We would like to thank Jonas Ries and the Ries lab for sharing imaging buffers for SMLM imaging. We also thank Yumi Kim for the C. elegans strain used in this protocol and Abby F. Dernburg for the chicken-anti-HTP-3 antibody. We thank Marko Lampe and Stefan Terjung from the Advanced Light Microscopy Facility at EMBL Heidelberg for their support in using the Olympus iXplore SPIN SR confocal microscope. This work was supported by the European Molecular Biology Laboratory and the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation - 452616889, SK). We acknowledge the access and services provided by the Imaging Centre at the European Molecular Biology Laboratory (EMBL IC), generously supported by the Boehringer Ingelheim Foundation.

1. Preparation of solutions and coverslips
NOTE: See the Table of Materials for details related to all materials and reagents and Table 1 for the composition of solutions used in this protocol.
<ol>
	<li>Poly-L-lysine-coated coverslips
	<ol>
		<li>Prepare 0.01% (w/v) poly-L-lysine (see Table 1).</li>
		<li>Wash a precision coverslip (24 mm diameter; 0.17 &#177; 0.005 mm, No. 1.5) in ethanol for 10-30 min. Rinse the coverslip with ddH2O to remove ethanol, and leave the coverslip to dry at room temperature.</li>
		<li>Plasma clean the coverslip using a plasma cleaner. 
		NOTE: Plasma cleaning increases the hydrophilicity of the coverslip and facilitates the following steps. If a plasma cleaner is not available, this step can be skipped, although this may require adjusting the volume and/or concentration of the poly-L-lysine solution. This modification has not been tested.</li>
		<li>Place one drop (120 &#181;L) of 0.01% (w/v) poly-L-lysine on the coverslip. Incubate for 10 min at room temperature.</li>
		<li>After incubation, rinse the coverslip in ddH2O and dry at room temperature. Store at 4 &#176;C up to 1 month.</li>
	</ol>
	</li>
	<li>F(ab&#39;)2 fragments conjugated with fluorescent organic dyes
	<ol>
		<li>Add in the following order to a PCR tube: 10 &#181;L of 0.6-0.7 mg/mL F(ab&#39;)2 fragment in PBS, 1 &#181;L of 0.1 M NaHCO3 (pH 8.3) and 1 &#181;L of 1 mM succinimidyl (NHS) ester reactive fluorophore in DMSO (molar ratio of F(ab&#39;)2:dye is ~1:17). Mix well by pipetting up and down.</li>
		<li>Incubate for 1 h at room temperature.</li>
		<li>Separate the F(ab&#39;)2 fragment from the remaining free reactive dye using a desalting column (7K MWCO) following the manufacturer&#39;s specifications. Use 1x PBS for equilibration of the column and elution of the labeled F(ab&#39;)2 fragment.</li>
		<li>Store the labeled F(ab&#39;)2 fragment at 4 &#176;C for up to 3 months. 
		NOTE: Storage times of longer than 3 months have not been tested.</li>
	</ol>
	</li>
</ol>
2. Dissection and fixation
NOTE: The dissection and fixation procedures are modified from previously recommended procedures16,40 to obtain optimal samples for super-resolution microscopy.
<ol>
	<li>Dissection
	<ol>
		<li>Pick age-matched C. elegans worms (grown at 20 &#176;C for this study) into a 30 &#181;L drop of EBTT (1x Egg buffer41 with 0.2% nonionic detergent, Table 1) on a coverslip (22 mm x 22 mm, No. 1). Place the cover slip on a glass slide for easier manipulation. Wash with 30 &#181;L of EBTT by pipetting up and down several times. Remove 30 &#181;L of the solution to leave a 30 &#181;L drop on the coverslip. 
		NOTE: Small amounts of nonionic detergent must be added to all solutions in which worms are pipetted to prevent the worms from sticking to the plastic tips.</li>
		<li>Use a scalpel blade to cut off the heads and/or the tails of the worms to extrude the gonad (Figure 2A).</li>
	</ol>
	</li>
	<li>Fixation
	<ol>
		<li>Pipet 30 &#181;L of fixative solution (Table 1) into the drop of the dissected worms and pipet up and down to mix. 
		NOTE: Pipetting up and down a few times may help to release more gonads.</li>
		<li>Fix for exactly 1 min after adding the fixative solution.</li>
		<li>Stop the fixation by transferring the worms into a PCR tube filled with TBST (Table 1). Transfer the worms in as little volume as possible (~15 &#181;L).</li>
		<li>Spin down the PCR tube on a mini benchtop centrifuge (2,000 &#215; g, 10 s). Remove the supernatant and wash 2x with 200 &#181;L of TBST each.</li>
		<li>Wash with 200 &#181;L of PBST (Table 1) for 5-10 min. Repeat steps 2.1.1 to 2.2.5 for up to four samples while keeping the dissected samples on ice. 
		NOTE: If processing more than four samples, proceed with steps 2.2.6 to 2.2.7&#160;after every four samples to ensure sample fixation remains consistent across all samples.</li>
		<li>Spin the dissected samples on a mini benchtop centrifuge (2,000 &#215; g, 10 s), remove the PBST, and add 50-100 &#181;L of cold methanol (-20 &#176;C). 
		CAUTION: Methanol is toxic. Wear protective equipment and avoid inhalation.</li>
		<li>Mix by pipetting up and down and leave the samples in methanol for 30-60 s. Wash the samples 2x in 200 &#181;L of PBST. 
		NOTE: If processing more than four samples, proceed with the dissection of the remaining samples (steps 2.1.1 to 2.2.7).</li>
		<li>Wash the samples a third time with 200 &#181;L of PBST.</li>
	</ol>
	</li>
</ol>
3. Antibody incubations
<ol>
	<li>Blocking
	<ol>
		<li>Block the samples in 1x Blocking Solution (Table 1) for 45-60 min at room temperature. 
		NOTE: The incubation time can range from 30 min at room temperature to several days at 4 &#176;C (testing was done up to 3 days).</li>
	</ol>
	</li>
	<li>Primary antibody solution
	<ol>
		<li>Dilute anti-HTP-3 (chicken42) and anti-HA (mouse) antibodies (or the antibodies of choice) to the working solutions (1:250 for SMLM and 1:1,000 for STED microscopy samples) in 1x Blocking Solution. 
		NOTE: Antibodies used to label SMLM samples are more concentrated than for STED samples since a higher labeling density is recommended for SMLM microscopy.</li>
		<li>Spin the samples on a mini benchtop centrifuge (2,000 &#215; g, 10 s), remove the blocking buffer, and add 30-50 &#181;L of the primary antibody solution. Incubate overnight at 4 &#176;C (preferred) or for 1-2 h at room temperature.</li>
		<li>After incubation, wash 3 x 5-15 min with PBST.</li>
	</ol>
	</li>
	<li>Working solution of F(ab&#39;)2 fragments conjugated to fluorescent dye
	<ol>
		<li>Dilute the labeled F(ab&#39;)2 fragments (step 1.2.4) to the working solutions (1:100 for SMLM and 1:1,000 for STED microscopy samples) in 1x Blocking Solution. 
		NOTE: For both super-resolution techniques, previously reported fluorophore pairs were used, namely AlexaFluor647/CF680 for SMLM and AlexaFluor594/Abberior STAR635P&#160;for STED. AlexaFluor647 and STAR645P were used to label anti-mouse (Fab&#39;)2 fragments to target the C-terminus of SYP-5, and CF680/AlexaFluor594-labeled anti-chicken (Fab&#39;)2 fragments to target HTP-3.</li>
		<li>Spin the samples on a mini benchtop centrifuge (2,000 &#215; g, 10 s), remove the PBST, and add 30-50 &#181;L of secondary antibody solution. Incubate for 30 min to 2 h at room temperature (preferred) or overnight at 4 &#176;C. Wash 3 x 5-15 min with PBST.</li>
	</ol>
	</li>
</ol>
4. Mounting samples on a coverslip
<ol>
	<li>Postfixation 
	NOTE: Process samples individually through steps 4.1.1-4.2.1.
	<ol>
		<li>Spin down the stained samples and remove the supernatant. Add 50 &#181;L of PBST0.2 and transfer the stained worms onto a 22 mm x 22 mm No. 1 coverslip. 
		NOTE: Use fresh PBST0.2 with 0.2% nonionic detergent (Table 1) for this step to prevent the worms from adhering to the coverslip.</li>
		<li>Pipet 5.7-6.3 &#181;L of postfixative solution onto a poly-L-lysine coverslip. 
		NOTE: Poly-L-lysine coverslips stored at 4 &#176;C should be brought to room temperature first.</li>
		<li>Pipet off the dissected worms in the same volume (5.7-6.3 &#181;L) and transfer into the drop of fixative on the poly-L-lysine coverslip (Figure 2B). 
		NOTE: In this and the following step, it is very important to retain the dissected tissue in the center of the poly-L-lysine-coated coverslip. This is especially important if mounting the samples in a custom holder to fit the custom-built SMLM microscope used here (see step 5.1, Figure 2B).</li>
		<li>Cover the sample with a small coverslip (12 mm diameter, Figure 2B). Remove excess liquid using a small piece of filter paper (Figure 2B). Fix for 3-5 min in a dark chamber.</li>
	</ol>
	</li>
	<li>&#34;Freeze-cracking&#34;
	<ol>
		<li>Freeze the samples by placing them on an aluminum block in dry ice (Figure 2B). 
		NOTE: The aluminum block must be well cooled in the dry ice before placing the samples on it. Proceed with postfixation of the remaining samples (steps 4.1.1 to 4.2.1). The sample needs to be on dry ice for at least 20 min or up to 1 h prior to the next step (4.2.2).</li>
		<li>Remove the smaller coverslip using a razor (Figure 2B). 
		NOTE: For STED, proceed to step 5.2.1. For SMLM, continue with step 4.2.3.</li>
		<li>Dip the coverslip into a 50 mL conical tube containing ice-cold PBS (preferred) or -20 &#176;C methanol for approximately 10 s. 
		NOTE: Temperature is a very important factor for this step. Therefore, use PBS that is freshly thawed or kept in an ice/ethanol bath.</li>
		<li>Place the coverslip into a well of a six-well plate filled with PBST buffer. Remove the PBST from the wells and add fresh PBS. Leave the samples in PBS for 5 min. 
		NOTE: Pipet the PBS onto the side of the well to avoid damaging and detaching the samples.</li>
		<li>Wash with fresh PBS and leave the samples at 4 &#176;C until imaging. 
		NOTE: Samples are stable for up to 2 weeks, but the best results are achieved if the samples are imaged within 2 days.</li>
		<li>Before imaging, assess the quality of the sample mounting under a stereo microscope. 
		NOTE: Successfully mounted germlines are stably attached with no discernable movement relative to the coverslip. Poorly attached germlines will flap in the buffer solution.</li>
	</ol>
	</li>
</ol>
5. Imaging
<ol>
	<li>Single-molecule localization microscopy 
	NOTE: Images were acquired at the EMBL Imaging Centre using a custom-built single-molecule localization microscope that was constructed around a custom body, as previously reported38,43, with the unique features specified in the Table of Materials; refer to&#160;https://www.embl.org/about/info/imaging-centre
	<ol>
		<li>Acquiring 3D bead calibration
		<ol>
			<li>Prepare a precision coverslip (24 mm diameter; 0.17 &#177; 0.005 mm, No. 1.5) with adherent 100 nm fluorescent beads as described previously38,44.</li>
			<li>Place the calibration sample from step 5.1.1.1 onto a sample holder.</li>
			<li>Add a drop of immersion oil on the clean 100x/1.5 oil objective and mount the calibration sample on the microscope.</li>
			<li>Within MicroManager 245,46, specify 15-20 positions in the calibration sample.</li>
			<li>Within the EMU plugin window47, set up the acquisition of a z-stack image for each of the positions from step 5.1.1.5. 
			NOTE: Here, a compound cylindrical lens provides the astigmatism required for 3D imaging, and 201 z-slices were acquired for each position spanning the range between -1 &#181;m to 1 &#181;m, with an increment of 10 nm. A 2 kW/cm2 640 nm laser illumination was used for 25 ms for each z-slice.</li>
			<li>Acquire the z-stack images of the 100 nm fluorescent beads through an identical optical path that will be used to acquire sample images in step 5.1.11.</li>
			<li>Using the super-resolution microscopy analysis platform (SMAP48), generate a cspline model of the experimental point spread function (PSF) that will be used to fit the 3D-SMLM data in step 5.1.13.</li>
		</ol>
		</li>
		<li>Prepare the sample holder. For the custom-built holder used here that uses a magnetic ring to create the imaging chamber (Figure 2B), wrap the magnetic ring with parafilm. 
		NOTE: Alternatively, a microscope slide with a concave depression cavity can be used to mount samples for microscopes with slide holders.</li>
		<li>Prepare 1 mL of imaging buffer44 (Table 1).</li>
		<li>Take one coverslip from step 4.2.6 and place it in the custom-made holder. Fix the coverslip in the holder with the parafilm-wrapped magnetic ring (step 5.1.2).</li>
		<li>Gently pipet the imaging buffer (step 5.1.3) in the chamber created by the magnetic ring on top of the sample (Figure 2B). Seal the chamber with a piece of parafilm.</li>
		<li>To mount the sample, add one drop of immersion oil on the clean 100x/1.5 oil objective. Without introducing any air into the immersion oil, gently place the sample holder with the mounted sample (step 5.1.5) onto the microscope stage. 
		NOTE: Before placing the sample on the microscope, clean the bottom of the coverslip with tissue and 70% ethanol.</li>
		<li>Using the EMU plugin window47 within MicroManager 245,46, move the piezo stage until the signal from the focus lock laser is detected at the quadrant photodiode (QPD). 
		NOTE: To maintain a fixed focus across the imaging time, focus locking is achieved by total internal reflection of a near-infrared fiber-coupled laser from the coverslip and subsequent height sensitive detection on a quadrant photodiode (QPD). The QPD signal provided closed-loop control of the objective lens piezo mount.</li>
		<li>Acquire a back focal plane image with a 640 nm excitation laser at low power (i.e., 1-5%) to confirm the absence of air bubbles in the immersion oil. 
		NOTE: Remove the sample from the stage if an air bubble is detected. Clean the bottom of the coverslip and the objective and repeat steps 5.1.6-5.1.8. Otherwise, proceed to lock the focus within the EMU software47.</li>
		<li>Localize the gonad tissue using the brightfield illumination. Using a low-intensity 640 nm illumination, focus on the section of the tissue that contains many SC stretches. 
		NOTE: Do not focus on structures that are more than 2 &#181;m from the coverslip. Do not use a higher laser power to locate the sample, as this may turn some fluorophores into a blinking state prematurely. Here, 1 kW/cm2 was used in rising mode with a pulse set to 1,000.</li>
		<li>Proceed to expose the sample with 640 nm illumination at high irradiance (27 kW/cm2) for ~30 s until an appropriate blinking rate is achieved (Supplementary Video 1).</li>
		<li>Acquire 200,000 frames with a 20 ms exposure time using the multidimensional acquisition tool in MicroManager 245,46.</li>
		<li>Meanwhile, set up the UV activation using the activation option of the EMU plugin38,47 to maintain the desired blinking rate. 
		NOTE: Use the UV laser at a 3 kW/cm2 irradiance in rising mode with maximum pulse length set to 10,000.</li>
		<li>Perform SMLM image reconstruction and postprocessing. 
		NOTE: To reconstruct images from raw SMLM data, refer to published methods. The data presented here was processed using the SMAP48,49 software. Super-resolved image reconstruction, channel assignment, drift correction, and filtering of localizations with poor localization precision and a maximum likelihood filter were performed in the SMAP48 software.</li>
	</ol>
	</li>
	<li>Stimulated emission depletion microscopy 
	NOTE: Images were acquired on the integrated STED microscopic system equipped with a white light laser, a 775 nm pulsed STED laser, and the FALCON Fluorescence Lifetime IMaging module (Table of Materials) at the EMBL Imaging Centre (https://www.embl.org/about/info/imaging-centre).
	<ol>
		<li>Place a 20 &#181;L drop of mounting medium (Table of Materials) onto a microscope slide. Take one coverslip from step 4.2.2 and place the sample gently onto the slide facing the mounting medium (Figure 2B). 
		NOTE: Avoid introducing air pockets within the mounting medium.</li>
		<li>Let the mounting medium cure overnight. 
		NOTE: Image the samples on the following day or keep them at 4 &#176;C until imaging.</li>
		<li>To mount the sample, add one drop of immersion oil on the coverslip of the sample from the step 5.2.2. Gently place the sample onto the microscope stage using a 100x/1.40 oil objective.</li>
		<li>Focus on the sample and locate the germline tissue using the bright-field illumination.</li>
		<li>Using the microscope software, specify the region of interest for which the TauSTED image will be acquired.</li>
		<li>Select the excitation lasers and their appropriate power used to excite the fluorophores used in the sample. 
		NOTE: Here, the 580 nm laser at 4% power was used to image AlexaFluor 594-conjugated F(ab&#39;)2 secondary antibody fragments, and 635 nm at 3% power to image STAR 635P-conjugated F(ab&#39;)2 fragments.</li>
		<li>Using the microscope software, select an appropriate STED depletion laser power and set up the image detection. 
		NOTE: Here, the 775 nm STED depletion laser power was set to 40%. The detector was used in counting mode with a gain value of 10 for photon detection, with a scan rate of 100 Hz, and at a pixel size of 17 nm. Four-line accumulation was used for TauSTED acquisition.</li>
	</ol>
	</li>
</ol>

Authors declare no conflicts of interest.

The ladder-like organization of the SC, which is essential for the correct recombination and segregation of homologous chromosomes, was first observed almost 70 years ago in electron microscopy3,4. While the overall organization of the SC is readily resolved in electron microscopy, the localization of individual components within this complex requires a more targeted approach. With its width of only ~100 nm, the substructure of the SC cannot be resolved by conventional fluorescence microscopy. However, super-resolution microscopy has become a major driver for novel discoveries on the structure and function of the synaptonemal complex16,19,24,25,26,27,28,29,30. To facilitate this research, we have demonstrated a mounting procedure that allows studying the architecture of the SC within C. elegans gonad tissue with SMLM and STED microscopy.
A critical step to optimize the resolution in SMLM imaging is directly cross-linking the germline tissue to a poly-L-lysine coated coverslip (step 4). The covalent attachment of the tissue to the coverslip is essential to reduce movements within the sample that would result in large drifts and make imaging over long periods of time for SMLM impossible. Additionally, even a suboptimal attachment that leaves the nuclei containing SCs at a distance from the coverslip leads to a significant drop in the achievable resolution resulting from spherical aberrations (Figure 4). Alternatively to the covalent attachment used here, stained germline tissue can also be immobilized between two sealed coverslips in a small drop of imaging buffer19,30. However, this immobilization method severely reduces the volume of imaging buffer in the sample from 1 mL used in the optimized protocol here to just a few &#181;L, which will result in an acidification of the imaging buffer and severely reduce the time for which the sample can be imaged38,53,54.
Long acquisition times for both SMLM and STED microscopy limit the use of these methods to imaging of chemically fixed samples. Here, paraformaldehyde fixation ensures that the structure of the SC is preserved during sample preparation and imaging. However, despite the precautions taken here to image the SC within intact tissue, the resulting structure of the SC after fixation is not necessarily identical to the structure in its native state within a living organism. Moreover, since a single image of the fixed SC represents a single &#34;snapshot&#34; of the biological structure, this approach remains blind to the dynamics of the native structure in vivo.
However, information on the dynamics and variability of macromolecular structures can also be obtained by acquiring not just a single but many &#34;snapshots&#34;. While this approach can resolve changes in the structure of the SC during pachytene19, there are several factors that limit the number of images that can be acquired from a single sample prepared using this protocol. First, the high laser powers used during image acquisition lead to permanent bleaching of the fluorophores and preclude imaging of adjacent regions of interest or multiple z-planes, thereby significantly reducing the number of images that can be acquired from a single sample. Second, the sample/tissue density on the coverslip prepared by this method is low, which significantly limits the number of images that can be acquired from a single coverslip. The low sample density also prohibits the use of automated image acquisition pipelines that helped shed light on other biological questions34,55,56,57,58,59. However, the sample density can be increased slightly by an experienced user.
The protocol presented here is optimized to obtain a high labeling density that is necessary to achieve optimal resolution in SMLM35. While previous protocols covalently attach the tissue to the coverslip before immunostaining16, this new protocol cross-links the tissue to the coverslip only after the samples were stained in solution. This modification allows the antibodies used for immunolabeling to freely access the tissue from all sides, while the covalent attachment of the tissue to the coverslip may restrict antibodies from reaching the nuclei closest to the coverslip, thereby reducing the degree of labeling. Together, the modifications described here improve the resolution from 40-50 nm (FRC resolution)16 to 30-40 nm (this protocol).
Importantly, while a high labeling density and a high concentration of antibodies is essential for SMLM, we found that better STED microscopy images are obtained using lower antibody concentrations (step 3). At a resolution of tens of nanometers, the size of the molecules used to label the protein of interest becomes increasingly important. We therefore employed F(ab&#39;)2 fragments that are half the size of full-length antibodies. The improvement in local contrast due to a smaller signal source, and therefore resolution gained by this modification compared to using full-length secondary antibodies, allowed the resolution of the two C-termini of SYP-5 within the central region by TauSTED, which are not resolved by conventional STED using full-length antibodies (16 and data not shown). We anticipate that this optimized protocol for imaging SCs in intact C. elegans germlines will facilitate investigating the structure-function relationship of the SC during meiosis.

Reducing the number of chromosomes by half during meiosis is key to generating healthy progeny in sexually reproducing organisms. To achieve this reduction in chromosome number, homologous chromosomes must pair and segregate during meiosis I. To ensure the accurate segregation of homologous chromosomes, germ cells undergo an extended prophase I, during which homologous chromosomes pair, synapse, and recombine to generate physical links between homologs1. The SC has emerged as the central structure that is key in regulating the correct progression through meiotic prophase2.
The SC is a complex whose general structure is evolutionarily conserved, even though there is little homology between its protein components. The SC was first identified in electron micrographs as a tripartite, ladder-like structure consisting of two lateral elements or axes, a central region formed by transverse filaments, and a central element3,4. Determining the organization of individual components within the complex is key to advancing our understanding of the SC&#39;s role during meiotic prophase.
The model organism C. elegans is ideally suited to study the structure and function of the SC since its germlines contain a large number of meiotic nuclei with fully assembled SCs5. Genetic and biochemical studies have revealed that the chromosome axes are formed by three distinct cohesin complexes6,7 and four HORMA domain proteins called HTP-1/2/3 and HIM-37,8,9,10,11 in C. elegans. In the central region of the SC, six proteins containing coiled-coil domains have been identified to date12,13,14,15,16,17. To bridge the distance between the two axes, SYP-1, -5, and -6 dimerize in a head-to-head manner (Figure 1), while three additional proteins stabilize their interaction in the central element16,17,18,19.
Obtaining detailed insight into the organization of these proteins is essential in understanding the SC&#39;s many functions during meiosis. Since the width of the central region of the SC is only ~100 nm, its substructure cannot be resolved by diffraction-limited fluorescence microscopy. However, visualizing components within a structure of this size is readily achievable by super-resolution microscopy. Indeed, structured illumination microscopy (SIM), expansion microscopy20, stimulated-emission depletion (STED) microscopy21, and single-molecule localization microscopy (SMLM)22,23 have emerged as essential tools to study the molecular architecture of the SC across species16,24,25,26,27,28,29,30.
To overcome the resolution limit, STED microscopy relies on overlaying the diffraction-limited spot of the emission light with a donut-shaped beam from the STED laser, which theoretically constricts the point spread function down to molecular dimensions31,32. However, the resolution that is practically achievable by STED within biological samples remains in the range of a few tens of nanometers in xy33.
Even higher resolution in biological samples can be obtained with SMLM techniques. SMLM harnesses the blinking properties of specific fluorophores to resolve objects at the sub-diffraction level by separating spatially overlapping fluorophores in time. The sample is then imaged repeatedly to capture different subsets of fluorophores. The position of the fluorophores within the sample is then determined by fitting the point spread function (PSF) to the obtained signals across all images, which can resolve structures down to 15 nm23,34.
Taken together, the localized images encode the positions of all the fluorophores. The resolution of SMLM is determined by the labeling density and the blinking characteristics of the fluorophore. According to the Nyquist-Shannon criterion, it is impossible to reliably resolve objects that are less than twice the average label-to-label distance. Thus, a high labeling density is needed for high-resolution imaging. For the SC in C. elegans, a high labeling density can be achieved by using epitope tags attached to specific sites of endogenous proteins using genome editing. The epitope tags can then be stained at a high density using specific monoclonal antibodies with high affinities19,30. At the same time, the on-cycle of individual fluorophores must be short enough to ensure that spatially overlapping fluorophores are not captured at the same time35.
Due to these two requirements, resolving the structure of big macromolecular complexes such as the SC requires imaging a sufficiently large number of images, and can thus take several hours. The pitfall of long imaging times is that samples tend to drift due to movement of the stage or small currents within the sample buffer; even small movements in the order of 10 nm are detrimental at nm resolution and must be corrected for. However, the drift correction methods commonly used are not robust enough to accurately overlay images of two channels imaged sequentially36. This is problematic because biological questions often ask for precise detection and localization of multiple targets within the same sample. To circumvent these issues, methods such as ratiometric imaging have been developed. Ratiometric imaging allows for the simultaneous imaging of multiple fluorophores with overlapping excitation and emission spectra, with a subsequent assignment of each detected signal to its respective fluorophore based on the ratio of intensities in spectrally distinct channels37,38.
Additionally, studying the organization of macromolecular complexes such as the SC calls for three-dimensional (3D) information. To achieve super-resolution in three dimensions (3D-SMLM), a cylindrical lens is incorporated in the optical path of the emitted light that distorts the shape of the PSF of a fluorophore depending on its distance from the focal plane. Hence, the precise position of a fluorophore in the z-plane can be extrapolated by analyzing the shape of its emission signal35,39. Combining these advances in SMLM allows for imaging of the 3D organization of macromolecular complexes, including the SC.

<table><tbody><tr><td>100x/1.5 oil objective</td><td>Olympus</td><td>UPLAPO100XOHR</td><td>UPLAPO100XOHR</td></tr><tr><td>2-mercaptoethylamine (MEA)</td><td>Sigma-Aldrich</td><td>30070-10G</td><td>Dissolved in MilliQ water to 5 M solution, pH&amp;nbsp; 8.7 adjusted with HCl. Aliquoted to a single-use volume, frozen, and kept at -80 &amp;deg;C.</td></tr><tr><td>Additional 640 nm booster laser</td><td>Toptica</td><td>IBEAM-SMART-640-S-HP</td><td></td></tr><tr><td>AlexaFluor 594, NHS ester</td><td>ThermoFischer Scientific</td><td>A37572</td><td>Dissolved in DMSO to 1 mM solution, aliquoted to single use volume, frozen and kept at -80 &amp;deg;C</td></tr><tr><td>AlexaFluor 647, NHS ester</td><td>ThermoFischer Scientific</td><td>A37573</td><td>Dissolved in DMSO to 1 mM solution, aliquoted to single use volume, frozen and kept at -80 &amp;deg;C</td></tr><tr><td>anti-HA</td><td>Thermo Fisher Scientific</td><td>2-2.2.14</td><td>Mouse monoclonal, 1:250 (SMLM), 1:1,000 (STED Microscopy)</td></tr><tr><td>anti-HTP-3</td><td>a gift from Abby F. Dernburg</td><td>MacQueen et al., 2005</td><td>Chicken polyclonal, 1:250 (SMLM), 1:1,000 (STED Microscopy)</td></tr><tr><td>&lt;em&gt;Caenorhabditis elegans&lt;/em&gt; strain&amp;nbsp; YKM349</td><td>a gift from Yumi Kim</td><td>Hurlock et al., 2020</td><td>syp-5(kim9[syp-5::HA]) I; meIs8[pie-1p::GFP::cosa-1, unc-119(+)] II</td></tr><tr><td>CF6680, NHS ester</td><td>Biotium</td><td>92139</td><td>Dissolved in DMSO to 1mM solution, aliquoted to single use volume, frozen and kept at -80 &amp;deg;C</td></tr><tr><td>Circular cover glass 12 mm No. 1</td><td>Menzel-Gl&amp;auml;ser; VWR</td><td>631-0713</td><td></td></tr><tr><td>Circular cover glass 24 mm No. 1.5</td><td>Carl Roth</td><td>PK26.1</td><td></td></tr><tr><td>Cylindrical lenses</td><td>Thorlabs</td><td>LJ1516RM-A, LK1002RM-A</td><td></td></tr><tr><td>Egg Buffer (10x)</td><td></td><td>Edgar 1995</td><td>250 mM HEPES, 1.18 M NaCl, 480 mM KCl, 20 mM EDTA, 5 mM EGTA, pH 7.4</td></tr><tr><td>Ethanol (absolute for analysis)</td><td>Merck</td><td>64-17-5</td><td></td></tr><tr><td>F(ab&amp;rsquo;)2 fragment anti-chicken IgY</td><td>Jackson Immunoresearch</td><td>AB_2340347</td><td>Donkey polyclonal, 1:100 (SMLM), 1:1,000 (STED Microscopy)</td></tr><tr><td>F(ab&amp;rsquo;)2 fragment anti-mouse IgG</td><td>Jackson Immunoresearch</td><td>AB_2340761</td><td>Donkey polyclonal, 1:100 (SMLM), 1:1,000 (STED Microscopy)</td></tr><tr><td>Fisherbrand Microscope slides T/F Ground 0.8-1.0 mm thick</td><td>Fisher scientific</td><td>7107</td><td></td></tr><tr><td>Gauge Worm Pick 30 diameter 0.254 mm - Iridium 10%</td><td>Kisker</td><td>789265</td><td></td></tr><tr><td>Glucose oxidase/Catalase enzyme mix (GlOX/Cat )</td><td>a gift from Jonas Ries</td><td>Hoess, Mund, Reitberger, &amp;amp; Ries, 2018</td><td>20x, 1916 U/mL glucose oxidase (Sigma G7141), 42350 U/mL catalase (Sigma C3155),&lt;br/&gt; 50 mM Tris-HCl pH 8.0, 51% glycerol, MilliQ water. Stored at -20 &amp;deg;C.</td></tr><tr><td>Imaging buffer base</td><td>a gift from Jonas Ries</td><td>Hoess, Mund, Reitberger, &amp;amp; Ries, 2018</td><td>50 mM Tris-HCl, pH 8.0, 10 mM NaCl, 10% D-Glucose.&lt;br/&gt; Aliquoted to a single-use volume (950 &amp;mu;L), frozen, and kept at -80 &amp;deg;C.</td></tr><tr><td>Invitrogen ProLong Glass Antifade Mountant</td><td>ThermoFischer Scientific</td><td>P36982</td><td></td></tr><tr><td>Leica Stellaris 8 STED FALCON</td><td>Leica</td><td>N/A</td><td>The microscope is equiped with the latest generation white light laser, a 775nm pulsed STED laser, the FALCON &lt;strong&gt;F&lt;/strong&gt;luorescence &lt;strong&gt;L&lt;/strong&gt;ifetime &lt;strong&gt;IM&lt;/strong&gt;aging module, HC PL APO CS2 100x/1.40 oil objective, and Leica HyD X detector. The system is capable of FLIM module enhanced Tau-STED which measures the specific fluorescence lifetime of a dye and is therefore capable of removing background signal based on differences in fluorescence lifetimes of the dyes, and dye conditions in the sample.&amp;nbsp; Additionally, the resolution is increased by accounting for the variation of fluorescence lifetimes in different areas of the depletion donut.</td></tr><tr><td>Longwave channel emission filter</td><td>AHF Analysentechnik</td><td>F47-702</td><td>700/100 nm bandpass</td></tr><tr><td>Methanol (absolute for analysis)</td><td>Merck</td><td>67-56-1</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma-Aldrich/Merck</td><td>S5761-500G</td><td>100 mM NaHCO&lt;sub&gt;3&lt;/sub&gt;, pH 8.3</td></tr><tr><td>Near-infrared fiber-coupled laser</td><td>Toptica</td><td>IBEAM-SMART-PT-CD</td><td>Custom Design, 808 nm - 75mW</td></tr><tr><td>Objective lens piezo mount (PIFOC )</td><td>Physik Instrumente</td><td>P-726.1.CD</td><td>100 &amp;micro;m travel range</td></tr><tr><td>Orca Fusion BT sCMOS camera</td><td>Hamamatsu</td><td>C15440-20UP</td><td></td></tr><tr><td>PCR tubes</td><td>Greiner Bio-One</td><td>673283</td><td>0.2 mL</td></tr><tr><td>Phosphate Saline Buffer (PBS 10x)</td><td></td><td>N/A</td><td>137 mM NaCl, 2.7 mM KCl, 10 mM Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 1.8 mM KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, pH 7.4</td></tr><tr><td>Pierce 16% formaldehyde (w/v), methanol free</td><td>ThermoFischer Scientific</td><td>28906</td><td>16% formaldehyde is transferred from the original glass ampule into 1.5 mL tube and kept at room temperature.</td></tr><tr><td>PlasmaPrep2 plasma cleaner</td><td>GaLa Instrumente GmbH</td><td>N/A</td><td></td></tr><tr><td>Poly-L-lysine hydrobromide</td><td>Sigma-Aldrich/Merck</td><td>P2636-25MG</td><td>0.1% w/v solution was prepared in Milli-Q water, and stored in aliquots at -20 &amp;deg;C.</td></tr><tr><td>Primary dichroic (illumination reflecting)</td><td>AHF Analysentechnik</td><td>F73-866S</td><td>quad bandpass @ 405, 488, 561, 640</td></tr><tr><td>Quadnotch filter</td><td>AHF Analysentechnik</td><td>F40-072</td><td>405/488/561/640 nm</td></tr><tr><td>Quadrant photodiode (QPD)</td><td>Laser Components</td><td>SD197-23-21-041, LC301DQD-PV</td><td></td></tr><tr><td>Razor blades</td><td>Apollo Herkenrath Solinger</td><td>N/A</td><td></td></tr><tr><td>Refractive beam shaper</td><td>AdlOptica</td><td>PiShaper 6_6_VIS</td><td></td></tr><tr><td>Roche Blocking Reagent</td><td>Roche</td><td>11096176001</td><td>10x solution was prepared according to recommendation. Frozen aliquots were stored at -20 &amp;deg;C.</td></tr><tr><td>Scalpel blade (Feather brand #11, No. 3)</td><td>Heinz Herenz Medizinalbedarf GmbH</td><td>1110911</td><td></td></tr><tr><td>Scalpel removal box</td><td>Fisher scientific</td><td>10002-50</td><td></td></tr><tr><td>Secondary dichroic (emission reflecting)</td><td>AHF Analysentechnik</td><td>F38-785S</td><td>750 nm longpass</td></tr><tr><td>Shortpass filter</td><td>Semrock</td><td>BSP01-785R-25</td><td>&amp;nbsp;750 nm</td></tr><tr><td>Shortwave channel emission filter</td><td>AHF Analysentechnik</td><td>F37-677</td><td>&amp;nbsp;676/37 nm bandpass</td></tr><tr><td>Single molecule localization microscope</td><td>EMBL Imaging Centre</td><td>Diekmann et al., 2020 with modifications</td><td>The microscope provides widefield epi-illumination via a single-mode fiber-coupled laser engine, additional booster laser, and refractive beam shaper to provide a uniform illumination field (Stehr et al, 2019). Widefield images are captured on a sCMOS camera&amp;nbsp; and appropriate relay optics for a system magnification of 61x and a pixel size of 106 nm.&lt;br/&gt; For ratiometric imaging of spectrally overlapping far-red dyes, an image splitter produces two spectrally distinct images on the camera (splitting dichroic: 665 nm long pass, shortwave channel emission filter: 676/37 nm bandpass, longwave emission filter: 700/100 nm bandpass. An additional 405/488/561/640 nm quadnotch filter and 750 nm shortpass filter are common to the two paths and provide additional laser blocking).&lt;br/&gt; A compound cylindrical lens provides the astigmatism required for 3D imaging.&lt;br/&gt; To maintain a fixed focus across acquisitions exceeding 2 hours in time (comprising 200 000 - 250 000 images), focus locking is achieved by total internal reflection of a near-infrared fiber-coupled laser&amp;nbsp; from the coverslip and subsequent height sensitive detection on a quadrant photodiode (QPD). The QPD signal provided closed-loop control of the objective lens piezo mount.&lt;br/&gt; For access to this microscope, refer to https://www.embl.org/about/info/imaging-centre or contact ic-contact@embl.de</td></tr><tr><td>Single-mode fiber-coupled multi-laser engine</td><td>Toptica</td><td>iCHROME MLE-LFA-HP</td><td>Provides widefield epi-illumination of 100 mW at 405, 488, 561, 640 nm</td></tr><tr><td>Splitting dichroic</td><td>AHF Analysentechnik</td><td>F48-665SG</td><td>665 nm long pass</td></tr><tr><td>Square cover glass 22 x 22 mm No.1</td><td>Menzel-Gl&amp;auml;ser; VWR</td><td>630-2882</td><td></td></tr><tr><td>STAR 635P, NHS ester</td><td>Abberior</td><td>ST635P-0002-1MG</td><td>Dissolved in DMSO to 1 mM solution, aliquoted to single use volume, frozen and kept at -80 &amp;deg;C</td></tr><tr><td>Stereo microscope Stemi 305 Stand K LAB</td><td>Zeiss</td><td>N/A</td><td></td></tr><tr><td>Tetramisole hydrochloride</td><td>Sigma-Aldrich/Merck</td><td>T1512-2G</td><td>1% (w/v) solution was prepared in Milli-Q water. Frozen aliquots were stored at -20 &amp;deg;C.&lt;br/&gt; Thawed aliquot was kept at 4 &amp;deg;C and used for several months.</td></tr><tr><td>TetraSpeck Microspheres</td><td>ThermoFischer Scientific</td><td>T7279</td><td>&amp;nbsp;0.1 &amp;micro;m, fluorescent blue/green/orange/dark red</td></tr><tr><td>Tris Saline Buffer (TBS 10x)</td><td></td><td>N/A</td><td>200 mM Tris-HCl, 1.5 M NaCl, pH 7.5</td></tr><tr><td>TWEEN 20</td><td>Sigma-Aldrich/Merck</td><td>P9416-50ML</td><td>Kept at room temperature in original packaging.</td></tr><tr><td>WormStuff worm pick</td><td>Kisker</td><td>789277</td><td></td></tr><tr><td>XY microscope stage</td><td>&amp;nbsp;Smaract</td><td>N/A</td><td>Custom Design</td></tr><tr><td>Zeba Micro Spin Desalting Column</td><td>ThermoFischer Scientific</td><td>89877</td><td>&amp;nbsp;7K MWCO, 75 &amp;micro;L</td></tr></tbody></table>

super-resolution microscopy of the synaptonemal complex within the caenorhabditis elegans germline

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures the interaction of homologous chromosomes is the assembly of the synaptonemal complex (SC) in meiotic prophase I. Even though there is little sequence homology between protein components within the SC among different species, the general structure of the SC has been highly conserved during evolution. In electron micrographs, the SC appears as a tripartite, ladder-like structure composed of lateral elements or axes, transverse filaments, and a central element. 
However, precisely identifying the localization of individual components within the complex by electron microscopy to determine the molecular structure of the SC remains challenging. By contrast, fluorescence microscopy allows for the identification of individual protein components within the complex. However, since the SC is only ~100 nm wide, its substructure cannot be resolved by diffraction-limited conventional fluorescence microscopy. Thus, determining the molecular architecture of the SC requires super-resolution light microscopy techniques such as structured illumination microscopy (SIM), stimulated-emission depletion (STED) microscopy, or single-molecule localization microscopy (SMLM). 
To maintain the structure and interactions of individual components within the SC, it is important to observe the complex in an environment that is close to its native environment in the germ cells. Therefore, we demonstrate an immunohistochemistry and imaging protocol that enables the study of the substructure of the SC in intact, extruded Caenorhabditis elegans germline tissue with SMLM and STED microscopy. Directly fixing the tissue to the coverslip reduces the movement of the samples during imaging and minimizes aberrations in the sample to achieve the high resolution necessary to visualize the substructure of the SC in its biological context.

To image the SC within the C. elegans germline tissue by SMLM, we have employed 2-color ratiometric 3D-SMLM to localize HTP-3, a component of the chromosome axes, and the C-terminus of the transverse filament SYP-5 endogenously tagged with a hemagglutinin (HA) tag. The location of both proteins within the SC of C. elegans was previously determined by other studies16,30.
To minimize light scattering and optical aberrations inherent in thick biological samples, we imaged the bottom-most z-section of meiotic nuclei that contain the SCs (Figure 3, yellow lines). For each acquired image, the piezo stage position of the imaging plane was marked relative to the piezo stage position when the objective was focused on the coverslip. This allowed the calculation of the piezo distance from the coverslip. Successfully mounted samples are stably attached close to the coverslip and retain the gonad shape (i.e., the tissue is not crushed between the two coverslips during the postfixation step). The quality of the sample mounting can readily be assessed under a stereo microscope since well-attached gonads do not show any movement in solution (step 4.2.6). Nevertheless, due to the stochasticity of the mounting process, the gonad tissue will not necessarily be laid out completely flat on the coverslip. Therefore, the bottom plane of the nuclei containing SCs may be found at varying distances relative to the coverslip within the same gonad.
To illustrate how the resolution changes depending on the attachment of the tissue to the coverslip, we acquired images at different piezo distances to the coverslip. To assess the quality of an individual image, Fourier ring correlation (FRC) curves50,51 were calculated and the resolution was determined using the FRCResolution plugin within the SMAP software48. Two representative nuclei extracted from two separate 3D-SMLM images taken at different distances to the coverslip are displayed in Figure 4. In SCs located close to the coverslip, the chromosome axes and the C-terminus of SYP-5::HA are well resolved in all three dimensions (Figure 4A, 0.8 µm from the coverslip). To resolve two structures separated by a given distance, the achieved FRC resolution generally must be smaller than half this distance in the axial resolution.
To separate the same structures laterally, even smaller FRC resolution values need to be achieved. Indeed, in samples that are located in close proximity to the coverslip, the FRC resolution is 38 nm for the AlexaFluor 647 channel and 34 nm for the CF680 channel, and thus well below the expected distance of 84 nm between the C-termini of SYP-516. This resolution therefore readily resolves the organization of the SC not only in frontal but also in lateral views (Figure 4B i,ii). By contrast, the resolution deteriorates in SCs located at a 5 µm distance from the coverslip due to light scattering and spherical aberrations (Figure 4B). The FRC resolutions at this distance drop to 47 nm (AlexaFluor 647) and 41 nm (CF680), which cannot fully resolve the C-termini of SYP-5. Since the optical aberrations impair the lateral resolution more severely than the axial resolution, HTP-3 and SYP-5 bands are no longer clearly resolved in the cross-section of the lateral view in samples located at a 5 µm distance from the coverslip (Figure 4B ii). Comparing the FRC resolution of images acquired at different piezo distances from the coverslip revealed that the imaged tissue should be no further than 2 µm from the coverslip (Figure 5). This result highlights the importance of correct execution of the postfixation step, during which the tissue must be successfully crosslinked to the poly-L-lysine coating of the coverslip.
To demonstrate the achievable resolution with another super-resolution technique, we also imaged SCs in fixed intact germline tissue with TauSTED microscopy. Figure 6A shows TauSTED images with the highest and lowest resolution achieved within this study as estimated from line profiles of the SC in frontal view (Figure 6B). In both nuclei, we could resolve the two localization bands of HTP-3 in the chromosome axes and the C-termini of SYP-5 in the central region, demonstrating that the resolution achievable in TauSTED using this optimized protocol is below 84 nm. Under optimal conditions (Figure 6A, top), we could resolve the C-termini in slightly tilted views of the SC that were separated by only 50 nm (Figure 6A, yellow rectangle and 6C).
<img alt="Homologous chromosome structure diagram, transverse filaments, axes; frontal, lateral, cross-sectional views." class="xfigimg" src="/files/ftp_upload/64363/64363fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Schematic of the organization of the synaptonemal complex in Caenorhabditis elegans. The cartoon shows a simplified structure of the SC in C. elegans bridging two homologous chromosomes (gray). The structure is shown in frontal, lateral, and cross-sectional views. Chromosome axes are displayed as red bars while transverse filaments are shown in cyan. Transverse filament proteins (SYP-1, 5, 6 in C. elegans) are oriented in a head-to-head manner (cyan ball-stick graphics) in the central region to bridge the distance between the two axes. The expected distances between axes and the C-termini of transverse filaments are indicated. Abbreviation: SC = synaptonemal complex. <a href="https://www.jove.com/files/ftp_upload/64363/64363fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Dissection and microscopy workflow diagram for C. elegans gonad preparation using SMLM and STED techniques." class="xfigimg" src="/files/ftp_upload/64363/64363fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Illustration of the sample preparation used in the study. (A) Young C. elegans adults are dissected at their head or tail (green, dashed lines) and processed as described in the protocol. (B) Individual steps of the method are indicated with graphics that are connected with gray arrows. Abbreviations: STED = stimulated-emission depletion; SMLM = single-molecule localization microscopy; PBS = phosphate-buffered saline. <a href="https://www.jove.com/files/ftp_upload/64363/64363fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Fluorescence microscopy image of cell nuclei arrangement; HTP-3, SYP-5:HA markers; xy, xz view." class="xfigimg" src="/files/ftp_upload/64363/64363fig03.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Location of the tissue section that can be observed by single molecule localization microscopy. MIP of a spinning disk confocal image of a whole mount C. elegans gonad. The tissue was stained for HTP-3 and the C-terminus of SYP-5 (SYP-5::HA), and the combined signal is shown in gray. Individual confocal images were stitched using the Grid/Collection stitching Fiji plugin52 to create an image of the whole gonad. The inset shows an xy view of the bottommost z-plane containing the SCs. The localization of this plane is shown in orthogonal views of the tissue section indicated by a rectangle in the MIP image of the gonad (yellow lines). Scale bars = 10 µm. Abbreviations: MIP = maximum intensity projection; SCs = synaptonemal complexes. <a href="https://www.jove.com/files/ftp_upload/64363/64363fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Chromosome imaging; fluorescence microscopy; piezo distance 0.8 vs 5.0 μm; cross-section views." class="xfigimg" src="/files/ftp_upload/64363/64363fig04.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: Single-molecule localization microscopy of HTP-3 and the C-termini of the SYP-5. (A,B) Left: SMLM images showing pachytene nuclei stained for HTP-3 (red) and the C-terminus of SYP-5 (SYP-5::HA, cyan) (scale bar = 1 µm). Center: Zoomed-in images of regions of interest that are indicated in A and B with corresponding cross-sectional views displayed below each image (i, ii; scale bar = 100 nm). The stretches of the SC within zoomed-in images are rotated to orient the chromosome axes parallel to the y-axis. Right: Graphical representation of the localization of the proteins of interest within the SC portraying the orientation of the SC in the zoomed-in regions displayed in the center of the figure. Abbreviations: SMLM = single-molecule localization microscopy; SC = synaptonemal complex. Raw data to reconstruct SMLM images are available through the BioStudies database60 (Accession ID: S-BIAD504). <a href="https://www.jove.com/files/ftp_upload/64363/64363fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="FRC vs. spatial frequency graph for AlexaFluor647 and CF680; piezo distance, FRC resolution inset." class="xfigimg" src="/files/ftp_upload/64363/64363fig05.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: The Fourier ring correlation resolution of single-molecule localization microscopy images depends on the distance of the imaged z-plane from the plane of the coverslip. Colored lines show FRC curves of images acquired at different distances (as depicted by the color bar) from the coverslip. The 1/7 threshold used to determine the FRC resolution is indicated by a black horizontal line. Insets show the dependence of the FRC resolution on the piezo distance from the coverslip. Plotting was performed by a custom-written R script (version 4.1.2, Supplementary File 1) in which original curves were smoothed with functions from the "ggplot2" package. Abbreviations: FRC = Fourier ring correlation; SMLM = single-molecule localization microscopy; SC = synaptonemal complex. Data for FRC curves and SMLM data are available through the BioStudies database60 (Accession ID: S-BIAD504). <a href="https://www.jove.com/files/ftp_upload/64363/64363fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Microscopy images with protein localization of HTP-3, SYP-5; fluorescence intensity graphs shown." class="xfigimg" src="/files/ftp_upload/64363/64363fig06.jpg" data-altupdatedat="1747911167000" data-alt="Figure 6"> 
Figure 6: Stimulated emission depletion microscopy enhanced by fluorescence lifetime-based information (TauSTED) resolves two localization bands for both HTP-3 and the C-terminus of SYP-5. (A) Two representative TauSTED images show pachytene nuclei stained for HTP-3 (red) and the C-terminus of SYP-5 (SYP-5::HA, cyan) with higher (top) and lower (bottom) structural definition (scale bar = 1 µm). The rectangles mark regions with the resolved C-termini of SYP-5 in frontal (white) and a slightly tilted view (yellow) of the SC. (B,C) Distribution of the HTP-3 (red) and the C-terminus of SYP-5 (cyan) signal resolved by TauSTED. Line profiles of regions of interest that contain the SC in frontal (B) or slightly tilted (C) views are shown as full lines with intensity normalized to the maximum value. Line profiles were generated using Fiji ImageJ. Dashed lines in B show the averaged data for each protein. The thick cyan line in C corresponds to the line profile with the shortest resolved distance between the C-termini of SYP-5. To determine the distances between the antibodies targeting specific proteins, the line profiles (n = 9 (B), n = 7 (C)) were fitted with double gaussians using a custom-written R script (version 4.1.2, Supplementary File 1). Mean distance ± standard deviation (B) and the range with minimum value highlighted in bold (C) are indicated on top of each graph, respectively. Abbreviations: STED = stimulated emission depletion microscopy; SC = synaptonemal complex. Displayed images and data points of plotted line profiles are available through the BioStudies database60 (Accession ID: S-BIAD504). <a href="https://www.jove.com/files/ftp_upload/64363/64363fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Composition of buffers and solutions used in this protocol. <a href="https://www.jove.com/files/ftp_upload/64363/63463_Table%201%20(1).xlsx" target="_blank">Please click here to download this Table.</a>
Supplementary Video 1: Single molecule localization microscopy acquisition. Video showing fluorophores blinking at an appropriate rate (50 frames are shown, scale bar = 5 µm, 20 ms/frame). <a href="https://www.jove.com/files/ftp_upload/64363/SupplementalVideo1.mov" target="_blank">Please click here to download this Video.</a>
Supplementary File 1: Data analysis script. <a href="https://www.jove.com/files/ftp_upload/64363/SupplementalFile1.Rmd" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline at JoVE.com

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

Super-resolution microscopy can provide a detailed insight into the organization of components within the synaptonemal complex in meiosis. Here, we demonstrate a protocol to resolve individual proteins of the Caenorhabditis elegans synaptonemal complex.

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures ...

super-resolution-microscopy-synaptonemal-complex-within

Nanyang Technological University

Research

JoVE Journal

Biology

2.5K Views.  European Molecular Biology Laboratory. Super-resolution microscopy can provide a detailed insight into the organization of components within the synaptonemal complex in meiosis. Here, we demonstrate a protocol to resolve individual proteins of the Caenorhabditis elegans synaptonemal complex. 

Video: Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

The C. elegans germline makes an excellent model for studying meiosis, in part due to the ease of conducting cytological analyses on dissected animals. Whole mount preparations preserve the structure of meiotic nuclei, and importantly, each gonad arm contains all stages of meiosis, organized in a temporal-spatial progression that makes it easy to identify nuclei at different stages. Adult hermaphrodites have two gonad arms, each organized as a closed tube with proliferating germline stem cells at the distal closed end and cellularized oocytes at the proximal open end, which join in the center at the uterus. Dissection releases one or both gonad arms from the body cavity, allowing the entirety of meiosis to be visualized. Here, a common protocol for immunofluorescence against a protein of interest is presented, followed by DAPI staining to mark all chromosomes. Young adults are immobilized in levamisole and quickly dissected using two syringe needles. After germline extrusion, the sample is fixed before undergoing a freeze crack in liquid nitrogen, which helps permeabilize the cuticle and other tissues. The sample can then be dehydrated in ethanol, rehydrated, and incubated with primary and secondary antibodies. DAPI is added to the sample in the mounting medium, which allows reliable visualization of DNA and makes it easy to find animals to image under a fluorescent microscope. This technique is readily adopted by those familiar with handling C. elegans after a few hours spent practicing the dissection method itself. This protocol has been taught to high-schoolers and undergraduates working in a research lab and incorporated into a course-based undergraduate research experience at a liberal arts college.

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

This is a commonly used method for C. elegans gonad dissection followed by freeze crack, which produces germline samples for immunofluorescence via antibody staining, or for simple DAPI staining to visualize DNA. This protocol has been successful for undergraduates in a research lab and in a course-based undergraduate research experience.

The C. elegans germline makes an excellent model for studying meiosis, in part due to the ease of conducting cytological analyses on dissected animals. ...

Genetics

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

The Caenorhabditis elegans (C. elegans) germline is used to study several biologically important processes including stem cell development, apoptosis, and chromosome dynamics. While the germline is an excellent model, the analysis is often two dimensional due to the time and labor required for three-dimensional analysis. Major readouts in such studies are the number/position of nuclei and protein distribution within the germline. Here, we present a method to perform automated analysis of the germline using confocal microscopy and computational approaches to determine the number and position of nuclei in each region of the germline. Our method also analyzes germline protein distribution that enables the three-dimensional examination of protein expression in different genetic backgrounds. Further, our study shows variations in cytoskeletal architecture in distinct regions of the germline that may accommodate specific spatial developmental requirements. Finally, our method enables automated counting of the sperm in the spermatheca of each germline. Taken together, our method enables rapid and reproducible phenotypic analysis of the C. elegans germline.

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

We present an automated method for three-dimensional reconstruction of the Caenorhabditis elegans germline. Our method determines the number and position of each nucleus within the germline and analyses germline protein distribution and cytoskeletal structure.

The Caenorhabditis elegans (C. elegans) germline is used to study several biologically important processes including stem cell development, apoptosis, and ...

Developmental Biology

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous system can be observed, with single cell resolution, in vivo. Here, we present an integrated protocol for the examination of neurodevelopment in C. elegans embryos. Our protocol combines imaging, lineaging and neuroanatomical tracing of single cells in developing embryos. We achieve long-term, four-dimensional (4D) imaging of living C. elegans&#160;embryos with nearly isotropic spatial resolution through the use of Dual-view Inverted Selective Plane Illumination Microscopy (diSPIM). Nuclei and neuronal structures in the nematode embryos are imaged and isotropically fused to yield images with resolution of ~330 nm in all three dimensions. These minute-by-minute high-resolution 4D data sets are then analyzed to correlate definitive cell-lineage identities with gene expression and morphological dynamics at single-cell and subcellular levels of detail. Our protocol is structured to enable modular implementation of each of the described steps and enhance studies on embryogenesis, gene expression, or neurodevelopment.

Here, we present a combinatorial approach using high-resolution microscopy, computational tools, and single-cell labeling in living C. elegans embryos to understand single cell dynamics during neurodevelopment.

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous ...

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this protocol focuses on how to image dividing neuroblasts, which are found underneath the epidermal cells and may be important for epidermal morphogenesis. Tissue formation is crucial for metazoan development&#160;and relies on external cues from neighboring tissues. C. elegans is an excellent model organism to study tissue morphogenesis in vivo due to its transparency and simple organization, making its tissues easy to study via microscopy. Ventral enclosure is the process where the ventral surface of the embryo is covered by a single layer of epithelial cells. This event is thought to be facilitated by the underlying neuroblasts, which provide chemical guidance cues to mediate migration of the overlying epithelial cells. However, the neuroblasts are highly proliferative and also may act as a mechanical substrate for the ventral epidermal cells. Studies using this experimental protocol could uncover the importance of intercellular communication during tissue formation, and could be used to reveal the roles of genes involved in cell division within developing tissues.

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes how to image dividing cells within a tissue in Caenorhabditis elegans embryos. While several protocols describe how to image cell division in the early embryo, this protocol describes how to image cell division within a developing tissue during mid late embryogenesis.

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this ...

Neuroscience

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

C. elegans is the premier system for the systematic analysis of cell fate specification and morphogenetic events during embryonic development. One challenge is that embryogenesis dynamically unfolds over a period of about 13 h; this half day-long timescale has constrained the scope of experiments by limiting the number of embryos that can be imaged. Here, we describe a semi-high-throughput protocol that allows for the simultaneous 3D time-lapse imaging of development in 80&#8211;100 embryos at moderate time resolution, from up to 14 different conditions, in a single overnight run. The protocol is straightforward and can be implemented by any laboratory with access to a microscope with point visiting capacity. The utility of this protocol is demonstrated by using it to image two custom-built strains expressing fluorescent markers optimized to visualize key aspects of germ-layer specification and morphogenesis. To analyze the data, a custom program that crops individual embryos out of a broader field of view in all channels, z-steps, and timepoints and saves the sequences for each embryo into a separate tiff stack was built. The program, which includes a user-friendly graphical user interface (GUI), streamlines data processing by isolating, pre-processing, and uniformly orienting individual embryos in preparation for visualization or automated analysis. Also supplied is an ImageJ macro that compiles individual embryo data into a multi-panel file that displays maximum intensity fluorescence projection and brightfield images for each embryo at each time point. The protocols and tools described herein were validated by using them to characterize embryonic development following knock-down of 40 previously described developmental genes; this analysis visualized previously annotated developmental phenotypes and revealed new ones. In summary, this work details a semi-high-throughput imaging method coupled with a cropping program and ImageJ visualization tool that, when combined with strains expressing informative fluorescent markers, greatly accelerates experiments to analyze embryonic development.

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

This work describes a semi-high-throughput protocol that allows simultaneous 3D time-lapse imaging of embryogenesis in 80&#8211;100 C. elegans embryos in a single overnight run. Additionally, image processing and visualization tools are included to streamline data analysis. The combination of these methods with custom reporter strains enables detailed monitoring of embryogenesis.

C. elegans is the premier system for the systematic analysis of cell fate specification and morphogenetic events during embryonic development. One ...

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

4D microscopy is an invaluable tool for unraveling the embryonic developmental process in different animals. Over the last decades, Caenorhabditis elegans has emerged as one of the best models for studying development. From an optical point of view, its size and transparent body make this nematode an ideal specimen for DIC (Differential Interference Contrast or Nomarski) microscopy. This article illustrates a protocol for growing C. elegans nematodes, preparing and mounting their embryos, performing 4D microscopy and cell lineage tracing. The method is based on multifocal time-lapse records of Nomarski images and analysis with specific software. This technique reveals embryonic developmental dynamics at the cellular level. Any embryonic defect in mutants, such as problems in spindle orientation, cell migration, apoptosis or cell fate specification, can be efficiently detected and scored. Virtually every single cell of the embryo can be followed up to the moment the embryo begins to move. Tracing the complete cell lineage of a C. elegans embryo by 4D DIC microscopy is laborious, but the use of specific software greatly facilitates this task. In addition, this technique is easy to implement in the lab. 4D microscopy is a versatile tool and opens the possibility of performing an unparalleled analysis of embryonic development.

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

Here, we present a protocol for preparing and mounting Caenorhabditis elegans embryos, recording development under a 4D microscope and tracing cell lineage.

4D microscopy is an invaluable tool for unraveling the embryonic developmental process in different animals. Over the last decades, Caenorhabditis elegans ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

Tracking Calcium in the Early Developmental Phases of C. elegans

In Vivo Visualization of Calcium Transients during Fertilization and Early Development in C. elegans

Calcium is an important signaling molecule during the oocyte-to-embryo transition (OET) and early embryogenesis. The hermaphroditic nematode Caenorhabditis elegans provides several unique advantages for the study of the OET as it is transparent and has an ordered gonad that produces one mature oocyte every ~23 min at 20 &#176;C. We have modified the genetically encoded calcium indicator jGCaMP7s to fluorescently indicate the moment of fertilization within a living organism. We have termed this reporter &#34;CaFE&#34; for Calcium during Fertilization in C. elegans. The CaFE reporter was&#160;engineered into a safe harbor locus in single copy, has no significant impact on physiology or fecundity, and produces a robust signal upon fertilization. Here, a series of protocols is presented for utilizing the CaFE reporter as an in vivo tool for dissecting the OET and embryogenesis. We include methods to synchronize worms, examine the effects of RNAi knockdown, mount worms for imaging, and to visualize calcium in oocytes and embryos. Additionally, we present the generation of additional worm strains to aid in this type of analysis. Demonstrating the utility of the CaFE reporter to visualize the timing of fertilization, we report that double ovulation occurs when ipp-5 is targeted by RNAi and that only the first oocyte undergoes immediate fertilization. Furthermore, the discovery of single-cell calcium transients during early embryogenesis is reported here, demonstrating that the CaFE reporter persists into early development. Importantly, the CaFE reporter in worms is simple enough to use for incorporation into course-based undergraduate research (CURE) laboratory classes. The CaFE reporter, coupled with the ordered gonad and ease of RNAi in worms, facilitates inquiry into the cell-cell dynamics required to regulate internal fertilization and early embryogenesis.

Here, we present protocols and tools for visualizing calcium during fertilization and early embryogenesis using a genetically encoded calcium reporter expressed in the germline of the model nematode C. elegans.

Calcium is an important signaling molecule during the oocyte-to-embryo transition (OET) and early embryogenesis. The hermaphroditic nematode ...

High-Resolution C. elegans Imaging Across All Larval Stages

Caenorhabditis elegans has become one of the most widely studied and best-understood animal models in biology. Three features are key to C. elegans' success as a model organism: its invariant cell lineage, transparency, and genetic tractability. These render it ideal for a diverse range of microscopy-based studies directly in vivo. Live C. elegans larvae and adults often need to be immobilized during image acquisition. Traditional immobilization methods adversely affect animal development, especially in time-lapse imaging applications. Here, a detailed setup and operation protocol for a novel microfluidic imaging method is introduced, which addresses the limitations associated with traditional agar-pad-based immobilization and other microfluidic strategies. This approach enables simultaneous live imaging across various larval stages while preserving worm orientation and identity over time. To achieve this, a microfluidic trap channel array is employed, with its geometry precisely designed to maintain a stable worm orientation while accommodating growth and molting. Immobilization is facilitated by an active hydraulic valve that applies pressure to secure worms against the cover glass solely during image acquisition. This design allows high-resolution imaging with minimal effects on worm viability or developmental timing.

This protocol describes microfluidic-based C. elegans time-lapse imaging across the entire post-embryonic development.

Caenorhabditis elegans has become one of the most widely studied and best-understood animal models in biology. Three features are key to C. elegans' ...

Imaging Dendritic Spines in Caenorhabditis elegans

Dendritic spines are specialized sites of synaptic innervation modulated by activity and serve as substrates for learning and memory. Recently, dendritic spines have been described for DD GABAergic neurons as the input sites from presynaptic cholinergic neurons in the motor circuit of Caenorhabditis elegans. This synaptic circuit can now serve as a powerful new in vivo model of spine morphogenesis and function that exploits the facile genetics and ready accessibility of C. elegans to live-cell imaging. 
This protocol describes experimental strategies for assessing DD spine structure and function. In this approach, a super-resolution imaging strategy is used to visualize the intricate shapes of actin-rich dendritic spines. To evaluate the DD spine function, the light-activated opsin, Chrimson, stimulates the presynaptic cholinergic neurons, and the calcium indicator, GCaMP, reports the evoked calcium transients in postsynaptic DD spines. Together, these methods comprise powerful approaches for identifying genetic determinants of dendritic spines in C. elegans that could also direct spine morphogenesis and function in the brain.

Imaging Dendritic Spines in Caenorhabditis elegans

Dendritic spines are important cellular features of the nervous system. Here live imaging methods are described for assessing the structure and function of dendritic spines in C. elegans.&#160;These approaches support the development of mutant screens for genes that define dendritic spine shape or function.

Dendritic spines are specialized sites of synaptic innervation modulated by activity and serve as substrates for learning and memory. Recently, dendritic ...

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

Podsumowanie

Przeglądaj więcej filmów

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

In Vivo Visualization of Calcium Transients during Fertilization and Early Development in C. elegans

High-Resolution C. elegans Imaging Across All Larval Stages

Imaging Dendritic Spines in Caenorhabditis elegans