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Optics Express

Optics Express

  • Editor: C. Martijn de Sterke
  • Vol. 20, Iss. 19 — Sep. 10, 2012
  • pp: 20998–21009
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Adaptive optics enables 3D STED microscopy in aberrating specimens

Travis J. Gould, Daniel Burke, Joerg Bewersdorf, and Martin J. Booth  »View Author Affiliations


Optics Express, Vol. 20, Issue 19, pp. 20998-21009 (2012)
http://dx.doi.org/10.1364/OE.20.020998


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Abstract

Stimulated emission depletion (STED) microscopy allows fluorescence far-field imaging with diffraction-unlimited resolution. Unfortunately, extending this technique to three-dimensional (3D) imaging of thick specimens has been inhibited by sample-induced aberrations. Here we present the first implementation of adaptive optics in STED microscopy to allow 3D super-resolution imaging in strongly aberrated imaging conditions, such as those introduced by thick biological tissue.

© 2012 OSA

1. Introduction

The recent development of sub-diffraction microscopy – or nanoscopy – is revolutionizing fluorescence imaging [1

1. D. Toomre and J. Bewersdorf, “A new wave of cellular imaging,” Annu. Rev. Cell Dev. Biol. 26(1), 285–314 (2010). [CrossRef] [PubMed]

]. Through either targeted or stochastic switching of fluorophores, resolution on the order of tens of nanometers or better is now achievable [2

2. S. W. Hell, “Microscopy and its focal switch,” Nat. Methods 6(1), 24–32 (2009). [CrossRef] [PubMed]

]. In thick samples, the targeted switching approach of stimulated emission depletion (STED) microscopy [3

3. S. W. Hell and J. Wichmann, “Breaking the diffraction resolution limit by stimulated emission: stimulated-emission-depletion fluorescence microscopy,” Opt. Lett. 19(11), 780–782 (1994). [CrossRef] [PubMed]

] has the advantage over widefield-based stochastic single-molecule switching techniques [4

4. E. Betzig, G. H. Patterson, R. Sougrat, O. W. Lindwasser, S. Olenych, J. S. Bonifacino, M. W. Davidson, J. Lippincott-Schwartz, and H. F. Hess, “Imaging intracellular fluorescent proteins at nanometer resolution,” Science 313(5793), 1642–1645 (2006). [CrossRef] [PubMed]

6

6. M. J. Rust, M. Bates, and X. Zhuang, “Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM),” Nat. Methods 3(10), 793–796 (2006). [CrossRef] [PubMed]

] that background fluorescence is suppressed through the inherent optical sectioning of a confocal laser scanning geometry. After initial successes in imaging thin samples such as single cells, STED microscopy has successfully been demonstrated to image tissue sections [7

7. J. B. Ding, K. T. Takasaki, and B. L. Sabatini, “Supraresolution imaging in brain slices using stimulated-emission depletion two-photon laser scanning microscopy,” Neuron 63(4), 429–437 (2009). [CrossRef] [PubMed]

,8

8. N. T. Urban, K. I. Willig, S. W. Hell, and U. V. Nägerl, “STED Nanoscopy of Actin Dynamics in Synapses Deep Inside Living Brain Slices,” Biophys. J. 101(5), 1277–1284 (2011). [CrossRef] [PubMed]

], nematodes [9

9. B. R. Rankin, G. Moneron, C. A. Wurm, J. C. Nelson, A. Walter, D. Schwarzer, J. Schroeder, D. A. Colón-Ramos, and S. W. Hell, “Nanoscopy in a living multicellular organism expressing GFP,” Biophys. J. 100(12), L63–L65 (2011). [CrossRef] [PubMed]

], and even living mice [10

10. S. Berning, K. I. Willig, H. Steffens, P. Dibaj, and S. W. Hell, “Nanoscopy in a living mouse brain,” Science 335(6068), 551 (2012). [CrossRef] [PubMed]

].

For thin samples, two 3D STED methods have been demonstrated thus far. The highest resolution has been achieved with a complex setup utilizing two opposing objective lenses [12

12. R. Schmidt, C. A. Wurm, S. Jakobs, J. Engelhardt, A. Egner, and S. W. Hell, “Spherical nanosized focal spot unravels the interior of cells,” Nat. Methods 5(6), 539–544 (2008). [CrossRef] [PubMed]

] which is not compatible with thick sample geometries. Alternatively, using a single objective, an annular phase filter featuring a central circle with a phase step of π radians (λ/2) has been applied [13

13. T. A. Klar, S. Jakobs, M. Dyba, A. Egner, and S. W. Hell, “Fluorescence microscopy with diffraction resolution barrier broken by stimulated emission,” Proc. Natl. Acad. Sci. U.S.A. 97(15), 8206–8210 (2000). [CrossRef] [PubMed]

]. This phase mask creates a ring-shaped focus with additional high-intensity lobes above and below the central minimum and thereby allows suppression of fluorescence emission in all directions around the center. Unfortunately, this approach is far more susceptible to aberrations than the helicoidal phase mask approach [11

11. S. Deng, L. Liu, Y. Cheng, R. Li, and Z. Xu, “Investigation of the influence of the aberration induced by a plane interface on STED microscopy,” Opt. Express 17(3), 1714–1725 (2009). [CrossRef] [PubMed]

] and, not surprisingly, no 3D STED imaging has been demonstrated to date in thick samples.

To realize 3D STED microscopy with the central λ/2 phase filter deep in samples, two approaches are conceivable. First, the tissue could be ‘cleared’ by embedding it in a refractive index-matching medium, but this approach is not compatible with living specimens. Second, adaptive optics (AO) could be employed. Adaptive elements such as deformable mirrors or spatial light modulators have been successfully implemented in a variety of microscopes to correct specimen-induced aberrations [14

14. M. J. Booth, M. A. Neil, R. Juskaitis, and T. Wilson, “Adaptive aberration correction in a confocal microscope,” Proc. Natl. Acad. Sci. U.S.A. 99(9), 5788–5792 (2002). [CrossRef] [PubMed]

17

17. X. D. Tao, O. Azucena, M. Fu, Y. Zuo, D. C. Chen, and J. Kubby, “Adaptive optics microscopy with direct wavefront sensing using fluorescent protein guide stars,” Opt. Lett. 36(17), 3389–3391 (2011). [CrossRef] [PubMed]

]. As aberrations are not predictable, but rather vary across different regions and depths in the specimen, the key to a successful implementation of AO is the use of feedback from the measured microscope focus quality into the aberration correction procedure. This approach distinguishes adaptive aberration correction from simple static correction methods, such as adjustment of the objective lens correction collar [8

8. N. T. Urban, K. I. Willig, S. W. Hell, and U. V. Nägerl, “STED Nanoscopy of Actin Dynamics in Synapses Deep Inside Living Brain Slices,” Biophys. J. 101(5), 1277–1284 (2011). [CrossRef] [PubMed]

]. These static methods cannot correct the complex aberrations that are actually encountered in biological microscopy [18

18. M. Schwertner, M. J. Booth, and T. Wilson, “Characterizing specimen induced aberrations for high NA adaptive optical microscopy,” Opt. Express 12(26), 6540–6552 (2004). [CrossRef] [PubMed]

]. Here we demonstrate the first implementation of AO in 3D STED microscopy using a novel image quality feedback metric and demonstrate resolution well below the diffraction limit in all three dimensions when imaging through optically aberrating specimens such as tissue.

2. Setup

In order to achieve 3D super-resolution, we adapted a custom-built STED microscope described previously [19

19. T. J. Gould, J. R. Myers, and J. Bewersdorf, “Total internal reflection STED microscopy,” Opt. Express 19(14), 13351–13357 (2011). [CrossRef] [PubMed]

] by implementing a central λ/2 phase mask in the depletion beam path. Figure 1
Fig. 1 Schematic of AO STED setup. FI - Faraday isolator; GLP - Glan laser polarizer; GB - glass block; DS - delay stage; AOM - acousto-optical modulator; PMF - polarization-maintaining fiber; SLM1/2 - spatial light modulators; PC - photonic crystal fiber; AOTF - acousto-optical tunable filter; PBS - polarizing beam splitter cube; λ/2 - half-wave plate; λ/4 - quarter-wave plate; DM1/2 - dichroic mirrors; F - bandpass filter; MMF - multimode fiber; APD - avalanche photodiode; OBJ - objective lens; xyz - 3 axes piezo sample stage. Insets show typical phase patterns for each SLM where phase contributions include a baseline flatness correction for the SLM (provided by the manufacturer), a circular blazed grating (for off-axis phase modulation) that defines the active area, correction for system-induced aberration, and a central λ/2 phase mask (SLM1 only).
illustrates the AO STED setup. The output from an 80 MHz mode-locked Ti:Sapphire laser (Chameleon Ultra II, Coherent) was passed through a Faraday isolator (Newport) and a half-wave plate before being split into two beam paths by a Glan laser polarizer (Newport). The transmitted beam was used for STED illumination. This beam was passed through a 19 cm long glass block, a delay stage for pulse delay adjustment, and an acousto-optical modulator (AA Optoelectronics) for laser power adjustment before it was coupled into a 100 m long polarization-maintaining single mode fiber (Fibercore). The glass block and single mode fiber served to stretch the pulses to a few hundred picoseconds. To generate synchronized excitation pulses, the beam reflected at the Glan laser polarizer was focused into a photonic crystal fiber (SCG-800, Newport). The white light spectrum output from this fiber was passed through an acousto-optical tunable filter (AOTF; AA Optoelectronics) for wavelength selection and power control and then coupled into a second polarization-maintaining single mode fiber (Thorlabs). At the output of each single mode fiber, the beams were collimated and reflected off of a liquid crystal spatial light modulator (SLM) placed in a plane conjugate to the objective pupil. Half- and quarter-wave plates were used to produce circular polarization of each beam in the sample. Additionally, a polarizing beam splitter cube was used in the STED beam path to clean up the beam polarization. The beams were then merged to a common path by a dichroic mirror before being focused by a 100x/1.4NA oil immersion objective lens (UPLSAPO 100XO/PSF, Olympus) mounted in a commercial microscope stand (IX71, Olympus). Samples were mounted to an xyz piezo stage (PINano, Physik Instrumente) for scanning. The fluorescence signal was collected by the objective, separated from laser light by the dichroic mirrors (FF01-740-Di01 and FF662-FDi01, Semrock), filtered by two bandpass filters (FF01-685/40, Semrock), and focused into a 62.5 µm core diameter (~0.64 Airy units) 50:50 signal-splitting multimode fiber (OZ Optics). Each output of the multimode fiber was attached to a single photon counting avalanche photodiode (ARQ-13-FC, Perkin Elmer). Image acquisition and instrument control was achieved using custom software written in Labview (National Instruments).

For imaging, the Ti:Sapphire laser was tuned to 770 nm for depletion and an excitation line at 633 nm was selected from the output of the photonic crystal fiber using the AOTF. Laser powers measured at the objective back aperture were 84-132 mW and 0.2-2 µW for depletion and excitation, respectively. Images (128x128 pixels) were acquired with a scan rate of 100 lines/s, a pixel size of 40 nm, and 1-3 frame accumulations (corresponding to 1.28-3.84 s/image). Lines were scanned unidirectionally using a sinusoidal velocity profile which resulted in a pixel dwell time of 21.5 µs at the scan center and approximately twice this value at the edges. Recorded pixel values were therefore normalized according the pixel dwell times such that the center pixel was divided by unity. Fitting of line profiles was performed using the software Imspector (Andreas Schönle, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany). Fits were performed either using a Lorentzian (for images taken after aberration correction) or a Gaussian function (for images taken without aberration correction since in this case the observed data did not fit a Lorentzian profile). 3D volume renderings were generated using the software Volocity (Perkin Elmer).

The STED phase mask as well as aberration correction were realized using the SLM (Hamamatsu X10468-02) placed in the depletion beam path (SLM1 in Fig. 1). The 800x600 pixels of the SLM can be addressed individually by connecting the SLM as an additional display device to the instrument PC. Phase modulation from 0 to 2π radians (and more) at the wavelengths used is achieved by varying the gray scale of the phase image fed to the SLM. Phase modulations significantly larger than 2π radians can be produced through phase-wrapping, whereby larger phase values are translated into the range 0 to 2π radians. Although aberrations affect the illumination, emission and depletion beam paths, the achievable resolution is dominated by the quality of the depletion focus. For this reason, aberration correction by the SLM solely in the depletion path is sufficient to significantly improve the image quality. Aberrations in the illumination and emission paths, however, influence the level of fluorescence excitation (by reducing the illumination intensity in the focus), the detection efficiency (by blurring the fluorescence focus on the confocal pinhole) and the background signal (through excitation and detection of fluorescence outside of the depletion region). Therefore, we included a second SLM (Hamamatsu X10468-01) to perform additional aberration correction in the excitation beam path (SLM2 in Fig. 1). Further improvement could in principle be attained with aberration correction in the emission path. However, the polarization dependence of the SLM limits its suitability for correction of fluorescence emission.

SLMs have been used previously in STED microscopy in a direct reflection configuration [20

20. G. Donnert, J. Keller, R. Medda, M. A. Andrei, S. O. Rizzoli, R. Lührmann, R. Jahn, C. Eggeling, and S. W. Hell, “Macromolecular-scale resolution in biological fluorescence microscopy,” Proc. Natl. Acad. Sci. U.S.A. 103(31), 11440–11445 (2006). [CrossRef] [PubMed]

,21

21. K. I. Willig, R. R. Kellner, R. Medda, B. Hein, S. Jakobs, and S. W. Hell, “Nanoscale resolution in GFP-based microscopy,” Nat. Methods 3(9), 721–723 (2006). [CrossRef] [PubMed]

]. However, this implementation requires near-perfect phase modulation of the light by the SLM. Instead, here we use an off-axis hologram which is beneficial because it separates the phase-modulated light from any directly reflected, unmodulated light [22

22. E. Auksorius, B. R. Boruah, C. Dunsby, P. M. Lanigan, G. Kennedy, M. A. Neil, and P. M. French, “Stimulated emission depletion microscopy with a supercontinuum source and fluorescence lifetime imaging,” Opt. Lett. 33(2), 113–115 (2008). [CrossRef] [PubMed]

]. In our setup an active pupil was defined on the SLM by creating such an off-axis hologram (Fig. 1 insets) – a blazed grating of circular outline that diffracts light off the principal reflection axis into the first diffraction order – and the diffracted light was focused into the sample. The remaining small non-diffracted fraction of the incident light (as well as light reflected from parts outside the active pupil) were simply reflected along the principal reflection axis and formed a secondary focus in the sample, much weaker than the depletion spot, approximately 3 µm away from the primary focus. This approach ensured that any residual light stemming from imperfect phase modulation could not interfere with the desired depletion focus and reduce its quality. We note that the non-diffracted light could also be blocked at an intermediate focal plane if necessary. Phase masks for creating the depletion focus were produced by adding the corresponding function to the SLM phase pattern. In this manner, phase masks can be added, modified and exchanged with considerable ease, compared to systems where manufactured phase plates are physically placed in the optical path.

The microscope employed an oil immersion lens of numerical aperture (NA) 1.4. Specimens were mounted in either aqueous media (phosphate buffered saline, PBS) of refractive index n = 1.34 or glycerol (n = 1.47). For the aqueous mounting media, the limiting system NA is restricted to the refractive index of the mounting medium due to total internal reflection at higher angles of incidence of the focusing beam. We therefore restricted the active aperture defined on the SLM to a size corresponding to an objective NA of 1.325. This enabled us, in effect, to mimic the performance of a water immersion objective lens of NA = 1.325, which is higher than commercially available objectives.

3. Aberration correction

The SLM enables adaptive aberration correction through the addition of further phase functions to the SLM [22

22. E. Auksorius, B. R. Boruah, C. Dunsby, P. M. Lanigan, G. Kennedy, M. A. Neil, and P. M. French, “Stimulated emission depletion microscopy with a supercontinuum source and fluorescence lifetime imaging,” Opt. Lett. 33(2), 113–115 (2008). [CrossRef] [PubMed]

]. In the system described here we employed a sensorless AO scheme to determine the aberrations induced by the system and specimen [23

23. M. Booth, “Wave front sensor-less adaptive optics: a model-based approach using sphere packings,” Opt. Express 14(4), 1339–1352 (2006). [CrossRef] [PubMed]

25

25. A. Facomprez, E. Beaurepaire, and D. Débarre, “Accuracy of correction in modal sensorless adaptive optics,” Opt. Express 20(3), 2598–2612 (2012). [CrossRef] [PubMed]

]. In sensorless AO, aberrations are estimated through analysis of a set of images that is obtained using a sequence of predetermined bias aberrations. This process requires 1) expansion of the aberrations using an appropriate set of modes, and 2) definition of an image quality metric that is maximized to find the optimal aberration correction.

We chose to model aberrations using a Zernike polynomial expansion modified to remove small image shifts introduced by displacement (tip, tilt, and defocus) modes [25

25. A. Facomprez, E. Beaurepaire, and D. Débarre, “Accuracy of correction in modal sensorless adaptive optics,” Opt. Express 20(3), 2598–2612 (2012). [CrossRef] [PubMed]

,26

26. A. Thayil and M. J. Booth, “Self calibration of sensorless adaptive optical microscopes,” J. Eur. Opt. Soc. 6, 11045 (2011). [CrossRef]

]. Table 1

Table 1. Definition of Zernike Modes, Zi

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lists the Zernike modes (following the numbering convention used by Neil et al. [27

27. M. A. Neil, M. J. Booth, and T. Wilson, “New modal wave-front sensor: a theoretical analysis,” J. Opt. Soc. Am. A 17(6), 1098–1107 (2000). [CrossRef] [PubMed]

]) used for aberration correction in this work. Zernike polynomial modes are widely used to model aberrations in adaptive optics. One reason for this is the orthogonality between the modes, which has useful practical consequences, such as the separation of displacement of the focus (by tip, tilt and defocus modes) and distortion of the focus (through higher order modes such as astigmatism, coma and spherical aberration). However, this separation of effects is only strictly true in the paraxial approximation and with a uniform pupil function. When using phase masks and high NA lenses in STED microscopy, these approximations are no longer valid. We observed experimentally that the zero intensity point of the depletion focus can shift laterally when, for example, Zernike coma is applied using the SLM. Similar axial shifts were observed when applying Zernike spherical modes. This behavior has been confirmed through theoretical modeling (data not shown). In order to remove these displacement effects, we measured the induced focal shifts for each Zernike mode using images of gold beads. Using a linear approximation of the zero shift, a proportionate amount of the tip, tilt, or defocus mode (as appropriate) was added to each applied Zernike mode to ensure that the intensity zero did not shift position. This procedure defined a modified set of “displacement-free” Zernike modes that was used for the subsequent experiments. Table 2

Table 2. Couplings Between Displacement and Aberration Zernike Modes

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lists the coupling between displacement and aberration modes used to generate displacement-free Zernike modes. Subsequent references to Zernike modes will hereafter represent displacement-free Zernike modes.

In practice, the characteristics that we desire from a STED image are high brightness combined with high resolution. We have therefore introduced a new metric that seeks to optimize both image brightness and image sharpness in a combined approach, which succeeds where either separate approach can fail. The combined metric was defined as
M=S+σβB[11+ek(SST)]
(1)
where B and S are the brightness and sharpness metrics, respectively. The expression in brackets in Eq. (1) is a logistic function that, in effect, introduces the contribution of the brightness metric B only when the sharpness metric S is above a particular threshold ST (typically chosen to be 90% of the peak sharpness). The parameter β was chosen empirically to balance the contributions from S and B and the parameter k was used to adjust the rate of transition of the logistic function in the region of SST. The constant σ was + 1 or −1 dependent upon whether the aberration mode in question produced a minimum or maximum in B for zero aberration magnitude.

The image brightness metric B was calculated as the sum of the pixel values in the confocal or STED image. The sharpness metric S was defined as the second moment of the image Fourier transform (FT) and calculated as
S=n,mμn,mI^n,m(n2+m2)/n,mI^n,m
(2)
where I^n,m is the discrete FT of the image,n=nN12, m=mM12,and n and m are the coordinates ranging from 0 to N-1 or M-1, respectively. The functionμn,m is a circular mask defined by
μn,m={1n2+m2w0n2+m2>w
(3)
where w is a radius (in units of pixels of the image FT) defined as the size of the field of view divided by an upper bound on the expected resolution (assumed to be 200 nm). This mask is used to cut off higher spatial frequencies that would otherwise cause S to be dominated by noise. The motivation for this choice of metric lies in the inverse relationship between the width of image features and the width of the corresponding image’s spatial frequency spectrum – sharp images lead to a broad image FT. As the second moment calculation of Eq. (2) is a measure of width of the image FT, it follows that a large value of S corresponds to a sharp image arising from a narrow PSF. Figure 2 shows comparisons of the three metrics B, S and M for selected Zernike modes for STED imaging (using a λ/2 phase mask) of 200 nm crimson beads (F8806, Invitrogen) attached to a coverglass and mounted in glycerol. Images were acquired in the xy (focal) plane. We demonstrate here the suitability of the metric M using images of fluorescent beads, however, the definition of M should mean that it is readily applicable to any sample that features structures smaller than the diffraction limit.

It is important to note that the 2nd and 3rd spherical modes lead to decreases in image brightness while the 1st spherical mode leads to a brightness increase when imaging with the central λ/2 phase mask (Fig. 2). These results reveal that manual adjustment of the objective correction collar to maximize image brightness cannot necessarily be applied to 3D STED microscopy as the collar adjusts a combination of these spherical aberration modes.

For our sensorless adaptive optics scheme, images (single accumulation) were acquired in either confocal (for excitation path correction) or STED (for depletion path correction) mode while adding a bias aberration, bZi, of a given mode Zi where b was a suitably chosen bias amplitude. Typically, b was chosen to cover the range of −1 to + 1 radian while acquiring a total of 5 to 7 images for each Zi (corresponding to ~6.4-9 s for each iteration of correction for each mode). For each image the appropriate metric was calculated (B for excitation path correction; M for depletion path correction) and then plotted as a function of the bias amplitude b as shown in Fig. 2. The correction aberration, acorr, was then estimated as the peak of the curve, which was calculated by fitting a quadratic function to the data. In cases where a quadratic approximation did not represent the data, peaks could be identified manually. The aberration acorrZi was then added to the SLM phase pattern, and this measurement and correction cycle was then repeated for each of the modes of interest. Aberrations due to coma, astigmatism, and trefoil could be assessed using a single iteration of our correction routine. Spherical modes were typically the dominant aberrations in the experiments conducted here and typically required 1-4 correction iterations to converge to optimal values. However, when results for correcting the excitation beam path were used as a starting point for correction of the depletion beam path 1-2 iterations were typically sufficient to assess the spherical aberrations. All AO STED images shown are of the same field of view used to perform aberration correction.

4. AO STED microscopy

As an initial implementation, system aberrations were corrected using images of fluorescent beads. For this purpose, a fluorescent bead sample was prepared by attaching 100 nm diameter crimson beads (C47248, Invitrogen) to a coverglass coated with poly-L-lysine and mounting the coverglass to a microscope slide with glycerol as the mounting medium. Optimization was performed first for the excitation beam path using the brightness metric (in confocal imaging mode) and then for the STED beam path using the combined metric defined in Eq. (1) (in STED imaging mode). Figure 3
Fig. 3 Adaptive correction of residual system aberrations. (A, B) STED PSF using the central λ/2 phase mask before (A) and after (B) correction for system aberrations with corresponding phase patterns used on SLM2. (C, D) xz STED images of 100 nm fluorescent bead attached to coverglass and imaged (C) before and (D) after correction of system aberrations. (E) Axial line profiles of pixels summed across horizontal dimension of dashed boxes in (C) and (D). Fitted curves give FWHM of 280 nm (Gaussian fit) for the uncorrected STED image and 140 nm (Lorentzian fit) when adaptive optics is used to compensate for system-induced aberrations in the STED beam path.
shows xz-images of fluorescent beads before and after aberration correction. It can be clearly seen that the aberrated depletion focus (PSF shown in Fig. 3(a)) had a detrimental effect on both the image intensity and resolution. By correcting system aberrations, the axial STED resolution could be improved by a factor of 2 from ~280 nm to ~140 nm while at the same time the signal intensity increased by a factor of ~1.6. This quality improvement can also be seen when measuring the depletion focus directly: Figs. 3(a) and 3(b) show the intensity distribution of the depletion focus measured in reflection mode by scanning a 150 nm diameter gold bead through the focal region without and with the obtained aberration correction, respectively. As shown in this figure, aberration correction leads to a far more symmetric depletion focus with a more pronounced intensity minimum.

To further test the correction procedure using aberrations encountered in biological specimens, 200 nm crimson beads were added on top of zebrafish retina sections attached to #1.5 coverglass and then mounted in PBS (to mimic the aqueous embedding conditions of live sample imaging) onto a microscope slide. Figures 5(a)
Fig. 5 AO STED images of fluorescent beads through zebrafish retina sections. (A-F) Results for beads imaged though ~14 µm of retina. Lateral and axial sections of a single fluorescent bead imaged in (A, D) confocal, (B, E) STED, and (C, F) AO STED show improvement in signal and resolution when adaptive aberration correction is applied to the depletion beam path. (G-L) Similar image sequences for beads imaged through ~25 µm of retina. Axial profiles of beads in AO STED images were ~208 nm and ~249 nm for (F) and (L), respectively. Color bar in (E) also applies to (B), (C), and (F). Color bar in (K) also applies to (H), (I), and (L). (M-O) Volume renderings for data shown in (A-F) for (M) confocal, (N) STED, and (O) AO STED data. (N) and (O) plotted on same color scale for comparison of signal.
-5(l) show xy and xz images of an individual bead after focusing ~14 µm or ~25 µm through the retina sections. In this application all aberration correction was performed solely on the STED beam path to demonstrate that correction of the depletion beam path is possible without prior knowledge of corrections made to the excitation path. These data also demonstrate that the quality of the depletion focus is the dominating factor in achieving sub-diffraction resolution in the STED images. While correction of relatively few aberration modes was sufficient to obtain ~200 nm and ~250 nm axial resolution for imaging depths of ~14 µm and ~25 µm, respectively, we expect that including higher order aberrations in the correction routine should further improve the STED resolution. Figures 5(m)-5(o) show volume renderings of the data depicted in Figs. 5(a)-5(f) and represent the first 3D super-resolution STED image recorded through scattering tissue.

We note that the STED images shown in Figs. 3-5 before and after aberration correction are of the same beads on which the correction routine was performed, thus demonstrating that photobleaching was not significant in these experiments. Labeling conditions in biological imaging applications are likely to use less photostable fluorophores, which in some instances may not tolerate the acquisition of 5-7 STED images per aberration mode. However, it has been demonstrated that only 2N + 1 images are required to correct for N aberration modes [25

25. A. Facomprez, E. Beaurepaire, and D. Débarre, “Accuracy of correction in modal sensorless adaptive optics,” Opt. Express 20(3), 2598–2612 (2012). [CrossRef] [PubMed]

] and we expect this approach to be compatible with methods presented here. Furthermore, applications otherwise inhibited by photobleaching may benefit from the implementation of triplet-state-relaxation during STED imaging [20

20. G. Donnert, J. Keller, R. Medda, M. A. Andrei, S. O. Rizzoli, R. Lührmann, R. Jahn, C. Eggeling, and S. W. Hell, “Macromolecular-scale resolution in biological fluorescence microscopy,” Proc. Natl. Acad. Sci. U.S.A. 103(31), 11440–11445 (2006). [CrossRef] [PubMed]

] either by increasing scan rates or using lower repetition rate lasers.

5. Summary

The SLM provides a convenient way to combine a STED phase mask and adaptive aberration correction in the same microscope. The alternative approach of using deformable mirrors would have the advantage of being able to correct all three (depletion, excitation, and emission) beam paths with the same device. However, unlike the SLM, the continuous reflective surface of mirror devices does not permit the generation of the STED phase masks, which require discontinuous phase jumps.

Aberrations were corrected using a sensorless AO scheme in which aberration modes were sequentially corrected through a process of image optimization. The definition of a new image quality metric, combining both image sharpness and brightness, was necessary to account for the way in which aberrations affect STED images. We have implemented the correction using images of beads through tissue specimens. We expect, however, that our correction scheme is applicable to any arrangement of fine scale objects.

The implementation of AO in STED microscopy is a critical step towards achieving 3D super-resolution inside strongly aberrating samples. This is especially important when imaging living samples where matching the refractive indices of immersion liquids and mounting media is not practical. In addition to the demonstrations of superresolution STED microscopy in tissue, there have recently been applications of stochastic switching-based nanoscopy approaches to the imaging of multi-cellular structures [28

28. F. Cella Zanacchi, Z. Lavagnino, M. Perrone Donnorso, A. Del Bue, L. Furia, M. Faretta, and A. Diaspro, “Live-cell 3D super-resolution imaging in thick biological samples,” Nat. Methods 8(12), 1047–1049 (2011). [CrossRef] [PubMed]

]. AO has also recently been used in this branch of nanoscopy to correct for instrument-induced aberrations [29

29. I. Izeddin, M. El Beheiry, J. Andilla, D. Ciepielewski, X. Darzacq, and M. Dahan, “PSF shaping using adaptive optics for three-dimensional single-molecule super-resolution imaging and tracking,” Opt. Express 20(5), 4957–4967 (2012). [CrossRef] [PubMed]

]. We expect that AO will be instrumental in extending correction of sample-induced aberrations to all superresolution modalities using an approach similar to that presented here. This implementation of AO has the potential to advance 3D diffraction-unlimited microscopy to a new imaging frontier in (living) tissue and intact specimens.

Acknowledgments

We thank Caixia Lv and David Zenisek for the preparation of the zebrafish tissue sections and Manuel Juette and Lena Schroeder for helpful discussions. This work was supported by the Wellcome Trust (095927/A/11/Z, 095927/B/11/Z) and the Kavli Foundation. T.J.G. was supported by the National Institute of General Medical Sciences (F32GM096859). M.J.B. was supported by the Engineering and Physical Sciences Research Council, UK (EP/E055818/1). JB discloses significant financial interest in Vutara, Inc.

References and links

1.

D. Toomre and J. Bewersdorf, “A new wave of cellular imaging,” Annu. Rev. Cell Dev. Biol. 26(1), 285–314 (2010). [CrossRef] [PubMed]

2.

S. W. Hell, “Microscopy and its focal switch,” Nat. Methods 6(1), 24–32 (2009). [CrossRef] [PubMed]

3.

S. W. Hell and J. Wichmann, “Breaking the diffraction resolution limit by stimulated emission: stimulated-emission-depletion fluorescence microscopy,” Opt. Lett. 19(11), 780–782 (1994). [CrossRef] [PubMed]

4.

E. Betzig, G. H. Patterson, R. Sougrat, O. W. Lindwasser, S. Olenych, J. S. Bonifacino, M. W. Davidson, J. Lippincott-Schwartz, and H. F. Hess, “Imaging intracellular fluorescent proteins at nanometer resolution,” Science 313(5793), 1642–1645 (2006). [CrossRef] [PubMed]

5.

S. T. Hess, T. P. Girirajan, and M. D. Mason, “Ultra-high resolution imaging by fluorescence photoactivation localization microscopy,” Biophys. J. 91(11), 4258–4272 (2006). [CrossRef] [PubMed]

6.

M. J. Rust, M. Bates, and X. Zhuang, “Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM),” Nat. Methods 3(10), 793–796 (2006). [CrossRef] [PubMed]

7.

J. B. Ding, K. T. Takasaki, and B. L. Sabatini, “Supraresolution imaging in brain slices using stimulated-emission depletion two-photon laser scanning microscopy,” Neuron 63(4), 429–437 (2009). [CrossRef] [PubMed]

8.

N. T. Urban, K. I. Willig, S. W. Hell, and U. V. Nägerl, “STED Nanoscopy of Actin Dynamics in Synapses Deep Inside Living Brain Slices,” Biophys. J. 101(5), 1277–1284 (2011). [CrossRef] [PubMed]

9.

B. R. Rankin, G. Moneron, C. A. Wurm, J. C. Nelson, A. Walter, D. Schwarzer, J. Schroeder, D. A. Colón-Ramos, and S. W. Hell, “Nanoscopy in a living multicellular organism expressing GFP,” Biophys. J. 100(12), L63–L65 (2011). [CrossRef] [PubMed]

10.

S. Berning, K. I. Willig, H. Steffens, P. Dibaj, and S. W. Hell, “Nanoscopy in a living mouse brain,” Science 335(6068), 551 (2012). [CrossRef] [PubMed]

11.

S. Deng, L. Liu, Y. Cheng, R. Li, and Z. Xu, “Investigation of the influence of the aberration induced by a plane interface on STED microscopy,” Opt. Express 17(3), 1714–1725 (2009). [CrossRef] [PubMed]

12.

R. Schmidt, C. A. Wurm, S. Jakobs, J. Engelhardt, A. Egner, and S. W. Hell, “Spherical nanosized focal spot unravels the interior of cells,” Nat. Methods 5(6), 539–544 (2008). [CrossRef] [PubMed]

13.

T. A. Klar, S. Jakobs, M. Dyba, A. Egner, and S. W. Hell, “Fluorescence microscopy with diffraction resolution barrier broken by stimulated emission,” Proc. Natl. Acad. Sci. U.S.A. 97(15), 8206–8210 (2000). [CrossRef] [PubMed]

14.

M. J. Booth, M. A. Neil, R. Juskaitis, and T. Wilson, “Adaptive aberration correction in a confocal microscope,” Proc. Natl. Acad. Sci. U.S.A. 99(9), 5788–5792 (2002). [CrossRef] [PubMed]

15.

N. Ji, D. E. Milkie, and E. Betzig, “Adaptive optics via pupil segmentation for high-resolution imaging in biological tissues,” Nat. Methods 7(2), 141–147 (2010). [CrossRef] [PubMed]

16.

M. Rueckel, J. A. Mack-Bucher, and W. Denk, “Adaptive wavefront correction in two-photon microscopy using coherence-gated wavefront sensing,” Proc. Natl. Acad. Sci. U.S.A. 103(46), 17137–17142 (2006). [CrossRef] [PubMed]

17.

X. D. Tao, O. Azucena, M. Fu, Y. Zuo, D. C. Chen, and J. Kubby, “Adaptive optics microscopy with direct wavefront sensing using fluorescent protein guide stars,” Opt. Lett. 36(17), 3389–3391 (2011). [CrossRef] [PubMed]

18.

M. Schwertner, M. J. Booth, and T. Wilson, “Characterizing specimen induced aberrations for high NA adaptive optical microscopy,” Opt. Express 12(26), 6540–6552 (2004). [CrossRef] [PubMed]

19.

T. J. Gould, J. R. Myers, and J. Bewersdorf, “Total internal reflection STED microscopy,” Opt. Express 19(14), 13351–13357 (2011). [CrossRef] [PubMed]

20.

G. Donnert, J. Keller, R. Medda, M. A. Andrei, S. O. Rizzoli, R. Lührmann, R. Jahn, C. Eggeling, and S. W. Hell, “Macromolecular-scale resolution in biological fluorescence microscopy,” Proc. Natl. Acad. Sci. U.S.A. 103(31), 11440–11445 (2006). [CrossRef] [PubMed]

21.

K. I. Willig, R. R. Kellner, R. Medda, B. Hein, S. Jakobs, and S. W. Hell, “Nanoscale resolution in GFP-based microscopy,” Nat. Methods 3(9), 721–723 (2006). [CrossRef] [PubMed]

22.

E. Auksorius, B. R. Boruah, C. Dunsby, P. M. Lanigan, G. Kennedy, M. A. Neil, and P. M. French, “Stimulated emission depletion microscopy with a supercontinuum source and fluorescence lifetime imaging,” Opt. Lett. 33(2), 113–115 (2008). [CrossRef] [PubMed]

23.

M. Booth, “Wave front sensor-less adaptive optics: a model-based approach using sphere packings,” Opt. Express 14(4), 1339–1352 (2006). [CrossRef] [PubMed]

24.

D. Débarre, E. J. Botcherby, T. Watanabe, S. Srinivas, M. J. Booth, and T. Wilson, “Image-based adaptive optics for two-photon microscopy,” Opt. Lett. 34(16), 2495–2497 (2009). [CrossRef] [PubMed]

25.

A. Facomprez, E. Beaurepaire, and D. Débarre, “Accuracy of correction in modal sensorless adaptive optics,” Opt. Express 20(3), 2598–2612 (2012). [CrossRef] [PubMed]

26.

A. Thayil and M. J. Booth, “Self calibration of sensorless adaptive optical microscopes,” J. Eur. Opt. Soc. 6, 11045 (2011). [CrossRef]

27.

M. A. Neil, M. J. Booth, and T. Wilson, “New modal wave-front sensor: a theoretical analysis,” J. Opt. Soc. Am. A 17(6), 1098–1107 (2000). [CrossRef] [PubMed]

28.

F. Cella Zanacchi, Z. Lavagnino, M. Perrone Donnorso, A. Del Bue, L. Furia, M. Faretta, and A. Diaspro, “Live-cell 3D super-resolution imaging in thick biological samples,” Nat. Methods 8(12), 1047–1049 (2011). [CrossRef] [PubMed]

29.

I. Izeddin, M. El Beheiry, J. Andilla, D. Ciepielewski, X. Darzacq, and M. Dahan, “PSF shaping using adaptive optics for three-dimensional single-molecule super-resolution imaging and tracking,” Opt. Express 20(5), 4957–4967 (2012). [CrossRef] [PubMed]

OCIS Codes
(180.2520) Microscopy : Fluorescence microscopy
(180.6900) Microscopy : Three-dimensional microscopy
(350.5730) Other areas of optics : Resolution
(110.1080) Imaging systems : Active or adaptive optics

ToC Category:
Microscopy

History
Original Manuscript: June 29, 2012
Revised Manuscript: July 31, 2012
Manuscript Accepted: August 2, 2012
Published: August 29, 2012

Virtual Issues
Vol. 7, Iss. 11 Virtual Journal for Biomedical Optics

Citation
Travis J. Gould, Daniel Burke, Joerg Bewersdorf, and Martin J. Booth, "Adaptive optics enables 3D STED microscopy in aberrating specimens," Opt. Express 20, 20998-21009 (2012)
http://www.opticsinfobase.org/oe/abstract.cfm?URI=oe-20-19-20998


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References

  1. D. Toomre and J. Bewersdorf, “A new wave of cellular imaging,” Annu. Rev. Cell Dev. Biol.26(1), 285–314 (2010). [CrossRef] [PubMed]
  2. S. W. Hell, “Microscopy and its focal switch,” Nat. Methods6(1), 24–32 (2009). [CrossRef] [PubMed]
  3. S. W. Hell and J. Wichmann, “Breaking the diffraction resolution limit by stimulated emission: stimulated-emission-depletion fluorescence microscopy,” Opt. Lett.19(11), 780–782 (1994). [CrossRef] [PubMed]
  4. E. Betzig, G. H. Patterson, R. Sougrat, O. W. Lindwasser, S. Olenych, J. S. Bonifacino, M. W. Davidson, J. Lippincott-Schwartz, and H. F. Hess, “Imaging intracellular fluorescent proteins at nanometer resolution,” Science313(5793), 1642–1645 (2006). [CrossRef] [PubMed]
  5. S. T. Hess, T. P. Girirajan, and M. D. Mason, “Ultra-high resolution imaging by fluorescence photoactivation localization microscopy,” Biophys. J.91(11), 4258–4272 (2006). [CrossRef] [PubMed]
  6. M. J. Rust, M. Bates, and X. Zhuang, “Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM),” Nat. Methods3(10), 793–796 (2006). [CrossRef] [PubMed]
  7. J. B. Ding, K. T. Takasaki, and B. L. Sabatini, “Supraresolution imaging in brain slices using stimulated-emission depletion two-photon laser scanning microscopy,” Neuron63(4), 429–437 (2009). [CrossRef] [PubMed]
  8. N. T. Urban, K. I. Willig, S. W. Hell, and U. V. Nägerl, “STED Nanoscopy of Actin Dynamics in Synapses Deep Inside Living Brain Slices,” Biophys. J.101(5), 1277–1284 (2011). [CrossRef] [PubMed]
  9. B. R. Rankin, G. Moneron, C. A. Wurm, J. C. Nelson, A. Walter, D. Schwarzer, J. Schroeder, D. A. Colón-Ramos, and S. W. Hell, “Nanoscopy in a living multicellular organism expressing GFP,” Biophys. J.100(12), L63–L65 (2011). [CrossRef] [PubMed]
  10. S. Berning, K. I. Willig, H. Steffens, P. Dibaj, and S. W. Hell, “Nanoscopy in a living mouse brain,” Science335(6068), 551 (2012). [CrossRef] [PubMed]
  11. S. Deng, L. Liu, Y. Cheng, R. Li, and Z. Xu, “Investigation of the influence of the aberration induced by a plane interface on STED microscopy,” Opt. Express17(3), 1714–1725 (2009). [CrossRef] [PubMed]
  12. R. Schmidt, C. A. Wurm, S. Jakobs, J. Engelhardt, A. Egner, and S. W. Hell, “Spherical nanosized focal spot unravels the interior of cells,” Nat. Methods5(6), 539–544 (2008). [CrossRef] [PubMed]
  13. T. A. Klar, S. Jakobs, M. Dyba, A. Egner, and S. W. Hell, “Fluorescence microscopy with diffraction resolution barrier broken by stimulated emission,” Proc. Natl. Acad. Sci. U.S.A.97(15), 8206–8210 (2000). [CrossRef] [PubMed]
  14. M. J. Booth, M. A. Neil, R. Juskaitis, and T. Wilson, “Adaptive aberration correction in a confocal microscope,” Proc. Natl. Acad. Sci. U.S.A.99(9), 5788–5792 (2002). [CrossRef] [PubMed]
  15. N. Ji, D. E. Milkie, and E. Betzig, “Adaptive optics via pupil segmentation for high-resolution imaging in biological tissues,” Nat. Methods7(2), 141–147 (2010). [CrossRef] [PubMed]
  16. M. Rueckel, J. A. Mack-Bucher, and W. Denk, “Adaptive wavefront correction in two-photon microscopy using coherence-gated wavefront sensing,” Proc. Natl. Acad. Sci. U.S.A.103(46), 17137–17142 (2006). [CrossRef] [PubMed]
  17. X. D. Tao, O. Azucena, M. Fu, Y. Zuo, D. C. Chen, and J. Kubby, “Adaptive optics microscopy with direct wavefront sensing using fluorescent protein guide stars,” Opt. Lett.36(17), 3389–3391 (2011). [CrossRef] [PubMed]
  18. M. Schwertner, M. J. Booth, and T. Wilson, “Characterizing specimen induced aberrations for high NA adaptive optical microscopy,” Opt. Express12(26), 6540–6552 (2004). [CrossRef] [PubMed]
  19. T. J. Gould, J. R. Myers, and J. Bewersdorf, “Total internal reflection STED microscopy,” Opt. Express19(14), 13351–13357 (2011). [CrossRef] [PubMed]
  20. G. Donnert, J. Keller, R. Medda, M. A. Andrei, S. O. Rizzoli, R. Lührmann, R. Jahn, C. Eggeling, and S. W. Hell, “Macromolecular-scale resolution in biological fluorescence microscopy,” Proc. Natl. Acad. Sci. U.S.A.103(31), 11440–11445 (2006). [CrossRef] [PubMed]
  21. K. I. Willig, R. R. Kellner, R. Medda, B. Hein, S. Jakobs, and S. W. Hell, “Nanoscale resolution in GFP-based microscopy,” Nat. Methods3(9), 721–723 (2006). [CrossRef] [PubMed]
  22. E. Auksorius, B. R. Boruah, C. Dunsby, P. M. Lanigan, G. Kennedy, M. A. Neil, and P. M. French, “Stimulated emission depletion microscopy with a supercontinuum source and fluorescence lifetime imaging,” Opt. Lett.33(2), 113–115 (2008). [CrossRef] [PubMed]
  23. M. Booth, “Wave front sensor-less adaptive optics: a model-based approach using sphere packings,” Opt. Express14(4), 1339–1352 (2006). [CrossRef] [PubMed]
  24. D. Débarre, E. J. Botcherby, T. Watanabe, S. Srinivas, M. J. Booth, and T. Wilson, “Image-based adaptive optics for two-photon microscopy,” Opt. Lett.34(16), 2495–2497 (2009). [CrossRef] [PubMed]
  25. A. Facomprez, E. Beaurepaire, and D. Débarre, “Accuracy of correction in modal sensorless adaptive optics,” Opt. Express20(3), 2598–2612 (2012). [CrossRef] [PubMed]
  26. A. Thayil and M. J. Booth, “Self calibration of sensorless adaptive optical microscopes,” J. Eur. Opt. Soc.6, 11045 (2011). [CrossRef]
  27. M. A. Neil, M. J. Booth, and T. Wilson, “New modal wave-front sensor: a theoretical analysis,” J. Opt. Soc. Am. A17(6), 1098–1107 (2000). [CrossRef] [PubMed]
  28. F. Cella Zanacchi, Z. Lavagnino, M. Perrone Donnorso, A. Del Bue, L. Furia, M. Faretta, and A. Diaspro, “Live-cell 3D super-resolution imaging in thick biological samples,” Nat. Methods8(12), 1047–1049 (2011). [CrossRef] [PubMed]
  29. I. Izeddin, M. El Beheiry, J. Andilla, D. Ciepielewski, X. Darzacq, and M. Dahan, “PSF shaping using adaptive optics for three-dimensional single-molecule super-resolution imaging and tracking,” Opt. Express20(5), 4957–4967 (2012). [CrossRef] [PubMed]

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