positive interac-
tion in a medium lacking histidine. Any background noise
because of basal level or leaky expression of HIS3 is
eliminated by including a competitive inhibitor 3-amino-
1,2,4-triazole (3AT) in the medium. Hence, a higher level
of expression of HIS3 is required for survival of yeast cell
in the medium. The higher level expression is only possible
in cells showing positive interaction and hence any false
Mol Cell Biochem (2012) 365:279–299 283
123
positives are eliminated [64]. LacZ is another reporter gene
which can be used in luciferase-based assays.
There are several modifications of the yeast one-hybrid
system. First, the one-hybrid system can be used to look for
interaction-defective proteins in which the reporter gene
codes for a toxic product. Thus, a positive interaction leads
to expression of the toxin and cell death while a lack of
interaction confers cell viability. This method is referred to
as the reverse one-hybrid assay. When used to screen an
AD fusion library of random mutations in the DNA-bind-
ing proteins, it can identify mutations that lead to disrup-
tion of a DNA–protein interaction [62]. It can also be used
to detect the therapeutic potential of drugs or other small
molecules that lead to disruption of DNA–protein interac-
tions which are involved in the onset of a disease. Like the
reverse two hybrid assay, the reverse one-hybrid assay can
be titrated to generate a range of DNA–protein affinities
[65]. Second, Y1H can be modified to screen various
binding sites in a bait sequence that binds to a known
DNA-binding protein. Third, Y1H can also be used to
screen for specific epitopes on a known DNA-binding
protein.
Phage display for DNA-binding proteins
Phage display refers to the method of expressing a peptide
or protein domain on a bacteriophage capsid by genetically
fusing its amino acid sequence to that of the coat proteins
encoded by the phage. A wide variety of proteins can be
expressed in this way yielding a pool of variants referred to
as a phage-display library. The proteins of interest can then
be selected from the library by affinity purification using an
appropriate ligand. The clones with the highest affinity for
the target ligand can be enriched by sequential rounds of
selection and amplified by passage through a bacterial host.
The identity of the selected clones can be obtained by
sequencing the phage genome thus giving complete infor-
mation about the protein of interest [66, 67]. Phage display
is also applied to map the DNA–protein interactions
because of the advantage of screening a large number of
protein variants simultaneously and also giving the com-
plete sequence information of the same [68–73].
In this modified version of phage display, phages
express a DNA-binding protein domain fused to its coat
protein. Affinity purification of a pool of clones expressing
various DNA-binding protein domains is carried out using
dsDNA oligos (with the binding sequence specific to a
protein) bound to a solid matrix. Enrichment and amplifi-
cation is done as in conventional phage-display experi-
ments and depending on the type of library screened, it
results in identification of the protein domains in the DNA-
binding protein that physically interact with the DNA
(Fig. 1e).
The choice of library to be screened is dictated by the
aim of the experiment. The most common types of phage-
display libraries are the random peptide libraries (RPL)
which are obtained by randomizing the DNA sequence at a
selected region of the gene encoding the DNA-binding
protein. This can be used to check which residue in the
region is involved in the interaction or for selecting rare
clones with enhanced function, or clones in which the
displayed domain has acquired a new function as a result of
mutation. Other libraries like cDNA libraries or genomic
libraries are used to determine the DNA-binding proteins
[67]. DNA oligos are prepared by annealing complemen-
tary oligonucleotides together and by biotinylating at least
one of the strands so that they can be bound to streptavidin-
coated matrices.
The procedure firstly entails the construction of an
appropriate phage-display library of DNA-binding proteins
according to the protocol previously described [66]. DNA
oligos bound to an appropriate matrix are then incubated
with the phages. The unbound phages are removed using
several rounds of washing. The bound phages are then
eluted and amplified by passage through a bacterial host.
These affinity-purified and amplified phages are then made
to undergo another round of affinity purification and then
again amplified. Several rounds of affinity purification
followed by amplification lead to the enrichment of the
phage clones expressing the DNA-binding protein domain
with maximum affinity to the DNA of interest.
Once the clones are selected and enriched, their binding
property is assayed using phage ELISA as a final confir-
mation. In this, the streptavidin-coated microtiter plates are
first coated with the biotinylated DNA oligos. Then the
enriched and amplified phages are allowed to bind to the
wells. The unbound phages are washed off and anti-phage
antibody conjugated with an enzyme is added. After
washing off the unbound antibodies the colour developing
solution carrying the substrate is added and reaction stop-
ped after a specified time. The intensity of colour devel-
oped is measured using plate reader spectrophotometer at
450 nm. Higher intensity indicates and confirms strong
interaction between the DNA oligo and the proteins dis-
played on those phage clones.
Proximity ligation assay (PLA)
PLA is used for ultrasensitive protein analysis for mea-
suring DNA–protein interaction. In this technique, direct
detection of proteins or DNA–protein interactions is pos-
sible and DNA representations of detected proteins are
created. Following this, the amplified oligonucleotides are
attached to specific protein-binding reagents (mono/poly-
clonal antibodies). One of the proximity probes is a partly
double-stranded oligonucleotide with a single-stranded
284 Mol Cell Biochem (2012) 365:279–299
123
extension. The other probe is an antibody directed against
the DNA-binding protein, and it has an attached DNA
strand with a free 50 end [74].
When two proximity probes recognize and bind the
same target molecule or a complex of two interacting target
molecules, the ends of their conjugated oligonucleotides
are joined by enzymatic ligation, assisted by the addition of
a connector oligonucleotide. The detected protein mole-
cules thus promote the ligation reactions by ensuring suf-
ficient proximity between the ends of the proximity probes’
oligonucleotide extensions [75].
This method is highly specific and sensitive for solution-
phase analysis of interactions. Large sets of proteins can be
analyzed in parallel. This assay can be a valuable tool to
characterize sequence specificity of DNA-binding proteins
and to evaluate the effect of polymorphism in transcription
factor binding sites [76].
In vivo techniques to map DNA–protein interactions
Several in vivo techniques have been developed to char-
acterize DNA–protein interactions. The requirement of an
in vivo technique is due to limits posed by in vitro
experiments. DNA inside a cell exists in a compact chro-
matin state with distinct properties from naked DNA, and
usually in vitro techniques do not faithfully replicate spe-
cific in vivo conditions.
Chromatin immunoprecipitation (ChIP)
Chromatin immunoprecipitation has proven to be an excel-
lent experimental method used to determine the in vivo
analysis of DNA–protein interactions [77]. The analysis of
specifically interacting proteins with regions of genome has
wide utility for screening the localization of post-transla-
tionally modified histones and histone variants in the genome
or for understanding transcriptional regulation of genetic
expression (by analyzing the chromatin-specific transcrip-
tion factors). The methodology of chip involves shearing of
protein associated chromatin into smaller fragments fol-
lowed by immunoprecipitating the DNA–protein complex
using protein-specific antibody. The isolated DNA–protein
complexes are then dissociated and the specifically enriched
DNA segment is analyzed using PCR amplification methods.
This is the approach in a classical antibody-based ChIP
format (Fig. 2a). Another format for conducting ChIP assays
is the antibody-free format by the use of HaloTag Technol-
ogy. This method involves the transfection of Halo tag
vectors containing halo tags fused to proteins of interest
followed by their expression in mammalian cell lines. The
cells are then cross-linked, lysed and sonicated and the
Fig. 2 In vivo techniques to study DNA–protein interactions
Mol Cell Biochem (2012) 365:279–299 285
123
DNA–protein complexes are captured onto a HaloLink
Resin. This is followed by the standard decross-linking,
DNA purification and PCR amplification of enriched DNA
[78–80]. In general, there are many alternatives to detect an
immunoprecipitated chromatin such as polymerase chain
reaction (PCR), quantitative PCR (qPCR), labelling and
hybridization to genome-wide or tiling DNA microarrays
(ChIP-on-chip), molecular cloning and sequencing, or direct
high-throughput sequencing (ChIP-seq) [81–87]. There are
several variations of ChIP assay.
X-ChIP
This method allows freezing of all DNA-associated proteins
by cross-linking using formaldehyde. Formaldehyde reacts
with primary amines located on amino acids and the bases on
DNA or RNA molecules, forming a covalent cross-link
between the specific proteins to the DNA on which they are
situated. Now the various DNA–protein complexes are iso-
lated by cell lysis and the crude cell extracts are sonicated to
shear the DNA to a smaller size. The protein–DNA complex
is immunoprecipitated and the DNA–protein cross-links are
reversed by heating. The proteins are then removed by
treatment with proteinase K. The DNA portion of the com-
plex is then purified and identified by PCR using specific
primers. The use of formaldehyde for cross-linking mini-
mizes nucleosome rearrangements and is an efficient method
to analyze proteins that are weakly or indirectly associated to
DNA. The use of formaldehyde as a crosslinking agent has
certain limitations like the short cross-linking arm of form-
aldehyde is not suitable for examining proteins that indi-
rectly associate with DNA, such as those found in larger
complexes. So, a variety of other long-range bifunctional
cross-linkers may have to be used in combination with
formaldehyde to detect such interactions [88]. The yield of
chromatin and its resolution may be less after sonication and
sometimes there are chances of epitope disruption [89, 90].
Native-ChIP (N-ChIP)
N-ChIP is a technique suited for natural DNA–protein
interactions where the proteins are tightly associated to
chromatin in their native state such as histones due to their
high-affinity for DNA. Hence, these interactions do not
require cross-linking with formaldehyde. Native chromatin
within a cell produces smaller fragments, by treatment with
micrococcal nuclease (MNase) which are then immuno-
precipitated using antibody specifically against the protein
of interest. Enzymatic digestion technique is mild and does
not result in loss of antibody epitope during immunopre-
cipitation yielding higher immunoprecipitation efficiencies
[91]. It also provides high resolution as it is possible to
produce single monosomes of about 175 base pairs.
However, the digestion by MNase is uneven, as the enzyme
favours certain areas of genome sequence more. To avoid
over represented or over looked data, X-chip should be
carried out as a comparative control [92]. Also nucleo-
somes may rearrange during digestion and this has to be
taken into consideration while performing N-ChIP.
Fast ChIP
As the name suggests, fast chip is a modification of the
chip technique for large cell numbers which reduces time
required for conventional ChIP assay and helps in elimi-
nating multiple tube transfers thereby preventing loss in
output. Conventional chip assays require a high cell num-
ber to begin with due to low recovery rate of cross-linked
DNA from total cellular DNA. Multiple washes during the
procedure may also cause loss of specific interactions.
Therefore, a technique that can reduce the time and chan-
ces of losing cells is favoured. In this modification of ChIP
assay, all the steps are similar. However, the cross-links are
reversed during 10 min incubation at 100�C in an ultra-
sonic bath, in the presence of Chelex-100, a resin that aids
in the extraction of DNA. After incubation, the tubes are
spun and DNA containing supernatant can be directly used
in PCR [93]. The limitation associated with the fast ChIP is
that it is suitable only for large cell samples.
Carrier ChIP
The carrier ChIP is based on immunoprecipitation from
very few cells up to 100 cells and is suited for examining
histone modifications associated with developmentally
regulated genes. Immunoprecipitation of such a small
amount of chromatin is facilitated by the addition of carrier
chromatin from Drosophila or any other species which is
evolutionarily distant from the species being investigated
to provide efficient precipitation of target chromatin [94].
Here native chromatin is partially digested using MNase
and immunoprecipitated using antibodies to modified his-
tones. The low amount of chromatin is detected by radio-
active PCR and phosphorimaging. This technique,
however, requires the primers to be designed with high
specificity to prevent any spurious amplification of carrier
DNA instead of the target chromatin.
Matrix ChIP
It is a microplate-based ChIP assay in which all the steps
are done in microplate wells without sample transfers [95].
In this method, antibodies immobilized with protein A/G
are coated into each well of a 96-well plate and further
processed. Hence, allowing 96 ChIP assays for histone and
various DNA-bound proteins, including transiently bound
286 Mol Cell Biochem (2012) 365:279–299
123
protein kinases, in a single run. It also allows maintaining
antibodies in correct orientation which enhances its binding
capacity [96].
ChIP-Chip
As the name suggests, ChIP-Chip is a technique that com-
bines ‘‘Chromatin Immunoprecipitation’’ with ‘‘Microarray
technology’’. It consists of labelling the immunoprecipitated
DNA fragments with a fluorescent dye such as Cy5 or Alexa
647 and combining it with the genomic DNA labelled with
Cy3 which serves as the reference DNA. This probe mixture
is then applied to the microarray chip ideally consisting of
whole genome and allowed to hybridize. The results of the
experiment signify the regions of the DNA enriched by
immunoprecipitation. Hence, the Chip data is obtained in the
form of one dimensional series of signals with peaks iden-
tifying the regions bound by the protein of interest [97]. Also,
since the exact location of each arrayed element is known, a
genome-wide map of DNA–protein interactions can be
constructed.
Various computational and mathematical models are
available which allow the analysis of regions bound by the
proteins [98]. CisGenome is one such software which ful-
fils almost all the needs of ChIP data analysis including
visualization, data normalization, peak detection, false
discovery rate computation, gene-peak association,
sequence and motif analysis. Many statistical approaches
have also been used for the analysis of ChIP data including
Hidden Markov Model, Welch’s t statistic method, and
titled model-based analysis of tiling-arrays (MAT), to
identify regions enriched by a transcription factor [99].
The ChIP–Chip technique offers several advantages
over traditional ChIP assays. First, it allows probing of a
large number of genomic regions in a single experiment,
eliminating bias and saving time. Second, commercially
available platforms can be used to study the localization of
protein binding dismissing the need of running expensive
large scale quantitative PCR assays. Third, it allows par-
allel analysis of different genes to be classified in various
classes which is further useful for their statistical com-
parison [100].
Since an ideal microarray covering all the human
chromosomes is not possible, this technique may be ben-
eficial if combined with other throughput technologies.
DIP-Chip
The modification of ChIP–Chip is DIP–Chip that over-
comes its limitations like interference of protein–protein
interactions and competitive binding in vivo. DIP–Chip is
more of an in vitro technique with results comparable to in
vivo assays. The procedure involves interaction of purified
and mechanically sheared genomic DNA with purified
protein of interest. The DNA–protein complexes are then
affinity-purified using appropriate resins. These affinity-
purified genomic fragments along with the whole genome
fragments are then amplified and fluorescently labelled
separately with different dyes for assessing their relative
abundance in the entire genome of the organism using
microarray. The samples are analyzed by comparative
hybridization to the DNA microarray that covers the entire
genome of the organism [101].
ChIP sequencing
ChIP sequencing combines the technique of chromatin
immunoprecipitation and DNA sequencing to identify the
binding sites of various protein factors co-precipitated
along with DNA fragments during ChIP [102]. For the
construction of ChIP-seq library, the ends of enriched DNA
fragments obtained by immunoprecipitation using con-
ventional ChIP protocol are blunted and phosphorylated
using T4 kinase. Following this Adenine is added using
Taq and an adapter is ligated to both the ends of the
fragment [103]. The library obtained is amplified by PCR
and DNA fragments of length 100–300 bp are selected and
sequenced. Finally the short sequenced fragments called
tags are analyzed computationally with the help of align-
ment tools using a particular genome as reference to
identify the enriched sites [104].
This technique has several advantages over ChIP–Chip
including low cost, lesser starting material and higher peak
resolution. However, it also has a number of issues which
need to be addressed. First, The ChIP-Seq tags represent
the ends of the enriched fragments and not the binding sites
of the protein factor. Moreover, the estimation of site to tag
distance is complicated. Second, no control samples are
sequenced deeply to check for the regional biases along the
genome arising on account of chromatin structure and copy
number variations [102]. Third, lack of advanced and user
friendly data analysis tools make the analysis of peaks
difficult.
However, ChIP-seq has been proved to be a potential
tool in the study of histone modifications, nucleosome
positioning and mapping of binding sites of various DNA-
binding proteins. Moreover, this strategy allows distin-
guishing alleles on the basis of difference in SNPs, which
would not have been possible using ChIP–Chip [103].
ChIP display
ChIP–Chip has been described as a potential method for
the identification of novel transcription factor binding sites
in the genome. But it suffers from severe limitations
including co-precipitation of non-specific DNA fragments
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123
which may sometime even overwhelm the specific ones
resulting in a strong background noise. To overcome this
problem, a new technique has been devised called ChIP
display. This technique is based on the principle of con-
centrating the target fragments via restriction digestion and
then scattering the precipitated non-specific DNA frag-
ments by partitioning the digested fragments into different
families. The partitioning is based on the identity of the
nucleotides at the end of these fragments [105]. Since all
the target fragments remain in the same family, the signal
is not eroded and is separated from the non-specific frag-
ments of different families.
ChIP display is a prospective tool for the reduction of
non-specific DNA precipitation. However, it suffers some
practical limitations. First, since non-specifically precipi-
tated DNA fragments can unexpectedly bind the protein in
vitro (but not in vivo), hence utility of this approach is
debatable. Second, ChIP display is not well suited for a
comprehensive analysis of target sequences for proteins
with a large number of genomic targets, such as GATA
proteins, histone deacetylases, polycomb proteins or for the
mapping of histone modifications [105]. It is better suited
for transcription factors with a more limited number of
targets.
Other ChIP variations
There are certain other categories of ChIP assay setups that
are classified based on different buffers used which affect
the purpose and efficiency of the immunoprecipitation such
as Quick and Quantitative ChIP (Q2 ChIP) and MicroChIP.
Q2 ChIP incorporates histone deacetylase inhibitor during
cross-linking which helps in elimination of non-specific
backgrounds and also has different elution buffers and
reduced time of protocol. MicroChIP is a miniaturized
ChIP protocol for 10,000 cells that has applicability in
genome-wide studies [106, 107].
DNA adenine methyltransferase identification
(DAMID)
DAMID is a novel methylation-based tagging technique
that has emerged as a powerful tool to study chromatin
interactions in vivo. It has been successfully used to gen-
erate genome-wide maps of several DNA-binding factors
including GAGA factors, Max family of transcription
regulators, coregulators and various other chromatin pro-
teins [108].
In this technique, the protein of interest is fused with a
bacterial DNA adenine methylase (DAM) which is a single
32 kDa polypeptide and methylates adenine at the sixth
position in the sequence GATC [109]. This methylation
causes few changes in the DNA topology and provides a
unique tagging system to mark the binding sites of specific
protein factors. This fusion protein is expressed in mam-
malian cells in low quantities by using a weak promoter
[108]. The binding of the fusion protein to the target site
results in the methylation of adenine nucleotides within the
DAM recognition sequence in close vicinity of the protein
target site. These methylated sequences are then cleaved by
DpnI enzyme to recover fragments containing regions
nearby or within the gene along with the target site itself.
Further, the fragments obtained may be analyzed by
quantitative PCR assay or subjected to microarray studies.
To overcome these effects of chromatin accessibility on the
level of methylation, a control experiment is run in parallel
which measures the methylation levels in the probed
sequences after the expression of dam [108].
DAMID has significant advantages over the conven-
tional ChIP technique. First, it does not use any cross-
linking agents to fix the chromatin and also eliminates the
use of protein-specific antibody. Hence, it provides a
simpler platform to study the binding properties of co-
factors and other proteins that bind indirectly to the DNA
[110]. Also, there are lesser chances of misidentification of
target sequences due to accidental cross-linking as in case
of ChIP. Second, it provides an easier way to study the
effects of mutations on the targeting specificity of the
protein of interest which is difficult to perform using
conventional ChIP assays [108].
The limitations of DAMID are that it requires dam to be
bound to the protein without inducing any changes in its
function. Also, this technique is unsuitable for the detection
of post-translational modifications, while ChIP successfully
detects histone modifications. It is a time consuming
technique as it involves expression of DAM-fusion protein
for several hours [108].
Chromosome conformation capture (3C)
and ChIP-loop assay
One of the key regulators of gene expression is spatial
organization of the eukaryotic genome. 3C is a novel
technique that is used to detect the frequency of interaction
between two genomic loci in the nuclear space. It is a
powerful tool to study the link between nuclear organiza-
tion and transcription regulation. This technique is carried
out by initially fixing the cells with formaldehyde which
helps in cross-linking of interacting segments of the gen-
ome via contacts between their DNA-bound proteins. The
resulting network of protein–DNA complexes is subjected
to restriction digestion followed by ligation at low DNA
concentration, such that the ligation between cross-linked
DNA fragments is favoured. After the reversal of cross-
links, the fragments are subjected to quantitative PCR to
further allow for the measurement of cross-linking
288 Mol Cell Biochem (2012) 365:279–299
123
frequency of the two specific restriction fragments
(Fig. 2c) [111].
Although ChIP and 3C operate through same basic
principle of cross-linking protein–DNA interactions but the
two techniques differ from each other in the information
they provide whereas ChIP provides information about
DNA-binding activity of a protein, 3C is used to study
interaction between two different genomic sites looped by
a protein factor.
To establish a link between 3C and ChIP a novel tech-
nique called ChIP-loop assay has been developed [112].
This technique allows the study of proteins mediating the
interaction between the two genomic loci, by combing the
two techniques. Initially, the cells are fixed by formalde-
hyde and the cross-linked chromatin purified from free
proteins by urea gradient centrifugation. This is followed
by restriction digestion of the purified cross-linked chro-
matin and precipitation by protein A/G beads and specific
antibodies. The precipitated chromatin is then allowed to
ligate and is further analyzed by quantitative PCR as in
standard 3C experiments [111]. Hence, ChIP-loop assay
helps in studying the proteins that are involved in organi-
zation of DNA loops to mediate genomic interactions. This
technique provides a better insight into interactions than 3C
and ChIP when used alone. However, the major concern is,
when the DNA is concentrated before ligation, it may lead
to formation of loops between bead-associated DNA
fragments. Hence the results obtained may not accurately
identify the loops of DNA fragments formed in the nuclear
space. This also makes the quantification of ligation
products very complicated [113]. Nevertheless, their
potential use in identifying proteins participating in long-
range interactions cannot be denied.
In silico tools for identification of DNA–protein
interactions
The computational and in silico approaches to identify
DNA–protein interactions are an important aspect of these
interactions. Diverse computational tools are freely avail-
able which are used to predict DNA–protein interactions.
Most of these are aimed at predicting the transcription
factor-based gene regulation.
TRANSFAC
TRANSFAC is a comprehensive knowledgebase contain-
ing eukaryotic gene regulation data from a wide variety of
eukaryotic organisms, ranging from yeast to humans. It
mainly comprises of data on transcription factors, their
experimentally proven binding sites, regulated genes and is
an extremely diverse tool for transcription factor (TF)
binding predictions. It has a broad compilation of binding
sites and allows the derivation of positional weight matri-
ces, which can be used with the available tools to search
DNA sequences. Several entries are grouped under differ-
ent tables of the TRANSFAC database. One of the features
is the assigning of a quality value to describe the confi-
dence with which an observed DNA-binding activity could
be assigned to a specific factor. Nucleotide weight matrices
are derived from a collection of binding sites for a factor,
and these matrices are used by the tool MatchTM to find
potential binding sites in uncharacterized sequences. Sev-
eral web programs are also available that utilize the
TRANSFAC database such as AliBaba2 which is a used for
predicting TF binding sites in an unknown DNA sequence
by utilizing the binding sites collected in TRANSFAC.
P-Match is another new tool for identifying transcription
factor binding sites in DNA sequences. It combines pattern
matching and weight matrix approaches to provide a high
accuracy of recognition.
TRANSFAC is maintained as a relational database, from
which public releases are made available via the web,
making it an easily accessible database. Several web-based
tools are linked to TRANSFAC and utilize its database to
perform unique computational functions [114, 115].
Identification of DNA-binding proteins (iDBPs) server
The iDBPs server was developed for the identification of
DNA-binding proteins with known three-dimensional
structure. In the first stage of classification, the functional
region of the protein is predicted using the PatchFinder
algorithm which searches for clusters or patches of evolu-
tionarily conserved residues on the protein surface. The
maximum-likelihood (ML) patches found by PatchFinder
often delineate the functional regions in proteins and spe-
cifically, the core of DNA-binding regions within DNA-
binding proteins [116]. The results are sent to the user which
includes the prediction score of the protein, the expected
sensitivity and the expected precision at this score cut-off.
DNA site prediction from a list of adjacent residues
(DISPLAR)
DISPLAR is a neural network method that predicts the
residues of a protein which interact with DNA, if the
structure of a protein known to bind DNA is provided.
Several inputs have to be provided to the neural network
including position-specific sequence profiles and solvent
accessibilities of each residue and its spatial neighbours.
The neural network is trained on known structures of
protein–DNA complexes. DISPLAR shows prediction
accuracy over 80% and coverage of over 60% of actual
DNA-contacting residues [117].
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FlyFactorSurvey
FlyFactorSurvey is a database of DNA binding specificities
for Drosophila TFs. It provides community access to over
400 recognition motifs and position weight matrices for
over 200 TFs, including many unpublished motifs. The
primary source of recognition motifs within FlyFactor-
Survey is TF binding site selections performed using the
bacterial one-hybrid system. Search tools and flat file
downloads are provided to retrieve binding site information
(as sequences, matrices and sequence logos) for individual
TFs, groups of TFs or for all TFs with characterized
binding specificity. Linked analysis tools allow users to
identify motifs within the database that share similarity to a
query matrix or to view the distribution of occurrences of
an individual motif throughout the Drosophila genome
[118].
YEAst search for transcriptional regulators
and consensus tracking (YEASTRACT)
YEASTRACT information system allows the identification
of potential transcription regulators. It is a database that
contains over 12,346 regulatory associations between
transcription factors and target genes in Saccharomyces
cerevisiae [119]. It also characterizes set of genes with
common expression profile obtained from microarray data
and searches for occurrence of candidate TF binding sites.
Multi-genome analysis of positions and patterns
of elements of regulation (MAPPER)
It is a search method that helps in identifying the TF
binding sites which is based on hidden Markov model
obtained from alignments of known sites. TF binding site
models can be used to align with the sites provided by the
TRANSFAC and other databases and then scan the
sequences of the human, mouse, fly, worms and yeast
genomes to identify the sites. It has a better specificity and
sensitivity than other similar computational models. A
sequence is uploaded as a query and then a model is built
by allowing multiple sequence alignment of binding sites
of the transcription factor [120].
Zinc finger binding site database (ZIFIBI)
It is a tool that helps in identifying the C2H2 zinc finger
transcription binding site in the cis regulatory regions of
the target genes. It makes use of the available data to
predict the interactions between the nucleotides and the
amino acids of the zinc finger domain of the protein. The
most probable state path is calculated using a hidden
Markov model [121].
Bioprospector
It helps in identifying regulatory sequence motifs in cis
region of target sequence by examining it in the same gene
expression pattern group. It is based on a C program and
uses Gibbs sampling strategy. The significance of each
motif is estimated using the Monte Carlo method. It has
been successful in identifying binding motifs for Saccha-
romyces cerevisiae Ras-related protein 1(RAP1), Bacillus
subtilis RNA polymerase, and Escherichia coli cyclic AMP
receptor protein (CRP) [122].
Bindn
It is a web-based tool that helps to predict the DNA and
RNA binding sites with the help of support vector
machines (SVMs). The SVM models are prepared using
three sequence features like side chain pKa values,
hydrophobicity index and molecular mass of an amino
acid. Thus, it helps to identify the functions of the binding
proteins based on primary sequence data [123].
Bindn?
Bindn? uses protein sequence features different from
Bindn to identify the binding sites in the sequences. It also
takes the support of the SVMs. The protein sequence fea-
tures used in this case are the biochemical property of the
amino acids and evolutionary information in terms of the
position-specific scoring matrix. The new descriptors used
in Bindn? have shown better performance, sensitivity and
specificity in comparison to the previous version [124].
DP-bind
It helps in predicting the binding sites of a protein by ana-
lyzing the amino acid sequence. It uses three support models
for predicting the sites: support vector machines, kernel
logistic regression and penalized logistic regression. Pre-
diction can be done using the input sequence alone or the
profile of evolutionary conservation of the input sequence.
The output of all the three models are used to provide a
combined and consensus result with high confidence [125].
PreDs
It is a web-based server that allows DNA-binding site
prediction on protein molecular surfaces. The molecular
surfaces of the proteins are generated with the help of
atomic coordinates that are available in a .pdb format. The
prediction is based on the evaluation of the electrostatic
potential, local and global curvature of the protein surface
[126].
290 Mol Cell Biochem (2012) 365:279–299
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ProNIT
It is a thermodynamic database that uses quantitative
binding data rather than just structural data. It contains
several parameters for analyzing the protein-nucleic acid
recognition like thermodynamic parameters, experimental
conditions and structural information of both the protein
and the DNA. It provides various sorting output options.
The thermodynamic parameters used are dissociation
constant, association constant, Gibbs free energy change,
enthalpy change and heat capacity change. A relational
database system combines all of this information to provide
flexible searching facilities [127].
Database for polyanion binding proteins (DB-PABP)
Polyanion binding proteins are diverse proteins that go and
interact with polyanions which are entities having multiple
negative charge. The various polyanions identified for such
interactions are actin, tubulin, DNA, heparin and heparin
sulphate. The database thus created is a comprehensive and
searchable database which has been manually curated. It
has been implemented as a MY SQL relational database.
The search is based on four criteria: protein names, poly-
anion names, source species and the methods used to dis-
cover the interactions [128].
DNAProt
It helps in identifying the DNA-binding proteins from the
protein sequence. It has considerably good accuracy in
distinguishing between the DNA-binding proteins and the
non-DNA-binding proteins by characteristically recogniz-
ing specific DNA chains. The random forest method is used
to identify the DNA-binding proteins [129].
Biophysical techniques as a potential tool for
DNA–protein interaction studies
Fluorescence-based techniques
Fluorescence is a form of luminescence caused by emission
of an electromagnetic radiation [130]. The simultaneous
absorption of two photons by an electron (two-photon
absorption) excites molecule from ground state to higher
energy (high frequency, low stability) state, leading to
emission of radiations [131]. This principle and its modifi-
cations are used to device different fluorescence detection
techniques, e.g. fluorescence spectroscopy, fluorescence
intensity, florescence depolarization, fluorescence resonance
energy transfer and fluorescence correlation spectroscopy. In
fluorescence-intensity distribution analysis fluorescence
intensity of a sample with a heterogeneous brightness profiles
is monitored by spatial brightness distribution and calculat-
ing theoretical photon count number distributions [132].
Capillary electrophoresis with laser-induced
fluorescence
Capillary electrophoresis coupled with laser-induced fluo-
rescence polarization is a hybrid approach to ultrasensitive
immunoassays [133]. Fluorescence polarization provides
additional information for identification of affinity com-
plexes. Protein–DNA interactions can be studied on the
basis of capillary electrophoretic (CE) separation of bound
from free fluorescent probe followed by detection with
laser-induced fluorescence polarization (LIFP) [134].
Changes in electrophoretic mobility and fluorescence
anisotropy upon complex formation can be monitored for
the determination of binding affinity and stoichiometry
[135]. There are two types of assays—Homogeneous and
heterogeneous. In the homogeneous assay, the free and
bound tracers are joined together and the fluorescence
polarization of the mixture is a quantitative measure of the
antibody-bound tracer. The heterogeneous assay involves a
baseline separation of the free and bound tracer using CE
with a phosphate running buffer. Results from both assays
suggest that the CE–LIFP approaches have a wider appli-
cation than the immunoassays based on either CE–LIF or
fluorescence polarization alone [136].
Narrow-bore capillaries provide high-speed, high reso-
lution separations and ultrasensitive detection in a minimal
sample detection volume. Increased detection limits,
enhanced identification capacity, potential for miniaturi-
zation, etc. also adds to its advantages. However, the free
and bound tracer may have similar electrophoretic mobil-
ities and thus cannot be separated, making the technique
inefficient in their identification and quantitation [137].
Time-resolved fluorescence depolarization
Time-resolved fluorescence depolarization (Anisotropy) is
a technique where a short pulse of vertically polarized light
is directed at the sample where the absorbed light prompts
the molecule to an excited singlet state [138]. After
vibrational relaxation, fluorescence light is emitted at lower
energy; if the molecule rotates during the time interval
between absorption and emission, there is a decrease in the
polarization with time that reflects a rate at which the
molecule rotates diffusionally [139].
Time-resolved fluorescence spectroscopy can be used to
analyze the interaction between proteins and DNA. Fluo-
rescence polarization anisotropy decay can be used as a
spectroscopic handle to scrutinize the interaction between
several site-specific DNA-binding proteins and their target
Mol Cell Biochem (2012) 365:279–299 291
123
DNA fragments. Solution conditions such as temperature,
pH, ionic strength, and the presence of effector molecules
can be varied and interaction can be studied [140].
Variety of DNA sequences can be tested, both for pre-
liminary experiments and for evaluating base sequence-
dependent effects. The assay is reversible which allows
manipulation of solution conditions so that the effects of
environment or effector molecules on complex formation
can be accessed directly. Also the rotational correlation
time directly measures molecular size and shape.
Double labelled native gel electrophoresis
and fluorescence-based imaging
Radiolabeled DNA gel mobility shift assay is modified to
incorporate an end-labelled DNA probe with a texas-red
fluorophore and a DNA-binding protein tagged with the
green fluorescent protein to monitor precisely DNA–pro-
tein complex by native gel electrophoresis [141]. This
method is applied to the DNA-binding proteins, demon-
strating that the method is sensitive, permits direct visu-
alization of both the DNA probe and the DNA-binding
protein, and enables quantitative analysis of DNA and
protein complex, and thereby an estimation of the stoi-
chiometry of protein-DNA binding [142].
Protein array method combining a near-infrared
fluorescence detection
The protein array methodology is used to study DNA–
protein and protein–protein interactions using probes
labelled with near-infrared fluorescence dyes (IRDye800)
with excitation characteristics near 700 or 800 nm detect-
ing signals from proteins immobilized on a nitrocellulose
membrane with a high sensitivity [143]. To study protein–
DNA binding, the membranes are incubated in a DNA-
binding buffer containing poly-dGdC and poly-dAdT or
sonicated salmon DNA at room temperature for 30 min.
Then, an Infra red Dye-labelled DNA probe is added to the
pre binding solution and incubated with a slow rotation at
room temperature or at 60�C. The membranes are washed
with PBS containing 0.1% Tween and then screened for the
detection of fluorescent signals by infrared Imaging sys-
tem. To perform protein–protein binding, the membranes
are incubated in the PBS solution with BSA at room
temperature and then with corresponding Cy5.5-labelled
protein in PBS containing 1% BSA and 0.1% Tween 20 at
room temperature for 1 h [144].
The fluorophores in protein array method with longer
wavelengths provide a high-signal-to-noise ratio that
decreases the background effect on membrane surfaces;
thereby increasing the sensitivity of the detection.
Fluorescence resonance energy transfer (FRET)
techniques
FRET is a non-radiative process whereby an excited donor
fluorophore transfers energy to a ground-state acceptor as a
result of a coupling of their transition dipoles. FRET pro-
vides structural and kinetic information of protein–DNA
interactions by preparation of dye-labelled nucleic acids
and proteins and increased optical sensitivity. The principle
of FRET relies on the site-specific labelling with a donor
and an acceptor dye, with FRET dyes in each interacting
partner (Intermolecular FRET) or both in the same bio-
molecule (Intramolecular FRET) (Fig. 3a, b). Direct opti-
cal excitation of the donor dye results in fast energy
transfer to the FRET acceptor, which emits fluorescence at
a longer wavelength [145, 146].
Intramolecular FRET assays, where both dyes are
located on the same biomolecule are extensively used to
monitor protein-induced conformational changes in the
DNA substrate and to determine the global structure and
assembly dynamics of a variety of nucleoprotein
complexes.
FRET technique relies on its continuous character, so
that the cleavage reaction can be monitored from the initial
steps in real-time with no need for extensive sample han-
dling [146].
FRET–FLIM in situ imaging for protein–DNA
interactions in the cell nucleus
This approach allows imaging of the in situ interaction
between a GFP-fusion protein and DNA in the cell nucleus,
using FRET [147]. A fluorescence lifetime imaging
microscopy (FLIM) is used as a reliable tool to detect
protein in contact with DNA. To develop a FRET-based
method to visualize DNA–protein interactions in situ, a
DNA-binding fluorescent dye that is suitable as FRET
acceptor if GFP is the donor must be used. The members of
the Sytox fluorescent dye family have a high-affinity for
nucleic acids and are available with a broad range of
excitation and emission spectra. Upon binding to DNA or
RNA, they show several hundred-fold enhancement of
fluorescence intensity [148].
Fluorescence lifetime measurements can be performed
by wide-field frequency-domain FLIM with Argon-ion
laser as an excitation source. Images at different phases can
be recorded at the image intensifier. Thereby, phase and
modulation depth-based lifetime of the emitted fluores-
cence can be calculated from the resulting set of images
[149].
FRET–FLIM in situ imaging for protein–DNA interac-
tions in the cell nucleus is a reliable and quantitative
method to measure FRET. It is a donor-selective FRET
292 Mol Cell Biochem (2012) 365:279–299
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method, which is not influenced by acceptor dye molecules
that are not involved in FRET.
Nuclear magnetic resonance
NMR is used to investigate the interactions of DNA with
proteins. NMR provides dynamic and structural infor-
mation on the changes in conformation and molecular
flexibility and enables formulation of mechanistic models
of DNA–protein interactions [150]. There are some
sample preparation steps that need to be followed. The
sample needs to be labelled and various strategies may
be employed. Either the protein is 15N or 13C labelled
while the DNA is unlabeled or vice versa. Sample pre-
cipitation needs to be taken care of as there is a strong
electrostatic interaction involved within the complex
[150].
The sample can be analyzed by chemical shift mapping
where hetero nuclear single quantum coherence (HSQC)
spectra of labelled molecule is analyzed separately for
bound and free state. Chemical shifts are sensitive to
changes in the chemical environment of the protein. The
DNA interaction with a protein alters chemical environ-
ment. Thereby, causing shift in the spectra as compared to
unbound molecules [151]. Cross-saturation experiments
can also be used to analyze DNA–protein, protein–protein
interactions and various binding surfaces of ligands on the
protein [152]. Another technique that is employed in NMR
is the solvent accessibility test which helps in the quanti-
tative analysis of the amide proton exchange rates of the
free and the bound protein. However, there are certain
intermolecular restraints to the NMR spectroscopy like
nuclear overhauser effect, residual dipolar couplings and
paramagnetic relaxation enhancement. These restraints
may hamper the precision and accuracy of the technique
and hence various modifications have been made which
can overcome these restraints [150].
Circular dichroism
Circular dichroism (CD) is a quantitative technique that
helps to identify the DNA–protein and protein–protein
interactions. It provides additional information about the
prosthetic groups, bound ligands and the co-factors
attached. It also helps to identify the conformational
change in protein molecules. There are signatures corre-
sponding to the particular interaction based on asymmetry
induced by the secondary structure of proteins. Thereby,
Fig. 3 Biophysical techniques to study DNA–protein interactions
Mol Cell Biochem (2012) 365:279–299 293
123
identifying the structure of bound protein and the possible
interactions involved [153, 154].
There are many variations to CD like the stopped flow
CD and the CD using synchrotron radiation [155]. In case
of nucleic acids-induced CD measures the asymmetry
among the bases. The bases as such are planar but there is
some amount of CD-induced due to the sugar present in the
backbone of the DNA. It is a powerful technique in ana-
lyzing the structural change with respect to factors like
temperature, ionic strength and pH. It helps in judging the
extent of interaction between the helices by analyzing the
melting of peptides [156].
The circular dichroism technique is considered to be a
better method than other techniques like nuclear magnetic
resonance (NMR) as it is faster, economical, uses a small
amount of sample to analyze and most of the sample can be
recovered for further analysis.
The limitations of CD are relatively low resolution
structural details and little information about the quaternary
structure of the protein [157].
Atomic force microscopy (AFM)
AFM is another powerful tool for imaging DNA–Protein
complexes at a single molecule level [158]. It allows to
characterize the mechanisms involved in DNA–protein
complex formation in different conditions with high reso-
lution. It quantitatively identifies protein position along
DNA molecules, DNA flexibility, curvature and confor-
mational change after protein binding.
AFM is operated in tapping mode which allows the
elimination of permanent shearing forces and causes less
damage to the sample surface, even with stiffer probes
[159]. Different components of the sample which exhibit
difference in adhesive and mechanical properties show a
phase contrast and therefore allow a compositional
analysis.
The potential of this technique for high-sensitivity, high-
throughput operation in fluid, and for force detection are
major considerations for its continued integration into
mainstream cellular and molecular analyses [160]. It uses
very small quantities (10-9 to 10-15) of DNA and proteins.
The technique has limitations when it is applied to
structural and functional studies of biomolecules, due to the
resolution limiting motion of DNA molecules. To over-
come this, the DNA must be tethered to the substrate sur-
face. Because of its flatness, mica is the most commonly
used substrate for DNA imaging [161, 162]. Also, large
DNA molecules remain difficult to be imaged by AFM
because of their tendency to aggregate. A modified method
is described by Lysetska et al. [163], to align long-DNA
fibres in a single direction on unmodified mica to facilitate
AFM studies.
Surface plasmon resonance (SPR)
SPR is a label-free optical technology and an emerging
alternative to the conventional in vitro techniques to study
DNA–protein interactions. It uses an evanescent wave
phenomenon to study changes in refractive index, occur-
ring close to the sensor chip surface, causing a shift in
plasmon resonance angle, detected by an imaging system.
The general principal that lies behind the working of
SPR is total internal reflection that occurs when a polarized
light travels through a medium of higher to lower refractive
index. When this occurs, the electromagnetic field com-
ponent penetrates over a short distance into the medium of
low refractive index resulting in the exponential attenua-
tion of the evanescent wave. If the interface is coated with
a thin layer of gold, then the projected beam at the given
angle will cause resonance coupling between light photons
and surface plasmons of gold as their frequencies match. A
change in the refractive index within the environment of
evanescent wave occurs due to the binding of DNA to
protein. Hence, a real-time measurement of biomolecular
interactions can be enabled by measuring the refractive
index changes corresponding to mass changes [164]. Many
advances have been done in this technique to study DNA–
protein interactions. A multistep chemical modification
procedure has been proposed to create DNA arrays on gold
surfaces specifically tailored for the study of protein–DNA
interactions [165].
To study DNA–protein interactions, DNA is immobi-
lized on the chip surface followed by a constant flow of
buffer over the surface (Fig. 3c). The protein analyte is
allowed to bind to the immobilized DNA and a change in
the position of reflected light minimum observed in terms
of resonance units (RUs) which are recorded and a sens-
ogram is generated. A sensogram is divided into four dif-
ferent phases: association phase, steady state or equilibrium
phase, dissociation phase and regeneration phase (Fig. 3d)
[166].
SPR offers a variety of advantages over other tech-
niques. First, the interaction can be monitored very accu-
rately in real-time. Since the change in refractive index
corresponds to a change in mass, this method can also yield
data on the stoichiometry of complexes in addition to
binding kinetics [164]. Second, simultaneous analysis of
multiple interaction partners can be seen. Third, it is a
label-free technology and optical radiation does not harm
the biomolecules.
Microcalorimetry
Being a non-invasive technique, microcalorimetry is a
potential technique to study the interactions and study of
biomolecules. It is the measure of calorimetry of small
294 Mol Cell Biochem (2012) 365:279–299
123
samples and relies on the similar basic principle of mea-
surement of heat energy changes occurring during any
physical or chemical processes.
For studying protein–DNA interactions, two most
commonly used microcalorimetric techniques are—differ-
ential scanning calorimetry (DSC) and isothermal titration
calorimetry (ITC). DSC measures the heat capacity profile
of proteins as a function of temperature during processes
like protein unfolding, thermal stability during complex
formation by measuring the differential heat energy chan-
ges between sample and reference cells [167]. A pair of
matched calorimetric cells (sample and reference cell)
enclosed in an adiabatic chamber and fitted with sensitive
thermocouple are used. Electronic/Computer controlled
feedback circuits are used to measure the differential
temperature lag between cells. ITC is used to study binding
proteins more directly by measuring not only the magni-
tude of the binding affinity but also the magnitude of the
two thermodynamic terms that define the binding affinity:
the enthalpy and entropy changes [168]. In a typical
experiment, a solution of a one biomolecule is titrated into
a solution of its binding partner and the heat released upon
their interaction is monitored over time. The temperature
dependence of enthalpy of binding can be used to calculate
the binding heat capacity [167].
Since microcalorimetry is not affected by the constraints
due to size and shape of molecule and does not require any
chemical modification or solid support, it has become an
invaluable resource in laboratories [169]. Also the high
sensitivity and its ability to analyse true binding affinities
by measuring heat changes and measure nanomolar to
picomolar binding constants (109 to 1012 M-1) using the
competitive binding technique makes it a promising tech-
nique in molecular biology.
Although ITC is particularly suitable to follow the
energetics of an association reaction between biomole-
cules, the combination of ITC and DSC provides a more
comprehensive description of the thermodynamics of an
associating system [170].
Conclusion
DNA–protein interactions are an integral component of
biological systems and their study is important for almost
all biological processes. Several techniques are available to
aptly determine these interactions and their understanding
is imperative. At the in vitro level, molecular biology-
based techniques such as footprinting assays, EMSA,
southwestern blotting, Y1H phage display and proximity
ligation assay (PLA) screen DNA–protein interactions
reliably. The highly dynamic in vivo tools of chromatin
immunoprecipitation and its variants, DNA adenine methyl
transferase identification (DAMID) and ChIP-loop assay
are robust techniques to characterize several DNA–protein
interactions in cells. In silico approaches have also evolved
drastically over the years to supplement the information
available to researchers. Various recent biophysical tech-
niques including fluorescence-based techniques, CD,
NMR, AFM, SPR and microcalorimetry have a great
potential for the detection of protein-based interactions.
Every technique is unique in its own way and serves a
unique purpose. As is evident, the current state of methods
leaves quite a lot to be desired. An ideal method would
require minimal cell numbers, able to detect rare interac-
tions with high specificity and sensitivity, easily modified
to quantify interactions and provide complete information
on either of protein or DNA, by themselves. Thus, the
above listed techniques will help researches to assess the
dynamics of DNA–protein interactions in cellular devel-
opment and disease progression.
Acknowledgments This study was supported by the research grant
awarded to Dr. Vibha Rani by the Department of Science and
Technology, Government of India (SR/FT/LS-006/2009: Sept 4,
2009). We acknowledge Jaypee Institute of Information Technology,
Deemed to be University for providing the infrastructural support.
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