The structural organization and dynamics of DNA are known to be of paramount importance in countless cellular processes, but capturing these events poses a unique challenge. Fluorescence microscopy is well suited for these live-cell investigations, but requires attaching fluorescent labels to the species under investigation. Over the past several decades, a suite of techniques have been developed for labeling and imaging DNA, each with various advantages and drawbacks. Here, we provide an overview of the labeling and imaging tools currently available for visualizing DNA in live cells, and discuss their suitability for various applications.

Introduction

Observing DNA inside cells has an increasing number of applications ranging from diagnostics [1], cancer and cell viability research [27] to more fundamental studies on chromatin structure and dynamics [811]. Fixed-cell measurements of genome-wide or chromosome-specific labeling can be combined with high-throughput or super-resolution imaging to obtain snapshots of various processes [10,1218]. In live cells, spatio-temporal tracking of labeled chromatin elements (e.g. telomeres and centromeres) has opened a window into the biophysical environment inside the nucleus [2,9,19].

Observing the positions of individual DNA loci in live cells can provide insights into chromatin dynamics, e.g. enhancer/promoter proximity [20], correlated loci motion [21] and global chromosome organization [22], and is complementary to the single time-point, DNA-interconnectivity mapping obtained via Chromatin Conformation Capture (3C) and related techniques [2327]. Such live-cell experiments require fluorescently labeling DNA loci, imaging and then analysis to determine the positions of the labeled objects. In this review, we discuss existing techniques and considerations for labeling and imaging DNA in cells, and then review several key applications.

DNA-labeling methods

Many methods have been developed for DNA labeling (Figure 1). While labeling and imaging in fixed cells is a relatively routine task, live-cell imaging, which is the focus of this review, introduces additional challenges. The choice of the fluorescent-labeling method is dictated mainly by two parameters: first, the desired level of sequence specificity: nonspecific, chromosome-specific, repetitive loci or unique (nonrepetitive) loci. Second, the scale of the assay: the number of loci and independent imaging channels required.

DNA-labeling methods.

Figure 1.
DNA-labeling methods.

(A) DNA-binding dyes with no sequence specificity can be used to visualize the genome via fluorescence. (B) DNA-binding proteins, such as histones, can be fused to fluorescent proteins. (C) By inserting an array of operators into the genome near a targeted locus, fluorescently labeled regulation machinery can be used to mark a locus. (D) The CRISPR system can be used to target a fluorescently labeled dCas9 to a sequence-specific region.

Figure 1.
DNA-labeling methods.

(A) DNA-binding dyes with no sequence specificity can be used to visualize the genome via fluorescence. (B) DNA-binding proteins, such as histones, can be fused to fluorescent proteins. (C) By inserting an array of operators into the genome near a targeted locus, fluorescently labeled regulation machinery can be used to mark a locus. (D) The CRISPR system can be used to target a fluorescently labeled dCas9 to a sequence-specific region.

For nonspecific-sequence applications, namely, fluorescently labeling the entire genome, several methods are available. One category of tools is DNA dye staining (Figure 1A), where molecules with a high affinity for DNA and no-to-little sequence preference can yield whole-genome fluorescence. These dyes attach to DNA by various mechanisms, including intercalating between base pairs (Figure 2A) [28], major-/minor-groove binding (Figure 2B) [29,30] and association with the phosphate backbone [31]. While some dyes, such as Hoechst and DRAQ5, are suitable for live-cell imaging, others are not, due to limited membrane permeability and cytotoxicity [32].

Two example DNA-binding dyes.

Figure 2.
Two example DNA-binding dyes.

(A) DRAQ5, a DNA-intercalating dye. (B) Hoechst, a groove-binding dye. Insets show schematics of the binding geometries. The main advantage of DNA dyes is that their molecular properties can be tuned with the full, synthetic-chemistry toolkit. For example, by covalently binding the stable, cell-permeable, silicone rhodamine (SiR) dye to the DNA-groove-binding Hoechst dye, Lukinavičius et al. [33] demonstrated live-cell, super-resolution microscopy of the DNA. This was largely made possible by using the longer-wavelength excitation and emission properties of SiR, thus avoiding the phototoxicity associated with blue illumination light, as well as the majority of the cellular autofluorescence. DNA-binding dyes can also be used to target conjugated proteins to the DNA by using the dye as a homing beacon. This strategy has been used to target regulatory machinery and labels required for further analysis by other complementary tools, such as mass spectrometry [34].

Figure 2.
Two example DNA-binding dyes.

(A) DRAQ5, a DNA-intercalating dye. (B) Hoechst, a groove-binding dye. Insets show schematics of the binding geometries. The main advantage of DNA dyes is that their molecular properties can be tuned with the full, synthetic-chemistry toolkit. For example, by covalently binding the stable, cell-permeable, silicone rhodamine (SiR) dye to the DNA-groove-binding Hoechst dye, Lukinavičius et al. [33] demonstrated live-cell, super-resolution microscopy of the DNA. This was largely made possible by using the longer-wavelength excitation and emission properties of SiR, thus avoiding the phototoxicity associated with blue illumination light, as well as the majority of the cellular autofluorescence. DNA-binding dyes can also be used to target conjugated proteins to the DNA by using the dye as a homing beacon. This strategy has been used to target regulatory machinery and labels required for further analysis by other complementary tools, such as mass spectrometry [34].

Unnatural nucleic acids offer a promising new way to bind fluorescent dyes directly to the chromosome with bio-orthogonal methods, such as click chemistry. Recently, organic fluorophore derivatives containing a triple bond were attached to azide-functionalized nucleotides in live cells [35,36].

An alternative to external labeling is making use of the cellular DNA-labeling machinery. Fluorescently labeled histones (Figure 1B) are chimeric proteins consisting of a histone (e.g. H2B, which is ubiquitous throughout the chromatin) fused to a fluorescent protein (FP) [3739]; specifically, a gene encoding for a fluorescent protein is added to the histone coding sequence, resulting in a fluorescently tagged, normally operating histone. Another option, which incorporates a synthetic fluorophore, is to add a motif-recognizing protein that can bind a fluorescent derivative (e.g. HaloTag [40] and PYP3R-tag [41]).

For sensing major cellular-level events, such as the cell cycle and cell division, or characterizing chromosomal territories, fluorescently tagged, DNA-binding proteins (Figure 1B) are applicable [9,42,43]. These proteins typically bind to DNA at specific motifs [44] or are recruited via secondary interactions with other DNA-binding proteins [45]. Locus-specific labeling can be performed by inserting artificial loci which contain motifs recognized by specific proteins. The ParB–INT-type system recruits ParB protein to the parS DNA motif (also called INT), where it clusters, but can be detached during transcription [46,47]. The repressor–operator assay similarly recruits protein to the operator motif; however, unlike ParB–INT interactions, the repressors do not cluster: one repressor recognizes one operator (Figure 1C) [21,4852]. To achieve a sufficiently high signal-to-background ratio to image DNA in live cells, at least several labels must be attached in close proximity; this is required to overcome the background of unbound fluorescent molecules and cellular autofluorescence.

Fluorophore-linked zinc finger proteins (ZFPs) [53,54] and transcription activator-like effectors (TALEs) [5558] are methods based on sequence-recognizing proteins. ZFs are a broad class of proteins capable of recognizing specific structures in DNA, RNA or other proteins [59]. DNA-binding ZFPs typically contain multiple domains to increase specificity and can be engineered to target any particular sequence [60]. TALEs consist of specific nucleotide-binding subdomains that can be combined to recognize any specific sequence [61]. To meet the aforementioned fluorescent-brightness requirement, it would be necessary to either express a library of specific DNA binders for adjacent sequences; therefore, these methods are practically applicable to loci containing repetitive sequences.

A novel, promising method for DNA labeling has been recently developed from the CRISPR–Cas9 system, termed CRISPR–dCas9 (Figure 1D) [6265]. Clustered, regularly interspaced, short, palindromic repeats (CRISPRs) are a specific locus on the bacterial genome that contains short sequences of viral origin [66,67]. CRISPR–Cas9 functions as an antiviral system in bacteria and archaea that recognizes and cleaves viral DNA [65,66,6873]; binding to DNA is facilitated by an incorporated target-complementary RNA strand. As these sequences are significantly easier to engineer compared with developing novel sequence-specific proteins, the CRISPR/dCas9 system has the potential to meet all of the requirements for large-scale, nonrepetitive labeling [11,63,74].

There are three components in the CRISPR–Cas9 complex that are responsible for sequence-specific targeting and cleavage of viral DNA (Figure 3). First, there is the enzyme Cas9, an endonuclease that causes a double-strand break in DNA [73,75]. Second, CRISPR RNA (crRNA) is transcribed from the CRISPR locus and contains both a common sequence (called the repeat sequence) and an antiviral DNA sequence that is responsible for the specificity to the target (called the spacer) [75]. Finally, trans-activating crRNA (tracrRNA) is transcribed separately from the crRNA and contains two parts: a strain-specific stem loop which binds to the Cas9, and an anti-repeat sequence which hybridizes the repeat sequence of the crRNA [76] (Figure 3B). To simplify the system, a synthetically designed linker loop has been used to combine the crRNA and tracrRNA activity into a single-guide RNA (sgRNA or gRNA) molecule [67]. For fluorescent labeling, rather than genome editing, a mutated Cas9 with deactivated nucleases is used, termed dead Cas9 (dCas9). While dCas9 is able to incorporate the gRNA to bind specific sequences, it does not cleave DNA [63,67].

Multicolor CRISPR–dCas9-based DNA labeling.

Figure 3.
Multicolor CRISPR–dCas9-based DNA labeling.

(A) Structures for three orthogonal CRISPR–dCas9 systems, each derived from a different organism: Streptococcus pyogenes (Sp), Streptococcus thermophiles (St1) and Neisseria meningitides (Nm). The fluorescently labeled constructs associate with their respective gRNA in a three-color assay, adapted from ref. [77]. (B) CRISPR-Display uses gRNA designed to contain additional stem loops (green dashed lines) which are recognized by fluorescently marked RNA-binding proteins, adapted from ref. [79]. (C) CRISPRainbow uses stem-loop labeling in a combinatorial fashion to increase the number of effective imaging channels, adapted from ref. [11].

Figure 3.
Multicolor CRISPR–dCas9-based DNA labeling.

(A) Structures for three orthogonal CRISPR–dCas9 systems, each derived from a different organism: Streptococcus pyogenes (Sp), Streptococcus thermophiles (St1) and Neisseria meningitides (Nm). The fluorescently labeled constructs associate with their respective gRNA in a three-color assay, adapted from ref. [77]. (B) CRISPR-Display uses gRNA designed to contain additional stem loops (green dashed lines) which are recognized by fluorescently marked RNA-binding proteins, adapted from ref. [79]. (C) CRISPRainbow uses stem-loop labeling in a combinatorial fashion to increase the number of effective imaging channels, adapted from ref. [11].

To fluorescently label DNA, the dCas9 protein or the gRNA must be attached to a fluorophore (Figures 1D and 3). Functionally, the gRNA stem loop is strain-specific, e.g. Streptococcus pyogenes (Sp) dCas9 only binds Sp-gRNA, and is orthogonal to the three other dCas9s that have been reported thus far [77,78]. This enables multicolor labeling by transfecting different dCas9–FP constructs, each with a unique gRNA [63,77] (Figure 3A). Existing studies have demonstrated three-color labeling for repetitive sequences [77] and dual-color labeling for nonrepetitive loci [63].

An alternative approach to the dCas9–FP fusion is to fluorescently tag the gRNA using proteins that are able to recognize and bind specific RNA structure motifs [79]. The CRISPR-display method uses modified gRNA containing additional stem loops (B) which are recognized by specific viral proteins. Three such RNA/protein systems have been used with CRISPR/dCas9: PP7 [11,79], MS2 [11,79] and BoxB/λN22 [11]. To enhance the brightness of each DNA-binding molecule, the gRNA can be extended to contain multiple stem loops (Qin et al. [80] demonstrated 16 loops). A related technique, CRISPRainbow, takes a combinatorial, multichannel approach using combinations of the three available loop-binding proteins to enable up to seven effective colors (Figure 3C) [11].

Imaging methods and considerations

To choose a suitable method for an imaging experiment (Figure 4), it is helpful to know the desired spatial and temporal resolutions [20,8184]. The confocal microscope, the workhorse microscope in biological laboratories [85], obtains an image by scanning a focused excitation laser spot over the sample (Figure 5A) and by collecting fluorescence with a photodetector after chromatically rejecting scattered excitation light. By placing a pinhole in the imaging path, out-of-focus light is rejected, enabling the confocal microscope to achieve z-sectioning as well as good lateral resolution, relative to widefield imaging. The scanning requirements of the configuration ultimately limit the rate at which a sample can be imaged, although technical advances such as spinning disc confocal microscopy have parallelized this process to significantly improve acquisition speed [86,87]. A comprehensive review for the theory of resolution in confocal microscopy is given by Inoué [88].

Spatial and temporal ranges for various biological processes and imaging techniques.

Figure 4.
Spatial and temporal ranges for various biological processes and imaging techniques.

(A) DNA-related biological processes take place over a broad range of timescales from microseconds to many hours. (B) Imaging techniques have a range of spatial and temporal limits. Circles represent the typical spatial and temporal capabilities of 2D imaging. Crosses show the decreased temporal and spatial resolution in 3D imaging (depth). Three-dimensional methods that involve scanning, i.e. confocal (yellow) or widefield-z-scanning (blue), sacrifice temporal resolution for 3D imaging. Scanning-free methods, e.g. PSF Engineering (green), do not sacrifice temporal resolution; however, in nearly all methods, the z-resolution is typically lower than that obtained for xy.

Figure 4.
Spatial and temporal ranges for various biological processes and imaging techniques.

(A) DNA-related biological processes take place over a broad range of timescales from microseconds to many hours. (B) Imaging techniques have a range of spatial and temporal limits. Circles represent the typical spatial and temporal capabilities of 2D imaging. Crosses show the decreased temporal and spatial resolution in 3D imaging (depth). Three-dimensional methods that involve scanning, i.e. confocal (yellow) or widefield-z-scanning (blue), sacrifice temporal resolution for 3D imaging. Scanning-free methods, e.g. PSF Engineering (green), do not sacrifice temporal resolution; however, in nearly all methods, the z-resolution is typically lower than that obtained for xy.

Optical microscopy modalities.

Figure 5.
Optical microscopy modalities.

The top row depicts a cell with fluorescently labeled loci in the nucleus being imaged with an inverted microscope. The bottom row shows the acquired images. The regions highlighted in red represent the section of an image acquired in a single snapshot. (A) A standard confocal microscope scans a diffraction-limited spot throughout the sample to acquire a 3D image. (B) Standard widefield imaging uses a collimated beam to illuminate the entire sample simultaneously. 3D information is obtained by adjusting the focus to scan vertically through the sample. (C) Particle tracking by PSF engineering modifies the shape of an image formed by a point source on the detector, which can be used to encode additional information, e.g. z-position or color in the resulting image. (D) Single-molecule localization microscopy (referred to as SMLM, STORM, dSTORM, PALM and fPALM) uses the on–off blinking characteristic of fluorescent molecules to capture random subsets of the emitter population within a sample during different frames. Each image is then processed and combined to create a final, high-resolution reconstruction.

Figure 5.
Optical microscopy modalities.

The top row depicts a cell with fluorescently labeled loci in the nucleus being imaged with an inverted microscope. The bottom row shows the acquired images. The regions highlighted in red represent the section of an image acquired in a single snapshot. (A) A standard confocal microscope scans a diffraction-limited spot throughout the sample to acquire a 3D image. (B) Standard widefield imaging uses a collimated beam to illuminate the entire sample simultaneously. 3D information is obtained by adjusting the focus to scan vertically through the sample. (C) Particle tracking by PSF engineering modifies the shape of an image formed by a point source on the detector, which can be used to encode additional information, e.g. z-position or color in the resulting image. (D) Single-molecule localization microscopy (referred to as SMLM, STORM, dSTORM, PALM and fPALM) uses the on–off blinking characteristic of fluorescent molecules to capture random subsets of the emitter population within a sample during different frames. Each image is then processed and combined to create a final, high-resolution reconstruction.

Widefield microscopy techniques (Figure 5B–D) are capable of faster image acquisition compared with their scanning-based counterparts. In short, an entire image is relayed onto a 2D detector, i.e. a camera. Axial (z) information is obtained by scanning the focus up and down. Relative to confocal imaging, there is a moderate degradation in the spatial resolution, ∼30%, due to the removal of the pinhole [88]. In epi-illumination (Figure 5B), a collimated beam of excitation light illuminates fluorophores in the entire volume of the sample. This approach is simple, but causes some increase in background due to out-of-focus emitters, unnecessary photobleaching and phototoxicity [89]. Light-sheet microscopy, which changes the illumination scheme to excite only a thin, flat section of the sample from the side, partially mitigates this problem. Various light-sheet modalities have been suggested over the years with moderate to very high degrees of implementation complexity [9094].

The main advantages of widefield over confocal microscopy are the imaging speed and the simultaneous acquisition of data; however, both benefits are largely mitigated if scanning is still required in the z-axis. Furthermore, performing imaging in a single plane of live cells also leaves out the potentially critical information about what is happening in the third dimension [95]; the nucleus can be quite thick (greater than several microns), and the motion of loci [9] can extend beyond the depth of field for high magnification/high numerical aperture objectives (which is only several hundred nanometers).

Point spread function (PSF) engineering has been employed to enable scan-free, three-dimensional, molecular orientation and multicolor imaging capabilities [21,96103]. In short, PSF engineering works by modifying the image formed on a detector. When light is collected from a point source (e.g. a fluorescently labeled DNA locus), it interacts with an additional optical element in the fluorescence detection path of the microscope, encoding additional information contained in the phase onto the resulting image [104]. Using concepts from estimation theory [105], it is possible to calculate the optimal PSF for a given parameter set of imaging conditions, including extended z-depth [99,100]. This improvement can be used to do scan-free, deep imaging without losing the z-information of the sample. A key requirement of this method is that the point sources be sufficiently separate (either spatially or chromatically) that they can be analyzed (Figure 5C).

Single-molecule localization microscopy solves the previously mentioned problem of label density by creating two populations of fluorescent labels: a set of actively emitting fluorophores that are sparse throughout the sample and a second, denser set of inactivated (off) molecules [106108]. By capturing a movie of stochastic switching events, it is possible to analyze the PSFs of many emitters that have been recorded at different times (Figure 5D). This technique is inherently slow because it requires sufficient sampling of many molecules to form an image. The key advantage is in the achieved spatial resolution, ∼10× better than the diffraction limit [109] (down to tens of nanometers), and the technique has been used to obtain high-resolution chromatin images [110]. Stimulated emission depletion (STED) is another super-resolution method based on a confocal configuration of the microscope [111,112] that has been used to image chromatin [113]. Unlike the blinking-based techniques, which require chemical photoswitching, STED images can be acquired at rates near those of canonical confocal microscopy while obtaining higher spatial resolution [114].

The relative tradeoffs of the imaging modalities are summarized in Figure 4B, where the typical spatio-temporal domains are shown in different colors. The circles represent the lateral and temporal resolution achievable in 2D imaging under normal conditions. The crosses show the approximate axial resolution as well as the associated temporal cost of imaging in three dimensions.

Example applications

In this section, we sample the application space of live-cell DNA imaging, by describing selected examples of reported studies. A summary of the key applications and suitable DNA-labeling methods for live-cell imaging are given in Table 1.

Table 1
Key application of DNA imaging in live cells
Application Suitable labeling method Reference 
Cell viability test DNA dye stain: label all DNA (intercalating dye) [32
Tracking all chromatin or chromosome territory dynamics DNA dye stain
Fluorescently labeled proteins: fluorescent histones, telomere-binding proteins
CRISPR–dCas9: targeting telomeres 
[9,37,41,115
Tracking cell cycle, DNA replication or cell division Unnatural nucleotides: bound to fluorescent probes
Fluorescently labeled proteins: fluorescently marked histones, PCNA and cell-cycle regulatory proteins
TALE: targeting centromeres, telomeres or satellites 
[35,42,43,56,57,116
Tracking specific loci dynamics in yeast cells Repressor–operator assay (e.g. lacO/lacI, tetR/tetO, etc.).
ParB–INT and related systems 
[21,46,48,49,51,117,118
Tracking of specific loci containing repetitive sequences Zinc finger protein (ZFP)
CRISPR–dCas9: targeting unique sub-telemetric satellites 
[11,53,77,119
Tracking of specific, nonrepetitive loci CRISPR–dCas9 methods [63,80,120
Application Suitable labeling method Reference 
Cell viability test DNA dye stain: label all DNA (intercalating dye) [32
Tracking all chromatin or chromosome territory dynamics DNA dye stain
Fluorescently labeled proteins: fluorescent histones, telomere-binding proteins
CRISPR–dCas9: targeting telomeres 
[9,37,41,115
Tracking cell cycle, DNA replication or cell division Unnatural nucleotides: bound to fluorescent probes
Fluorescently labeled proteins: fluorescently marked histones, PCNA and cell-cycle regulatory proteins
TALE: targeting centromeres, telomeres or satellites 
[35,42,43,56,57,116
Tracking specific loci dynamics in yeast cells Repressor–operator assay (e.g. lacO/lacI, tetR/tetO, etc.).
ParB–INT and related systems 
[21,46,48,49,51,117,118
Tracking of specific loci containing repetitive sequences Zinc finger protein (ZFP)
CRISPR–dCas9: targeting unique sub-telemetric satellites 
[11,53,77,119
Tracking of specific, nonrepetitive loci CRISPR–dCas9 methods [63,80,120

Non-sequence-specific DNA labeling

This group of applications focuses on major cellular events. These include: nonchromosome-specific characterization of replication foci dynamics [116]; differences in loci replication fork dynamics between DNA replication and DNA damage repair by fluorescently labeling the proliferating cell nuclear antigen (PCNA) [36,43]; tracking all phases of the cell cycle, using four-color imaging of fluorescently labeled histones (H1.0), cell-cycle factors (SLBP, cdt1) and PCNA [42] and tracking cancer-cell migration, also using fluorescently labeled histones (H2B) [121]. Specific sequence-binding proteins can also be used for more general purposes, by targeting recurring sequences, such as those in the centromeres, telomeres and major satellites (highly repetitive sequence of noncoding DNA). For example, TALE was used to track the cell-division phases [56] and late-replication timing of major satellite loci [57], and fluorescently labeled, telomere-binding proteins were used to track the diffusion dynamics of chromosome territories in live human cells [9].

CRISPR–dCas9 has been used to study the dynamics of telomeres in plant cells [115], and to image specific chromosomes in mouse cells, by targeting the satellite regions [122]. As described previously, the orthogonal dCas9s, derived from different species, can be used to uniquely label multiple specific chromosomes simultaneously (Figure 3A) [77]. Using a combinatorial strategy, six chromosomes were imaged simultaneously by labeling the gRNA (Figure 3C) [11].

Locus-specific labeling

To track processes such as locus-specific chromatin reorganization, specific loci have been labeled and imaged. One pertinent process, DNA repair, is known to play a major role in cancer suppression [123], immunology [124] and neurodegenerative disorders [125127]. Yeast are an especially convenient model organism for this process because they undergo spontaneous changes between genotypes via DNA break and repair, analogous to mechanisms in mammals [46,128]. Several studies have shown this unique event using the ParB–INT [46] system and the repressor–operator assay [49,129,130].

The repressor–operator assay has been used for numerous studies on chromatin dynamics [22,46,48,131,132]. This method, in combination with PSF engineering, has been used also to correlate loci motion [21] and to quantify inter-locus distance upon transcription [117]. While methods using inserted repetitive sequences for imaging are routine in yeast and Drosophila [133] and are suitable in plant cells [134], there has only recently been a report in mammalian cells [135].

Labeling methods for endogenous sequences in specific loci depend on whether or not these loci contain repetitive sequences (several examples below). When the target locus contains repetitive sequences: ZFP, TALE and CRISPR methods may be used; for example, labeling specific repetitive sequences in the 5S rRNA gene was reported using ZFP [53] as well as CRISPR–dCas9 [78]. A combinatorial, two-color assay was used to simultaneously image three loci via two dCas9s (Sp–dCas9–FP and Sa–dCas9–FP) and gRNAs targeting the repetitive sequences in the Muc4 and 5S rRNA genes [78]. Another two-color assay used a CRISPR-display-like method (gRNA labeling) to bind and track the repetitive sequences in the IgH (immunoglobulin heavy chain) and Akap6 genes on mouse chromosome 12 [122].

A highly challenging application is the labeling and imaging of nonrepetitive loci in mammalian cells. There have been two approaches to solving this problem, both based on CRISPR methods which use an array of adjacent target sites. One uses fluorescently labeled dCas9 [63]; the second extends the gRNA by adding stem loops which are then recognized by labeled, RNA-binding proteins [120,136]. Recently, the two approaches have been combined for simultaneous, multicolor imaging of active and nonactive loci [80].

Summary

With the advent of novel microscopy and labeling methods in recent years, live-cell chromatin imaging is emerging as a uniquely powerful tool for elucidating dynamic processes which are otherwise extremely difficult to observe, due to their associated spatio-temporal scales. Such dynamic imaging completes the static and typically population-based picture obtained by 3C and related approaches. Future development of new labeling strategies and imaging methods will enable researchers to move toward a holy grail of DNA imaging: time-resolved, 3D positions of the entire genome, with information on sequence and epigenetic modification [137].

Abbreviations

     
  • (s)gRNA

    (sequence) guide RNA

  •  
  • 3C

    Chromatin Conformation Capture

  •  
  • CRISPR

    Clustered, Regularly Interspaced, Short, Palindromic Repeats

  •  
  • crRNA

    CRISPR RNA

  •  
  • dCas9

    dead Cas9

  •  
  • FP

    fluorescent protein

  •  
  • IgH

    immunoglobulin heavy chain

  •  
  • Nm

    Neisseria meningitides

  •  
  • PCNA

    proliferating cell nuclear antigen

  •  
  • PSF

    point spread function

  •  
  • Sp

    Streptococcus pyogenes

  •  
  • St1

    Streptococcus thermophiles

  •  
  • TALE

    transcription activator-like effectors

  •  
  • tracrRNA

    trans-activating CRISPR RNA

  •  
  • ZFP

    zinc finger proteins

Author Contribution

All authors contributed to writing the manuscript.

Funding

L.E.W. and Y.S. are supported by the Zuckerman Foundation. Y.S. is supported in part by a Career Advancement Chairship from the Technion, Israel Institute of Technology.

Competing Interests

The Authors declare that there are no competing interests associated with the manuscript.

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