Miniature scanning light-sheet illumination implemented in a conventional microscope

Living cells are highly dynamic systems responding to a large variety of biochemical and mechanical stimuli over minutes, which are well controlled by e.g. optical tweezers. However, live cell investigation through fluorescence microscopy is usually limited not only by the spatial and temporal imaging resolution but also by fluorophore bleaching. Therefore, we designed a miniature light-sheet illumination system that is implemented in a conventional inverted microscope equipped with optical tweezers and interferometric tracking to capture 3D images of living macrophages at reduced bleaching. The horizontal light-sheet is generated with a 0.12 mm small cantilevered mirror placed at 45° to the detection axis. The objective launched illumination beam is reflected by the micro-mirror and illuminates the sample perpendicular to the detection axis. Lateral and axial scanning of both Gaussian and Bessel beams, together with an electrically tunable lens for fast focusing, enables rapid 3D image capture without moving the sample or the objective lens. Using scanned Bessel beams and line-confocal detection, an average axial resolution of 0.8 µm together with a 10-15 fold improved image contrast is achieved.


Introduction
The detailed investigation of living cells represents one of the biggest challenges in optics, since cellular structures scale down to molecular dimensions with dynamics on millisecond timescales [1]. In addition, all live-cell studies based on fluorescence microscopy are limited by the lifetime of the fluorophores and their excited states. Fluorophores are destroyed after about 10 5 -10 6 excitations, leading to exponential blackout (bleaching) and hence to the end of the imaging experiment. In addition, the fluorophores' excited state lifetime of some nanoseconds limit the number of emitted photons per time and therefore restricts the signalto-noise ratio in the image at short integration times. A minimal integration time, on the other side, is necessary to achieve maximal temporal resolution in live cell imaging [2].
Light-sheet microscopy (LSM) [3,4] is known to reduce photobleaching significantly [5], since only fluorophores in the focal plane are excited, but not those off-axis contributing to unnecessary background signals. In addition, LSM scans 3D objects plane-wise [3,6] or linewise [7,8] and is therefore significantly faster than point-scanning microscopy.
Light-sheet microscopy has also been combined with optical tweezers to mount the whole sample by optical forces [17] or to probe the mechanical properties of epithelial cell junctions in embryos [18]. However, most of these schemes are significantly different to conventional inverted micro as in Photoni trapped nearb In this stu is suitable fo objective lens

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The optical c illumination p changes. The before illumin behind the ax (SM, Newpor of focal lengt lens L2 and th oscopes, which c Force Micro by living cells a udy, we present or most invert s confines the u up scheme ope has two hi omat 63x/1.0) o aced on the co els as shown in α ≈60°) is form s with 0.5-3 µm ight and light s n tracking of th geometry of th inside the s in this configu s is used for bo zontal beam, w n from a small lder. As shown is deflected b s. 2D images ined by vertica le lens. 3D images were recorded in 2 steps. First, the illumination beam was scanned in xdirection during the integration time of the camera. The focus of the ETL was adjusted to image the illuminated plane onto the camera. Second, the illumination beam was shifted by Δz in horizontal z-direction to illuminate the next sample plane in a vertical distance Δy = Δz, through deflection of the 45° micro mirror (see inset of Fig. 2). Simultaneously, the focal power of the ETL was adjusted to image the next illumination plane. This process was repeated to obtain a 3D image stack. The axial scanning range (y-direction) was limited by the tuning range of the ETL, which limits the height of 3D images to 20 µm. To investigate different imaging modalities, we used line-confocal [25] and wide-field detection for both Gaussian and Bessel beams

Materials and methods
Mechanical parts required for precise placement of the mirror were custom designed using CAD software considering the space constraints of the PFM. As shown in Fig. 3(a), the fixed part was designed with 3 v-groves at 120° to each other to enable kinematic mounting. The removable part was assembled from 3 parts to provide various degrees of freedom to adjust the mirror during alignment and calibration. While the first part manufactured from Aluminum with 3 threaded holes aligned with v-groves of the fixed part, the second part was made of brass with v-groves running along the sides. This part glides over 4 screws from the sides, pushed by springs from the back against 2 screws at the front. Finally, the micro fabricated mirror was glued to the end of the third part using polymer based adhesive. This arrangement allows all 6 degrees of freedom to be adjusted, which is useful for precise lightsheet alignment. The design is stable with a mirror placement precision of a few micrometers, when the removable part was taken off for sample or coverslip replacement. However, it should be noted that adjustments are not independent of each other, resulting in some extra calibration time.
Mirrors were fabricated with in-house cleanroom processing using polished Silicon wafer (surface roughness < 50 nm rms). The dimensions of the mirror were constrained by the optical tweezers configuration, which should not be obstructed by light-sheet imaging. During initial experiments, two types of mirrors were designed with a width of 120 µm to enable maximum volume imaging. The type 1 mirror was fabricated with dicing of the Aluminum coated wafer as strips of mirror with width 120 µm as shown in Fig. 3(b). Due to their size, type 1 mirrors were difficult to bond to the mirror holder shown in Fig. 3(a). Therefore, a type 2 mirror with broader mirror base (500 µm) was developed. However, the fabrication involved more steps including (i) oxide and nitride deposition, (ii) photolithography, (iii) reactive ion etching (RIE), (iv) resist stripping, (v) KOH etching, (vi) evaporation and (vii) dicing as shown in Fig. 3(b).
The initial calibration of the setup was performed by imaging free floating fluorescence beads in water. The position of the micro mirror and the tilt of the scan mirror were adjusted to get the sharp image throughout the field of view. After calibration, living cells were placed in the sample chamber. Due to inherent beam spreading, a portion of the sample close to the coverslip was not illuminated. This made it necessary to lift the cells off the coverslip by some 50 µm. Therefore, cells were grown on a drop of matrigel on the coverslip that acted as support structure to lift the sample and enable illumination of entire cells. Images were captured with rolling shutter mode of the sCMOS camera (Hamamatsu Orca flash 4.0 V2) with a slit width of 4 pixels for line confocal detection [25,26] and 400 pixels for nonconfocal detection. 3D image stacks (scanning in y-direction) were recorded with the tunable lens. With the 1.2 NA objective lens and the relay lenses L4 and L5, image stacks up to a depth of 20 µm were recorded. The operation of the microscope was controlled by custom software routines written in Python. The laser with wavelength of 491 nm was used for excitation as it was optimal for imaging macrophages labeled with LifeAct GFP.  image mammalian breast cancer cells in a cluster (Fig. 5(c)). The shape and the nuclei of the propidium iodide stained cells were visible although the sample was highly scattering.

Scanned Bessel beams
The achromatic lens was replaced by the axicon (see Fig. 2) to enable light-sheet illumination with scanned Bessel beams for further improvement in axial resolution. The line-confocal detection mode implemented in the sCMOS camera and in our self-written software suppresses the background fluorescence generated by the surrounding concentric ring system. The maximum projection of 190 nm beads diffusing through the beam ( Fig. 6(a)) displays the surrounding concentric ring system [25,27], which is characteristic to Bessel beams. The difference between line confocal and non-confocal detection is shown in Fig. 6(b, c) with xy cross-sections of fixed beads imaged with our miniature scanned Bessel beam light-sheet. The axial PSFs fitted with Gaussian function (Fig. 6(d)) clearly indicate the improvement in axial resolution with line confocal-detection. It can be seen that both the lateral and axial FWHM of the bead images (approximated PSF) hardly spread along the propagation direction of the Bessel beam ( Fig. 7(a, b)). In addition, the mean non-confocal FWHM of 1.01 ± 0.08 µm µm was decreased by 23% to a 0.78 ± 0.07 µm mean FWHM in line-confocal mode. This improvement is also validated in the MTFs averaged over 512 slices (Fig. 7(c, d)) and the corresponding line scans in axial and lateral directions.
The lateral widths of the bead images (FWHM = 0.42µm) are always broader than of a PSF alone. Furthermore, a small broadening is caused by spherical aberrations induced by a slight refractive index mismatch of the gel embedding the beads, which were in a distance of 50-100 µm to the coverslip.   To demonstrate the live cell imaging capability of our miniature light-sheet system, live J774 mouse macrophages (labelled with LifeAct-GFP) grown on a Matrigel were imaged with Bessel beam illumination and line-confocal detection. The images shown in Fig. 8(a, b) were deconvolved with Microvolution (Microvolution, llc, USA) to demonstrate the near maximal possible quality. The xz-cross-sections at different heights show multiple cellular protrusions with very low background. In Fig. 8(c, d) the deconvolution effect is shown by the maximum projections of unprocessed and deconvolved images. Further comparison of an unprocessed image section from our setup with commercial microscopy techniques such as Epi-fluorescence or Confocal Spinning Disc (Fig. 9) shows the capability of our custom built Mini Bessel light-sheet setup. Our solution clearly outperforms epi-fluorescence, but cannot compete with confocal spinning disc, which is however not applicable in combination with PFM. Fig. 9. Comparison of unprocessed xy-cross-sections from 3D images of different J774 mouse macrophages recorded with our miniature Bessel beam light-sheet setup, with conventional Epi-fluorescence and with commercial Confocal Spinning disc microscopy.
A comparable miniature reflected light-sheet microscope (RLSM) [11], used an AFM cantilever and an opposing lens displaced relative to the objective lens. However, this prevents a combination with other optical techniques based on opposing objective lenses with a common optical axis. While Gebhardt et al. [11] introduced RLSM for single molecule tracking with reduced background, 3D imaging was not demonstrated, which is a key application in LSM. Furthermore, we do not need an opposing lens, since our micromirror is mounted kinematically on the stage.

Summary and conclusion
In this study we have presented a miniature light-sheet illumination system, which can be combined with an inverted high-NA light microscope and an optical tweezers setup, without the need for any microscope reconstruction. It consists of a cantilevered micro mirror, which is positioned by a standard kinematic mounting with baseplate fixed on the microscope frame.
In our 3D LSM setup, we applied a reflecting area of 0.12 mm × 3 mm is sufficient to scan both Gaussian beams and Bessel beams through the focal plane of a NA = 1.2 water immersion lens using standard galvanometric scan mirrors. The same scan mirrors are used to displace the resulting light-sheet also in axial direction, thus keeping the setup relatively simple. The resulting scan volume was about ΔxΔyΔz ≈100 µm × 50 µm × 60 µm, the detection volume currently limited to Δy = 20µm due to the focal range of the electro optical tunable lens (ETL). Living cells can be placed inside or on top of gels of different mechanical/biochemical properties, which were mounted directly on a standard coverslip. Gels should not be too viscous to allow displacement of the cantilevered micro mirror.
Our design motivation was to avoid any displacement of the cell on the coverslip during recordings to enable simultaneous experiments with optical tweezers. Here, a constant and stable trapping focus must hold a trapped particle (or bacteria) at a well-defined position relative to the cell periphery. In our application, the design geometry of the micro mirror was further constrained by a water dipping lens opposing the objective lens at a fixed distance, thus enabling fast 3D particle position tracking. However, another decisive advantage of not displacing the sample in axial direction, but of moving the scanned light-sheet up and down is imaging speed. Although not shown in this study, this allows 2D image acquisition at several hundred Hz [24] by refocusing the fluorescence light emitted inside the light-sheet onto the camera with an ETL. In this way 3D image stacks can be recorded sufficiently fast to investigate the complex dynamics of cells, in particular of cellular protrusions.
We could show that scanned Gaussian and scanned Bessel beams in combination with line-confocal detection achieve a resolution of 0.42 µm laterally and 0.78 µm axially, while providing a more than 10-fold improved image contrast in comparison to conventional fluorescence microscopy. The images of living mouse macrophages, cells that have a rather dense cell cortex, are of high quality and provide excellent sectioning, as demonstrated in this study. Although experiments with optical tweezers have not been performed in this study, mounting living cells on top of the gel and approaching an optically trapped bead to the cell periphery is straightforward. We strongly believe that add-on systems to standard microscopes -similar to our presented system -will have great impact on future 3D life cell imaging applications, especially when speed and reduced fluorophore bleaching are of significant relevance.

Funding
German Research Foundation (DFG); Albert Ludwigs University Freiburg Open Access Publishing Fund.