Innovative sample preparation using alcohol dehydration and high refractive index medium enables acquisition of two-channel super-resolution 3D STED image of an entire oocyte

Abstract

Super-resolution (SR) microscopy is a cutting-edge method that can provide detailed structural information with high resolution. However, the thickness of the specimen has been a major limitation for SR methods, and large biological structures have posed a challenge. To overcome this, the key step is to optimise sample preparation to ensure optical homogeneity and clarity, which can enhance the capabilities of SR methods for the acquisition of thicker structures. Oocytes are the largest cells in the mammalian body and are crucial objects in reproductive biology. They are especially useful for studying membrane proteins. However, oocytes are extremely fragile and sensitive to mechanical manipulation and osmotic shocks, making sample preparation a critical and challenging step. We present an innovative, simple and sensitive approach to oocyte sample preparation for 3D STED acquisition. This involves alcohol dehydration and mounting into a high refractive index medium. This extended preparation procedure allowed us to successfully obtain a unique two-channel 3D STED SR image of an entire mouse oocyte. By optimising sample preparation, it is possible to overcome current limitations of SR methods and obtain high-resolution images of large biological structures, such as oocytes, in order to study fundamental biological processes. Lay Abstract: Super-resolution (SR) microscopy is a cutting-edge tool that allows scientists to view incredibly fine details in biological samples. However, it struggles with larger, thicker specimens, as they need to be optically clear and uniform for the best imaging results. In this study, we refined the sample preparation process to make it more suitable for SR microscopy. Our method includes carefully dehydrating biological samples with alcohol and then transferring them into a mounting medium that enhances optical clarity. This improved protocol enables high-resolution imaging of thick biological structures, which was previously challenging. By optimizing this preparation method, we hope to expand the use of SR microscopy for studying large biological samples, helping scientists better understand complex biological structures.

Keywords: 3D STED; alcohol dehydration; high refractive index medium; large biological objects; oocyte; sample preparation; super‐resolution.

FIGURE 1

Schematic representation of the final mounting workflow for the standard protocol and extended protocol . (A) Standard protocol: The sample (represented by the red circle) is transferred from a water‐based buffer through a series of glycerol solutions, starting at 5% and increasing to 70%, with a 10 min incubation at each step to equilibrate the gradient between the glycerol solution and the sample's surrounding environment. The process is finalised by mounting the sample in Vectashield mounting medium (Vector Laboratories, Burlingame, CA, USA). (B) Extended protocol: The sample is first transferred from the water‐based buffer to a 50% ethanol solution, followed by 96% ethanol for dehydration. The sample is then moved through a 50% TDE in ethanol mixture to 100% TDE, with a 20min incubation at each step. Finally, the sample is mounted in AD‐MOUNT C mounting medium (ADVI, Ricany, CZ).

FIGURE 2

Comparison of signal intensity differences and fading in deeper layers, as demonstrated in confocal images of oocytes prepared using the standard protocol and the extended protocol . The deeper part of the object refers to layers situated farthest from the objective, represented on the right side of the ZY orthogonal views, labelled as the ‘top’. The regions of the oocyte closest to the objective are labelled as the ‘bottom’. (A, B) Representative confocal sections in XY and ZY (orthogonal) layers, along with XY maximum projection images, of oocytes prepared using the (A) standard protocol and the (B) extended protocol. The ‘glow’ lookup table was used to intuitively visualise changes in signal intensities: green indicates zero values; dark red, red, yellow, and white represent the dynamic range; and blue highlights maximal values (image saturation). The scale bars represent 10 µm. In the standard protocol (A), spherical aberration results in signal fading and structure smearing at the top of the object. In contrast, the extended protocol (B) demonstrates minimal spherical aberration, with the clearing effect preserving signal intensities, resulting in almost no visible fading and a fairly homogeneous structure shape in the top of the specimen. (C, D) Charts visualising the normalised signal intensity ‘I _{( N )}’ on the Y‐axis against the distance ‘d’ into the sample on the X‐axis, representing plot profiles from the ZY orthogonal views of confocal images. Chart (C) shows the plot for the standard protocol, while (D) shows the plot for the extended protocol. In both (A) and (B), signal intensities were measured from the area under the light red‐grey rectangle in the YZ orthogonal projections. Area measurements were used instead of simple line profiles to minimise noise in the plotted data. (E) The chart represents the average decrease in intensities ‘I _{( N )}’ in the top area, normalised to the intensities in the bottom area. The average and standard deviation are plotted, with corresponding values shown for oocytes processed using the standard protocol (StP) and the extended protocol (ExP).

FIGURE 3

Confocal images of an oocyte with adhered 80 nm gold beads prepared using the extended protocol . The images demonstrate the appropriate formation of the 2D STED and 3D STED depletion laser patterns at depths ranging from a few micrometres to approximately 80 µm. (A) Maximum intensity projection of the oocyte in confocal images along the XY and XZ axes. The grey channel represents the fluorescence signal of the counterstaining used to visualise the oocyte membrane, while the ‘Red Hot’ lookup table represents the visualisation of the adhered 80 nm gold beads, acquired in reflection mode. The scale bar represents 10 µm. (B) Confocal sections showing the oocyte membrane in the grey channel, with gold beads used to visualise the PSFs of the excitation laser (green) and the depletion laser (red). The XY confocal section shows the lateral co‐alignment, while the XZ confocal scan shows the axial co‐alignment of the excitation and depletion lasers. The positions relative to the objective are labelled as ‘bottom’ or ‘top’. The labels ‘2D STED’ and ‘3D STED’ indicate the settings of the microscope STED module, with the corresponding depletion laser PSF shape formed on the gold beads. The scale bar represents 1 µm.

FIGURE 4

Confocal and 3D STED images of oocytes prepared using the standard protocol and the extended protocol . The figure shows a sequence of images for each protocol: The first row presents a simple XZ orthogonal scan, while the second row shows a single XY section at the top of the oocyte. The third row visualises a single XY section at the bottom of the oocyte, and the fourth row displays a simple XZ orthogonal scan of the bottom region. The method used for image acquisition is labelled as either ‘confocal’ or ‘3D STED’. The scale bar represents 10 µm. (A) The standard protocol yields images with standard confocal quality at the bottom, suitable for 3D STED super‐resolution imaging. However, at the top region, the signal intensity is weak and fades, resulting in unsuccessful 3D STED imaging. To highlight the weak signals and demonstrate this issue, we nonsystematically increased the intensities of the images within the white‐bordered squares located at the top. (B) The extended protocol produces confocal and 3D STED super‐resolution images, with fully comparable quality at both the bottom and top, with no significant fading of signal intensities.

FIGURE 5

The image resolution achieved with 2D STED approach within the oocyte sample. (A) Comparison between the confocal (labelled as ‘confocal’) and 2D STED (labelled as ‘2D STED’) acquisition methods at the bottom and top positions of the oocyte. The scale bar represents 10 µm in the overview image and 1 µm in the zoomed‐in area. (B) A table quantifying the mean and standard deviation of the resolution, as calculated by the FRC method, for images acquired from the bottom and top regions of the specimen.

FIGURE 6

The visualisation of the final deconvolved two‐channel 3D STED super‐resolution volume reconstructed image of the entire mouse oocyte. This figure provides an overview of the oocyte and reveals detailed microvillar structures visualised with immunofluorescent staining of Juno (green) and co‐localised CD9 protein (red) in the zoomed‐in area. (A) 3D maximum projection of the full z‐stack of the entire oocyte using 3D STED microscopy, and (B) a close‐up of a selected area from the entire oocyte image. The scale bars represent (A) 10 µm and (B) 5 µm. (C) A representative maximum intensity projection of 10 selected optical sections around the equatorial layer of the oocyte in the super‐resolution 3D STED image. The scale bar represents 5 µm.