Nanoscopic Cellular Imaging: Confinement Broadens Understanding

Nanoscopic Cellular Imaging: Confinement Broadens Understanding Stephen A. Lee,†,§ Aleks Ponjavic,‡,§ Chanrith Siv,†,§ Steven F. Lee,*,‡ and Julie S. Biteen*,† †

Department of Chemistry, University of Michigan, Ann Arbor, Michigan 48109, United States Department of Chemistry, Cambridge University, Cambridge CB2 1EW, United Kingdom

‡

ABSTRACT: In recent years, single-molecule fluorescence imaging has been reconciling a fundamental mismatch between optical microscopy and subcellular biophysics. However, the next step in nanoscale imaging in living cells can be accessed only by optical excitation confinement geometries. Here, we review three methods of confinement that can enable nanoscale imaging in living cells: excitation confinement by laser illumination with beam shaping; physical confinement by micron-scale geometries in bacterial cells; and nanoscale confinement by nanophotonics. KEYWORDS: super-resolution microscopy, single-molecule imaging, nanophotonics, light-sheet microscopy, bacterial biophysics, optical sectioning, plasmonic enhancement

T

methods, the location of each molecule is determined by analyzing the detected image. The localization precisionor spatial uncertainty on the position of the object11in singlemolecule imaging is fundamentally limitless; this precision and thus the achievable spatial resolution is only practically limited by the number of photons detected above background.18 Modern dyes have excellent fluorescence quantum yields, good absorption cross sections, and can be pumped at power densities approaching saturation.19 Efficient detectors, high numerical aperture (NA) objective lenses, and sharp filters have improved SMF detection. However, out-of-focus background limits the contrast of the fluorescence signal, particularly in biological specimens, which are prone to intrinsic background fluorescence.20 In the presence of background emission, the ability to localize single molecules precisely is degraded. Thus, although fluorescence microscopy is generalizable to a wide variety of samples, thick samples have evaded good superresolution measurements due to this intrinsic fluorescent background and concerns about phototoxicity.21 These technological limitations have made the extension of SMF imaging to tissues and organisms difficult. Thus, SMF imaging must be improved by reducing the fluorescence background, and here we contrast the advantages and disadvantages of three orthogonal styles of confinement (Figure 1 and Table 1): (1) excitation confinement through instrument innovation at the micron scale, (2) physical confinement in bacterial cells at the

here exists a fundamental mismatch between the limited spatial resolution of a light microscope and the nanoscale world of cellular biophysics. As a result, though the mysteries of subcellular biology have been investigated indirectly by biochemical approaches, the ability to directly observe intracellular processes remained elusive for many years. More recently, super-resolution innovations such as single-molecule fluorescence (SMF) microscopy and stimulated emission depletion (STED) microscopy, combined with a growing interest in developing the methods of physical chemistry for biology, have imparted an ability to detect, probe, and understand the processes occurring in cells.1 In particular, the development of SMF microscopy has enabled the direct visualization of dynamics,2−4 stoichiometry,5,6 and organization7,8 inside of cells. In this short review, we focus on confinement, a theme that has greatly increased the variety of biological specimens and the level of detail that can be probed by single-molecule superresolution microscopy. Other approaches, like STED microscopy, as well as other aspects of super-resolution imaging, such as the labeling, sample preparation, and microscopy innovations that have increased resolution, speed, and performance, have been reviewed extensively elsewhere.9−12 SMF superlocalization microscopy permits single-molecule tracking,13−15 and super-resolution approaches like photoactivated localization microscopy (PALM)16 and stochastic optical reconstruction microscopy (STORM)17 overcome the diffraction limit in a densely labeled sample by detecting low concentrations of reporter molecules, trading off temporal resolution for increased spatial resolution. In all of these single-molecule © 2016 American Chemical Society

Received: April 29, 2016 Accepted: September 7, 2016 Published: September 7, 2016 8143

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153

Review

www.acsnano.org

Review

ACS Nano

Figure 1. Confinement in cellular microscopy. (a) Typical relative reduction in background signal for a modern high numerical aperture objective lens as a function of axial confinement. (b−f) Modes of confinement and their relative scales.

Table 1. Properties of Excitation Confinement in Common Biophysical Imaging Techniques strategy

a

technique

scale (nm)

typical power densities (kW/cm2)

Evanescence

TIRF

∼200

0.01−10

Scanning

Plasmonics Confocal

∼10 ∼300

0.1−10 102−105

STEDa

40−70

102−104

Physical

Bacteria

1000−5000

0.1−10

Light-sheet

HILO

5000−10 000

0.1−10

SPIM Beam Shaping

500−5000 ∼300

0.1−10 0.1−1

advantages Simple implementation; High contrast; High speed Best potential contrast No surface requirement; Medium contrast Greatly improved contrast Small sample; Simple implementation Simple implementation; No surface requirement; High speed Improved contrast Even better contrast; Large field of view

disadvantages Restricted to interfaces Requires metal nanoparticles Slow due to point scanning; Prone to phototoxicity and photobleaching High phototoxicity and photobleaching; Slow due to scanning Limited systems; Sample size limitation; Intracellular labeling with dyes is difficult Low contrast; Small field of view due to aperture Optical access issues Expensive and complicated optics

For comparison, this refers to continuous excitation STED; in the case of pulsed STED the instantaneous power is far higher than the quoted value.

submicron scale, and (3) field confinement though plasmonic interactions at the nanoscale. The intent of this review is not to be exhaustive but rather to introduce each confinement style and give examples of major recent breakthroughs enabled by the technology.

optical axis.22 This probe volume is then scanned across the sample to build up an image. Scanning can be performed both in two23 and three24 dimensions. Still, while confocal confinement achieves diffraction-limited spatial resolution as well as background suppression, scanning decreases the practical temporal resolution relative to widefield illumination, worsens photobleaching,25,26 and increases phototoxicity.27 Despite these drawbacks, video-rate confocal SMF imaging has been achieved by removing the confocal pinhole to eliminate the need to scan, replacing the photodiode with a CCD array, and using optical sectioning to ensure sufficient contrast. Such video-rate confocal imaging has measured singlemolecule membrane receptor diffusion on the cell surface28 and resolved transient single-molecule dynamics in the nuclear pore complex (Figure 2a,b).29 Stimulated Emission Depletion (STED) Microscopy (Laterally 10−100 nm). STED microscopy increases the

EXCITATION CONFINEMENT Limiting the light-exposed sample volume enables singlemolecule fluorescence imaging (Figure 1a). In particular, we focus here on axial confinement (along the optical axis), which improves SMF imaging by reducing unwanted background light and increasing the image contrast. The techniques used for such optical confinement are summarized in Figure 1b−f and Table 1. Confocal Microscopy (200−500 nm). In confocal microscopy, a physical pinhole limits the collection volume radius to ∼200 nm in the focal plane and ∼500 nm along the 8144

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153

Review

ACS Nano

Figure 2. (a) Translocation of transport receptor importin β (red) through the nuclear pore complex (green) acquired with high-speed confocal microscopy. Time in seconds and scale bar is 1 μm.29 (b) Localization density map constructed from results in (a) with an overlay of the nuclear pore complex structure (yellow). Reproduced from ref 29. Copyright (2010) National Academy of Sciences. (c) HILO microscopy excitation geometry. Reprinted from ref 48 by permission from Macmillan Publishers Ltd: Nature Methods 5, 159, copyright (2008). (d) 3D diffraction-limited (left) and 3D super-resolution images (right) of the bacterial cytoskeleton (red) and membrane (gray) of Caulobacter crescentus, acquired using HILO and a double helix point spread function. Inset: white-light image of the same cell. Reproduced from ref 8. Copyright (2011) National Academy of Sciences. (e) Geometry used for SMF light-sheet microscopy. Reproduced from ref 56. Creative Commons. (f) Export of mRNA (top-white, bottom-red) across the nuclear envelope (bottom-green) observed by light-sheet microscopy. Reproduced from ref 57. Copyright (2012) National Academy of Sciences.

coating,43 the substrate may interact with extracellular membrane proteins. For example, the human T-cell receptor is partially immobilized on polylysine-coated surfaces.44 In certain cases, the surface constraint of TIRF can be relaxed,45 but these approaches, although impressive, are limited to specific cell shapes. Highly Inclined and Laminated Optical Sheet (HILO) Microscopy (5−10 μm). Illuminating the sample through the objective lens below the critical angle for TIRF produces a sheet of light inclined away from the coverslip surface.46,47 This method of excitation confinement is referred to as inclined or oblique illumination,11 or sometimes “pseudo-TIRF”.46 Adding an aperture in the excitation beam path produces a laminated thin optical sheet to further reduce the sheet thickness to ∼5− 10 μm (Figure 2c).48 This HILO microscopy enables SMF imaging as deep as ∼20 μm into cells and tissue. HILO has been used to study the dynamics and stoichiometry of ATP binding proteins on the apical surface of cells6 and to enable intracellular super-resolution imaging with nanopipette delivery.49 Furthermore, the cytoskeletal structure of the Caulobacter crescentus bacterium was resolved by combining HILO and a 3D super-resolution double-helix point spread function microscope (Figure 2d).8 Light-Sheet Microscopy (0.5−5 μm). Instead of throughobjective widefield epi-illumination, excitation light can be introduced through a second objective lens perpendicular to the optical axis of the collection objective. Typically, a cylindrical lens is used to create a lateral “sheet” in the sample plane, with an axial depth of ∼0.5−5 μm, limited by the NA of the second objective lens (Figure 1c).50 In most cases, the

resolution and contrast of confocal microscopy by using a depletion laser to reduce the effective confocal volume by providing a depletion zone about the beam profile (Figure 1b).30,31 With STED, lateral resolutions of ∼2 nm have been achieved for nitrogen vacancies in diamond,32 and lateral resolutions as good as ∼40−70 nm are typically achieved in biological samples.33−35 STED can be done with high label densities, and single-molecule STED imaging has been demonstrated on static fluorophores.30 Unfortunately, the phototoxicity of STED is even larger than that of confocal microscopy due to the ∼GW/cm2 STED beam power densities. This issue can be somewhat alleviated with time gating36 or reversible saturable optical fluorescence transitions.37 Evanescent Illumination: Total Internal Reflection Fluorescence (TIRF) Microscopy (100−300 nm). Nonpropagating evanescent waves are created when light undergoes total internal reflection at a boundary. Because evanescent waves decay nonlinearly within ∼200 nm, TIRF illumination confines excitation to the sample surface, drastically reducing unwanted background fluorescence, and improving contrast (Figure 1e).38 Thus, though SMF detection has been possible for decades,39,40 TIRF microscopy was the major breakthrough that enabled SMF imaging in biological samples,38 and TIRF has become one of the most common and important techniques for SMF imaging in live cells.41 The major limitation of TIRF is that it practically constrains imaging to surfaces. This drawback is important because firm substrates, such as glass coverslips, can deleteriously affect soft, flexible biological membranes.42 Furthermore, even when such a nonphysiological surface is coated with a soft, biologically inert polyelectrolyte 8145

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153

Review

ACS Nano

most cases, physical confinement within these small cells enables high-contrast single-molecule detection and tracking inside bacterial cells without the need for 3D optics or excitation confinement.70−72 Because bacterial cell biology is accessible with the current state-of-the-art SMF microscopy, important biological processes can already be understood in bacterial cells. This increased understanding can be translated to eukaryotic systems as improved methods developmentfor instance, the excitation confinement approaches described abovecontinues. On the other hand, the physical geometry of a bacterium produces important challenges for visualizing fluorescent molecules inside living bacteria: for instance, due to a small total volume, barriers to dye entry, and possible artifacts induced by fluorescence labeling.73,74 Emerging solutions to the difficulties of subcellular imaging inside bacterial cells include a high-throughput method to introduce fluorescently labeled DNA and proteins into cells with electroporation75 and adapting image correlation approaches to highly confined volumes.76 Single-Molecule Tracking in Bacteria. Fluorescent protein fusions are the most widely used labels for fluorescence imaging in living bacterial cells. Highly specific fusions can be genetically encoded, and photoactivatable fluorescent proteins77 can be switched from a dark state to a fluorescent state one at a time to enable single-particle tracking superresolution imaging (sptPALM)78 of molecules in living bacterial cells to directly observe protein dynamics and understand protein function. Recently, the spatial distribution of translational and free ribosomal subunits measured by sptPALM in Escherichia coli (Figure 3a) gave rise to a model of fast translation initiation of nascent mRNA transcripts that was not previously revealed with electron or fluorescence microscopy.73 Single-molecule localization and tracking during amino acid starvation in E. coli uncovered that during stringent response, RelA synthesizes hyper-phosphorylated GTP/GDP while bound to the 70S ribosome (Figure 3b).79 As well, two-color sptPALM in Bacillus subtilis led to the discovery that the highly conserved DNA mismatch repair protein MutS can only bind mismatches near the replisome and demonstrated that mismatch detection increases the MutS speed in vivo (Figure 3c).80 E. coli and B. subtilis are model bacteria for single-molecule imaging, but there is increasing research interest in studies of pathways in pathogenic and commensal bacteria. An emerging approach to single-molecule tracking in the heterogeneous cellular environment is a single-step analysis based on the cumulative probability distribution (CPD) of mean square displacements from all trajectories.81 With this approach, singlemolecule investigations of cholera toxin regulation82 in Vibrio cholerae showed how proteins regulate cholera toxin expression while remaining localized to the cell inner membrane during DNA binding and transcription activation.83 The community is becoming increasingly aware of the importance of creating genetic fusions at the native promoter to ensure that native function and expression levels are maintained. Recently, CPD investigations of the virulence regulation protein TcpP labeled with the photoactivatable fluorescent protein PAmCherry at the native chromosomal locus in V. cholerae revealed a faster diffusing population that was not previously captured with ectopic expression (red curve in Figure 3d).84 Importantly, based on endogenously expressed TcpP-PAmCherry the underlying mechanism of membrane-

decreased axial thickness in this single-plane illumination microscopy (SPIM) improves imaging contrast relative to HILO microscopy. The thickness of a light sheet can vary across the image, but zoom lenses51 or two-photon excitation52 produce more uniform illumination. Furthermore, inverted SPIM (iSPIM) enables diffraction-limited 3D imaging of large samples based on placing the excitation and acquisition objective lenses in a perpendicular configuration on separate piezo scanners.53 Though multiple planes are required for 3D reconstruction, 3D imaging can be done at high speeds by continuously switching excitation and acquisition between the objective lenses in dual-view iSPIM.54 SPIM has been used to study transient transcription factor binding in the cell nucleus,55 to image single RNA molecules inside cells,56 and to elucidate the mechanism of nuclear transport at a single-molecule level.57 SPIM can resolve the axial position of single molecules to the scale of the sheet thickness (Figure 2e,f). SPIM can also be combined with 3D localization methods such as the doublehelix point spread function for 3D intracellular diffusion measurements.58 However, light-sheet microscopy imaging poses some engineering challenges, as it is difficult to introduce perpendicular light in close proximity to the acquisition objective lens, due to the fact that high NA objectives have short working distances. These issues of access can be overcome with microscopic mirrors in reflected light-sheet microscopy,55 and in single-objective SPIM.59 Beam Shaping (300 nm). Most light-sheet microscopes use a standard TEM00 (Gaussian-like) transverse mode laser. Beam shaping can be combined with light-sheet microscopy to produce even thinner sheets. For instance, a Bessel beam can be formed using holography or an axicon lens, yielding a beam of constant radius over much larger distances.60 Bessel beams have the major advantage of being reconstructive, which limits aberrations caused by scattering and enables imaging deep into tissue. Furthermore, the combination of Bessel beams and structured illumination makes it possible to create very thin beams that surpass the diffraction limit (∼300 nm).60 By quickly dithering this beam, a virtual 2D light sheet can be created; this Bessel beam plane illumination provides superior optical sectioning compared to the standard light sheet.61 Alternatively, curved Airy laser beams can be created using holography to form a special interference pattern.62 Light-sheet microscopy with Airy beams retains the long field of view offered by Bessel beam light sheets while improving image contrast.63 Alternatively, beam scanning can be entirely avoided by directly creating a “lattice” light sheet.64 By combining lattice light-sheet microscopy with structured illumination, it is possible to perform high-speed super-resolution imaging of an entire cell.65 State of the art optical sectioning methods and lattice light-sheet microscopy have also recently been combined to study the dynamics of transcription factor proteins in stem cells at a single-molecule level demonstrating the extensive development and power of excitation confinement.66

PHYSICAL CONFINEMENT WITHIN A BACTERIAL CELL Microscopy in Bacterial Cells (1−5 μm). Excitation confinement enables SMF imaging in thick cells and tissues, but most bacteria are less than 2 μm thick.67 This small size confines fluorescent probes to a thickness that is comparable to the depth of field of a high NA microscope objective lens, thereby eliminating out-of-focus fluorescence (Figure 1f). Though the excitation confinement schemes described in the previous section can be used for imaging in bacteria,68,69 in 8146

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153

Review

ACS Nano

rates, simple growth media, and established genetics make these organisms ideal to understand life’s essential processes. Our understanding of basic bacterial cell physiology, as well as extensions to human health and disease, have benefited tremendously from super-resolution imaging in living bacterial cells,69−72 and it is expected that these advances will transfer to more complicated eukaryotic cells in the coming years.

ELECTROMAGNETIC FIELD CONFINEMENT WITH PLASMONICS Nanophotonics (10−100 nm). Nanotechnology solutions which can complement super-resolution imaging are appearing. The surface plasmon resonances that arise from collective oscillations of free electrons in metals are widely used spectroscopic sensors with sensitivity to refractive index change on a substrate.85 Further confining plasmons to subwavelength sized metal nanoparticles produces localized surface plasmon resonances which act as nanoantennas to enhance scattering process efficiency by concentrating the electromagnetic field, and to increase the absorption and emission of fluorescent dyes by increasing the local density of states.86,87 Nanoparticle plasmonics has found a home in cellular analysis with a host of different applications such as electrochemical sensing, organelle imaging, biosensing, and photothermal imaging.88,89 Though only molecules that are close to a nanoscale feature are enhanced, recent advances have used holography to modulate hotspot positioning in real time, permitting real-time superresolution surface-enhanced Raman scattering (SERS).90 Furthermore, plasmonic materials can confine light for selective and enhanced fluorescence excitation and emission to advance biological imaging.91 Excitingly, plasmonic nanoparticles can improve the brightness and photostability of probes in fluorescence imaging.92 These advantages, which are greatest for low quantum yield emitters, have been observed on the single-molecule level with over 1000-fold enhancements,93 and can also be extended to increase fluorescence brightness for higher quantum yield labels like fluorescent proteins.94 As plasmon-coupled fluorescence emerges for enhanced superresolution imaging, much of the current research on plasmoncoupled fluorescence is focused on studying the physics governing the plasmon-dye system.95 Examinations of the

Figure 3. Single-particle tracking of protein dynamics in bacteria reveals biological mechanisms. (a) Trajectories of individual free and mRNA-bound mEos2-labeled ribosomal subunits in E. coli show the degree of exclusion of bound subunits from the nucleoid. Reproduced from ref 73. Copyright (2014) National Academy of Sciences. (b) Experimental trajectories for RelA-mEos2 in E. coli measure the dynamics of the stringent response.79 (c) The mismatch repair protein MutS-PAmCherry explores the B. subtilis cell, but accumulates near the replisome. Reproduced from ref 80. Copyright (2015) National Academy of Sciences. (d) The virulence regulation protein TcpP-PAmCherry diffuses throughout the V. cholerae cellular membrane, and single-step analysis reveals three modes of motion (red, green, and blue curves).84 All scale bars: 1 μm.

bound transcription regulation in V. cholerae can now be investigated. Overall, bacteria are among the most successful and widespread organisms on Earth, and their rapid growth

Figure 4. (a) Configuration for imaging mobile lipids in a CHO cell membrane via excitation by the confined field about a bowtie nanoaperture (left) and representative fluorescence time trace (right). Reproduced from ref 99. Copyright (2015) American Chemical Society. (b) Configuration for imaging TcpP-PAmCherry in the periplasm of V. cholerae above a gold nanotriangle substrate (left) and associated histograms of fluorescence intensities with and without the plasmonic substrate (right). Reproduced from ref 91. Copyright (2016) American Chemical Society. 8147

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153

Review

ACS Nano

Figure 5. Single-molecule imaging of microbial community members. (a) A microbiota is grown in a culture flask, (b) an aliquot of the coculture is flowed through a nanofluidic device capable of separating and immobilizing bacteria by channel closing, and (c) labeled molecules within individual cells are imaged in situ using SMF microscopy.

spectral dependence, emission spectrum reshaping, distance dependence, and emission patterns have all improved our understanding of plasmon-enhanced fluorescence.87,94−98 Live-cell fluorescence imaging achieves excitation confinement and emission enhancement based on plasmonic substrates directly underneath the cell (Figure 1d). For instance, confined light delivered via bowtie nanoapertures (BNAs) was used to image single fluorescent membrane lipids in CHO cells.99 These BNAs increased confinement and electric field density to produce an array of pinholes for multiplexed confocal imaging. The volumetric restriction obtained more accurate diffusion coefficients and thus a more precise model of membrane fluidity (Figure 4a). In a widefield plasmon-enhanced SMF approach, nanosphere lithography was used to fabricate large arrays of gold nanotriangle to plasmonically enhance the fluorescence of the V. cholerae virulence regulator TcpPmCherry.91 The nanotriangles doubled the mCherry brightness in living cells (Figure 4b). These 2-fold enhancements can be further improved by substrate optimization; enhancements in excess of 15 times are observed in vitro.95 Overall, plasmonics stands to fundamentally improve super-resolution by addressing SMF limitations through excitation confinement, enhanced electric field intensities, and increased local density of states.

Figure 6. Single-cell analysis on a plasmonic substrate within a microfluidic channel will permit active control of the cellular environment. Two intracellular fluorescent proteins (red and green) couple to the plasmonic substrate for plasmon-enhanced two-color single-molecule imaging. (Inset) Electric field enhancement above each plasmonic nanotriangle (red: high field).

As microscopes become more complicated, more intricate sample geometries are required. Taking advantage of the freedom afforded by light-sheet microscopy to image single molecules in cells physically suspended away from a surface requires immobilization in space using methods such as gelation107 and optical trapping.108 It will be crucial to verify that cells remain unperturbed by establishing the influence that such immobilization methods have on the resting cell state.109 Ultimately, leaving the surface has made it possible to study both live and synthetic tissue at the single-molecule level; the combination of optical trapping110 or 3D-printed tissues111 with off-surface optical sectioning provides a unique platform for investigating interactions between cells. Application of these biophysical tools will lead to a wealth of discoveries about how signaling works on an intercellular scale. Direct Observations of Multiple Components in Bacterial Cells. In parallel with technological development focused on imaging thicker and more complex biological samples, bacterial subcellular biology will remain an important target for SMF imaging experiments because these accessible, controllable systems provide natural confinement to the micron scale without complex optics. On the other hand, continuing to answer problems of increasing complexity about subcellular bacterial biophysics will require approaches that advance the state of the art beyond investigations of single proteins in isolated cells.

FUTURE OUTLOOK: TOWARD ANSWERING INCREASINGLY COMPLEX QUESTIONS Improving Microscopy with Optical Solutions. Advanced technologies will continue to augment advanced microscopy by improving contrast and optimizing optical sectioning while still escaping the coverslip surface. The combination of light-sheet microscopy with two-photon excitation or STED microscopy has potential to greatly improve SMF contrast.100,101 Still, excitation confinement can only improve contrast up to a point; at this limit it becomes more beneficial to improve the efficiency of light collection. All microscopes suffer from noise and imaging artifacts due to optical imperfections; adaptive optics can correct these systematic aberrations:102 both holography103 and deformable mirrors104 have improved imaging through wavefront correction. Correcting aberrations becomes increasingly important in multicellular and deep-tissue imaging, where variations in refractive indices cause significant light scattering. Adaptive optics, which can correct for scattering even when using lightsheet microscopy,105 will therefore become indispensable. Alternative solutions to increased hardware complexity will simplify data acquisition. For instance, software-based information retrieval algorithms generate optimal optical sectioning with a “virtual light-sheet”.106 8148

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153

Review

ACS Nano

(Figure 5). This sample connects a microbial community coculture137 to a nanofluidic sorting mechanism138 that prepares the sample for imaging. A more elaborate experiment might include tissue sectioning or imaging through mice139 or rabbits140 to understand how a microbiota interacts with a host.141 Overall, with the use of microfluidics, researchers can multiplex studies on individual cells in a variety of environments and with different stimuli, and extend single-cell measurements to the regime of cellular communities. Finally, looking forward, plasmonics will allow a variety of spectroscopies to be performed on the subcellular level to obtain dynamic chemical and structural information. Additionally, plasmonic nanoparticle arrays will improve the resolution of fluorescence microscopy to the order of the size of the emitter and reduce photobleaching for monitoring real-time protein dynamics.94 For instance, we envision combining plasmonics and nanofluidics to produce a platform for confined, physiological, enhanced imaging in bacteria (Figure 6). Overall, as technologies improve, all of the tools discussed here can be combined to reach higher levels of understanding about fundamental, subcellular biology. Through innovative combinations of confinement approaches, imaging modalities, and nanotechnologies, we will finally close the mismatch between the spatial resolution of light microscopy and the nanoscale world of cellular biophysics to enable a wealth of discoveries.

While most single-molecule work has focused on proteins, for which labeling schemes are more well developed,9 improving the level of subcellular complexity accessible to SMF imaging requires the development of labeling and imaging methods that simultaneously and directly visualize the complex interactions between DNA, RNA, proteins, and lipids in live cells. The CRISPR/Cas9 system is now a near-ubiquitous tool for genome editing in biology.112 Excitingly, fluorescent protein fusions to the endonuclease-defective mutant dCas9 create an irreversibly bound probe capable of superlocalizing a specific chromosomal locus.113 Ongoing research will continue to improve the sgRNA specificity114,115 and will provide multiple dCas9 probes for multicolor imaging.116,117 To visualize RNA, single-molecule fluorescence in situ hybridization (smFISH) targets mRNA transcripts with oligonucleotide fluorescent probes which is used to monitor transcriptional regulation.118 Super-resolution imaging based on smFISH permits precise quantification of gene expression even in bacteria,119 and can be extended for efficient multiplexing.120−122 Another important frontier in bacterial super-resolution imaging is the ability to combine chemical sensing with imaging,123 thereby merging high spatial resolution and accurate chemical specificity. pH- and force-sensing dyes can also be incorporated into single-molecule experiments,124 combining imaging with vibrational spectroscopy permits noninvasive chemical identification,125,126 and complementing nondestructive optical imaging with matrix-assisted laser desorption ionization (MALDI) mass spectrometry imaging maps molecular distributions with high specificity.127 Nanotechnology Solutions to Widen the Scope of Super-Resolution Imaging. Advances in nanotechnology, microfluidics,128,129 and plasmonics will continue to increase opportunities for SMF imaging of cells. Recent advances in fluorescence activated cell sorting (FACS) are improving bacterial cell sorting,130 which is vitally important for studying heterogeneous cell populations. Microfluidic devices allow single cell manipulation and analysis131 to enable multiple experiments on a single cell. In addition to sorting, microfluidics have made physiologically relevant conditions much more accessible. For instance, gradient hydrogels produced by a microfluidic mixing system enabled a study of hematopoietic stem cell differentiation as a function of niche cell concentration.132 As an alternative to chip-based microfluidics, individual cells identified by optical microscopy can be selected, manipulated, and sorted with a modified atomic force microscope with a microchanneled cantilever in fluidic force microscopy.133 Furthermore, though single-cell super-resolution experiments are common, bacterial cells do not exist in isolation; rather it is becoming increasingly obvious that most bacteria function as a part of a microbiome. We envision single-molecule imaging as a bridge for the gap between community-level observations and nanoscale biophysics. On one hand, tools like optical sectioning will find their place in studies of thick biofilms. On the other hand, 2D bacterial biofilm models134,135 will provide confinement in the axial direction while maintaining intercellular connectivity. Alternatively, sample confinement can make single cells behave as if they were in communities: for instance, volumetrically confined single Pseudomonas aeruginosa cells will undergo autoinduced quorum sensing, in an effective community of one.136 We envision a simple apparatus for studying protein function at the single-molecule level in single cells from a microbiome

AUTHOR INFORMATION Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. Author Contributions §

SAL, AP, and CS contributed equally to this work.

Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENTS This work was generously supported by funding from the National Science Foundation (CAREER Award CHE1252322) and the Engineering and Physical Sciences Research Council (EP/L027631/1). We would like to thank the Royal Society for the University Research Fellowship of SFL (UF120277). GLOSSARY Super-resolution imaging, Light microscopy with a spatial resolution smaller than the submicron diffraction limit of light; Single-molecule fluorescence (SMF) microscopy, Sensitive fluorescence imaging attained by detecting and precisely localizing fluorescent emitters; Optical sectioning, Decreasing out-of-focus excitation by illuminating a thin focal plane within a thick sample; Photoactivatable labels, Dyes or fluorescent proteins that can be irreversibly switched from a dark (nonemissive) state to a fluorescent state; Localized surface plasmon resonance, Confined collective free electron oscillation that concentrates the local electromagnetic field and increases the local density of states; Microbiome, Microbial cell community living within a certain environment. REFERENCES (1) Moerner, W. E. Nobel Lecture: Single-Molecule Spectroscopy, Imaging, and Photocontrol: Foundations for Super-Resolution Microscopy. Rev. Mod. Phys. 2015, 87, 1183−1212. 8149

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153

Review

ACS Nano (2) Yildiz, A.; Forkey, J. N.; McKinner, S. A.; Ha, T.; Goldman, Y. E.; Selvin, P. R. Myosin V Walks Hand-Over-Hand: Single Fluorophore Imaging with 1.5-nm Localization. Science 2003, 300, 2061−2065. (3) Drbal, K.; Moertelmaier, M.; Holzhauser, C.; Muhammad, A.; Fuertbauer, E.; Howorka, S.; Hinterberger, M.; Stockinger, H.; Schütz, G. J. Single-Molecule Microscopy Reveals Heterogeneous Dynamics of Lipid Raft Components upon TCR Engagement. Int. Immunol. 2007, 19, 675−684. (4) Hern, J. A.; Baig, A. H.; Mashanov, G. I.; Birdsall, B.; Corrie, J. E. T.; Lazareno, S.; Molloy, J. E.; Birdsall, N. J. M. Formation and Dissociation of M1Muscarinic Receptor Dimers Seen by Total Internal Reflection Fluorescence Imaging of Single Molecules. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 2693−2698. (5) Dunne, P. D.; Fernandes, R. A.; McColl, J.; Yoon, J. W.; James, J. R.; Davis, S. J.; Klenerman, D. DySCo: Quantitating Associations of Membrane Proteins Using Two-Color Single-Molecule Tracking. Biophys. J. 2009, 97, L5−L7. (6) Nagata, K. O.; Nakada, C.; Kasai, R. S.; Kusumi, A.; Ueda, K. ABCA1 Dimer-Monomer Interconversion during HDL Generation Revealed by Single-Molecule Imaging. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 5034−5039. (7) Douglass, A. D.; Vale, R. D. Single-Molecule Microscopy Reveals Plasma Membrane Microdomains Created by Protein-Protein Networks that Exclude or Trap Signaling Molecules in T Cells. Cell 2005, 121, 937−950. (8) Lew, M. D.; Lee, S. F.; Ptacin, J. L.; Lee, M. K.; Tweig, R. J.; Shapiro, L.; Moerner, W. E. Three-Dimensional Superresolution Colocalization of Intracellular Protein Superstructures and the Cell Surface in Live Caulobacter Crescentus. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, E1102−E1110. (9) Fernández-Suárez, M.; Ting, A. Y. Fluorescent Probes for SuperResolution Imaging in Living Cells. Nat. Rev. Mol. Cell Biol. 2008, 9, 929−943. (10) Sahl, S. J.; Moerner, W. Super-Resolution Fluorescence Imaging with Single Molecules. Curr. Opin. Struct. Biol. 2013, 23, 778−787. (11) Deschout, H.; Zanacchi, F. C.; Mlodzianoski, M.; Diaspro, A.; Bewersdorf, J.; Hess, S. T.; Braeckmans, K. Precisely and Accurately Localizing Single Emitters in Fluorescence Microscopy. Nat. Methods 2014, 11, 253−266. (12) Liu, Z.; Lavis, L.; Betzig, E. Imaging Live-Cell Dynamics and Structure at the Single-Molecule Level. Mol. Cell 2015, 58, 644−659. (13) Vrljic, M.; Nishimura, S. Y.; Moerner, W. E. Single-Molecule Tracking. Methods Mol. Biol. 2007, 398, 193−219. (14) Braeuchle, C.; Lamb, D. C.; Michaelis, J. Single Particle Tracking and Single Molecule Energy Transfer; Wiley-VCH: Weinheim, 2010. (15) Kusumi, A.; Tsunoyama, T. A.; Hirosawa, K. M.; Kasai, R. S.; Fujiwara, T. K. Tracking Single Molecules at Work in Living Cells. Nat. Chem. Biol. 2014, 10, 524−532. (16) Betzig, E.; Patterson, G. H.; Sougrat, R.; Lindwasser, O. W.; Olenych, S.; Bonifacino, J. S.; Davidson, M. W.; Lippincott-Schwartz, J.; Hess, H. F. Imaging Intracellular Fluorescent Proteins at Nanometer Resolution. Science 2006, 313, 1642−1645. (17) Rust, M. J.; Bates, M.; Zhuang, X. Sub-Diffraction-Limit Imaging by Stochastic Optical Reconstruction Microscopy (STORM). Nat. Methods 2006, 3, 793−795. (18) Thompson, R. E.; Larson, D. R.; Webb, W. W. Precise Nanometer Localization Analysis for Individual Fluorescent Probes. Biophys. J. 2002, 82, 2775−2783. (19) Osborne, M. A. Small Things Bright and Beautiful: Single Molecule Fluorescence Detection; Imperial College Press: London, 2007; pp 283−312. (20) Benson, R. C.; Meyer, R. A.; Zaruba, M. E.; McKhann, G. M. Cellular Autofluorescence − Is It Due to Flavins? J. Histochem. Cytochem. 1979, 27, 44−48. (21) Wäldchen, S.; Lehmann, J.; Klein, T.; van de Linde, S.; Sauer, M. Light-Induced Cell Damage in Live-Cell Super-Resolution Microscopy. Sci. Rep. 2015, 5, 15348.

(22) Palmer, R. J.; Sternberg, C. Modern Microscopy in Biofilm Research: Confocal Microscopy and Other Approaches. Curr. Opin. Biotechnol. 1999, 10, 263−268. (23) Revankar, C. M.; Cimino, D. F.; Sklar, L. A.; Arterburn, J. B.; Prossnitz, E. R. A Transmembrane Intracellular Estrogen Receptor Mediates Rapid Cell Signaling. Science 2005, 307, 1625−1630. (24) Monks, C. R.; Freiberg, B. A.; Kupfer, H.; Sciaky, N.; Kupfer, A. Three-Dimensional Segregation of Supramolecular Activation Clusters in T Cells. Nature 1998, 395, 82−86. (25) Bernas, T.; Zarebski, M.; Cook, R. R.; Dobrucki, J. W. Minimizing Photobleaching during Confocal Microscopy of Fluorescent Probes Bound to Chromatin: Role of Anoxia and Photon Flux. J. Microsc. 2004, 215, 281−296. (26) Segers-Nolten, G. Scanning Confocal Fluorescence Microscopy for Single Molecule Analysis of Nucleotide Excision Repair Complexes. Nucleic Acids Res. 2002, 30, 4720−4727. (27) Dobrucki, J. W.; Feret, D.; Noatynska, A. Scattering of Exciting Light by Live Cells in Fluorescence Confocal Imaging: Phototoxic Effects and Relevance for FRAP Studies. Biophys. J. 2007, 93, 1778− 1786. (28) Lee, J.; Miyanaga, Y.; Ueda, M.; Hohng, S. Video-Rate Confocal Microscopy for Single-Molecule Imaging in Live Cells and Superresolution Fluorescence Imaging. Biophys. J. 2012, 103, 1691−1697. (29) Ma, J.; Yang, W. Three-Dimensional Distribution of Transient Interactions in the Nuclear Pore Complex Obtained from SingleMolecule Snapshots. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 7305− 7310. (30) Kasper, R.; Harke, B.; Forthmann, C.; Tinnefeld, P.; Hell, S. W.; Sauer, M. Single-Molecule STED Microscopy with Photostable Organic Fluorophores. Small 2010, 6, 1379−1384. (31) Klar, T. A.; Hell, S. W. Subdiffraction Resolution in Far-Field Fluorescence Microscopy. Opt. Lett. 1999, 24, 954−956. (32) Wildanger, D.; Patton, B. R.; Schill, H.; Marseglia, L.; Hadden, J. P.; Knauer, S.; Schonle, A.; Rarity, J. G.; O’Brien, J. L.; Hell, S. W.; Smith, J. M. Solid Immersion Facilitates Fluorescence Microscopy with Nanometer Resolution and Sub-Angstrom Emitter Localization. Adv. Mater. 2012, 24, OP309−OP313. (33) Willig, K. I.; Kellner, R. R.; Medda, R.; Hein, B.; Jakobs, S.; Hell, S. W. Nanoscale Resolution in GFP-Based Microscopy. Nat. Methods 2006, 3, 721−723. (34) Harke, B.; Keller, J.; Ullal, C. K.; Westphal, V.; Schönle, A.; Hell, S. W. Resolution Scaling in STED Microscopy. Opt. Express 2008, 16, 4154−4154. (35) Mueller, V.; Ringemann, C.; Honigmann, A.; Schwarzmann, G.; Medda, R.; Leutenegger, M.; Polyakova, S.; Belov, V. N.; Hell, S. W.; Eggeling, C. STED Nanoscopy Reveals Molecular Details of Cholesterol- and Cytoskeleton-Modulated Lipid Interactions in Living Cells. Biophys. J. 2011, 101, 1651−1660. (36) Vicidomini, G.; Moneron, G.; Han, K. Y.; Westphal, V.; Ta, H.; Reuss, M.; Engelhardt, J.; Eggeling, C.; Hell, S. W. Sharper Low-Power STED Nanoscopy by Time Gating. Nat. Methods 2011, 8, 571−573. (37) Hofmann, M.; Eggeling, C.; Jakobs, S.; Hell, S. W. Breaking the Diffraction Barrier in Fluorescence Microscopy at Low Light Intensities by Using Reversibly Photoswitchable Proteins. Proc. Natl. Acad. Sci. U. S. A. 2005, 102, 17565−17569. (38) Axelrod, D.; Burghardt, T. P.; Thompson, N. L. Total Internal Reflection Fluorescence. Annu. Rev. Biophys. Bioeng. 1984, 13, 247− 268. (39) Moerner, W. E.; Kador, L. Optical Detection and Spectroscopy of Single Molecules in a Solid. Phys. Rev. Lett. 1989, 62, 2535−2538. (40) Orrit, M.; Bernard, J. Single Pentacene Molecules Detected by Fluorescence Excitation in a P-Terphenyl Crystal. Phys. Rev. Lett. 1990, 65, 2716−2719. (41) Mattheyses, A. L.; Simon, S. M.; Rappoport, J. Z. Imaging with Total Internal Reflection Fluorescence Microscopy for the Cell Biologist. J. Cell Sci. 2010, 123, 3621−3628. (42) Yeung, T.; Georges, P. C.; Flanagan, L. A.; Marg, B.; Ortiz, M.; Funaki, M.; Zahir, N.; Ming, W.; Weaver, V.; Janmey, P. A. Effects of 8150

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153

Review

ACS Nano Substrate Stiffness on Cell Morphology, Cytoskeletal Structure, and Adhesion. Cell Motil. Cytoskeleton 2005, 60, 24−34. (43) Elbert, D. L.; Herbert, C. B.; Hubbell, J. a. Thin Polymer Layers Formed by Polyelectrolyte Multilayer Techniques on Biological Surfaces. Langmuir 1999, 15, 5355−5362. (44) James, J. R.; Mccoll, J.; Oliveira, M. I.; Dunne, P. D.; Huang, E.; Jansson, A.; Nilsson, P.; Sleep, D. L.; Gonc, C. M.; Morgan, S. H.; Felce, J. H.; Mahen, R.; Fernandes, R. A.; Carmo, A. M.; Klenerman, D.; Davis, S. J. The T Cell Receptor Triggering Apparatus is Composed of Monovalent or Monomeric Proteins. J. Biol. Chem. 2011, 286, 31993−32001. (45) Sako, Y.; Minoghchi, S.; Yanagida, T. Single-Molecule Imaging of EGFR Signalling on the Surface of Living Cells. Nat. Cell Biol. 2000, 2, 168−172. (46) Cui, B.; Wu, C.; Chen, L.; Ramirez, A.; Bearer, E. L.; Li, W.; Mobley, W. C.; Chu, S. One at a Time, Live Tracking of NGF Axonal Transport Using Quantum Dots. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 13666−13671. (47) Konopka, C. A.; Bednarek, S. Y. Variable-Angle Epifluorescence Microscopy: A New Way to Look at Protein Dynamics in the Plant Cell Cortex. Plant J. 2008, 53, 186−196. (48) Tokunaga, M.; Imamoto, N.; Sakata-Sogawa, K. Highly Inclined Thin Illumination Enables Clear Single-Molecule Imaging in Cells. Nat. Methods 2008, 5, 159−161. (49) Hennig, S.; van de Linde, S.; Lummer, M.; Simonis, M.; Huser, T.; Sauer, M. Instant Live-Cell Super-Resolution Imaging of Cellular Structures by Nanoinjection of Fluorescent Probes. Nano Lett. 2015, 15, 1374−1381. (50) Voie, A. H.; Burns, D. H.; Spelman, F. A. Orthogonal-Plane Fluorescence Optical Sectioning: Three-Dimensional Imaging of Macroscopic Biological Specimens. J. Microsc. 1993, 170, 229−236. (51) Ritter, J. G.; Spille, J.; Kaminski, T.; Kubitscheck, U. A Cylindrical Zoom Lens Unit for Adjustable Optical Sectioning in Light Sheet Microscopy. Biomed. Opt. Express 2011, 2, 185−193. (52) Cella Zanacchi, F.; Lavagnino, Z.; Faretta, M.; Furia, L.; Diaspro, A. Light-Sheet Confined Super-Resolution Using Two-Photon Photoactivation. PLoS One 2013, 8, e67667. (53) Wu, Y.; Ghitani, A.; Christensen, R.; Santella, A.; Du, Z.; Rondeau, G.; Bao, Z.; Colon-Ramos, D.; Shroff, H. Inverted Selective Plane Illumination Microscopy (iSPIM) Enables Coupled Cell Identity Lineaging and Neurodevelopmental Imaging in Caenorhabditis Elegans. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 17708−17713. (54) Wu, Y.; Wawrzusin, P.; Senseney, J.; Fischer, R. S.; Christensen, R.; Santella, A.; York, A. G.; Winter, P. W.; Waterman, C. M.; Bao, Z.; Colón-Ramos, D. A.; McAuliffe, M.; Shroff, H. Spatially Isotropic Four-Dimensional Imaging with Dual-View Plane Illumination Microscopy. Nat. Biotechnol. 2013, 31, 1032−1038. (55) Gebhardt, J. C.; Suter, D. M.; Roy, R.; Zhao, Z. W.; Chapman, A. R.; Basu, S.; Maniatis, T.; Xie, X. S. Single-Molecule Imaging of Transcription Factor Binding to DNA in Live Mammalian Cells. Nat. Methods 2013, 10, 421−426. (56) Ritter, J. G.; Veith, R.; Veenendaal, A.; Siebrasse, J. P.; Kubitscheck, U. Light Sheet Microscopy for Single Molecule Tracking in Living Tissue. PLoS One 2010, 5, e11639. (57) Siebrasse, J. P.; Kaminski, T.; Kubitscheck, U. Nuclear Export of Single Native mRNA Molecules Observed by Light Sheet Fluorescence Microscopy. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 9426−9431. (58) Yu, J.; Cao, B.; Li, H.; Yu, B.; Chen, D.; Niu, H. Improved Localization Accuracy in Double-helix Point Spread Function Superresolution Fluorescence Microscopy Using Selective-plane Illumination. Proc. SPIE 2014, 9230, 92300N. (59) Galland, R.; Grenci, G.; Aravind, A.; Viasnoff, V.; Studer, V.; Sibarita, J. 3D High- and Super-Resolution Imaging Using SingleObjective SPIM. Nat. Methods 2015, 12, 641−644. (60) Durnin, J.; Miceli, J. J.; Eberly, J. H. Comparison of Bessel and Gaussian Beams. Opt. Lett. 1988, 13, 79−80. (61) Planchon, T. A.; Gao, L.; Milkie, D. E.; Davidson, M. W.; Galbraith, J. A.; Galbraith, C. G.; Betzig, E. Rapid Three-Dimensional

Isotropic Imaging of Living Cells Using Bessel Beam Plane Illumination. Nat. Methods 2011, 8, 417−423. (62) Siviloglou, G.; Broky, J.; Dogariu, A.; Christodoulides, D. N. Observation of Accelerating Airy Beams. Phys. Rev. Lett. 2007, 99, 213901. (63) Vettenburg, T.; Dalgarno, H. I. C.; Nylk, J.; Coll-Llado, C.; Ferrier, D. E. K.; Cizmar, T.; Gunn-Moore, F. J.; Dholakia, K. LightSheet Microscopy Using an Airy Beam. Nat. Methods 2014, 11, 541− 544. (64) Chen, B. C.; Legant, W. R.; Wang, K.; Shao, L.; Milkie, D. E.; Davidson, M. W.; Janetopoulos, C.; Wu, X. S.; Hammer, J. A., 3rd; Liu, Z.; English, B. P.; Mimori-Kiyosue, Y.; Romero, D. P.; Ritter, A. T.; Lippincott-Schwartz, J.; Fritz-Laylin, L.; Mullins, R. D.; Mitchell, D. M.; Bembenek, J. N.; Reymann, A. C.; Bohme, R.; Grill, S. W.; Wang, J. T.; Seydoux, G.; Tulu, U. S.; Kiehart, D. P.; Betzig, E. Lattice LightSheet Microscopy: Imaging Molecules to Embryos at High Spatiotemporal Resolution. Science 2014, 346, 1257998. (65) Gao, L.; Shao, L.; Higgins, C.; Poulton, J.; Peifer, M.; Davidson, M.; Wu, X.; Goldstein, B.; Betzig, E. Noninvasive Imaging Beyond the Diffraction Limit of 3D Dynamics in Thickly Fluorescent Specimens. Cell 2012, 151, 1370−1385. (66) Liu, Z.; Legant, W. R.; Chen, B. C.; Li, L.; Grimm, J. B.; Lavis, L. D.; Betzig, E.; Tjian, R. 3D Imaging of Sox2 Enhancer Clusters in Embryonic Stem Cells. eLife 2014, 3, e04236−e04236. (67) Koch, A. L. What Size Should a Bacterium be? A Question of Scale. Annu. Rev. Microbiol. 1996, 50, 317−348. (68) Reyes-Lamothe, R.; Sherratt, D. J.; Leake, M. C. Stoichiometry and Architecture of Active DNA Replication Machinery in Escherichia Coli. Science 2010, 328, 498−501. (69) Tuson, H. H.; Biteen, J. S. Unveiling the Inner Workings of Live Bacteria Using Super-Resolution Microscopy. Anal. Chem. 2015, 87, 42−63. (70) Cattoni, D.; Fiche, J.; Nöllmann, M. Single-Molecule SuperResolution Imaging in Bacteria. Curr. Opin. Microbiol. 2012, 15, 758− 763. (71) Robinson, A.; van Oijen, A. M. Bacterial Replication, Transcription and Translation: Mechanistic Insights from SingleMolecule Biochemical Studies. Nat. Rev. Microbiol. 2013, 11, 303−315. (72) Gahlmann, A.; Moerner, W. E. Exploring Bacterial Cell Biology with Single-Molecule Tracking and Super-Resolution Imaging. Nat. Rev. Microbiol. 2014, 12, 9−22. (73) Sanamrad, A.; Persson, F.; Lundius, E. G.; Fange, D.; Gynna, A. H.; Elf, J. Single-Particle Tracking Reveals that Free Ribosomal Subunits Are Not Excluded from the Escherichia Coli Nucleoid. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 11413−11418. (74) Tuson, H. H.; Aliaj, A.; Brandes, E. R.; Simmons, L. A.; Biteen, J. S. Addressing the Requirements of High Sensitivity Single-Molecule Imaging of Low-Copy Number Proteins in Bacteria. ChemPhysChem 2016, 17, 1435−1440. (75) Crawford, R.; Torella, J.; Aigrain, L.; Plochowietz, A.; Gryte, K.; Uphoff, S.; Kapanidis, A. Long-Lived Intracellular Single-Molecule Fluorescence Using Electroporated Molecules. Biophys. J. 2013, 105, 2439−2450. (76) Rowland, D. J.; Tuson, H. H.; Biteen, J. S. Resolving Fast, Confined Diffusion in Bacteria with Image Correlation Spectroscopy. Biophys. J. 2016, 110, 2241−2251. (77) Wang, S.; Moffitt, J. R.; Dempsey, G. T.; Xie, X. S.; Zhuang, X. Characterization and Development of Photoactivatable Fluorescent Proteins for Single-Molecule-Based Superresolution Imaging. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 8452−8457. (78) Manley, S.; Gillette, J. M.; Patterson, G. H.; Shroff, H.; Hess, H. F.; Betzig, E.; Lippincott-Schwartz, J. High-Density Mapping of SingleMolecule Trajectories with Photoactivated Localization Microscopy. Nat. Methods 2008, 5, 155−157. (79) Li, W.; Bouveret, E.; Zhang, Y.; Liu, K.; Wang, J. D.; Weisshaar, J. C. Effects of Amino Acid Starvation on RelA Diffusive Behavior in Live Escherichia Coli. Mol. Microbiol. 2016, 99, 571−585. (80) Liao, Y.; Schroeder, J. W.; Gao, B.; Simmons, L. A.; Biteen, J. S. Single-Molecule Motions and Interactions in Live Cells Reveal Target 8151

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153

Review

ACS Nano Search Dynamics in Mismatch Repair. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, E6898−E6906. (81) Schütz, G. J.; Schindler, H.; Schmidt, T. Single-Molecule Microscopy on Model Membranes Reveals Anomalous Diffusion. Biophys. J. 1997, 73, 1073−1080. (82) Matson, J. S.; Withey, J. H.; DiRita, V. J. Regulatory Networks Controlling Vibrio Cholerae Virulence Gene Expression. Infect. Immun. 2007, 75, 5542−5549. (83) Haas, B. L.; Matson, J. S.; DiRita, V. J.; Biteen, J. S. SingleMolecule Tracking in Live Vibrio Cholerae Reveals that ToxR Recruits the Membrane-Bound Virulence Regulator TcpP to the toxT Promoter. Mol. Microbiol. 2015, 96, 4−13. (84) Siv, C.; DiRita, V. J.; Biteen, J. S., unpublished data. (85) Nguyen, H. H.; Park, J.; Kang, S.; Kim, M. Surface Plasmon Resonance: A Versatile Technique for Biosensor Applications. Sensors 2015, 15, 10481−10510. (86) Willets, K. A. Probing Local Electromagnetic Field Enhancements on the Surface of Plasmonic Nanoparticles. Prog. Surf. Sci. 2012, 87, 209−220. (87) Anger, P.; Bharadwaj, P.; Novotny, L. Enhancement and Quenching of Single-Molecule Fluorescence. Phys. Rev. Lett. 2006, 96, 113002. (88) Chen, X.; Wang, J.; Guo, Y.; Xu, L.; Pei, R. Cell Imaging Applications of Fluorescent Gold and Silver Clusters. Sci. Adv. Mater. 2015, 7, 2123−2133. (89) Langer, J.; Novikov, S. M.; Liz-Marzan, L. M. Sensing Using Plasmonic Nanostructures and Nanoparticles. Nanotechnology 2015, 26, 322001. (90) Ertsgaard, C. T.; McKoskey, R. M.; Rich, I. S.; Lindquist, N. C. Dynamic Placement of Plasmonic Hotspots for Super-Resolution Surface-Enhanced Raman Scattering. ACS Nano 2014, 8, 10941− 10946. (91) Flynn, J. D.; Haas, B. L.; Biteen, J. S. Plasmon-Enhanced Fluorescence from Single Proteins in Living Bacteria. J. Phys. Chem. C 2016, DOI: 10.1021/acs.jpcc.5b08049. (92) Aslan, K.; Gryczynski, I.; Malicka, J.; Matveeva, E.; Lakowicz, J. R.; Geddes, C. D. Metal-Enhanced Fluorescence: An Emerging Tool in Biotechnology. Curr. Opin. Biotechnol. 2005, 16, 55−62. (93) Kinkhabwala, A.; Yu, Z.; Fan, S.; Avlasevich, Y.; Mullen, K.; Moerner, W. E. Large Single-Molecule Fluorescence Enhancements Produced by a Gold Bowtie Nanoantenna. Nat. Photonics 2009, 3, 654−657. (94) Donehue, J. E.; Wertz, E.; Talicska, C. N.; Biteen, J. S. PlasmonEnhanced Brightness and Photostability from Single Fluorescent Proteins Coupled to Gold Nanorods. J. Phys. Chem. C 2014, 118, 15027−15035. (95) Wertz, E.; Isaacoff, B. P.; Flynn, J. D.; Biteen, J. S. SingleMolecule Super-Resolution Microscopy Reveals how Light Couples to a Plasmonic Nanoantenna on the Nanometer Scale. Nano Lett. 2015, 15, 2662−2670. (96) Fu, B.; Flynn, J. D.; Isaacoff, B. P.; Rowland, D. J.; Biteen, J. S. Super-Resolving the Distance-Dependent Plasmon-Enhanced Fluorescence of Single Dye and Fluorescent Protein Molecules. J. Phys. Chem. C 2015, 119, 19350−19358. (97) Ringler, M.; Schwemer, A.; Wunderlich, M.; Nichtl, A.; Kuerzinger, K.; Klar, T. A.; Feldmann, J. Shaping Emission Spectra of Fluorescent Molecules with Single Plasmonic Nanoresonators. Phys. Rev. Lett. 2008, 100, 203002. (98) Bharadwaj, P.; Anger, P.; Novotny, L. Nanoplasmonic Enhancement of Single-Molecule Fluorescence. Nanotechnology 2007, 18, 044017. (99) Flauraud, V.; van Zanten, T. S.; Mivelle, M.; Manzo, C.; Garcia Parajo, M. F.; Brugger, J. Large-Scale Arrays of Bowtie Nanoaperture Antennas for Nanoscale Dynamics in Living Cell Membranes. Nano Lett. 2015, 15, 4176−4182. (100) Friedrich, M.; Gan, Q.; Ermolayev, V.; Harms, G. STEDSPIM: Stimulated Emission Depletion Improves Sheet Illumination Microscopy Resolution. Biophys. J. 2011, 100, L43−L45.

(101) Friedrich, M.; Harms, G. S. Axial Resolution Beyond the Diffraction Limit of a Sheet Illumination Microscope with Stimulated Emission Depletion. J. Biomed. Opt. 2015, 20, 106006−106006. (102) Booth, M.; Andrade, D.; Burke, D.; Patton, B.; Zurauskas, M. Aberrations and Adaptive Optics in Super-Resolution Microscopy. Microscopy 2015, 0, 1−11. (103) Kim, M. K. Adaptive Optics by Incoherent Digital Holography. Opt. Lett. 2012, 37, 2694−2694. (104) Fernandez, E.; Artal, P. Membrane Deformable Mirror for Adaptive Optics: Performance Limits in Visual Optics. Opt. Express 2003, 11, 1056−1069. (105) Bourgenot, C.; Saunter, C. D.; Taylor, J. M.; Girkin, J. M.; Love, G. D. 3D Adaptive Optics in a Light Sheet Microscope. Opt. Express 2012, 20, 13252−13252. (106) Palayret, M.; Armes, H.; Basu, S.; Watson, A. T.; Herbert, A.; Lando, D.; Etheridge, T. J.; Endesfelder, U.; Heilemann, M.; Laue, E.; Carr, A. M.; Klenerman, D.; Lee, S. F. Virtual-’Light-Sheet’ SingleMolecule Localisation Microscopy Enables Quantitative Optical Sectioning for Super-Resolution Imaging. PLoS One 2015, 10, e0125438−e0125438. (107) Pampaloni, F.; Berge, U.; Marmaras, A.; Horvath, P.; Kroschewski, R.; Stelzer, E. H. K. Tissue-Culture Light Sheet Fluorescence Microscopy (TC-LSFM) Allows Long-Term Imaging of Three-Dimensional Cell Cultures Under Controlled Conditions. Integr. Biol. 2014, 6, 988−998. (108) Yang, Z.; Piksarv, P.; Ferrier, D. E. K.; Gunn-Moore, F.; Dholakia, K. Macro-Optical Trapping for Sample Confinement in Light Sheet Microscopy. Biomed. Opt. Express 2015, 6, 2778−2778. (109) del Rosal, B.; Haro-Gonzalez, P.; Ramsay, W. T.; Maestro, L. M.; Santacruz-Gomez, K.; Iglesias-de la Cruz, M. C.; Sanz-Rodriguez, F.; Chooi, J. Y.; Rodriguez-Sevilla, P.; Choudhury, D.; Kar, A. K.; Garcia-Sole, J. G.; Patterson, L.; Jaque, D. Heat in Optical Tweezers. Proc. SPIE 2013, 8810, 88102A. (110) Oddos, S.; Dunsby, C.; Purbhoo, M. A.; Chauveau, A.; Owen, D. M.; Neil, M. A. A.; Davis, D. M.; French, P. M. W. High-Speed High-Resolution Imaging of Intercellular Immune Synapses Using Optical Tweezers. Biophys. J. 2008, 95, L66−L68. (111) Villar, G.; Graham, A. D.; Bayley, H. A Tissue-Like Printed Material. Science 2013, 340, 48−52. (112) Jinek, M.; Chylinski, K.; Fonfara, I.; Hauer, M.; Doudna, J. A.; Charpentier, E. A Programmable Dual-RNA−Guided DNA Endonuclease in Adaptive Bacterial Immunity. Science 2012, 337, 816−821. (113) Chen, B.; Gilbert, L. A.; Cimini, B. A.; Schnitzbauer, J.; Zhang, W.; Li, G. W.; Park, J.; Blackburn, E. H.; Weissman, J. S.; Qi, L. S.; Huang, B. Dynamic Imaging of Genomic Loci in Living Human Cells by an Optimized CRISPR/Cas System. Cell 2013, 155, 1479−1491. (114) Taylor, D. W.; Zhu, Y.; Staals, R. H.; Kornfeld, J. E.; Shinkai, A.; van der Oost, J.; Nogales, E.; Doudna, J. A. Structural Biology. Structures of the CRISPR-Cmr Complex Reveal Mode of RNA Target Positioning. Science 2015, 348, 581−585. (115) Wu, X.; Kriz, A. J.; Sharp, P. A. Target Specificity of the CRISPR-Cas9 System. Quant. Biol. 2014, 2, 59−70. (116) Ma, H.; Naseri, A.; Reyes-Gutierrez, P.; Wolfe, S. A.; Zhang, S.; Pederson, T. Multicolor CRISPR Labeling of Chromosomal Loci in Human Cells. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 3002−3007. (117) Deng, W.; Shi, X.; Tjian, R.; Lionnet, T.; Singer, R. H.; CASFISH. CRISPR/Cas9-Mediated in Situ Labeling of Genomic Loci in Fixed Cells. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 11870−11875. (118) Femino, A. M.; Fay, F. S.; Fogarty, K.; Singer, R. H. Visualization of Single RNA Transcripts. Science 1998, 280, 585−590. (119) Skinner, S. O.; Sepulveda, L. A.; Xu, H.; Golding, I. Measuring mRNA Copy Number in Individual Escherichia Coli Cells Using Single-Molecule Fluorescent in Situ Hybridization. Nat. Protoc. 2013, 8, 1100−1113. (120) Lubeck, E.; Cai, L. Single-Cell Systems Biology by SuperResolution Imaging and Combinatorial Labeling. Nat. Methods 2012, 9, 743−748. 8152

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153

Review

ACS Nano (121) Lubeck, E.; Coskun, A. F.; Zhiyentayev, T.; Ahmad, M.; Cai, L. Single-Cell in Situ RNA Profiling by Sequential Hybridization. Nat. Methods 2014, 11, 360−361. (122) Chen, K. H.; Boettiger, A. N.; Moffitt, J. R.; Wang, S.; Zhuang, X. Spatially Resolved, Highly Multiplexed RNA Profiling in Single Cells. Science 2015, 348, aaa6090. (123) Biteen, J. S.; Blainey, P. C.; Cardon, Z. G.; Chun, M.; Church, G. M.; Dorrestein, P. C.; Fraser, S. E.; Gilbert, J. A.; Jansson, J. K.; Knight, R.; Miller, J. F.; Ozcan, A.; Prather, K. A.; Quake, S. R.; Ruby, E. G.; Silver, P. A.; Taha, S.; van, d. E.; Weiss, P. S.; Wong, G. C. L.; Wright, A. T.; Young, T. D. Tools for the Microbiome: Nano and Beyond. ACS Nano 2016, 10, 6−37. (124) Schaferling, M. Nanoparticle-Based Luminescent Probes for Intracellular Sensing and Imaging of pH. Wiley Interdiscip. Rev. Nanomed. Nanobiotechnol. 2016, 8, 378−413. (125) Watanabe, K.; Palonpon, A. F.; Smith, N. I.; Chiu, L. D.; Kasai, A.; Hashimoto, H.; Kawata, S.; Fujita, K. Structured Line Illumination Raman Microscopy. Nat. Commun. 2015, 6, 10095. (126) Cheng, J.; Xie, X. S. Vibrational Spectroscopic Imaging of Living Systems: An Emerging Platform for Biology and Medicine. Science 2015, 350, 1054. (127) Hanrieder, J.; Phan, N. T.; Kurczy, M. E.; Ewing, A. G. Imaging Mass Spectrometry in Neuroscience. ACS Chem. Neurosci. 2013, 4, 666−679. (128) Bell, L.; Seshia, A.; Lando, D.; Laue, E.; Palayret, M.; Lee, S. F.; Klenerman, D. A Microfluidic Device for the Hydrodynamic Immobilisation of Living Fission Yeast Cells for Super-Resolution Imaging. Sens. Actuators, B 2014, 192, 36−41. (129) Zhou, Y.; Basu, S.; Wohlfahrt, K. J.; Lee, S. F.; Klenerman, D.; Laue, E. D.; Seshia, A. A. A Microfluidic Platform for Trapping, Releasing and Super-Resolution Imaging of Single Cells. Sens. Actuators, B 2016, 232, 680−691. (130) Chen, C. H.; Cho, S. H.; Chiang, H.; Tsai, F.; Zhang, K.; Lo, Y. Specific Sorting of Single Bacterial Cells with Microfabricated Fluorescence-Activated Cell Sorting and Tyramide Signal Amplification Fluorescence in Situ Hybridization. Anal. Chem. 2011, 83, 7269− 7275. (131) Shields, C. W.; Reyes, C. D.; Lopez, G. P. Microfluidic Cell Sorting: A Review of the Advances in the Separation of Cells from Debulking to Rare Cell Isolation. Lab Chip 2015, 15, 1230−1249. (132) Mahadik, B. P.; Wheeler, T. D.; Skertich, L. J.; Kenis, P. J. A.; Harley, B. A. C. Microfluidic Generation of Gradient Hydrogels to Modulate Hematopoietic Stem Cell Culture Environment. Adv. Healthcare Mater. 2014, 3, 449−458. (133) Stiefel, P.; Zambelli, T.; Vorholt, J. A. Isolation of Optically Targeted Single Bacteria by Application of Fluidic Force Microscopy to Aerobic Anoxygenic Phototrophs from the Phyllosphere. Appl. Environ. Microbiol. 2013, 79, 4895−4905. (134) Sadanandan, S. K.; Baltekin, O.; Magnusson, K. E. G.; Boucharin, A.; Ranefall, P.; Jalden, J.; Elf, J.; Wahlby, C. Segmentation and Track-Analysis in Time-Lapse Imaging of Bacteria. Selected Topics in Signal Processing, IEEE Journal of 2016, 10, 174−184. (135) Liu, J.; Prindle, A.; Humphries, J.; Gabalda-Sagarra, M.; Asally, M.; Lee, D. D.; Ly, S.; Garcia-Ojalvo, J.; Suel, G. M. Metabolic CoDependence Gives Rise to Collective Oscillations within Biofilms. Nature 2015, 523, 550−554. (136) Boedicker, J. Q.; Vincent, M. E.; Ismagilov, R. F. Microfluidic Confinement of Single Cells of Bacteria in Small Volumes Initiates High-Density Behavior of Quorum Sensing and Growth and Reveals its Variability. Angew. Chem., Int. Ed. 2009, 48, 5908−5911. (137) Ze, X.; Duncan, S. H.; Louis, P.; Flint, H. J. Ruminococcus Bromii is a Keystone Species for the Degradation of Resistant Starch in the Human Colon. ISME J. 2012, 6, 1535−1543. (138) Cheng, M. C.; Leske, A. T.; Matsuoka, T.; Kim, B. C.; Lee, J.; Burns, M. A.; Takayama, S.; Biteen, J. S. Super-Resolution Imaging of PDMS Nanochannels by Single-Molecule Micelle-Assisted Blink Microscopy. J. Phys. Chem. B 2013, 117, 4406−4411.

(139) Bolea, I.; Gan, W. B.; Manfedi, G.; Magrane, J. Imaging of Mitochondrial Dynamics in Motor and Sensory Axons of Living Mice. Methods Enzymol. 2014, 547, 97−110. (140) Abran, M.; Stahli, B. E.; Merlet, N.; Mihalache-Avram, T.; Mecteau, M.; Rheaume, E.; Busseuil, D.; Tardif, J. C.; Lesage, F. Validating a Bimodal Intravascular Ultrasound (IVUS) and NearInfrared Fluorescence (NIRF) Catheter for Atherosclerotic Plaque Detection in Rabbits. Biomed. Opt. Express 2015, 6, 3989−3999. (141) Earle, K.; Billings, G.; Sigal, M.; Lichtman, J.; Hansson, G.; Elias, J.; Amieva, M.; Huang, K.; Sonnenburg, J. Quantitative Imaging of Gut Microbiota Spatial Organization. Cell Host Microbe 2015, 18, 478−488.

8153

DOI: 10.1021/acsnano.6b02863 ACS Nano 2016, 10, 8143−8153