fluorescence imaging for bacterial cell biology: from localization to dynamics, from ensembles to...

MI68CH24-Carballido-Lopez ARI 9 June 2014 17:0

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Fluorescence Imaging forBacterial Cell Biology: FromLocalization to Dynamics,From Ensembles to SingleMoleculesZhizhong Yao and Rut Carballido-LopezINRA, UMR1319 Micalis, F-78350 Jouy-en-Josas, France;email: [email protected]

Annu. Rev. Microbiol. 2014. 68:459–76

The Annual Review of Microbiology is online atmicro.annualreviews.org

This article’s doi:10.1146/annurev-micro-091213-113034

Copyright c© 2014 by Annual Reviews.All rights reserved

Keywords

protein subcellular localization, superresolution, single molecule, MreB,bacterial cytoskeleton

Abstract

Fluorescent proteins and developments in superresolution (nanoscopy) andsingle-molecule techniques bring high sensitivity, speed, and one order ofmagnitude gain in spatial resolution to live-cell imaging. These technolo-gies have only recently been applied to prokaryotic cell biology, revealing theexquisite subcellular organization of bacterial cells. Here, we review the par-allel evolution of fluorescence microscopy methods and their application tobacteria, mainly drawing examples from visualizing actin-like MreB proteinsin the model bacterium Bacillus subtilis. We describe the basic principles ofnanoscopy and conventional techniques and their advantages and limitationsto help microbiologists choose the most suitable technique for their biolog-ical question. Looking ahead, multidimensional live-cell nanoscopy com-bined with computational image analysis tools, systems biology approaches,and mathematical modeling will provide movie-like, mechanistic, and quan-titative description of molecular events in bacterial cells.

459

Review in Advance first posted online on June , 2014. (Changes may still occur before final publication online and in print.)

Changes may still occur before final publication online and in print

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Fluorescence:absorption of energyupon illumination,followed by emissionof light with a longerwavelength than theincident light (Stokesshift)

Contents

THE MAKING OF BACTERIAL CELL BIOLOGY:THE HISTORICAL PERSPECTIVE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 460

CONSTRUCTING FLUORESCENCE FUSION PROTEINS . . . . . . . . . . . . . . . . . . . 461CONVENTIONAL FLUORESCENCE MICROSCOPY:

ADVANCES AND MISSTEPS. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 462Fluorescence Microscopy Setups . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 462Protein Subcellular Localization: The Standard Practice . . . . . . . . . . . . . . . . . . . . . . . . . . 464Protein Subcellular Dynamics: The Emerging Trend . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 464

SUPERRESOLUTION APPROACHES: BEYOND THEDIFFRACTION LIMIT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 467Structured Illumination Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 467Stimulated Emission Depletion Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 468Point Localization–Based Superresolution Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . 469Single-Molecule Tracking: One Molecule Over Time . . . . . . . . . . . . . . . . . . . . . . . . . . . . 471

CONCLUDING REMARKS: FUTURE CHALLENGESAND OPPORTUNITIES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 472

THE MAKING OF BACTERIAL CELL BIOLOGY:THE HISTORICAL PERSPECTIVE

Nearly 25 years ago, the landmark discovery that the bacterial tubulin homolog FtsZ polymerizesinto the cytokinetic Z ring at the division site revealed that bacterial proteins, like their eukaryoticcounterparts, display specific subcellular locations that are coupled to their specific function (5).With the advent of fluorescent proteins (FPs) and imaging platforms, characterization of proteinsubcellular localization in bacteria has become widespread. These studies, including the break-through discovery that the bacterial actin homolog MreB assembles into filamentous structuresalong the sidewall that control cell shape (38), greatly expanded our understanding of the archi-tecture of the bacterial cell. Bacteria are no longer considered simple bags of enzymes withoutinternal organization; instead, they are recognized as highly organized cells with modular func-tionalities, sometimes even compartmentalized (44, 61). The field of prokaryotic cell biology hasgrown explosively in the last few years and holds exciting potential for new developments in thefuture.

Understanding the genotype-phenotype relationship, or in other words how linear DNA codesfor 3-D structures with specific functions, remains one of the fundamental questions in modernmicrobiology. Despite great progress in genome sequencing, the function of many proteins re-mains elusive, mainly owing to the lack of clear phenotypes. Even for extensively studied proteinswith an established function such as FtsZ or MreB, the detailed molecular mechanisms underlyingtheir function remain unknown. To bridge the gap between genetic information and cellular func-tion, it is imperative to characterize where, when, and how molecular events take place in the cell.Fluorescence microscopy enables characterization of the spatial localization of bacterial proteinsin their native environment with nanometer precision and characterization of their dynamics withhigh temporal resolution.

In this review, we summarize conventional light microscopy techniques and their applicationsto study the subcellular localization and dynamics of proteins in bacterial cells. Then we focuson modern superresolution microscopy (nanoscopy) methods, which overcome the diffraction

460 Yao · Carballido-Lopez


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limit (∼250 nm) of conventional light microscopy. Last, we discuss open questions associatedwith these new methods and their impact on the future of prokaryotic cell biology. Our goal isnot a comprehensive review of the ever-expanding nomenclature of available techniques and allreported applications in bacteria; rather, we focus on key physical concepts and landmark stud-ies that illustrate generalities. The evolution of fluorescence microscopy in bacterial cell biologyis best illustrated by studies of the actin-like MreB cytoskeleton over the last decade, partic-ularly in the gram-positive model bacterium Bacillus subtilis, from which we draw most of ourexamples. Comprehensive discussion of individual topics can be found in other review articles(7, 30, 65, 72, 90).

CONSTRUCTING FLUORESCENCE FUSION PROTEINS

A typical fluorescence microscopy experiment requires the construction of a functional FP fusionand the choice of the suitable microscopy configuration. The ideal fluorophore is the brightest,most stable, monomeric FP derivative. For colocalization experiments, compatible fluorophoresshould be chosen so that their excitation and emission spectra do not overlap, to maximize signalwhile minimizing bleed-through artifacts. Wild-type green fluorescent protein (GFP) is a weakdimer (85) that can cause artificial clustering of fusion proteins through the formation of dimersand higher-order oligomers (48). Therefore the use of monomeric variants, which can be obtainedby introducing point mutations at the GFP dimer interface (48), is highly recommended.

Fluorescent fusion proteins can differ from their cognate wild-type protein in two respects.First, the fluorescent fusion may not be functional, or it may be only partially functional, likeFP fusions to FtsZ, which are temperature sensitive in most species. Complementation testsmust be carried out to show that the fusion protein can replace the native protein in the cell.Some FP tags interfere with the function of the protein but do not prevent its normal targetingand can be used in merodiploid strains. It is often useful to insert a flexible polypeptide linkerbetween the FP and the target protein to circumvent steric problems, promote proper folding,and increase the functionality of the fusion. Functionality may also determine the choice of C-or N-terminal or even sandwich fluorescent fusion proteins. For example, extended filamentoushelical structures are formed by MreB in Escherichia coli when yellow fluorescent protein (YFP)is fused to its N-terminus. These were shown to be an artifact due to the FP tag, as native(untagged) MreB and a functional sandwich (in frame hybrid) fusion to mCherry (MreB-RFPSW)(2) do not form helices (83). Second, the fluorescent fusion protein may not be expressed at wild-type levels. Fluorescent fusions (in particular when proteins are tagged at the N-terminus) havebeen widely expressed from inducible promoters, often as an extopic copy in the chromosomeor from a plasmid. Different concentrations of inducer should then be tested to be as close aspossible to native expression levels, and under- and overexpression effects on cell physiologyand/or protein localization ruled out. Examples of overexpression artifacts in B. subtilis includethe stable accumulation of the cell division inhibitors MinC and MinD at the cell poles (54, 55),instead of at new and constricting septa (25), and changes in the localization of the sporulationlandmark protein SpoIIQ (8). Constitutive expression from inducible promoters may also maskpossible cell cycle–dependent changes in localization patterns. Thus, the ideal fluorescent fusionprotein can replace the wild-type protein and is at the native locus under wild-type regulatorycontrols. Even fusions such as these must be treated with caution, as the FP tag may still affect thelevels, stability, or other subtle properties of the target protein. For all these reasons, subcellularprotein localization in live cells is sometimes difficult to prove independently. When necessary,complementary immunoelectron or immunofluorescence microscopy should be carried out infixed cells to rule out possible artifacts.

www.annualreviews.org • Fluorescence Imaging in Bacteria 461


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a b cEpi TIRF(θ > θc)

HILO(0º < θ < θc)

θ

d Bottom Middle Top

Co

nfo

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θθ

2 μm

2 μm

Figure 1MreB of Bacillus subtilis visualized by conventional (diffraction-limited) microscopy methods. Depending onthe region of the cell, different illumination methods using wide-field illumination (a–c) or point illumination(d ) can be used. Blue regions within the cell indicate areas under illumination; green circles indicate excitedfluorophores and open circles nonexcited fluorophores. B. subtilis cells expressing functional GFP-MreBwere grown to early exponential phase, and the same cell was simultaneously visualized by conventionalepifluorescence, TIRF, HILO, and LSCM on a Zeiss microscope (Elyra PS1 LSM780). (a–c) Depth ofpenetration: epifluorescence > HILO > TIRF. θ, incident light angle (black arrows); θc, critical angle (bluearrows). In epifluorescence (a) θ ≈ 0◦, and light propagates through the sample; all parts and depths of thecell are simultaneously excited, giving a low signal-to-noise ratio. In TIRF (b) light is completely reflected(θ > θc) and generates an exponentially decaying evanescent wave (near-field illumination) that selectivelyilluminates molecules close to the cell surface. In HILO (c) light enters the sample as a collimated highlyinclined beam (0◦ < θ < θc), going deeper into the cell than in TIRF microscopy but remaining shallowrelative to epifluorescence. (d ) LSCM provides high contrast and allows optical sectioning of the sample.The bottom, middle, and top parts of the cell were visualized. Abbreviations: Epi, epifluorescence; GFP,green fluorescent protein; HILO, highly inclined laminated optical sheet; LSCM, laser scanning confocalmicroscopy; TIRF, total internal reflection fluorescence.

Wide-field: mode ofillumination withsimultaneous, evenexcitation across theentire sample (incontrast to pointillumination inconfocal microscopy)

CONVENTIONAL FLUORESCENCE MICROSCOPY:ADVANCES AND MISSTEPS

Fluorescence Microscopy Setups

Conventional wide-field epifluorescence and laser scanning confocal microscopy (LSCM) are thetraditional modes of fluorescence microscopy. In epifluorescence (epi-, from ancient Greek επi,“upon”), the entire sample on the optical path of the incident light (from a light source) is simulta-neously excited, and the emitted light (fluorescence) is detected throughout the sample in a large,unfocused manner (Figure 1a). Classical light sources are mercury or xenon arc lamps, which pro-vide high-intensity illumination throughout the visible spectrum (390–700 nm). Different filtersets then select excitation and emission spectral windows characteristic of each fluorophore. Al-ternatively, high-power light-emitting diodes (LEDs) can serve as light sources. Individual LEDssupply the optimum excitation wavelength band for fluorophores spanning the ultraviolet (12–390 nm), visible, and near-infrared (700–2,500 mm) regions, providing high specificity and con-trast. They do not heat the sample, and they provide stable illumination through the entire lifetimeof the lamp (over 30,000 h versus ∼150–300 h for halogen and arc lamps) and are cheaper. Finally,



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NA: numericalaperture, n·sinθ (n,refractive index of theimaging medium; θ,half-aperture angle ofthe objective). NA <

1.5 (θ < 70◦) in mostobjectives

Deconvolution:mathematical methodthat removes blurringfrom out-of-focussignal in fluorescencemicroscopy images

Near-field: mode ofillumination based onevanescent waves(nonpropagatinglight), which vanishwithin onewavelength; used inTIRF microscopy

TIRF: totalinternal reflectionfluorescence; light iscompletely reflectedwhen striking amedium boundarywith lower refractiveindex (e.g., from glassto water) with incidentangle > θc

θc: critical angle atwhich light striking aboundary from high(n1) to lower (n2)refractive index mediaexits tangentially tothe interface [θc =sin−1 (n2/n1)]

Photobleaching:photochemicalreaction (e.g.,photo-oxidation) thatirreversibly abolishesfluorescence offluorophore; alsoreferred to as fading

wide-field illumination can also be achieved with a laser. As with LED light sources, excitationfilters are not needed, but each excitation wavelength required needs its corresponding—andcostly—laser. Regardless of the light source, epifluorescence microscopy suffers from low signal-to-noise ratio owing to fluorescence coming from above and below the focal plane. In contrast,confocal microscopy uses point illumination (a laser beam passes through an aperture and is thenfocused by an objective lens into a small focal volume), scanning across the sample and a pinholein front of the detector to keep only an in-focus signal (hence the appellation confocal: con-, fromthe Latin cum, “with, together” the focal plane). LSCM therefore produces images with enhancedcontrast and resolution compared to epifluorescence in samples that are thicker than the focalplane, allowing optical sectioning (Figure 1d ). However, given that bacterial cells (∼0.5–1 μmwide) are only slightly thicker than the focal plane [∼0.5 μm; defined by the wavelength of emittinglight (λ) and the numerical aperture (NA)], and improved contrast is to the detriment of fluores-cence intensity (much of the light is blocked at the pinhole) and temporal resolution (scanning isrequired because only one point is illuminated at a time), LSCM offers no particular advantagesover conventional epifluorescence microscopy for bacterial studies at the subcellular level. Becauseepifluorescence systems are less expensive and easier to use, epifluorescence has been the method ofchoice in most bacterial cell biology laboratories. Optical sectioning combined with image restora-tion using deconvolution (6, 80) and deblurring methods can greatly improve the signal-to-noiseratio in wide-field epifluorescence and LSCM images and generate high-quality 3-D images.

Two related so-called near-field techniques, total internal reflection fluorescence (TIRF) mi-croscopy and highly inclined laminated optical sheet (HILO), both compatible with wide-fieldepifluorescence setups, have been recently applied to bacterial cells. TIRF microscopy is a sensitivemethod that allows visualization of fluorescently labeled molecules specifically at the cell surface.It exploits the unique property of the exponentially decaying evanescent wave created at the in-terface of two media of different refractive indices (sample/water and coverslip) when a specimenis exposed to laser illumination at or above an incident critical angle (θc), at which total internalreflection occurs. Fluorophores are selectively excited within a couple of hundred nanometersfrom the coverslip (i.e., near field) (Figure 1b). Thus, TIRF enables a high signal-to-noise ratioand increased temporal resolution, which limits photobleaching. To date, TIRF microscopy hasbeen used to image MreB and other components of the cell wall elongation machinery (15, 21),MinC (25), and membrane-associated flotilling and dynein orthologs (13, 14) in B. subtilis, to im-age cytochrome bd-I complexes (50) and phage shock protein PspA in E. coli (40), and to examinethe adhesion of Caulobacter crescentus cells to surfaces (51).

HILO, also known as quasi-, pseudo- or near-TIRF microscopy, takes a middle ground betweenTIRF and wide-field epi-illumination (75, 84). Slightly varying the laser light incident angle awayfrom total internal reflection to a subcritical angle increases the penetration depth of illuminationso that it is deeper in the cell than would be the case with TIRF, but still shallower than withepifluorescence (Figure 1c). Fluorescence signal intensity increases in the cytoplasmic volumeclose to the membrane whereas the high background scatter responsible for the poor contrast ofepi-illumination is still reduced, enabling increased temporal resolution, as in TIRF (84). The firstHILO application to bacteria explored the dynamics of the SpoIIIE DNA pump at the middleof the septum of sporulating B. subtilis cells (19). Although epifluorescence microscopy initiallysuggested that SpoIIIE assembles as a ring at constricting sporulation septa (78), the increasedtemporal resolution of HILO revealed that these rings represented the blurred motion of dynamicfoci imaged with long exposure times relative to the rate of movement (19). Two more recentstudies used HILO to reveal more challenging clusters formed by Clp proteases and cell wallprecursor enzymes in the cytoplasm (48, 73), highlighting the potential of this illumination modeto unravel cytoplasmic organization.



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Applying these wide- and near-field illumination methods to bacterial cells has led to severalbreakthroughs but also created some controversies. A decade ago, MreB proteins were first in-ferred to form extended filamentous helical structures from epifluorescence studies (38). However,TIRF microscopy recently revealed that in exponentially growing cells MreBs do not form heli-cal structures; instead, they form patches that exhibit circumferential movement around the cellperiphery (15, 21). Variation in penetration depth and exposure time together with differencesin the growth conditions, fusion proteins (and their levels) used (see above), and postacquisitiontreatment (e.g., deconvolution) may explain the observed differences (10). Figure 1 shows a pair ofexponentially growing B. subtilis cells expressing a functional GFP-MreB fusion imaged simulta-neously by conventional epifluorescence, TIRF microscopy, HILO, and LSCM. Because of the in-creased depth of field, when imaged by epifluorescence the pattern of membrane-associated MreBpatches appears as diffuse and elongated patches around the cell periphery, with significant back-ground (Figure 1a). Similar patterns are observed by HILO (Figure 1c) and LSCM (Figure 1d,middle plane), albeit with reduced background signal. However, TIRF microscopy reveals dis-crete patches along the longitudinal axis of the cell and virtually no signal around the periphery(Figure 1b). Because of the geometry of rod-shaped cells (think of them as cylinders) the sectionthat is illuminated by the evanescent wave only includes the middle section of the cylinder but notthe sidewalls. Consistently, the TIRF image and the image of the bottom plane (sample/coverslipinterface) of the cell obtained by LSCM are very similar. Whereas these techniques have similarlateral (x, y) resolution (Table 1), TIRF microscopy provides the best signal-to-noise ratio andimproved axial (z) and temporal resolution.

Protein Subcellular Localization: The Standard Practice

Given the small size of bacterial cells and the rapid diffusion of proteins in the bacterial cyto-plasm (1–10 μm2/s; significantly slower than in eukaryotic cells, ∼27 μm2/s) (16, 17) and in thecytoplasmic membrane (0.1–1 μm2/s) (60), a free-diffusing soluble protein can move across a bac-terial cell within ∼100 ms and a membrane protein can diffuse across a cell membrane within∼1 s. Thus, diffusing proteins appear as randomly distributed in their corresponding subcellularcompartment. However, proteins display a variety of specific subcellular localizations in bacteria.Protein anchors, geometric or chemical cues (e.g., anionic phospholipids or substrate availability),and regulated or stochastic self-assembly in scaffold cellular structures target proteins to one ormore subcellular locations in a dynamic manner (69, 72, 79).

Some membrane and membrane-associated cytoplasmic proteins localize at specific sites ofthe cell envelope: for example, FtsZ and division and sporulation proteins at future or ongoingdivision sites, MreB and components of the cell wall elongation machinery at distinct foci throughthe sidewalls, and chemoreceptors or the DNA uptake machinery during transformation at the cellpoles. Soluble proteins are believed to mostly diffuse randomly in the cytoplasm, but some formgradients throughout the cell, others localize to or are excluded from the bacterial chromosome,and others form cytoplasmic clusters (56, 72). Yet accumulating evidence in recent years indicatesthat compartmentalization or organelles are prevalent in the prokaryotic world in the form ofprotein-bound and lipid-bound metabolic channels and organelles in the cytoplasm (73, 76, 77).

Protein Subcellular Dynamics: The Emerging Trend

In the late 1990s, early landmark studies on the cell cycle dependence of the localization of FtsZ andother division proteins such as the Min system introduced the concept of spatiotemporal proteinlocalization in bacterial cell biology (55, 70). Similar to their eukaryotic counterparts, bacterial



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Table 1 Comparison of fluorescence microscopy techniquesa

Diffraction-limited Superresolution

Far-field Near-field Far-field

ConfocalWide-field

(EPI) HILO TIRF SIM STED Point-basedx,y-resolutionb

(nm)

180–250 200–250 200–250 200–250 100–130 ∼50 ∼20

z-resolutionb

(nm)500–700 500–700 500–700 ∼100 250–340 ∼100 ∼50

Speed + ++ ++ ++ + + −Contrast ++ − + ++ + ++ ++Live imaging ++ ++ ++ ++ + ++ −Cell region All All Cytosol Surface All All AllPhototoxicity Medium Medium Medium Low Medium High HighMain pros Optical

sectioningSensitive High

contrastFast, sensitive All probes No data

processingSimple setup

Main cons Weakerintensity

Poorcontrast

Uncertaindepth

Surface only Slow, dataprocessing

Complexsetup

Slow,processing

Selected ref.for bacteria

9, 21, 71 38, 56, 70 19, 48, 73 13, 14, 15, 21,40, 51

63, 81 71 17, 18, 19, 62,86, 87, 88

aAbbreviations: ++, very good; +, good; −, poor; EPI, epifluorescence; HILO, highly inclined laminated optical sheet; SIM, structured illuminationmicroscopy; STED, stimulated emission depletion; TIRF, total internal reflection fluorescence.bLateral (x,y) and axial (z) resolution vary depending on the fluorophore (excitation wavelength, λ) used, the biological sample, and objective and specificexperimental conditions. Spatial resolution continuously improves owing to technical innovation. Users should determine spatial resolution for their ownsystem.

Kymograph:graphicalrepresentation ofchange in spatialposition over time

proteins can exhibit cell cycle–dependent localization and/or undergo directed or random motionin the cell during growth. Time-lapse studies and depletion strains can be used to determine thetiming and hierarchy of assembly of protein complexes; e.g., the divisome (22, 36).

Using time-lapse TIRF microscopy, our lab and others recently showed that MreB patchesmove processively along putative tracks perpendicular to the cell axis (15, 21), revealing for thefirst time restricted movement of bacterial proteins in a directed manner. MreB patches couldbe tracked at the single particle level and their speed quantified using kymographs (Figure 2a).Time-lapse colocalization experiments showed that MreB isoforms and other morphogeneticfactors involved in sidewall elongation move together across the cell at the same speed (15). Interms of experimental design, illumination of the biological sample (i.e., exposure time) should beminimized in time-lapse experiments to reduce photobleaching and achieve maximal duration ofobservation. Moreover, frame rate must be chosen carefully to match the timescale of the observeddynamics. In addition, robust imaging analysis tools, ideally aided by automated computation, arerequired to extract quantitative data from these time-lapse images (26).

Kinetics of diffusive proteins in live bacterial cells can also be monitored and quantified bydifferent techniques originally invented to study protein dynamics in eukaryotic cells, includingfluorescence recovery after photobleaching (FRAP), fluorescence correlation spectroscopy (FCS),and single-molecule tracking (SMT, see below) (58). FRAP utilizes laser light to photochemicallyand irreversibly inactivate the FP tag within a subcellular region of interest. The characteristic timeover which fluorescent molecules from the unbleached region of the cell occupy the photobleached



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θθ

t (15

s)

t (6

0 s

)

TIRF Max Kymographsa

bBefore bleaching Bleaching

After bleaching Recovery

Time (min)

Inte

nsi

ty (

a.u

.)

Unbleached area

Whole cell

Bleached area

200 6040

2 μm

–30’’ t0

1’ 15’

1 μm

Figure 2Dynamics of MreB in Bacillus subtilis visualized by TIRF microscopy and FRAP. (a) Motile MreB patches inan exponentially growing cell imaged by TIRF microscopy. (left to right) Single frame of the time-lapseTIRF microscopy, maximum intensity projection of all frames, corresponding kymograph, andrepresentative intensity-colored kymograph trace. MreB patches move processively along tracksperpendicular to the cell’s longitudinal axis. Track orientation is extracted from maximum projections, asindicated by the white arrows. Patch speed can be obtained from kymographs (v = tan θ). Color map ofrepresentative kymograph trace (inset) represents MreB patch fluorescence intensity during its movementacross the cell. (b) Slow turnover of GFP-Mbl filamentous structures imaged by FRAP. A region (outlined inred ) of a slow-growing cell was bleached, and time-lapse images were acquired to simultaneously monitorfluorescence in the bleached region, in a nonbleached region of the same cell, and throughout the whole cell.The experiment was performed in the presence of chloramphenicol, which blocks de novo protein synthesis,to exclude fluorescence recovery due to the synthesis of new GFP molecules. Averaged fluorescence intensityin the three regions was plotted as a function of time. The image sequence shows the GFP images 30 sbefore photobleaching (-30′ ′), the high-intensity laser of the selected region covering approximately half ofthe cell (t0), and recovery after photobleaching at the times indicated. The decay rate of fluorescenceintensity in the unbleached area (blue) was directly proportional to the fluorescence recovery in the bleachedarea (red ), until the same fluorescence was reached in all regions, with a recovery half-time (t1/2) of ∼8 min,indicating continuous exchange of molecules. Abbreviations: FRAP, fluorescence recovery afterphotobleaching; GFP, green fluorescent protein; TIRF, total internal reflection fluorescence.

region determines the diffusion coefficient. FCS monitors the fluorescence fluctuations of highlydiluted fluorophores within a laser focal point. Autocorrelation analysis of such fluctuation can befitted according to different models to obtain the diffusion coefficient as well (58).

In addition to localizing dynamically in the cell, protein complexes may exhibit internal dy-namics. FRAP has also been used to study dynamic turnover within cytoskeletal elements. The Zring was shown to be extremely dynamic, continually remodeling itself with a half-time of 30 s(82), whereas filamentous structures formed by the MreB isoform Mbl in slowly growing B. sub-tilis cells exhibited a half-time of recovery of about 8 min (Figure 2b) (9). Consistent with this



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Abbe diffractionlimit: maximumresolution ofconventional lightmicroscopy due to thediffraction of light,roughly half of thelight wavelength

PSF: point-spreadfunction, blurred 3-Delliptical patternproduced by a pointsource of light

Moire fringe:interference patterncommonly created bysuperimposition of twogrids displaced orrotated a small amountfrom one another

Fouriertransformation:a reversiblemathematicaltransformationbetween spatial andfrequency space; oftenused to analyzeperiodicity

low rate of turnover in MreB assemblies, when individual GFP-MreB or GFP-Mbl patches werepartially bleached in exponentially growing cells, no fluorescence recovery occurred during theirrapid movement across the cell (∼10 s) (15). Conversely, when a whole cell was bleached with theexception of a single GFP-Mbl patch using an inverse FRAP protocol (also known as fluorescenceloss in photobleaching, FLIP), the signal from this patch did not decrease during its movementacross the cell. These FRAP experiments indicate that MreB subunits do not turn over withinone patch during its motion across the cell, which implies that patch movement is not driven bypolymerization/depolymerization (treadmilling) of the actin-like MreB as expected. Thus, quanti-tative analysis of protein dynamics can provide important insights into the mechanism underlyingprotein function.

SUPERRESOLUTION APPROACHES: BEYOND THEDIFFRACTION LIMIT

Regardless of the technique, spatial resolution in conventional fluorescence microscopy isultimately limited by the wavelength of light. Lateral and axial resolution in optical microscopesare limited by the Abbe diffraction limit: ∼200–300 nm in the lateral dimension and ∼500–700 nmin the axial dimension [λ/(2NA) and λ/(2NA)2, respectively], owing to the 3-D diffraction patternof light emitted from a point source, the point-spread function (PSF). In the context of thisdiscussion, this means that all fluorescently tagged proteins and protein complexes (1–10 nm) showup as blurred focal spots that are ∼250 nm in diameter (and ∼600 nm long in the axial dimension),and thus that two molecules within this distance cannot be separated using either a wide-fieldor a confocal fluorescence microscope. Fluorescence microscopy methods that push or break thediffraction limit of resolution have seen explosive growth in the last decade and revolutionizedsubcellular imaging in eukaryotic systems (30). These superresolution nanoscopy techniqueshave just launched in bacterial studies. A typical bacterium (1–4 μm) is not much larger than thediffraction limit, therefore presenting a challenge and becoming a benchmark for imaging innova-tion (33). In this section we provide a quick overview of the principles, advantages, and drawbacksof the commercially available techniques from a microbiologist’s perspective, and we give thefew known examples of their application to visualize bacterial structures (also summarized inTable 1).

Structured Illumination Microscopy

Structured illumination microscopy (SIM) (27) uses classical wide-field illumination combinedwith signal acquisition through a grid that generates interference patterns when rotated andtranslated across the sample (Figure 3a). The signal variations between the different imagesare analyzed for each pixel to reconstruct in silico a higher-resolution 2-D image (27). Typically,three grid orientations (rotations) and three to five grid positions (translations) are used, resultingin 9–15 images per final SIM-reconstructed image of a given x,y plane (47). The information inthe moire fringes is extracted through Fourier transformation between spatial space and frequencyspace and used to generate an image with ∼twofold gain in lateral resolution (47). By enhancinglateral resolution in each plane of a z stack (3-D SIM), SIM can also provide higher-resolutioninformation of 3-D structures. Recently, 2-D SIM was used to visualize MreB structures (71)(Figure 3b) and SpoIIQ foci (20) in B. subtilis, and 3-D SIM revealed that FtsZ is heterogeneouslydistributed in the Z ring instead of forming a continuous ring structure as traditionally assumed(81) (Figure 3c). 4-D SIM (time-lapse 3-D SIM) additionally showed fluorescence fluctuationsin the discontinuous Z ring, indicating constant exchange between cytoplasmic and polymerized



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ba TIRF TIRF-SIM Intensity c

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Figure 3Structure illumination microscopy (SIM). (a) Phase and angle shifts during SIM image acquisition. SIM relies on moving a fine patternof grids across the sample to generate moire fringes, followed by mathematical processing and deconvolution to enhance resolution andcreate a single high-resolution image. (b) Comparison of TIRF and TIRF-SIM images of the same Bacillus subtilis cell expressingGFP-MreB. The line intensity profile shows that TIRF-SIM displays a twofold increase in lateral resolution. (c) 3-D SIM image ofFtsZ-GFP in B. subtilis revealed the bead-like, discontinuous structure of the Z ring. Other abbreviations: GFP, green fluorescentprotein; TIRF, total internal reflection fluorescence. Images were kindly provided by A. Rohrbach (b) and E. Harry (c).

FtsZ, as previously shown by FRAP (82). Finally, two-color, 3-D SIM provided further insightinto the recruitment of SpoIIIE to the Z ring during the early steps of septation in dividing andsporulating cells (18).

A major advantage of SIM is that it is compatible with conventional (nonswitchable) fluo-rophores (e.g., GFP) and can easily image multiple dyes. However, the high number of acquisi-tions required for every resolved image (9–15 × number of z stacks) is a technical limit to imagingsparse proteins and very dynamic structures in live cells. Fluorophores must be sufficiently abun-dant and photostable to support repeated illumination without significant photobleaching, andthe observed structures must stay steady during the whole acquisition process to be properly re-constructed. Also, SIM-reconstructed images often contain artifacts, observed as residual stripepatterns, that result from imprecise instrumental hardware, bleaching, and/or incorrect experi-mental setups that cause spherical aberration, such as using media or oil of the wrong refractiveindex, or coverslips of the wrong thickness.

Stimulated Emission Depletion Microscopy

Based on LSCM, stimulated emission depletion (STED) microscopy (31, 46) fluorescently ac-tivates a spot and then shrinks its diffraction-limited focus by using a second doughnut-shapedlaser that selectively switches off fluorophores in the periphery, so that only photons from thecentral (subdiffraction) hole are collected (Figure 4a,b). Theoretically speaking, the resolution ofSTED could be unlimited if the power of the depletion laser beam were increased. In practice,photobleaching (because of constant excitation of the fluorophore) and phototoxicity (due to thegreater excitation energy applied over time) limit the range of power usable for the high-intensitydepletion laser. Currently, STED reaches three- to tenfold improvement in lateral resolution(25–80 nm) (30). MreB assemblies were recently imaged by STED in live B. subtilis cells (63).



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a c

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Confocal STED

Confocal

STED

hvexc.

hvSTED

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Spontaneousrelaxation

Figure 4Stimulated emission depletion (STED) microscopy. (a) Jablonski energy diagram depicting the transitionbetween electronic states of a fluorophore in STED during stimulated emission. In addition to the excitationlaser (in blue; e.g., 640 nm), a second red-shifted, doughnut-shaped STED laser (in red; e.g., 740 nm) inducesstimulated emission, before spontaneous fluorescence relaxation occurs ( green arrow), which effectivelydepletes regular emission of the fluorophores (excepting those in the doughnut hole). Photons released fromstimulated emission (in red ) are identical to the STED laser and are filtered out. (b) The effective PSF ofconfocal and STED microscopy is visualized in blue. In STED, the doughnut-shaped depletion laser (in red )shrinks the confocal PSF, sharpening the remaining fluorescence spot to a size much smaller than thediffraction-limited focus obtained by confocal microscopy. (c) Comparison of confocal and STED images ofthe same Bacillus subtilis cell expressing GFP-MreB. Other abbreviations: GFP, green fluorescent protein;PSF, point-spread function. Image kindly provided by P. Graumann.

Photoactivation:irreversible process bywhich a moleculebecomes fluorescentupon a conformationalchange achieved byirradiation with a pulseof light

Under the growth conditions used, filamentous MreB structures were observed across the widthof the cell with a resolution of 50 nm (Figure 4c). STED can use conventional fluorophoresand does not require data processing, and image acquisition can be very fast for small fields ofview (but it is relatively slow for large fields). Drawbacks are mainly related to the high level ofphotobleaching/phototoxicity and the complexity of the optical system.

Point Localization–Based Superresolution Microscopy

Photoactivation localization microscopy (PALM) (4, 32) and stochastic optical reconstructionmicroscopy (STORM) (74) combine wide-field (generally TIRF) laser illumination with theso-called pointillism strategy. The difference between PALM and STORM is that they rely ongenetically encoded photoactivatable/photoconvertible proteins or on photoswitching syntheticdyes, respectively, to let individual fluorophores go on and off stochastically (4, 28, 64, 74).



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a b cActivation/conversion

Excitation

Localization

Reconstructions/sqrt (N)

Figure 5Point localization–based superresolution microscopy. (a) Centroid determination. Least-squares fit to a Gaussian approximation of thePSF is applied to individual single-molecule events to obtain precise centroid position. Localization accuracy is determined by thestandard deviation of the original PSF (s) and number of photons from each fluorophore (N), roughly 10 nm for 1,000 photonscollected. (b) Schematic illustration of PALM/STORM principle. Iterative cycles of photoactivation/conversion with a first laser (often∼405 nm) followed by excitation by a second laser (typically 488–561 nm) and localization of relatively few, well-separatedfluorophores within densely labeled structures. The final image is reconstructed by compiling mathematically determined centroidpositions from all imaging cycles. (c) The principle of PALM/STORM is similar to that of pointillism paintings, such as La Tour Eiffelby Georges Seurat. Abbreviations: PALM, photoactivation localization microscopy; PSF, point-spread function; STORM, stochasticoptical reconstruction microscopy.

Pointillism:a strategy to achievesuperresolution bymerging allsingle-moleculepositions; also knownas localization-basedsuperresolutionmicroscopy; includesPALM and STORM

Photoswitching:reversiblelight-inducedconformational changeof a molecule thatchanges itsfluorescenceproperties

Thus, otherwise spatially overlapping images of individual molecules are temporally separated(Figure 5b). The centroid of each diffraction-limited spot can then be determined by applyinga least-square fit to a Gaussian approximation of the PSF (Figure 5a). Thousands of framesare acquired from which sparsely distributed centroids are localized and summed to generate asuperresolution image, like in impressionist pointillism paintings (Figure 5c). The precision ofcentroid localization is proportional to the number of photons received, and thus lateral resolutionis also theoretically unlimited. Current setups allow ∼20- to 30-nm lateral resolution. Recentdevelopments based on aberrated and/or engineered PSF imaging allow a huge improvementin axial resolution too (up to ∼50–80 nm), in 3-D PALM/STORM experiments (34, 41, 67) aswell as in 3-D STED (29). However, the resolution of the reconstructed images heavily dependson the total number of events recorded, which depends on the number (labeling density) andphotostability of fluorophores in the cell. PALM/STORM setups are relatively simple (andthus cheaper than SIM or STED systems), but powerful computers and software are needed tohandle and process the enormous amount of data generated. Different analysis algorithms canoutput different results, rendering image interpretation a difficult task. Application of PALMand STORM to bacteria is becoming increasingly popular. PALM has been used to studyclustering of chemotaxis receptors in the poles of E. coli cells (24) and recruitment and assemblyof SpoIIIE in sporulating B. subtilis cells (18, 19), whereas STORM has been used to study thenucleoid-associated protein H-NS (88) and cell wall synthesis in E. coli (86).



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a b c

MS

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d

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Excitation

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Tracking

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Figure 6Point localization–based single-molecule tracking (SMT). (a) Principle of SMT. Few, well-separatedfluorophores (ideally a single fluorophore) are expressed or activated/converted, followed by pulses ofexcitation. Sequential localization of each subcellular position and centroid determination with nanometricprecision produces the trace of each fluorophore. (b–d ) PALM tracking and analysis of single fluorescent-labeled DNA polymerase I (Pol-PAmCherry) molecules in live Escherichia coli cells. (b) Micrographsrepresent (from top to bottom) maximum-intensity projection in a representative cell; PSF of a single Pol-PAmCherry molecule; representative tracks of diffusing Pol (blue) and bound Pol (red ) on a transmitted lightimage; and all Pol tracks in the representative cell. (c) MSD analysis for Pol in live cells. Shape of the MSDcurve indicates confinement of Pol within the nucleoid. Effective diffusion coefficient can be obtained byfitting the MSD curve to confined diffusion models. (d ) Cumulative distributions of the diffusion step lengthbetween consecutive localization for Pol in live cells (blue) and fixed (red ) cells. Cumulative distributions shifttoward larger steps with increasing diffusion coefficient D. Other abbreviations: MSD, mean squareddisplacement; PALM, photoactivation localization microscopy; PSF, point-spread function. Modified andreprinted, with permission, from Reference 87.

Single-Molecule Tracking: One Molecule Over Time

Like PALM and STORM, SMT uses single-molecule centroid determination with nanometerprecision as the underlying principle. However, whereas ensemble techniques reconstruct denselylabeled structures by adding many single-molecule centroids, SMT investigates the dynamics of asingle molecule over time, holding the potential of untangling short-lived, intermediate states anddynamic events at the single-cell level. Expression of the fluorescent fusion protein or activationenergy of a photoactivable fluorophore is minimized to limit the number of excited single fluo-rophores in the cell. Then multiple localization steps of individual fluorophores are detected untilphotobleaching. Centroid positions of each position are localized and added to generate single-molecule trajectories (37, 53) (Figure 6a,b). Further quantitative analysis can provide diffusioncharacteristics and kinetics; e.g., diffusion coefficients (Figure 6c,d). SMT datasets can easily reach∼10,000 frames (for ∼1,000 single molecules) over the course of several minutes. Sophisticatedimaging analysis tools are therefore needed to extract and organize meaningful data (37).

So far, only a few proof-of-concept studies have used SMT in bacterial cells. Pioneering workby Kim et al. showed that two subpopulations of MreB exist in C. crescentus; one exhibited fastmotion in the cytoplasm, characteristic of Brownian motion, and the other displayed slow, directedmovement in the membrane (43). A similar study found two heterogeneous subpopulations of FtsZmolecules with distinct diffusion dynamics in E. coli (62). More recently, SMT was applied to study



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Microfluidic device:microcham-ber/microchannelimmobilizing cells anddelivering smallvolumes of fluids; idealfor time-lapsesingle-cell analysisduring growth andupon dynamicperturbations

diffusive and bound states of DNA repair enzymes (87) (Figure 6b–d) and of RelA, a key mediatorin stringent response (17) in E. coli. Altogether, these studies reveal that localization and dynamicsof a given protein can be highly heterogeneous at the single-molecule level in a given cell. Lookingahead, we expect SMT to illustrate that there is no average molecule, potentially refuting resultsof bulk assays, much as single-cell experiments debunked the myth of the average cell (90).

CONCLUDING REMARKS: FUTURE CHALLENGESAND OPPORTUNITIES

Microbiology is in transition from a discipline that is preoccupied with assigning functions toindividual proteins or genes to one that is trying to grasp the complexity of molecules that interactand self-assemble to form functional modules and establish long-range orders. This requirescareful examination of the intracellular localization and dynamics of individual proteins and proteincomplexes. One of the most exciting prospects is the possibility of following, in real time, thedynamics at play in recruiting, assembling, maintaining, and ultimately disassembling bacterialsubcellular structures (49). The superresolution techniques described in this review have nowpassed the proof-of-principle test in bacterial cell biology. In the next few years, TIRF, HILO,SIM, STED, PALM/STORM, SMT, and techniques that combine multiple and new modalitieswill be widely used in the field. However, despite the unprecedented improvement in resolution,limitations still exist regarding imaging speed, sensitivity, field and depth of view, instrumentationcomplexity, and fluorophore versatility (Table 1), and thus these techniques are not always feasibleor optimal for live-cell imaging. Microbiologists should balance the strength and limitation of eachtechnique before applying it. In general, they should consider factors such as the abundance andthe dynamics of their protein of interest, what resolution is sufficient, what fluorophores work intheir system, etc. (12). It is recommended to use multiple methods and to interpret each result inlight of the limitations and strengths of each method. Caution is also needed when interpretingthe observed structures to ensure that no artifacts are created, in particular by techniques basedon image reconstruction such as SIM and PALM/STORM.

Looking to the future, we anticipate technological innovation and new questions and conceptswill be addressed. First, superresolution/single-molecule methods demand further advances. Forexample, technical development is still needed to visualize intracellular single molecules in 4-D(SMT in 3-D). A possibility also lies in multicolor and live pointillism techniques, currently lim-ited by the slow acquisition time needed in PALM/STORM in commercially available platforms(39). Next-generation fluorescent probes and labeling strategies well suited for bacterial super-resolution imaging studies will probably be in play soon. A different direction of innovation takesadvantage of genome-wide fluorescent fusion libraries (11, 45, 56, 89) and high-throughput auto-mated microscopy to carry out image-based screens. New genes modulating subcellular structures,such as FtsZ-binding proteins, were successfully identified by this method in C. crescentus (23).It would also be important to develop high-throughput methods for studying a large number ofsingle-molecule events in a large set of experimental conditions or to screen libraries of compoundsin search of new antibacterial molecules.

Conceptually, three avenues of investigation are likely to yield exciting results. First, study ofthe establishment, maintenance, and regulation of subcellular patterns such as small membranemicrodomains (52), cytoskeletal scaffolds (57), and multiprotein factories (35, 42) should berevisited with new microscopy methods. Ideally, these studies will be carried out at the single-celllevel in real time in well-controlled and easily adjusted environments using microfluidic devices.Second, shared as well as distinct features in different model microbial organisms need to beinvestigated and, when appropriate, compared with their counterparts in eukaryotes (35, 66).



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For example, extensive knowledge about actin dynamics in eukaryotic systems might provideinvaluable hints as to how it is regulated in bacteria in response to diverse environments (3, 59).Last, major advances will require concerted interdisciplinary effort. Fluorescence microscopyin combination with powerful genetics, genomics, and biochemistry techniques available inseveral model organisms and in combination with systems and synthetic biology approaches,physics, material science, and mathematical modeling will provide the intellectual framework forunderstanding how a set of molecules self-assemble and build a cell (1, 68).

DISCLOSURE STATEMENT

The authors are not aware of any affiliations, memberships, funding, or financial holdings thatmight be perceived as affecting the objectivity of this review.

ACKNOWLEDGMENTS

We apologize to all researchers whose relevant studies could not be included here because ofspace constraints. We thank Arnaud Chastanet for his comments on the manuscript. We alsothank Michael Gue and the French Team of Zeiss for providing assistance for Figure 2. Work inthe R.C.L. laboratory is supported by a starting grant from the European Research Council (ERC-StG 311231), the French National Research Agency (ANR-12-ISV3-0004-01, ANR-12-BSV3-0021-02), and the EMBO Young Investigator program (EMBO YIP 2259). Z.Y. is supported bya long-term postdoctoral fellowship from the Human Frontier Science Program Organization.

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fluorescence imaging for bacterial cell biology: from localization to dynamics, from ensembles to...

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