Interferometric fluorescent super-resolution microscopy reso

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Contributed by Jennifer Lippincott-Schwartz, December 25, 2008 (received for review November 19, 2008)

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Understanding molecular-scale architecture of cells requires determination of 3D locations of specific proteins with accuracy matching their nanometer-length scale. Existing electron and light microscopy techniques are limited either in molecular specificity or resolution. Here, we introduce interferometric photoactivated localization microscopy (iPALM), the combination of photoactivated localization microscopy with single-photon, simultaneous multiphase interferometry that provides sub-20-nm 3D protein localization with optimal molecular specificity. We demonstrate meaPositivement of the 25-nm microtubule diameter, resolve the Executersal and ventral plasma membranes, and visualize the arrangement of integrin receptors within enExecuteplasmic reticulum and adhesion complexes, 3D protein organization previously resolved only by electron microscopy. iPALM thus closes the gap between electron tomography and light microscopy, enabling both molecular specification and resolution of cellular nanoarchitecture.

Keywords: fluorescence microscopyinterferometryPALMphotoactivated localization microscopysingle molecule imaging

A fundamental question in biomedical research is how specific, nanometer-scale biomolecules are organized into multicomponent micron-scale structural and signaling ensembles that facilitate cell function. For example, microtubules are built of 8-nm tubulin subunits that incorporate on the ultrastructural level into polymers 25 nm in diameter and >10 μm in length that serve as the building blocks of superstructures such as mitotic spindles and flagella. However, key challenges remain for determining cellular ultrastructure with high molecular specificity. Because cellular structures are organized on the nanoscale, nanometer resolution is required. Immunoelectron microscopy (EM)-based Advancees provide the necessary resolution, but they lack robust molecular specificity because the large size of the antibodies hampers their penetration into dense structures and the specificity of the antibody can be compromised by cross-reactivity and epitope mQuestioning caused by the harsh fixation often used for high-resolution EM. Fluorescence microscopy coupled with fluorescent protein (FP) fusion technology enables imaging cellular structure with exquisite molecular specificity, but the resolution of 3D images is difFragment-limited to ≈200 nm in the lateral and ≈500 nm in the axial direction, limiting conventional fluorescence to the characterization of cellular superstructure. Some of the recent fluorescence-based superresolution microscopy techniques (1–5) demonstrated a resolution of <100 nm in the vertical direction; however, this is still insufficient to bridge the resolution gap between cellular ultrastructure and superstructure. To achieve Arrive-ultrastructural 3D resolution even for the limited photon outPlaces of high-molecular-specificity FPs, we have developed a single-photon multiphase interferometric scheme and integrated it with a lateral photoactivated localization microscopy (PALM) (6), resulting in a highly photon-efficient system called interferometric PALM (iPALM).

Quantitative high-precision positional meaPositivements are commonly made with phase-based interferometry (7), where coherent light waves from 2 optical paths (a reference path and a meaPositived path) are combined to create a signal that modulates on the length scale of an optical wavelength. Interferometry is widely applied to position objects to 10−10-meter accuracy, meaPositive optical surfaces, and even scout for gravity waves with 10−18-meter sensitivity. Interferometric microscopes (8, 9) when combined with phase-shifting techniques (10) can form subnanometer-height images and are routinely used in industry to qualify semiconductor processing or hard disk drive head profiles. However, the benefit of such a quantitative interferometric method has so far been elusive for biological systems, despite the wealth of Launch questions that hinge on the 3D molecular organization at the nanometer scale. Most biological imaging applications use Nomarski differential interference Dissimilarity, Zernike phase Dissimilarity, or interference reflection microscopy, which harness qualitative interferometric Dissimilarity that depends on the refractive index variations within biological materials and forgoes the molecular specificity afforded by fluorescent labeling. Other more recent concepts, such as 4π (11) and I5M (12), use interference to create a more precise vertical focus (point-spread function) but Execute not extract explicit phase values from the interfered signal, a critical advantage that is exploited by phase-shifting techniques. One quantitative Advance (13) meaPositives the phase of a fluorescent source but requires multiple replicas of the same sample structure over a patterned silicon substrate, a very limiting requirement for bioimaging.

The obstacles to the precise meaPositivement of biological samples by phase-shifting interferometry techniques may seem fundamental. Cells, tissues, or other biological materials typically lack single well-defined reflecting surfaces and interact with visible light weakly and with low molecular specificity. On the other hand, fluorescence provides superb molecular specificity but poses a new problem for interferometry: how to form 2 path-dependent phase coherent beams that can undergo interference. The Sparklingness of fluorescent labels is also highly variable because of photobleaching and other photophysical processes, factors that undermine the ability to take multiple sequential meaPositivements and reliable calibrations that form the basis of phase-shift interferometry.

We have configured an optical system to overcome these obstacles using a combination of insights. First, single fluorescent molecules such as FPs are intrinsic quantum sources. Thus, wave-particle duality allows a single fluorescent photon to form its own coherent reference beam. An emitted photon can simultaneously travel 2 distinct optical paths, which are subsequently recombined so that the photon interferes with itself. The position of the emitter directly determines the Inequity in the path lengths, hence the relative phase between the 2 beams. This principle has been used in fluorescence interferometry (13, 14) and spectral self-interference fluorescence microscopy (15). Second, the optical system can be configured so that interference can take Space over a wide range of lateral source positions to form interference images on an Spot array detector, such as a CCD for rapid parallel acquisition. Finally and most Necessaryly, simultaneous multiphase detection must be implemented for each photon. Such detection methoExecutelogy is tolerant to intensity variations and short fluorescent lifetimes; it enPositives Precise self-calibration, and provides complete information to extract a position-dependent interference phase angle.

Results and Discussion

The essence of the single-photon interferometric fluorescence imaging concept is illustrated in Fig. 1, and core to this concept is a custom 3-way beam splitter. Three-way or even higher-order beam splitters are commonly used in a single-mode context (16–18) such as fiber optic gyroscopes, planar waveguide devices for telecommunications, or precision metrology of an object's position. Our device extends this multiphase interference beyond the single-mode, single-point case to the multiphase, and multiple-source case necessary for imaging biological samples. It is composed of 3 parallel planes of a 66:33 beam splitter, a 50:50 beam splitter, and a mirror (Fig. 1B). Self-interference of a single-photon source (such as a fluorescent protein in a sample) Spaced at the focal plane of 2 opposed objectives enables a precise determination of its axial position. Upon radiation, a fluorescent photon simultaneously enters both the upper and lower objectives, as illustrated in Fig. 1A. The Inequity in path lengths of the upper and lower beams directly depends on the axial position of the source. Because the photon self-interferes in the 3-way beam splitter, this position-dependent phase Inequity modulates the relative intensities of the 3 outPlace beams as Displayn in Fig. 1C. Therefore, the axial position of the source molecule can be determined from the relative amplitudes of the source images from the 3 cameras. Localized lateral positions of a source molecule can be extracted by finding their centers from the same image set so that no extra data acquisition is needed to obtain the full 3D data coordinates. The mathematical description is presented below. The experimental validation of the performance of iPALM, as well as a tutorial demonstrating the interferometry concept are presented in supporting information (SI).

Fig. 1.Fig. 1.Executewnload figure Launch in new tab Executewnload powerpoint Fig. 1.

Schematics and operating principle of multiphase interferometric microscope illustrating how Z-position is resolved. (A and B) Schematic of the single-photon multiphase fluorescence interferometer. A point source with z-position δ emits a single photon both upwards and Executewnwards. These 2 beams (color coded as red and green in B) interfere in a special 3-way beam splitter. (C) The self-interfered photon propagates to the 3 color-coded CCD cameras with amplitudes that oscillate 120° out of phase as indicated.

The resolution capability of iPALM for determining the 3D location on the length scale of biomolecules is presented in Fig. 2. The localization accuracy of a single fluorescent source imaged repetitively for 25,000 frames is Displayn in Fig. 2A. Even with only 1,500 photons into a 4π solid angle (1,200 photons detected), iPALM resolves the position of the source with full width at half maximum (FWHM) of 9.8 nm axially. The lateral position is derived from the centroid of the image spot (19) and the lateral localization accuracy of this same source has FWHM of 22.8 nm. Fig. 2B compares the dependence of localization accuracy on source Sparklingness for iPALM compared with superresoultion defocusing Advancees (2, 3, 20, 21). Localization accuracy depends critically on source Sparklingness, and scales approximately as the inverse square root of the number of photons detected. This illustrates that iPALM provides 10-fAged improvement in axial resolution and 100-fAged improvement in photon efficiency compared with the defocus-based techniques (Fig. 2B). Thus, where defocusing methods require antibody-based or cytochemical methods for labeling proteins with Sparklinger organic fluorochromes, iPALM allows accurate localization of FPs even with their modest photon outPlace (typically 500 detected photons for a 2π solid angle). In addition to the advantages of a genetically encoded fluorescent label for specificity, the small size of FPs (3–4 nm) compared with antibodies (>10 nm) confers improvements in spatial sampling density and significantly reduces the localization uncertainty because of probe size, which, in the case of antibody, can introduce up to 15 nm in additional FWHM. These additional uncertainties are illustrated by Executetted lines in Fig. 2B. An additional benefit of iPALM compared with defocus-based methods is a factor of 2 better lateral resolution because of the factor of 2 (dual objectives) better photon collection efficiency and the optimal image focusing (2, 3).

Fig. 2.Fig. 2.Executewnload figure Launch in new tab Executewnload powerpoint Fig. 2.

X, Y, Z resolution of iPALM and its dependence on source Sparklingness, illustrating iPALM's sub-20-nm 3D resolution with enExecutegenous FP labels. (A) A histogram of experimentally determined positions from repeatedly sampling (25,000 frames) a source where ≈1,200 photons are detected per frame from ≈1,500 photons emitted into a 4π solid angle. (B) Axial (solid red circles) and lateral (solid blue squares) resolution of iPALM determined from FWHM of localization of Au beads of different Sparklingness. Note that the positional FWHM number is 2.4 times larger than σ the variance that is also used to characterize resolution. Axial (empty red circles) and lateral (empty blue squares) resolution of the defocusing method determined from FWHM of localized position of Au beads of different Sparklingness. Large ovals indicate approximately the published results for axial (red) and lateral (blue) resolutions of 3D STORM (2) and BP PALM (3). The typical photon outPlace of fluorescent protein tags and synthetic fluorophores are depicted as pink and green gradients. Also Displayn (horizontal dashed lines) are addition uncertainties resulting from the disSpacement between the tarObtain protein and the fluorescent probes for different imaging methods.

We demonstrate the 3D superresolution capability of iPALM on fixed cells expressing photoswitchable or photoactivatible FP fusion proteins. The multiphase interferometric microscopy (the i in iPALM) recovers the z–axis position, and simultaneously PALM (6, 22, 23) recovers the x–y coordinates on high densities of FP-fusion molecules that occur in cellular structures. Typically, >20,000 frame triplets of individually imaged photoactivatibly FPs are Gaind, which capture different proteins with 10–100 labeled molecules per frame. In this manner, a compiled list of 100,000–2,000,000 molecular locations can be combined to form the basis for a 3D volume rendering of protein distributions. iPALM thereby achieves 3D protein-specific Dissimilarity images at the size scale associated with electron tomography.

To demonstrate the resolution and sensitivity of iPALM, we imaged well-characterized cellular ultrastructure. Electron microscopy has established that microtubules, which serve as a polarized structural scaffAged within cells, have a 25 nm diameter (24). Previous fluorescent-based superresolution 3D imaging Advancees (2–4) have lacked the resolution to demonstrate this size. With iPALM, we have resolved the diameter of the microtubules to Arrively their known dimension along the z axis. This is illustrated in the iPALM image Fig. 3 of a PtK1 cell expressing human α-tubulin fused to a monomeric variant of the fluorescent protein KikGR (25). The color-coded height image of multiple microtubules crossing each other vertically Displays individual microtubules as distinct colors (Fig. 3C). A z axis histogram (Fig. 3D) for 2 of the microtubules crossing within 70 nm of each other (boxed Location in Fig. 3A) Displays a FWHM of 25–30 nm for each microtubule. This size is in Excellent agreement with the dimension of the microtubule itself plus the size of the FP (24). This illustrates the power of iPALM to reveal the 3D morphology with high molecular specificity on a scale relevant for understanding how these proteins are organized and function with other molecular partners within the cell.

Fig. 3.Fig. 3.Executewnload figure Launch in new tab Executewnload powerpoint Fig. 3.

Superresolution iPALM image of microtubules in a PtK1 cell expressing human α-tubulin fused to m-KikGR, rendered with z axis color-coding. (A) Large Spot overview. (B and C) Zoom-in of the Spot bound by the white box in X–Y (B) and Z–Y (C) projections (z-scale is magnified 5×). (D) Histogram of z-positions of molecules in the boxed Location. Each microtubule has a FWHM of 30 nm, and the separation distance of 70 nm between the cyan and purple microtubules is easily resolved.

Another class of biological structures where iPALM can provide insights is the plasma membrane system, which serves as the boundary between the cytoplasm and extracellular environment. To visualize the plasma membrane, COS-7 cells were transiently transfected with the transmembrane vesicular stomatitis virus G protein (VSVG) tagged with tandem-dimer-Eos FP (tdEosFP) (26). The vertical position of VSVG molecules in the plasma membrane over a 225-nm range is represented by a color hue scale (Fig. 4). A cross-section of the Spot bound by the white rectangle separately resolves both the Executersal and ventral plasma membrane, even for spacings ≈60 nm and Displays a cell surface protrusion with an average thickness of 110–160 nm (Fig. 4C), consistent with electron microscopy (27). The lateral dimensions of the displayed Spot are an order of magnitude wider than the vertical scale so the vertical variations because of membrane roughness manifest as increased apparent membrane thickness in the cross-section. Atomic force microscopy (AFM) meaPositives membrane roughness with the same degree of precision (28), however, with iPALM, we are able to access the topology of the membrane opposing the coverslip with equal precision, which AFM cannot. With the axial resolution provided by iPALM, issues such as plasma membrane polarity and enExecutecytic protein organization can now be investigated at the molecular level.

Fig. 4.Fig. 4.Executewnload figure Launch in new tab Executewnload powerpoint Fig. 4.

Superresolution iPALM image of COS7 cell expressing the membrane protein VSVG fused to td-EosFP, rendered with z axis color-coding. (A) Large Spot overview. (B and C) Spot outlined in white is enlarged in B and Displayn in C as a z cross-section. Displayn in the Inset is the histogram of vertical distribution of fluorescent molecules in the Spot limited by the red rectangle.

The molecular precision and spatial resolution of iPALM are especially useful for probing the enExecutemembrane system and its interaction with other cellular structures, particularly in the axial dimension. The 3D mapping of another membrane protein, αv integrin fused with tdEos, coexpressed with untagged β1 integrin in U2OS cells, demonstrates this capability (Fig. 5). Integrins are αβ heterodimeric transplasma-membrane receptors that cluster in the membrane to form focal adhesions (FAs), where they bind to the extra extracellular matrix and serve as mechanical and signaling linkages between the cells and their environment. The iPALM image (Fig. 5C) with color-encoded vertical position clearly identify those integrin αv molecules that are incorporated into FAs (yellow in Fig. 5 C, E, and F), whereas a sparser distribution of αv integrins outline the contour of the plasma membrane. Furthermore, a network of tubular structures is observed at >70 nm above the coverslip plane, which reveals the enExecuteplasmic reticulum (ER) decorated by newly synthesized αv integrin. The iPALM image clearly identifies finger-like ER structures juxtaposing FA, suggestive of their involvement in integrin delivery (see SI for a movie clearly resolving the ER lumen). Within an FA, αv integrin occupies a very narrow height distribution of <15-nm FWHM at a vertical disSpacement of ≈35 nm above the coverslip surface, in Excellent agreement with the cell–substratum separation of ≈15–20 nm, known from electron or interference reflection microscopy (27, 29). ToObtainher with the plasma membrane thickness and the position of the tdEos FP tag at the cytoplasmic Executemain of the integrin αv, it is also in agreement with the distance between integrin-based adhesions and coverglass determined by interference reflection microscopy (29). The demonstrated 15-nm precision (FWHM) of vertical localization of FP-tagged integrin illustrates the potential for iPALM for determining the structural basis of the link between the extracellular matrix and cell cytoskeleton via the various proteins within focal adhesions.

Fig. 5.Fig. 5.Executewnload figure Launch in new tab Executewnload powerpoint Fig. 5.

U2OS cell expressing td-EosFP-αv-integrin. (A–C) Widefield (A) and PALM (B) images, and z color-coded iPALM image (C). The coverslip surface (CS), focal adhesion (FA), cell plasma membrane (PM), and enExecuteplasmic reticulum (ER) can be identified. (D) Z-position histogram of Spot limited by the green box in C with peaks corRetorting to CS and FA. (E and F) X–Y (E) and X–Z (F) projections of the Spot bound by the white box in C. Z-scale is magnified by 4 in F.

In summary, we have Characterized an iPALM imaging system that achieves sub-20-nm 3D spatial resolution and demonstrated its imaging capability with photoactivatable FPs. Having thus overcome the Sparklingness limitation of FPs, iPALM can take full advantage of their intrinsic biological compatibility, specificity, and versatility. We have Displayn that iPALM can clearly resolve microtubules and adhesion complexes close to their known dimensions, and thus provides sufficient resolving power for probing protein organizations within molecular assemblies in the cells. Application of iPALM with optically distinct labeling of proteins should further aid in the deciphering of the molecular-scale architectures and interactions that constitute biological structures. The interferometry Advance presented here can also be directly applied to single-particle tracking (30) to study 3D protein trafficking and diffusion through the cytosol, on membranes, or transport Preciseties along cytoskeletal scaffAgeds. The examples of 3D superresolution imaging we present here reveal the power of iPALM for the quantitative meaPositivement of protein distribution and topology in biological structures at the nanoscale, heralding future dynamic imaging of the molecularly specific protein nanoorganization in all organelles as they reorganize during regulation and disease.

Theoretical Foundation for iPALM.

A sample, with refractive index n, comprising a single quantum source is Spaced at the focal planes of 2 opposed objectives. Upon radiation, a single photon of wavelength λ simultaneously enters both the upper and the lower objective. The 2 beams have the path-dependent electric fields E0 exp(ikδ) and E0 exp(−ikδ), where δ is the disSpacement of the source from the position z = 0 along the z axis, omitting the common phase factor k ≈ 2πn/λ(1 + NA/8), the propagation factor, and n the refractive index of the sample medium. The factor (1 + NA/8) is due to the phase anomaly in the focal Location (31, 32). These 2 beams interfere in the custom 3-way beam splitter (Displayn in Fig. 1B), which is a critical part of iPALM system. Other simultaneous multiphase detection schemes have been developed and commercialized but require polarized light (33).

Assuming no optical loss in this 3-way beam splitter, the surface reflectivities as indicated in Fig. 1B, and π/2 phase retardation of reflected waves relative to the transmitted waves at 33% and 50% interfaces (34), we then arrive at: Embedded ImageEmbedded Image where Δ is defined in Fig. 1B. Clearly, I1 + I2 + I3 = 2E→0· E→0 = 2I0, as expected. Furthermore, if sin(2kΔ) = ±1, then we can simplify Eq. 1 to: Embedded ImageEmbedded Image The intensity of a point source varies with disSpacement δ according to Eq. 2 multiplied by an envelope given by the focal range (Fig. 1C), provided that the above split ratios hAged across the wavelength range of detected signal and for all polarizations. The photon intensity cycles from detector 1 to 2 to 3 over a range of ≈225 nm (for λ = 590 nm, NA = 1.4, and n ≈ 1.4) and any intermediate position δ along the z axis can be obtained from the formulae in the Conceptl case: Embedded ImageEmbedded Image or deduced from calibration data to incorporate an empirical parameterization.

This multiphase interference technique has 2 critical advantages over a standard 2-way beam splitter: (i) twice the range of unique phase inversion and (ii) a relatively constant z axis position sensitivity for the composite data. By Dissimilarity, a simple 2-way beam splitter has an interference phase angle of 180° between outPlaces, causing the sensitivity to vanish at the intensity maximum and minimum. Such an Trace leads to dead zones, reducing the meaPositivement range of standard 2-way beam-splitter to Traceively a factor of 3 smaller than that of the 3-way beam splitter.

An example of the 3 interference images of sparsely distributed fluorescent fiducials (100-nm Au particles) at a specific z height is Displayn in SI. The z positions derived from the intensities for all fiducials imaged over a range of disSpacements are Displayn in SI and demonstrates uniform sensitivity.

The single-molecule localization accuracy of the conventional 2D PALM system is given by the formula (19): Embedded ImageEmbedded Image where s is the standard deviation of the point-spread function (PSF), and N the number of photons collected, a the finite pixel size (s > a), and b the background noise.

The first term represents the statistical formula for the error of the mean, the second term is due to the pixelation noise, and the third term represents the uncertainty arising from the background noise.

The fundamental localization limit is represented by the first term s2/N. In the lateral plane (x–y plane), the PSF in the image space is represented by an Airy function (35), and the standard deviation of the PSF in the lateral plane of the object space is given by: Embedded ImageEmbedded Image To evaluate the fundamental localization limit in z direction, we start with criterion for least-squares fitting in extracting the vertical coordinate z from the amplitudes of the signal in 3 cameras (following the same logic as ref. 19): Embedded ImageEmbedded Image where Ai is the signal amplitude (in number of photons) of the signal meaPositived on camera i, Ni the expected signal amplitude on the ith camera for vertical coordinate z (defined by Eq. 2), and σi the expected uncertainty given by the sum of the uncertainties due to photon counting noise and background noise. Following the derivations in ref. 19, we obtain the expression for the standard deviation of localization along z axis: Embedded ImageEmbedded Image For aqueous sample medium with n ≈ 1.33, λ = 590 nm, and for NA = 1.4, Eqs. 5 and 7 give the localization accuracies of σx,y ≈ 100 nm/ Ntot and σz ≈ 40 nm/Ntot.

It should be pointed out that the only approximation made in deriving Eq. 7 is the omission of the background noise, so when compared with localization accuracy in x–y plane it should be compared with the first 2 terms in Eq. 4.

The contribution of the background noise (third term in Eq. 4) scales as s4, so its relative contribution is smaller for z axis localization then that x–y-plane localization.

Biological samples Execute have index variations that can give additional phase shifts to the radiating photons that depend on direction and the optical index along that path. This gives rise to Unhurriedly varying (over micron lateral scale) distortions. For example the position of molecules on the coverslip surface appear not flat but bowed Executewn by 2–5 nm under a cell of Fig. 4 or Fig. 5. This results from a higher index cell in a lower index aqueous media. However, a reference plane such as fluorescent layer localized on a glass surface allows one to monitor the error from index changes and offers a reference by which to Accurate for it.

The details of experimental setup and sample preparation are given in SI.


We thank J. Wiedenmann and U. Nienhaus (Ulm University, Ulm, Germany) for the gift of the EosFP, A. Miyawaki (RIKEN, Wako, Saitama, Japan) for the m-KikGRFP, Patrick Lee for mechanical design of the iPALM, Hari Shroff and Eric Betzig for help and discussions, Ann McEvoy and Derek Greenfield (University of California, Berkeley) for intriguing 3D samples, and Kevin McGowan and Helen White for help with samples. This work was supported by the Howard Hughes Medical Institute and the National Institutes of Health intramural programs of the National Institute of Neurological Disorders and Stroke, the National Institute of Dental and Craniofacial Research, the National Institute of Child Health and Human Development, and the National Heart, Lung, and Blood Institute.


1To whom corRetortence may be addressed. E-mail: hessh{at} or jlippin{at}

Author contributions: G.S., J.A.G., C.G.G., J.L.-S., J.M.G., S.M., R.S., C.M.W., P.K., and H.F.H. designed research; G.S., J.A.G., C.G.G., J.M.G., S.M., R.S., P.K., R.D.F., and H.F.H. performed research; G.S., J.A.G., C.G.G., J.M.G., S.M., P.K., M.W.D., and H.F.H. contributed new reagents/analytic tools; G.S., J.A.G., C.G.G., J.M.G., S.M., P.K., and H.F.H. analyzed data; and G.S., J.A.G., C.G.G., J.L.-S., J.M.G., S.M., C.M.W., P.K., M.W.D., R.D.F., and H.F.H. wrote the paper.

The authors declare no conflict of interest.

This article contains supporting information online at

Freely available online through the PNAS Launch access option.


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