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Edited by Jennifer Lippincott-Schwartz, National Institutes of Health, Bethesda, MD, and approved January 7, 2013 (received for review September 25, 2012)
Abstract
A recent key requirement in life sciences is the observation of biological processes in their natural in vivo context. However, imaging techniques that allow fast imaging with higher resolution in 3D thick specimens are still limited. Spinning disk confocal microscopy using a Yokogawa Confocal Scanner Unit, which offers high-speed multipoint confocal live imaging, has been found to have wide utility among cell biologists. A conventional Confocal Scanner Unit configuration, however, is not optimized for thick specimens, for which the background noise attributed to “pinhole cross-talk,” which is unintended pinhole transmission of out-of-focus light, limits overall performance in focal discrimination and reduces confocal capability. Here, we improve spinning disk confocal microscopy by eliminating pinhole cross-talk. First, the amount of pinhole cross-talk is reduced by increasing the interpinhole distance. Second, the generation of out-of-focus light is prevented by two-photon excitation that achieves selective-plane illumination. We evaluate the effect of these modifications and test the applicability to the live imaging of green fluorescent protein-expressing model animals. As demonstrated by visualizing the fine details of the 3D cell shape and submicron-size cytoskeletal structures inside animals, these strategies dramatically improve higher-resolution intravital imaging.
When imaging the 3D interior of living organisms, fast confocal fluorescence microscopy is essential in monitoring dynamic life processes by collecting optically sectioned images over time. One of the main challenges to achieving high-speed acquisition has been to use a multipoint scanning strategy using a spinning disk scanner containing a set of confocal pinholes. This overcomes the speed-limiting disadvantage in single-beam laser scanning microscopy (LSM). The Confocal Scanner Unit (CSU) developed by Yokogawa Electric is one of the most advanced spinning disk systems (1, 2) (Fig. S1A).
The Yokogawa CSU consists of two disks, each containing ∼20,000 microlenses and pinholes, arranged in a spiral array of equal pitch (3) (Fig. 1 A and B). The microlenses are positioned to increase the amount of light transmitted through the disk. The light shone from a beam-expanded laser onto the first disk is focused by each microlens onto the corresponding pinhole of the second disk to generate a parallelized multibeam (Fig. S1A). During imaging of emitted fluorescence at each illumination point, each pinhole then acts as a detection pinhole and rejects out-of-focus signals, and the points of fluorescence merge into a 2D image that can be collected with a digital camera as the disk rotates.
Fig. 1.
Quantitative analysis of the pinhole cross-talk. (A) Brief CSU specifications used in this study (left column) and throughput of laser light across the CSU models (right column). The power of the 488-nm Ar-ion laser and the mode-locked Ti:Sa laser at 950 nm were measured at, before, and after each CSU model, and the throughput of laser light was calculated. (B) Pinhole pattern on the CSU disk. The pinholes are arranged in an equal-pitch spiral pattern. (C and D) Quantification of pinhole cross-talk by sea response analysis (20). Fluorescence sea responses were obtained by collecting z-stack images of a dye solution layer with different thicknesses (10, 30, and 80 μm) using three different CSU models with 1PE or 2PE (C), and the proportion of the background noise intensity to total fluorescence intensity (FI) is plotted in D.
Cell biologists have noted that spinning disk confocal (SDC) microscopy with CSU significantly reduces radiation damage, compared with conventional point-scanning LSM and allows high-resolution imaging using a low-noise and high-dynamic-range detector such as a charge-coupled device (CCD) camera, as opposed to the noisier photomultiplier tubes (PMTs) used for LSM (2, 4⇓⇓–7). As a result, this system has been found to be indispensable for biological cell studies. Recent progress in light microscopy has included the achievement of imaging large specimens such as a whole embryo and separating nanostructures at resolution of less than 100 nm by combining optics technologies including planar illumination and superresolution techniques, respectively (8⇓⇓–11; reviewed in refs. 12⇓–14). However, light-sheet microscopy for large specimens is not intended for higher-resolution imaging and not yet available for routine laboratory use, whereas satisfactory results obtained using superresolution techniques are limited to very thin specimens. The SDC method using a CSU is, on the other hand, well suited to a wide range of applications in biological studies using a variety of sample formats. The simple interface can be easily affixed to any upright or inverted microscope in combination with a variety of devices, such as separate stimulation lasers, without special expertise.
The multipoint scanning strategy, however, suffers an intrinsic problem. Fluorescence emissions originating from remote focal planes and out-of-focus scattering pass through adjacent pinholes, increasing the background signal haze that obscures the image (Fig. S2). This phenomenon, called “pinhole cross-talk” (1, 2, 15), makes it difficult to apply the SDC to thick specimens such as tissues and animals, which produce a large amount of out-of-focus background noise.
In this study, we sought to improve SDC microscopy by eliminating pinhole cross-talk (Fig. S2). It is expected that, if the interpinhole distance is increased, the amount of pinhole cross-talk will be reduced, in proportion to the distance, but at the cost of overall power. If the generation of out-of-focus light is prevented by two-photon excitation (2PE), which excites fluorophores only in the focal plane (16, 17), the major contributor of background noise will be removed. Here, we evaluate the effect of increased interpinhole distance and the use of 2PE and test its applicability to the live-imaging of green fluorescent protein (GFP)-expressing model animals.
Results
Construction and Evaluation of a 2PE SDC Microscope.
The current CSU model, CSU-X1, contains 250-μm-diameter microlenses and 50-μm-diameter pinholes with 250-μm pitch (Fig. 1 A and B and Fig. S1B). The pinhole size has been designed for high magnification with high numerical aperture (NA) objectives set close to 1 Airy unit. To adapt 2PE, we modified the CSU configuration: the diameter of the microlenses was increased to 580 μm, and the interpinhole distance was increased to 580 μm (CSU-MP series). The larger microlens size was used to increase the throughput and concentration of excitation light (Fig. 1A, right column) to increase two-photon absorption efficiency. In addition, to evaluate the effect of the pinhole size, we constructed two pinhole array disks containing 55-μm- and 100-μm-diameter pinholes, respectively.
We first evaluated optical performance by point spread function (PSF) analysis (Fig. S3). The 2PE improved axial resolution by ∼8% over 1PE, while providing ∼4% less lateral resolution, as theoretically expected (18, 19). Next, we quantified the pinhole cross-talk by measuring the “sea response,” in which a densely fluorescent thick layer of solution was imaged by z-scanning to detect the axial response (20) (Fig. 1 C and D). The sea response is sensitive to secondary contributions from above and below the focal plane that are superimposed on the sectioned image. It is, therefore, a stringent test of the axial sectioning property. As shown in Fig. 1D, in the conventional 1PE-X1 setting, the background signal level reaches as high as >60% of total detected signals when sample thickness is increased. With larger interpinhole distance, the background level decreased to less than 40%, in proportion to the pinhole size. The 2PE significantly reduced the background noise; however, ∼6% transmission was detectable in 2PE-X1. In 2PE-MP models, the background noise was less than 0.5%.
To demonstrate the benefit of 2PE in the imaging of a bulky object, we imaged a pollen grain of ∼30 μm diameter that emits stable auto-fluorescence (Fig. 2 and Fig. S4). In 2PE, the signal contrast was much higher than that in 1PE, especially in the deepest region. The surface rendering demonstrated that with 2PE-MPφ55, details were visible even on the far side of the pollen grain along the observation axis, whereas with the 1PE-X1 setting, the far side was buried in the background haze (Fig. 2C and Movie S1).
Fig. 2.
Comparison of the optical sectioning capability of 1PE and 2PE-SDC for a bulky object. (A) Image of a pollen grain collected using CSU model X1 with 1PE (1PE-X1) and CSU model MPφ55 with 2PE (2PE-MPφ55) with an sCMOS camera. Voxel pitch (xyz): 56 × 56 × 200 nm, 194 stacks; depth: 0–38.8 μm. The images of the same pollen grain are shown in x–z slices. For the full data set, see Fig. S4. (B) Fluorescence intensity (FI) profiles along the line (seven-pixel width) indicated in A. Black and red lines represent profiles for 1PE-X1 and 2PE-MPφ55, respectively. (C) Surface-rendered representation of the pollen grain. The threshold level was determined so as to display the same equatorial diameter. The entire surface of the 2PE image can be rendered, whereas with 1PE, high background noise interferes with the surface rendering, especially at the far side. Also see Movie S1. (Scale bars: 10 μm.)
Application of CSU Models to Animals Expressing GFP-Fusion Proteins.
We examined the application of the CSU models to the imaging of GFP-expressing animals. In advance, we confirmed that the GFP-photobleaching rate for 2PE with CSU is comparable to that of conventional two-photon-excited fluorescence laser scanning microscopy (2PE-LSM) (16, 17, 21, 22) (Fig. S5). First, we observed a slice of a whole mouse embryo expressing GFP-fused histone H2B (H2B-GFP), which accumulates in the nucleus (Fig. 3 A and B, Fig. S6, and Movie S2). Dramatic improvements made by 2PE for optical sectioning were apparent in x–z slice views: in 2PE, the shape of the nuclei was visible at a depth of 100 μm, whereas with 1PE, the contrast was completely lost at a depth of ∼30 μm, and accumulated background noise overwhelmed the localized signals in regions deeper than 40 μm. Next, we observed embryos of the fruit fly Drosophila expressing membrane-GFP, which are distributed along plasma membranes (Figs. 3C and Fig. S7). In Fig. 3D, x–y plane images of a membrane-GFP–expressing fly embryo at z = 80 μm collected under different conditions are compared. The signal-to-background ratio (SBR) is more than 35 times larger in 2PE than in 1PE. The advantage of 2PE was apparent in high-contrast images, precisely visualizing the shape of cells in deeper regions (Fig. 3E and Movie S3). Because the transparency is higher for this fly sample than for H2B-GFP–expressing mouse embryo, with 1PE-MPφ55, fine structures are visible up to a depth of ∼50 μm in the x–z slice view (Fig. S7). SBR values at z = 80 μm for all modalities are shown in Fig. S8C. Comparison between 1PE-MPφ100 and 1PE-MPφ55 or 2PE-MPφ100 and 2PE-MPφ55 shows that the smaller pinhole size has the effect of reducing background noise both in 1PE and 2PE. Although the pinhole-size effect was small in the sea response (Fig. 1 C and D), in tissues, fluorescence photons may have scattered multiple times and were prone to generate a diffuse background. From these observations, we conclude that the 2PE-MPφ55 achieved the highest level of confocality with sufficient signal intensity.
Fig. 3.
Comparable images of GFP-expressing animals obtained with different modalities. (A) x–z slices constructed from z-stack images of forebrain neuroepithelial cells in a slice of a fixed mouse whole embryo expressing H2B-GFP, which localizes in nuclei. The z-stack images were collected from the sample surface to a region 100 μm deep using different modalities as indicated in the figures with an iXon EM-CCD camera. Voxel pitch (xyz): 183 × 183 × 500 nm, 200 stacks; depth: 0–100 μm. Also see Movie S2. (Scale bar: 20 μm.) (B) Fluorescence-intensity profiles along the line (seven-pixel width) shown in the images in A. The profiles were normalized to 1.0 at the first peak from the sample surface. (C) Orthogonal representation of a fixed whole Drosophila embryo expressing membrane-GFP, which localizes along entire plasma membranes, collected with an sCMOS camera. Voxel pitch (xyz): 112 × 112 × 400 nm, 200 stacks; depth: 0–80 μm. The head part of a stage 17 embryo was optically sectioned from the lateral surface (Upper) to the midline (Lower). For a schematic diagram of Drosophila embryo, see Fig. 4B. With 2PE, the detailed cellular structures are visible in regions 80 μm deep. (Scale bar: 10 μm.) (D) x–y plane images of Drosophila foregut expressing membrane-GFP at z ∼ 80 μm. The signal-to-background ratio calculated using the central 260 × 260-pixel areas of the pictures is indicated. (Scale bars: 10 μm.) Dorsal, left; anterior, down. (E) Surface-rendering representation of eight cells located at a depth of 65–80 μm of the Drosophila embryo shown in C. The surfaces of cells having distinct shape can be traced. Also see Movie S3. Note that in all 2PE images, only the central areas ∼40 μm in diameter were excited because of the lack of power of the mode-locked Ti:Sa laser used to excite the GFP. For the full data set for A and D, see Figs. S6 and S8, respectively. Also see Fig. S7.
Fig. 4.
Visualization of submicron-size structures inside living fly embryos using 2PE. (A) Subcellular distributions of microtubules (green) and a microtubule plus-end-binding protein EB1 (magenta) in cultured HeLa cells. The images were collected by structured illumination microscopy. Microtubules are filaments with 25-nm diameter. EB1 binds to growing ends of microtubules in comet shapes. The dimensions of an EB1 comet are about 25 nm in width and up to ∼500 nm in length. The Inset is the 2.5× magnified image of the boxed area. (Scale bar: 10 μm.) (B) Schematic diagram of Drosophila embryo at approximately stage 13. The guts are composed of tubular epithelium. An x–y section plane was observed using different modalities as indicated in the figures. (C) x–y plane images of a living Drosophila embryo expressing EB1-GFP at z ∼ 25 μm from the surface collected using an sCMOS camera. The horizontal sections of a hindgut were observed from the dorsal side. EB1-GFP comets are visible with 2PE (arrows), whereas with 1PE, they are difficult to detect because of high background noise. Also see Movie S4. (Scale bar: 20 μm.) (D) Series of time-lapse images of the boxed area in C. Individual comets moving from the apical to basal side are indicated with arrowheads and arrows. (Scale bar: 5 μm.) The time scale is seconds:milliseconds. (E) EB1-GFP tracking result superimposed on the first time-lapse image with lower brightness. The tracking paths and the last tracked positions are indicated with lines and dots, respectively. (Scale bar: 10 μm.)
We also compared the images collected with the 2PE-MPφ55 setting equipped with a Scientific Complementary Metal Oxide Semiconductor (sCMOS) camera and a conventional single-beam 1PE-LSM or 2PE-LSM, for which the signals are detected using PMTs (Fig. S9). Images of a fly embryo expressing membrane-GFP collected with 2PE-MPφ55 have less noise and display fine detail, especially in the x–z slice view, compared with images by 1PE-LSM or 2PE-LSM, demonstrating the advantage of low-noise digital imaging. Improvement of axial resolution with pinholes (Fig. S3) also contributes to providing x–z cross-section images with high image contrast.
Imaging Submicron-Size Structures Inside Living Animals.
We attempted to detect submicron-size structures inside living animals. End-binding (EB)1 is a microtubule associating protein that accumulates specifically at the growing microtubule end in a comet shape with a 25-nm short axis and a posteriorly elongated tail of up to ∼500 nm (23) (Fig. 4A). GFP-fused EB1 (EB1-GFP) has been widely used to monitor microtubule growth dynamics as a microtubule-growth marker (24). We attempted to visualize EB1-GFP in a fruit fly embryo in its 13th developmental stage (Fig. 4B). At ∼25-μm depth in the epithelial cell monolayer constructing a hindgut, with 1PE, the EB1-GFP comet was hardly detectable because of high haze levels, but with 2PE, moving comets were visible (Fig. 4 C–E and Movie S4). In addition to unidirectional movement of the EB1-GFP comets from the apical to basal side, which is a common feature of microtubule organization in polarized epithelial cells, a unique accumulation of EB1 near the apical surface was observed, indicating frequent microtubule nucleation. In unfertilized mouse oocytes (Fig. 5A), in which most microtubules are present in the meiotic spindle apparatus arrested at metaphase of second meiosis (Fig. 5B), the 2PE method visualized the EB1-GFP comet inside thick spindle microtubule bundles (Figs. 5C and Movie S5), demonstrating dynamic microtubule turnover even in the arrested spindle. In the two-cell stage of Caenorhabditis elegans embryo, in which microtubules are visualized by GFP–β-tubulin, the ends of individual microtubules are precisely visualized with 2PE, whereas with 1PE, the contrast was lost in the high level of noise (Fig. 5D and Movie S6).
Fig. 5.
Visualization of submicron-size structures inside living mouse oocytes and C. elegans embryos using 2PE. (A) Schematic diagram of mouse oocyte. The spindle apparatus structure was observed through the cytoplasm. (B) x–y image slices of a living mouse oocyte expressing EB1-GFP at z ∼ 64 μm, collected with an iXon EM-CCD camera. (Scale bars: Left, 20 μm; Right, 5 μm). (C) High-magnification views of spindle poles in a living mouse oocyte expressing EB1-GFP at z ∼ 64 μm, collected with an sCMOS camera. Arrowheads indicate individual EB1-GFP comet. EB1-GFP tracking results are presented at the bottom as in Fig. 4E. Also see Movie S5. (Scale bar: 5 μm.) (D) z-sectioning images of living C. elegans embryos expressing GFP-β-tubulin collected at z = 0.1-μm intervals using an sCMOS camera and maximum projections of 130 sections. Voxel pitch (xyz): 75 × 75 × 100 nm, 130 stacks; depth: 0–13 μm. Magnified images of boxed areas are shown on the right. With 2PE, the precise position of an individual microtubule is visualized (arrowheads). Also see Movie S6. (Scale bars: Left, 5 μm; Right, 1 μm.)
Discussion
In this study, we demonstrated that elimination of pinhole cross-talk dramatically improves the optical sectioning capability of SDC microscopy. The wider interpinhole distance effectively reduced the pinhole cross-talk in proportion to the distance. The introduction of 2PE significantly reduced the background noise, while maintaining sufficient signal intensity. Although multipoint scanning 2PE microscopy without pinhole detection can provide a measure of an optical sectioning property (25), the introduction of pinhole detection can eliminate fluorescence photons that scatter in the specimen and cause haze in images (26) and improve axial resolution as theoretically predicted (18, 19). Indeed, the results obtained in this study demonstrate that a smaller pinhole size improves image contrast when observing thick tissues. In combination with digital camera devices, this method achieves high-speed confocal imaging of the inside of thick specimens with remarkably low noise and high dynamic-range information. Thus, the CSU models described here, especially when using 2PE, are potentially of great use in the life sciences.
It should be noted, however, that with configurations using detector pinholes, deeper imaging at the millimeter level would be difficult, unlike conventional single-beam 2PE-LSM that collects a large pool of scattered light from a single source without a pinhole (16, 17). In addition, because photobleaching in the focal plane is greater for 2PE than 1PE, and lateral resolution is higher in 1PE than in 2PE, 2PE is considered to be unsuitable for imaging thin specimens. The 1PE-SDC with wider interpinhole distance will have benefit for imaging of relatively thin (less than ∼20 μm) or transparent specimens [e.g., those treated with tissue-clearing reagent after fixation (27)]. Nevertheless, 2PE-SDC microscopy will contribute greatly to the advancement of biological science, in which monitoring the behavior of fast-moving fine structures inside the cells of living tissue and animals (intravital imaging) is crucial for the understanding of life processes. This approach can fill the gap between sheet-light microscopy optimized for a large specimen and superresolution microscopy. Not only for live imaging, this method will be a great tool for the collection of a large number of optical sections throughout bulky specimens to reconstitute 3D views with higher resolution.
In putting the system to practical use, the available output laser power is the limiting factor for the illumination area and the maximum imaging depth. Even with the two-photon lasers having the highest levels of output power currently available, we could trigger two-photon absorption for only ∼40-μm-diameter areas of GFP-expressing animals (less than 10% of the effective frame size with a 60× objective; Fig. S1B). Lasers with roughly 5–10 times the power would be required. The higher laser power is also required to excite red fluorescent proteins, which can be excited at wavelengths between 1,000 and 1,200 nm, at which the output power of the existing mode-locked laser decreases to ∼15% of peak power. Following the development of the next generation of high-power but low-cost lasers for SDC microscopy, the most optimal system will have a large impact on the broad field of biology.
Materials and Methods
SDC Microscope Systems.
A schematic diagram of the setup is presented in Fig. S1. The SDC microscope system was based on an IX81 inverted microscope (Olympus) equipped with a silicone immersion objective UPLSAPO60XS (NA = 1.3) moved by a piezo actuator with a feedback loop (Physik Instrumente). The CSU models (Yokogawa) used in this study are listed in Fig. 1A, left column. In addition to the existing model CSU-X1, two models described here, having larger microlens diameters and interpinhole distances with different pinhole diameters (CSU-MPφ55 and CSU-MPφ100), were evaluated. In the CSUs, a dichroic mirror (700- to 1,100-nm band pass; Yokogawa) was installed. In the standard CSU models, the disks were treated with an antireflection (AR) coating (400–700 nm), but the CSU-MP models were used without an AR coating to minimize loss of the excitation light. The transmittance of laser light through CSU is shown in Fig. 1A, right column. Excitation light was directly introduced to the CSU by air propagation. As a light source, for 2PE, mode-locked Ti:Sa lasers (Chameleon Vision II: 80 MHz, 140 fs, 1.35 W at 920 nm; Chameleon Ultra II: 80 MHz, 140 fs, >1.6 W at 920 nm; Coherent) were used. The incident beam power was adjusted using a Glan-laser polarizer with an AR coating (1,050–1,620 nm; Thorlabs), and the laser beam width was controlled by a beam expander consisting of a concave and a convex lens pair with an AR coating (650–1,050 nm; T = 94%). For 1PE, an Ar-ion laser (488 nm; >10 mW; Melles Griot) was used. The incident beam power was adjusted using neutral density filters and the laser beam width was controlled by a beam expander consisting of a concave and a convex lens pair with an AR coating (400–700 nm). The laser beams were introduced into the CSU using broad band dielectric mirrors (750–1,100 nm; >99% reflectivity in infrared; Thorlabs) in the optical path of the Ti:Sa laser and silver-coated mirrors (>98% reflectivity in infrared; Thorlabs) in the optical paths of both the Ti:Sa laser and Ar-ion laser. Using these optics, the system achieved ∼83% throughput of the Ti:Sa laser, with power up to 1.56 W, at the entrance of the CSU. The images were acquired using a water-cooled EM-CCD camera (iXon+ DV897; Andor), an EM-CCD camera (Luca; Andor), or an sCMOS camera (Neo; Andor). The fluorescence signals were detected through an infrared ray cut filter (FF01-680/SP; Semrock) and an emission filter for GFP (FF01-525/45; Semrock) inserted before the cameras. The system was controlled with iQ2 software (Andor).
Commercial Microscope Systems.
Conventional single-beam 1PE-LSM imaging was performed using an LSM780 (Carl Zeiss) equipped with an Axio Observer.Z1 inverted microscope with a C-Apochromat 63×/1.20 NA water immersion objective, a multi-line Ar laser (458, 488, 514 nm), and a highly sensitive gallium arsenide phosphide (GaAsP) photomultiplier (PMT) detector. A conventional single-beam 2PE-LSM FV-1000 MPE (Olympus) was equipped with a BX61 upright microscope, a silicone immersion objective UPLSAPO60XS (NA = 1.3), MaiTai DeepSee laser (Spectral Physics), and GaAsP PMT detector. Structured illumination microscopy was performed using a ELYRA PS.1 (Carl Zeiss) equipped with an Axio Observer.Z1 inverted microscope with an αPlan-APOCHROMAT 100×/1.46 NA oil immersion objective, four diode laser lines (405, 488, 561, 642 nm), an EM-CCD camera iXon885 (Andor), and system control software ZEN 2010 C.
PSF and Sea Response Measurement.
Specimens were prepared using No.1S coverslips (Matsunami) and slide glasses (Matsunami). Three-dimensional image stacks of fluorescent beads (PS-Speck Microscope Point Source Kit; Component B; 174 nm diameter; excitation/emission: 505/515 nm; Invitrogen) were z-scanned at 50-nm steps for 9.7 μm using a Luca camera. The PSF width [full width at half-maximum (FWHM)] was calculated by Gaussian approximations (28, 29). The sea response, which describes the signal generated by a fluorescent half-space moving along the optic axis, was measured as described previously (20) using a 50 μM solution of Rhodamine 6G (Tokyo Chemical Institute) in methanol–glycerol mixed solvent (6:4; vol./vol.) with thicknesses of 10, 30, and 80 μm. The layers of dye solutions were z-scanned at 1-μm steps for 200 μm including under- and over-focus positions using an sCMOS camera. The average fluorescence intensity in each stack was obtained from a 100 × 100-pixel area at the center of the image.
Biological Sample Preparation and Imaging.
A HeLa cell line expressing GFP–α-tubulin (clone 1E10) was described previously (30). For immunostaining of microtubules and EB1 in HeLa cells, HeLa cells cultured on coverslips (No.1S; Matsunami) were fixed and stained as described previously (30) using a mouse anti-tubulin antibody conjugated with FITC (Sigma) and a rabbit anti-EB1 antibody (Abcam). GFP-fused histone H2B (H2B-GFP) transgenic heterozygous mice (R26-H2B-EGFP; accession no. CDB0238K; www.cdb.riken.jp/arg/reporter_mice.html) (31) were provided by the CDB Laboratory for Animal Resources and Genetic Engineering. Slices of H2B-GFP mice at embryonic day 10.5 were prepared as follows. Whole embryos were fixed in 4% (vol/vol) paraformaldehyde overnight and embedded in a 4% (vol/vol) agarose gel (low-melting-point agarose; Invitrogen). Sagittal slices (200 μm thick) were then cut using a vibratome (LinierSlicer PRO10; DOSAKA) and mounted in Prolong Gold antifade reagent (Invitrogen). GFP-fused mouse EB1 (EB1-GFP) (23) was expressed in mouse oocytes by mRNA injection as described previously (32). To maintain a living mouse oocyte during imaging, a CO2 incubator (Tokai Hit) was used. Fruit fly Drosophila embryos carrying ubi-EB1:GFP transgene (33) were dechorionated with 50% (vol/vol) bleach. Dechorionated embryos were mounted on a glass-base dish (Iwaki) with glue and covered with water. Fly embryos carrying the ubi-GFP:CAAX transgene (membrane-GFP) (34) were dechorionated with 50% bleach and fixed by 4% paraformaldehyde in PBS for 30 min at room temperature, followed by devitellinization in methanol. After washing with 0.2% Tween-20 and 0.2% Triton X-100 in PBS, embryos were mounted in a Vectashield mounting medium (Vector Laboratories). The C. elegans strain used was AZ235, which expresses GFP-fused tubulin in embryonic cells (35). Embryos were dissected from a gravid hermaphrodite cultured using the standard method at 22–25 °C (36) and mounted on a multiwell test slide (MP Biomedicals) with M9 buffer. Coverslips were sealed with nail polish.
Image Analysis and Presentation.
The signal intensity analysis and manual tracking were performed using iQ software (Andor) or ImageJ software. The statistical analysis was performed using Microsoft Excel or R software. The statistical significance of the differences was assessed in a two-sample t test. The SBR was obtained according to:
where I is average intensity. In the presentation of comparative raw image sets, image brightness was normalized with the average fluorescence intensity obtained from the top 1% of bright pixels after subtracting the average background intensity detected without a specimen. Image presentation by orthogonal slice and surface rendering was performed using Imaris software (Andor/Bitplane).
Acknowledgments
We thank Coherent Japan Inc., which rented us a Chameleon Vision II and a Chameleon Ultra II laser. We thank Mr. Yoshiharu Saito and Mr. Yuichiro Imai (Olympus Corp.) and Mr. Yasuhiko Sato (Carl Zeiss Microscopy Co., Ltd.) for providing valuable advice and technical support in the construction of the microscope system and in the operation of the Imaris software, respectively. Imaging experiments using commercial microscopes and image analysis were performed at the RIKEN Center for Developmental Biology (CDB) Imaging Facility. The conventional two-photon scanning images were collected at the CDB 4D tissue analysis unit using an FV-1000 MPE (Olympus). We also thank Dr. Miho Ohsugi (The University of Tokyo) for a plasmid construction to generate EB1-GFP mRNA; Dr. Hiroyuki Ohkura (University of Edinburgh) for the fly strain expressing EB1-GFP; Ms. Emiko Maekawa (RIKEN CDB) for technical assistance; and the Caenorhabditis Genetics Center, which is funded by the National Institutes of Health National Center for Research Resources, for the nematode strain. This work was supported by the RIKEN CDB Director’s Fund. Y.M.-K. was also funded by the Kurata Memorial Hitachi Science and Technology Foundation, the Japan Society for the Promotion of Science Funding Program for Next Generation World-Leading Researchers, the Takeda Science Foundation, and the Uehara Memorial Foundation.
Footnotes
- ↵1To whom correspondence should be addressed. E-mail: y-kiyosue{at}cdb.riken.jp.
Author contributions: Y.M.-K. designed research; T.S., K.Y., T.K., S.H., A.S., D.K., F.M., J.T., S.O., H.N., Y.K., T.M.W., K.F., and Y.M.-K. performed research; T.S. and Y.M.-K. analyzed data; and Y.M.-K. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1216696110/-/DCSupplemental.
References
Improving spinning disk confocal microscopy
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