We thank the TisBio facilities and the PLBS platform for providing the technical environment conducive to achieving this work. This work was supported by grants from the CNRS and the Minist&#232;re de l&#39;Enseignement Sup&#233;rieur et de la Recherche. We would like to thank Dr. Fran&#231;ois Foulquier, Dr. Zo&#233; Durin, Mrs. Doroth&#233;e Vicogne, and Mrs. C&#233;line Schulz for stimulating discussions and for providing us with the Fibroblast 533T cell line and the primary antibody GM130.

1. Cell seeding
NOTE: Carry out the next steps under sterile conditions under a laminar flow hood. This method can be applied to any of the cell lines used in the present work (HeLa, MCF7, primary fibroblasts), or to most adherent cell line models commonly used in research20,24,25.
<ol>
	<li>Grow cells in DMEM high glucose medium supplemented with 10% fetal bovine serum (FBS) in a T75 flask at 37 &#176;C under a 5% CO2 atmosphere.</li>
	<li>Once the cells have reached full confluency, remove the cell culture medium and wash the cells with 4 mL of PBS. 
	NOTE: PBS solutions used in this step and all subsequent washing steps must be made sterile by filtration through a 0.2 &#181;m PTFE membrane prior to use.</li>
	<li>Incubate the cells for 5 min at 37 &#176;C in 2 mL of 1x trypsin-EDTA solution containing 0.5 g/L of trypsin, to release them from the bottom of the flask.</li>
	<li>Add 8 mL of DMEM high glucose medium containing 10% FBS, and mix thoroughly to make sure every single cell is detached from the bottom of the flask.</li>
	<li>Transfer the cells to a 15 mL conical tube and centrifuge for 5 min at 200 &#215; g. In the meantime, add one coverslip (diameter 12 mm, thickness 130-160 &#181;m) to each well of a 12-well plate. 
	NOTE: The coverslip thickness should be adjusted to match the requirements of the microscope used in subsequent imaging steps.</li>
	<li>Remove the supernatant after centrifugation and resuspend the pellet in 1 mL of cell culture medium.</li>
	<li>Determine the average number of cells/mL by hemocytometry across at least two measurements.</li>
	<li>Dilute the cell suspension to a concentration of 300,000 cells per mL in cell culture medium, and seed 1 mL of the suspension into each well.</li>
	<li>Allow the cells to settle for 24 h in DMEM high glucose medium containing 10% FBS at 37 &#176;C under a 5% CO2 atmosphere before the next steps.</li>
</ol>
2. Chloroquine treatment (optional)
NOTE: Steps 2.1-2.8 illustrate how to use the protocol on cells that are treated with an external reagent (inhibitor, effector, drug), using chloroquine as an example. Skip these steps for cells untreated with external reagents.
<ol>
	<li>Weigh the correct amount of chloroquine diphosphate salt (CQ) to prepare a sufficient volume of a 10 mM stock solution. 
	NOTE: Carry out the next steps under sterile conditions under a laminar flow hood.</li>
	<li>Solubilize the solid CQ in the desired amount of deionized water for a final concentration of 10 mM.</li>
	<li>Filter the solution through a 0.2 &#181;m filter.</li>
	<li>Aliquot 1 mL of the 10 mM stock solution in 1.5 mL microcentrifuge tubes and store them at -20 &#176;C.</li>
	<li>Wash the cell sample prepared in step 1.9 with 1 mL of PBS.</li>
	<li>Add 100 &#181;L of 10 mM CQ in 9.9 mL of DMEM high glucose medium containing 10% FBS, for a final concentration of 100 &#181;M of CQ.</li>
	<li>Incubate cells for 8 h in 1 mL of a medium supplemented with 100 &#181;M of CQ at 37 &#176;C under a 5% CO2 atmosphere.</li>
	<li>Wash the cells 3x with 1 mL of PBS.</li>
</ol>
3. Metabolic oligosaccharide engineering
<ol>
	<li>Prepare a 5 mM stock solution of ManNAz (or ManNAc for the negative control condition) in DMEM high glucose medium containing 10% FBS. 
	NOTE: This stock solution should be of sufficient volume such that each well can be treated with 1 mL of sugar analog solution. Carry out the next steps under sterile conditions under a laminar flow hood.</li>
	<li>Filter the solution through a 0.2 &#181;m filter and store in 1 mL aliquots at -20 &#176;C until required.</li>
	<li>Prepare a medium supplemented with 500 &#181;M of ManNAz or ManNAc by diluting a 1 mL stock solution aliquot prepared in step 3.2 with 9 mL of DMEM high glucose medium containing 10% FBS.</li>
	<li>Incubate the cells prepared in step 1.9 (for experiments on untreated cells) or in step 2.8 (for experiments on CQ-treated cells) in 1 mL of the medium supplemented with 500 &#181;M ManNAz (or 500 &#181;M ManNAc for the control group) for 24 h at 37 &#176;C under a 5% CO2 atmosphere.</li>
</ol>
4. Fixation and permeabilization
NOTE: All these steps are carried out under a fume hood.
<ol>
	<li>Wash the samples with 1 mL of PBS.</li>
	<li>Fix the cells on each coverslip by adding 500 &#181;L of 5% paraformaldehyde. Let it rest for 15 min at room temperature.</li>
	<li>Wash 3x with 1 mL of PBS.</li>
	<li>Prepare a custom humid chamber using a sufficiently large opaque box (e.g., a polystyrene box) with a lid. Place a wet piece of blotting paper at the bottom of the box, and then lay a layer of parafilm on top of it. 
	NOTE: The humid chamber should completely protect the sample from outside light sources when closed.</li>
	<li>Place the coverslips on the parafilm, cells facing up. 
	NOTE: Label the parafilm with the name or reference of the samples in advance.</li>
	<li>Carry out cell permeabilization using 200 &#181;L of 0.5% (v/v) Triton X-100 in PBS for 15 min at room temperature.</li>
	<li>Wash 3x with 200 &#181;L of PBS.</li>
</ol>
5. Fluorescence labeling
<ol>
	<li>CuAAC labeling of tagged sialylated glycans 
	NOTE: The CuAAC reaction buffer must be freshly prepared prior to use, and the reactants should be added in the specified order, with sodium ascorbate being added last. In oxygenated aqueous solutions, Cu(I) ions formed by the reduction of Cu(II) with ascorbate will gradually re-oxidize to Cu(II), leading to a decrease in ascorbate concentration over time. To maintain result reproducibility, the buffer should not be stored for more than a few days at 4 &#176;C and is preferably used on the same day.
	<ol>
		<li>To label the azide-modified sialylated glycans engineered in section 3, prepare a CuAAC reaction buffer consisting of 5 &#181;M AlexaFluor 488 alkyne (AF488Alk), 150 &#181;M CuSO4, 300 &#181;M 2-(4-((bis((1-(tert-butyl)-1H-1,2,3-triazol-4-yl)methyl)amino)methyl)-1H-1,2,3-triazol-1-yl)acetic acid (BTTAA), 0.1 M K2HPO4, and 2.5 mM sodium ascorbate in deionized water. Prepare enough solution to apply 200 &#181;L to each coverslip.</li>
		<li>Add 200 &#181;L of CuAAC buffer on each coverslip to initiate the reaction. Make sure the sample is homogeneously covered with the drop, and let it react for 45 min at room temperature, protected from the light.</li>
		<li>Wash 3x with 200 &#181;L of PBS to stop the reaction and remove excess probe and reagents.</li>
	</ol>
	</li>
	<li>Nucleus staining
	<ol>
		<li>For nuclear labeling with Hoechst 33342, dilute a 10 mg/mL stock solution at 1:2000 in PBS. 
		NOTE: Equivalent nuclear counterstains such as 4&#39;,6-diamidino-2-phenylindole (DAPI) may be used to the same effect.</li>
		<li>Cover the cells with 200 &#181;L of this solution and incubate for 5-10 min at room temperature, protected from the light.</li>
		<li>Wash 3x with 200 &#181;L of PBS.</li>
	</ol>
	</li>
	<li>Immunofluorescence staining
	<ol>
		<li>Prepare a Bovine Serum Albumin (BSA) blocking buffer containing 0.1 g of gelatin, 1 g of BSA, 1 mL of FBS, and 49 mL of PBS for 50 mL of buffer.</li>
		<li>Incubate the samples for 1 h at 4 &#176;C in 200 &#181;L of BSA blocking buffer. Protect the sample from light.</li>
		<li>For immunofluorescence staining of the Golgi apparatus using a purified mouse anti-GM130 antibody (epitope recognized: Rat GM130 aa. 869-982 | RRID: AB_398142), dilute the primary antibody at 1:100 in the prepared BSA blocking buffer. 
		NOTE: This step can be adapted to immunofluorescence staining targeting other organelles by using any relevant primary antibody, in which case the dilution factor should be adapted as per the manufacturer&#39;s guidance.</li>
		<li>Add 70 &#181;L of the solution containing the antibody to the coverslips and incubate for 1 h at 4 &#176;C, protected from light.</li>
		<li>Wash 3x with 200 &#181;L of PBS.</li>
		<li>Dilute the fluorescent secondary antibody (Alexa Fluor 546 anti-mouse IgG) at 1:600 in BSA blocking buffer and add 100 &#181;L on each coverslip.</li>
		<li>Incubate the samples for 1 h at room temperature, protected from light.</li>
		<li>Wash 3x with 200 &#181;L of PBS.</li>
		<li>Transfer the coverslips to a 6-well plate. Store the samples in 2 mL of PBS at 4 &#176;C for a few days, protected from light. 
		NOTE: it is advised to always prepare a replicate exclusively for visualization preExM as a reference for comparison purposes before and after expansion and to ensure the labeling pattern has not been distorted after expansion.</li>
	</ol>
	</li>
</ol>
6. Preexpansion imaging of samples
NOTE: The following steps are dependent on the bioimaging equipment used and may need to be adjusted to the requirements of the machine. Please follow the rules of the local laboratory or bioimaging platform. Steps 6.1-6.13 here are carried out on a laser scanning confocal microscope.
<ol>
	<li>Turn on the microscope and ensure that light sources (lasers) are warmed up. Open the bioimaging acquisition software.</li>
	<li>Select the laser excitation wavelengths corresponding to the fluorophores to create the channels (e.g., 405 nm for nuclear staining, 488 nm for AF488Alk-labeled sialylated glycans, and 546 nm for immunofluorescence labeling). Activate the lasers accordingly.</li>
	<li>Place and secure a 32 mm diameter coverslip (130-170 &#181;m thickness) in a holder adapted to the microscope.</li>
	<li>Place the sample prepared in step 5.3.9, cells facing down, on the 32 mm coverslip. Add a drop of deionized water to prevent the sample from drying out.</li>
	<li>Select an appropriate objective lens, here a 63x oil immersion objective with a 1.4 numerical aperture or equivalent. 
	NOTE: The highest numerical aperture provides the best resolution. When using an oil immersion objective, remember to add a drop of oil between the objective and the coverslip.</li>
	<li>Place the holder on the microscope stage above the objective.</li>
	<li>Use brightfield or low laser intensity fluorescence mode to locate an area of interest in the sample.</li>
	<li>Set the focus.</li>
	<li>Visualize the cells in the software in live fluorescence mode. Set the laser intensity for each channel. Start with low laser power to avoid photobleaching. Slowly increase the laser power and adjust the detector gain and offset to amplify the signal without introducing excessive noise as needed to visualize the fluorescence clearly without overexposure. 
	NOTE: Parameters should be adjusted to minimize both background noise and avoid signal saturation. It is crucial to maintain consistent settings across all samples in a series of experiments, including negative controls.</li>
	<li>Set image parameters such as pinhole size (e.g., 1 Airy Unit), scan speed (e.g., 8), zoom factor (e.g., 1), range of brightness values (e.g., 16 bits), and averaging number (e.g., 2), mode (e.g., line), and method (e.g., mean) as needed to reach the desired observation. 
	NOTE: These values are system- and sample-dependent and should be optimized prior to acquisition. Pinhole size depends on the numerical aperture, magnification, and wavelength: decreasing it may improve resolution at the expense of signal-to-noise ratio. It is advised to set scan speed to fast values (corresponding to small dwell time per pixel) to avoid photobleaching.</li>
	<li>Perform the acquisition and save data in the desired format with proper metadata for future reference.</li>
	<li>Once the acquisition is over, put the sample back in the 6-well plate by adding a small amount of deionized water to facilitate the transfer with a pair of tweezers. 
	NOTE: Step 6.12 is only carried out if expansion of this particular sample is needed. Otherwise, dispose of the sample and use other replicates for the following steps.</li>
</ol>
7. Expansion microscopy protocol
NOTE: The samples are protected as much as possible from the light to avoid photobleaching. The anchoring and gelation steps are carried out under a fume hood.
<ol>
	<li>Anchoring
	<ol>
		<li>Incubate the samples with N-acryloylsuccinimide (NAS) at a concentration of 3.2 mg/mL in PBS on a mechanical shaker for 1 h at room temperature. Prepare a sufficient amount of solution to add 500 &#181;L of solution per well (stock solution: 16 mg/mL NAS in DMSO). 
		NOTE: NAS reacts with the lysine residues and N-termini of proteins to graft an acryloyl group, enabling the proteins to anchor to the hydrogel in subsequent steps.</li>
		<li>Wash 3x with 1 mL of PBS.</li>
	</ol>
	</li>
	<li>Gelation
	<ol>
		<li>Prepare a monomer solution composed as follows: in 50 mL of 1x PBS (pH 7.4), add 5.843 g of NaCl, 1.25 g of acrylamide, 0.075 g of bis-acrylamide, and 4.313 g of sodium acrylate. Prepare enough solution to apply 70 &#181;L to each coverslip. Freeze 1 mL aliquots at least 24 h before gelation. 
		CAUTION: The monomers are toxic; they must be handled with care with appropriate protection equipment and weighed under a fume hood.</li>
		<li>On a piece of parafilm, a 70 &#181;L drop of monomer solution is placed, followed by the addition of 1.4 &#181;L of 10% (v/v) N,N,N&#8242;,N&#8242;-tetramethylethylenediamine (TMEDA), then 1.4 &#181;L of 10% (w/v) Ammonium persulfate (APS), which are mixed into the drop. 
		NOTE: The crosslinker reagents must be added in this specific order, TMEDA first, then APS.</li>
		<li>Quickly, place the coverslip carefully on the solution with the cells facing down.</li>
		<li>Allow the solution to polymerize for 1 h at room temperature in a humid chamber.</li>
	</ol>
	</li>
	<li>Digestion
	<ol>
		<li>Prepare a digestion buffer solution composed of 1x TAE buffer, 0.5% (v/v) Triton X-100, 0.8 M guanidine HCl, and 8 U/mL proteinase K. 
		NOTE: The digestion buffer must be freshly prepared and preincubated for 30 min at 37 &#176;C before use.</li>
		<li>Add 1 mL of digestion buffer to the hydrogel formed in step 7.2. Allow the digestion to proceed for 3 h at 37 &#176;C on a mechanical shaker.</li>
		<li>Remove the digestion buffer, and gently wash the gel 3x with 2 mL of deionized water. 
		NOTE: The hydrogel will slightly expand and detach itself from the cover slip.</li>
	</ol>
	</li>
	<li>Expansion
	<ol>
		<li>Leave the hydrogel to expand in 3 mL of deionized water for 2 h. Change the water every 30 min.</li>
	</ol>
	</li>
</ol>
8. Post expansion imaging of samples
<ol>
	<li>Place a 32 mm diameter coverslip (130-170 &#181;m thickness) and secure it in on a holder.</li>
	<li>Cut out a piece of the hydrogel and place it on the cover slip, with the cells at the bottom of the gel. 
	NOTE: The excised pieces should be large enough to provide adequate surface area for observation but not too large, as this could make handling the gel difficult and increase the risk of breakage (recommended size: 5 to 8 mm). Place enough hydrogel pieces to cover the entire surface of the coverslip; this will help secure the pieces in place and prevent any drift during imaging. In case of drifting issues, precoating the coverslips with poly-D-lysine might prove advantageous6.</li>
	<li>Place the holder on the microscope stage above the objective.</li>
	<li>Perform the image acquisition on a Laser Scanning Confocal Microscope with an oil immersion objective as described in step 6.</li>
</ol>
9. Expansion Factor (EF) calculation in ImageJ
<ol>
	<li>Open the imaging data in the ImageJ freeware or equivalent.</li>
	<li>If working on multiple channels image, split the channels and work only on the nucleus channel by navigating to Select Image | Color | Split channels.</li>
	<li>To measure the nucleus diameter, highlight the signal corresponding to the nucleus by determining a threshold by clicking on Select Image | Adjust | Threshold&#8230;. Choose a method in the default dropdown menu that best fits the image (e.g., Otsu), and adjust the threshold until the shape of the nucleus is clearly cut out from the background. 
	NOTE: The threshold setting needs to be adjusted for each image as one setting might not fit all images or every sample.</li>
	<li>Set the measurement to determine the Feret diameter by clicking on Select Analyze | Set measurements&#8230;. From the menu, select Feret diameter and adjust the decimal if necessary.</li>
	<li>To begin the analysis of the image, click on Select Analyze | Analyze particles to set the size of the particles; for the nucleus, choose 5-infinity &#181;m&#178; | Display results | Add to manager | Exclude on edges &#38; Overlay. Start the analysis.</li>
	<li>As the analysis begins, multiple windows will appear. In the window results, look for the maximum Feret diameter of each nucleus in the column Feret. Copy-paste the raw data to a spreadsheet and calculate the average nucleus size for preexpanded and expanded samples. 
	NOTE: Make sure that the regions of interest (ROIs) corresponding to each nucleus follow the entire periphery of the nucleus.</li>
	<li>Calculate the expansion factor (EF) as follows: 
	EF = <img alt="Equation 1" src="/files/ftp_upload/67479/67479eq01.jpg" style="margin: auto;" /> 
	NOTE: The EF factor determines how much the sample has been expanded.</li>
</ol>

Expansion Microscopy for Visualizing Intracellular Sialylated Glycans via MOE

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The authors have no competing financial interests or other conflicts of interest.

The present CuAAC labeling protocol does not include aminoguanidine in the reaction buffer. Since it is aimed at visualizing intracellular glycoconjugates, it is performed on cells that are fixed after the metabolic incorporation step, to avoid any cytotoxicity issue and improve uptake of the catalytic system. The use of aminoguanidine is typically recommended for cell-surface labeling of living cells to prevent side reactions between dehydroascorbate and arginine, histidine, and lysine residues of proteins28. These modifications could indeed affect protein function and potentially cause perturbations or cytotoxicity. Another important consideration for the CuAAC reaction buffer is that the complexation of copper ions by BTTAA is rapid but not immediate. To ensure efficient labeling of the modified sugars, we recommend preparing the solution at least 30 min before starting the labeling process, to ensure reproducibility of results by novice users.
Any confocal microscope may be used for acquisition. As the sample expands, so does the observable field. To capture an entire cell, performing a tile scan of the area with a 10% overlap between images is recommended. Although it is possible to conduct a tile scan and a z-stack simultaneously, we suggest opting for a smaller tile scan in that case. This is because the process is time-consuming and generates a substantial amount of raw data, leading to intensive data processing. It is advisable to separate both methods, creating a z-stack only for areas of interest, with parameters consistent with the preexpansion z-stack. As no antifading mounting medium is used on expanded gels, we advise setting acquisition parameters accordingly to minimize photobleaching (fast scan speed, low laser intensities).
Successful expansion experiments typically lead to an expansion factor of 4 to 5, which corresponds to a volumetric increase between 64x and 125x. Since the same number of fluorophore molecules is spread over a much larger volume after expansion, a major disadvantage of ExM is an unavoidable reduction in signal intensity. To be able to observe the fluorescent signal after expansion, the gain and/or the intensity of the excitation laser can be greatly increased depending on the fluorophore. The pinhole opening might be modulated as well. In addition to this purely physical dilution effect on the fluorescence signal, there are other possible causes of signal loss.
Following labeling, fixation, and bioconjugation of molecular anchors, the gelation (protocol step 7.2) aims to encapsulate the sample in a polyelectrolyte hydrogel. Such polymers are composed of three monomers: acrylamide, bis-acrylamide, and sodium acrylate. The latter gives the gel its expanding property due to its ionic nature; in fact, this polymer can absorb several times its weight in water by exchanging sodium ions for water molecules29. The crosslinking of the gel is a radical reaction catalyzed by TMEDA and APS. It is noteworthy that a cause of further signal reduction is the partial degradation of organic fluorophores during this radical-induced polymerization. Various dyes are affected at varying degrees; for example, cyanine dyes are almost completely destroyed during expansion, exhibiting a poor signal retention of 0.3-0.4 %30,31, while Alexa Fluor 488 dyes allow retention of ~57% of the signal intensity post ExM4. For a summary of the signal retention of various organic fluorophores and fluorescent proteins, see the review by Wen et al.31.
In addition, biomolecule retention may also be impacted by specific steps of the protocol. For example, glycan chains situated on protein fragments that are not anchored to the hydrogel may be washed out during digestion, further decreasing the signal intensity (Figure 4). Digestion employs at least one detergent32, such as Triton X-100, with or without proteinase K4,5,6,31, to disrupt membranes and protein structures like the cytoskeleton. The digestion time and the composition of the digestion buffer may be adjusted to best preserve certain fluorophores and biomolecules. The digestion parameters (protocol step 7.3) should, therefore, be optimized for the type of sample under study in terms of composition, concentration, and duration, to obtain the best balance between homogenization to ensure isotropic expansion and retention of biomolecules and probes. In certain cases, the type of digestion buffer can be completely modified for milder conditions, like in Ultrastructure-ExM where the sample&#39;s proteins are denaturized in SDS at 95 &#176;C before the gel expansion26,27.
When possible, the lower signal post ExM may be simply addressed by adjusting the power of the excitation source and/or the gain of the detector camera during the acquisition of the imaging data without the need for specific probes. In cases where the signal-to-noise ratio would be insufficient post ExM, additional signal retention and/or amplification strategies have been developed by various laboratories, for example, utilizing trifunctional molecules that allow anchoring of the fluorophores themselves directly to the polymer matrix6,14. As an alternative solution to address these challenges, staining of the sample may be done after the expansion since the gel is sufficiently permeable. However, this prevents the observation of the sample before expansion. This approach has recently been reported for immunostaining33, where specimens are first expanded isotropically and then stained with antibodies. The authors demonstrated that this may lead to improved fluorescence brightness and epitope accessibility, while reducing linkage errors that impede localization accuracy when combining ExM and super-resolution techniques.
Alternative strategies to thwart the diminution of fluorescence intensity have been explored. One example is to use polymer-based fluorophores instead of conventional fluorescent molecules. Liu et al. developed fluorescent polymer dots (Pdots)34 that are much brighter than conventional fluorophores, allowing for better signal-to-noise ratio. Pdots can be functionalized with an anchor and an antibody. In another example, Click Expansion Microscopy (ClickExM)6 introduced a universal strategy to enhance signal retention with an indirect biotin-streptavidine strategy, using a fluorescent streptavidine that bears an anchor. This allows direct grafting of the fluorescent probes to the polymer frame. Similarly, Molecule Anchorable Gel-enabled Nanoscale In-situ Fluorescence microscopY (MAGNIFY)35 heavily relies on biomolecule anchoring to increase fluorescent signal retention. Contrary to ClickExM, which bonds a specific molecule to the gel, MAGNIFY uses methacrolein to crosslink a broad range of biomolecules to the frame of the hydrogel.
Another possible limitation of ExM in terms of data acquisition on the microscope is the necessity to image an expanded sample over a larger physical area, which significantly increases the time required to capture the entire sample. Since the expansion process enlarges the sample, achieving high-resolution imaging necessitates capturing more image tiles to cover the same biological region, resulting in prolonged acquisition times. In this regard, for a number of applications, ExM would not be considered as a replacement for classical confocal microscopy but as a complementary technique for comprehensive and detailed imaging.
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/67479/67479fig07.jpg" /> 
Figure 7: Comparison of preExM versus postExM 3D imaging of human primary fibroblast 533T cells. Cells were incubated with 500 &#181;M ManNAz for 24 h and labeled with Alexa Fluor 488 alkyne by CuAAC (green color). Co-localization staining of the Golgi apparatus (red color) was carried out with a mouse anti-GM130 primary antibody and an anti-mouse Alexa Fluor 546-labeled secondary antibody, and nuclei were stained with Hoechst 33342 (blue color). (Left) (A) Z-stack and (B,C) 3D surface rendering of cells visualized preExM. (Right) Cells were submitted to anchoring, gelation, digestion, and expansion as described, (D) prior to z-stack acquisition. (E) 3D surface rendering of the signals clearly illustrates the resolution increase that is particularly valuable when detecting intricate intracellular substructures. 3D reconstruction and surface rendering were created using signal intensity. Scale bars = 20 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67479/67479fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
ExM has emerged as an innovative technique for achieving nanoscale imaging without actually surpassing the diffraction limit, allowing for the visualization of details from biological samples that would otherwise be overlooked in standard protocols6. When using a ~250 nm diffraction-limited equipment (as is usually the case for routine confocal microscopes equipped with green lasers), an expansion factor of 4.5x typically leads to a lateral resolution close to 60 nm. This is a very significant improvement when studying subcellular structures such as the Golgi apparatus or vesicles, which can be exploited in 2D imaging but also by generating 3D images from z-stack acquisition (Figure 7). To harness the full power of expansion microscopy, the attained resolution may be further improved by combining ExM with super-resolution microscopy techniques such as STED or STORM, which is outside the scope of this article.
The field of Expansion Microscopy is witnessing very rapid growth and significant advancements, making it an exciting and dynamic area of research. Boyden and co-workers introduced Expansion Microscopy in 2015, achieving approximately 70 nm resolution with confocal microscopes by using gel-anchorable labels to retain the geometry of biomolecules in the polymer matrix. Following the initial interest, the need for custom-made oligonucleotide-antibody conjugates drove the scientific community to develop new tools and approaches for this method. Various strategies for directly anchoring proteins to the gel using commercially available agents have been developed, allowing the use of fluorescently labeled antibodies or proteins, and recent developments have further simplified Expansion Microscopy and broadened its application to various biomolecules without needing separate anchoring steps.
Achieving MOE labeling with higher resolution through ExM is particularly useful for studying the spatial distribution of sialylated glycans routinely without the need for super-resolution microscopes that are costly and difficult to access. For example, sialoglycans on the plasma membrane have been visualized with this method, revealing their distribution in microvilli and other cellular structures6. The ability to visualize intracellular glycans with improved resolution is even more crucial to study their biosynthesis, trafficking, and recycling8,13,14,15 and decipher their role in pathologies such as metabolic diseases or cancers. Combining ExM and bioorthogonal chemistry will offer a unique opportunity to better understand their localization and involvement in cellular pathways, organelle function, and interactions that are key to cell regulation and disease mechanisms. This will undoubtedly improve our understanding of the role played by sialic acids in various pathologies in the future.

Fluorescence microscopy, while widely used for labeling and visualizing specific molecules within cells, is inherently limited in resolution by Abbe&#39;s diffraction limit of light1, which restricts the ability to distinguish between objects closer than approximately 200-250 nm. This limitation arises from the wave nature of light and the numerical aperture of the microscope&#39;s objective lens, introducing a challenge when imaging subcellular structures. Overcoming these limitations provides better insights into certain biological processes at a nanometric scale.
To surpass the diffraction limit of light, super-resolution microscopy techniques such as STORM (Stochastic Optical Reconstruction Microscopy) and STED (Stimulated Emission Depletion) have been developed2,3. STORM relies on the stochastic activation of fluorophores, allowing only a sparse subset to be imaged at any given time. This enables the precise localization of individual fluorophores, which allows the reconstruction of a high-resolution image. STED, on the other hand, improves resolution by using a depletion laser to selectively quench fluorescence around the periphery of the excitation spot, effectively narrowing the point spread function.
These approaches contrast with widefield or confocal microscopy, where all fluorophores are simultaneously detected, resulting in an image that combines all diffraction patterns and prevents the distinction between close-by individual fluorophores, leading to a loss of resolution. However, these super-resolution methods require very specific light sources, equipment, sample preparation, and/or fluorophores, making these technologies costly, difficult to access in most laboratories, and challenging to implement in routine experiments. These constraints have led the scientific community to search for alternative solutions to achieve higher resolution, that would be compatible with readily available microscopy equipment and routine staining protocols. In 2015, a method to circumvent the limitations of optical microscopy by physically expanding the sample was developed by Boyden and co-workers, called Expansion Microscopy (ExM)4.
ExM is a three-step method that provides nanoscale details of biological samples without surpassing the diffraction limit of light (Figure 1). Instead, it uses conventional diffraction-limited microscopes to image samples that have been physically magnified in an isotropic manner. The first step, called gelation, consists of embedding the biological sample, typically fixed cells or tissues, in a swellable polyelectrolyte hydrogel based on sodium acrylate and acrylamide. The biological sample then undergoes enzymatic treatment to partially degrade certain components such as proteins or membranes, and break down dense cellular structures to ensure cells can expand uniformly. This step, called digestion, helps achieve this by homogenizing the sample&#39;s structural components with regard to mechanical properties, preventing differential expansion that could lead to distortion of the sample. Finally, in the last step, called expansion, the hydrogel-embedded and digested sample is placed in deionized water, causing it to swell. This technology results in the sample expanding by a linear magnification factor of approximately 4-5x in every dimension, allowing for the visualization of fine cellular details using standard microscopy techniques. Given the isotropic nature of the expansion, the biological sample enlarged within the gel matrix retains its three-dimensional geometrical details and the spatial relationships between its various structural components. Spatial information is, therefore, preserved upon swelling, while the distance between the gel-anchored fluorescent labels or biomolecules increases uniformly in all directions. This allows for better separation of signals, resulting in enhanced resolution.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67479/67479fig01.jpg" /> 
Figure 1: Overview of the ExM Protocol. (a) Gelation: The biological sample is anchored and embedded within a swellable polyelectrolyte hydrogel. (b) Digestion: The mechanical properties of the sample are homogenized through the use of enzymes and detergents, breaking down proteins and lipid membranes that would otherwise restrict the expansion and create distortions. (c) Expansion: The hydrogel is immersed in deionized water, causing it to expand isotropically. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67479/67479fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Since 2015, ExM has seen significant technological improvements in preserving spatial information. In particular, various molecular anchors have been designed to covalently attach biomolecules directly to the swellable hydrogel5,6. In this protocol, we used N-acryloxysuccinimide (NAS), which reacts with the free amine groups of biomolecules (typically, lysine residues or N-terminal positions of proteins) to bioconjugate an acryloyl group to the cell&#39;s proteins, including glycoproteins. This acryloyl group then reacts with other monomers through cross-linking reactions during gelation, anchoring these biomolecules directly to the polymer.
ExM is compatible with a wide range of labeling methods, including Metabolic Oligosaccharide Engineering (MOE). MOE is a powerful tool that enables the labeling of glycans by incorporating analogs of metabolic precursors equipped with a bioorthogonal chemical handle7. These analogs, known as chemical reporters, integrate into metabolic pathways without causing toxicity. In the MOE approach, reporter monosaccharides are metabolically processed into activated nucleotide sugars and then transferred to nascent glycoconjugates. Our primary focus is on the study of sialylation, particularly the intracellular dynamics and trafficking of sialylated N-glycoproteins, which is crucial in the context of health and disease due to its roles in cell-cell interactions, immune regulation, and development. Abnormal sialylation is implicated in diseases like cancer8,9,10,11, infectious diseases12, and genetic disorders13,14,15, making it a key target for therapeutic development and biomarker discovery. Understanding sialylation enhances insights into glycobiology and disease mechanisms.
Sialylation can be probed using MOE with analogs of N-acetylneuraminic acid (Neu5Ac), the most abundant sialic acid in humans, or with analogs of N-acetylmannosamine (ManNAc), a metabolic precursor of Neu5Ac, bearing a bioorthogonal handle8,16. ManNAc is converted into Neu5Ac in the cytosol, then activated into cytidine-5&#8242;-monophospho-N-neuraminic acid (CMP-Neu5Ac) in the nucleus. Once activated into a nucleotide sugar, sialyl transferases in the Golgi apparatus transfer Neu5Ac units to the terminal positions of growing glycan chains (Figure 2). Following the metabolic incorporation of unnatural ManNAc derivatives, the tagged sialylated glycans can be covalently linked to a fluorophore bearing a reactive group complementary to the reporter handle through bioorthogonal click chemistry. This allows for the direct observation of glycoconjugates in vivo or ex vivo.
For most monosaccharide reporters, a peracetylated form is required to cross the plasma membrane via passive diffusion. However, both peracetylated and unprotected reporters have been shown to efficiently probe sialylation16. Unprotected sialylation reporters are able to enter cells by active transport mechanisms, namely pinocytosis for Neu5Ac analogs, and a yet unidentified transporter for ManNAc. While unprotected sugars require a higher concentration (typically 100-500 &#181;M) to achieve comparable effects, once inside the cell, they can directly enter the sialic acid metabolic pathway. In contrast, peracetylated sugars need to be fully deacetylated by intracellular non-specific esterases before becoming metabolically active. Although they can be used at lower concentrations (typically 10-50 &#181;M), incomplete deacetylation may interfere with enzyme activity or lead to the incorporation of partially acetylated sialic acid analogs, potentially skewing downstream analysis. Additionally, the release of acetic acid may affect the pH locally, potentially impacting cellular function. Chen and colleagues have also demonstrated that per-O-acetylated sugars react with free cysteine residues in proteins via a non-enzymatic mechanism, leading to off-target incorporation and increased non-specific signal17,18. In the present protocol, we therefore employ unprotected ManNAc reporters.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67479/67479fig02.jpg" /> 
Figure 2: Metabolic oligosaccharide engineering and labeling of sialic acids. UDP-GlcNAc is converted to ManNAc by UDP-GlcNAc 2-epimerase domain of GNE/MNK in the cytosol. ManNAc is then phosphorylated in the cytosol by ManNAc 6-kinase domain of GNE/MNK to form ManNAc-6-phosphate. N-acetylneuraminate synthase catalyzes the condensation of ManNAc-6-P with phosphoenolpyruvate to produce Neu5Ac-9-phosphate, which is subsequently dephosphorylated by sialic acid phosphatase to yield Neu5Ac. Neu5Ac can also be supplied by the salvage pathway via endocytosis and lysosomal recycling8. After transport to the nucleus, it is converted to CMP-Neu5Ac by CMP-sialic acid synthetase. In the Golgi apparatus, CMP-NeuAc is the substrate of sialyltransferases that introduce a Neu5Ac moiety at the terminal positions of glycans on maturing glycoconjugates, which are eventually expressed at the cell membrane or secreted. ManNAz chemical reporters bearing a bioorthogonal handle can penetrate the cell through an unidentified active transporter and enter the metabolic pathway. The tagged Neu5Az units incorporated in glycans after biosynthesis are then labeled through CuAAC-mediated conjugation of a fluorescent probe. Abbreviations: NAc = N-acetyl; UDP-GlcNAc = Uridine diphosphate N-acetylglucosamine; ManNAc = N-acetylmannosamine; GNE = UDP-GlcNAc 2-epimerase; MNK = ManNAc 6-kinase; ManNAc-6-P = ManNAc-6-phosphate; NANS = N-acetylneuraminate synthase; PEP = phosphoenolpyruvate; Neu5Ac = N-acetylneuraminic acid; Neu5Ac-9-P = Neu5Ac-9-phosphate; NANP = sialic acid phosphatase; CMP = cytidine-5&#8242;-monophosphate; CMAS = CMP-sialic acid synthetase; STs = sialyltransferases; ManNAz = N-azidoacetylmannosamine; Neu5Az = N-azidoacetylneuraminic acid. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67479/67479fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Among the possible bioorthogonal reactions, we focused our interest on the Copper-catalyzed Azide-Alkyne Cycloaddition (CuAAC)16,19. According to our experience, CuAAC is the best option for labeling intracellular glycans in fixed cells. This reaction is well-established, thoroughly studied in bioorthogonal chemistry, and has been standardized for use in complex biological environments, providing a solid foundation of knowledge and optimized protocols. Its fast kinetics indeed offer high reaction efficiency and specificity, and it involves azide and alkyne groups that are easy to synthesize, stable, absent from living systems, and inert toward native biomolecules, making it ideal for fixed-cell applications where copper toxicity is not an issue. Strain-promoted Alkyne-Azide Cycloaddition (SPAAC)20,21, while avoiding copper toxicity, has slower kinetics and bulkier probes, which tends to lead to higher background signal and decreased signal-to-noise ratio for intracellular applications due to hydrophobic trapping, whereas Inverse Electron Demand Diels-Alder (IEDDA)22,23, though fast and copper-free, involves more complex probe synthesis and requires bulkier reporter groups that are yet to be fully characterized in MOE applications. Since ExM exclusively requires fixed cells, CuAAC offers a robust and efficient solution for bioorthogonal labeling.
This paper presents a protocol for the visualization of intracellular sialylated glycoproteins in cells, combining metabolic labeling, bioorthogonal click chemistry, and expansion microscopy. In the experimental procedures described in this paper, we utilize N-azidoacetylmannosamine (ManNAz) as the chemical reporter, and CuAAC ligation of small organic fluorophores is performed prior to the expansion procedure. Glycoproteins are anchored to the hydrogel with NAS prior to gelation, digestion, and expansion. The MOE labeling of sialic acids can be associated with immunofluorescence approaches for co-localization assessment, as exemplified here with a mouse anti-GM130 primary antibody localized in the cis-Golgi apparatus. This protocol can be applied to cells in their physiological state, or to cells that have been subjected to chemical treatment. To illustrate this, chloroquine was used to inhibit lysosomal function, which affects the processing and trafficking of glycoproteins within cells. Nuclear staining is used as a landmark, not only to localize cells but also as an indicator of the quality of the ExM process. The expansion factor can indeed be measured by comparing the size of the nucleus pre-expansion (preExM) and post-expansion (postExM).

<table><tbody><tr><td>(+) Sodium L-ascorbate</td><td>Sigma Aldrich</td><td>11140</td><td></td></tr><tr><td>12 well cell culture plate</td><td>Corning</td><td>3513</td><td></td></tr><tr><td>Acrylamide</td><td>Sigma Aldrich</td><td>A8887</td><td></td></tr><tr><td>Acrylic acid N-hydroxysuccinimide ester&amp;nbsp;</td><td>Sigma Aldrich</td><td>A8060</td><td></td></tr><tr><td>Alexa Fluor 488 alkyne&amp;nbsp;</td><td>Jena Bioscience</td><td>CLK-1277-5</td><td></td></tr><tr><td>Alexa Fluor 546 goat anti-mouse IgG</td><td>Invitrogen</td><td>A11003</td><td></td></tr><tr><td>Amonium persulfate</td><td>Sigma Aldrich</td><td>9913</td><td></td></tr><tr><td>Bis-Acrylamide</td><td>Sigma Aldrich</td><td>146072</td><td></td></tr><tr><td>BSA</td><td>Sigma Aldrich</td><td>A7906</td><td></td></tr><tr><td>BTTAA</td><td>Jena Bioscience</td><td>CLK-067-100</td><td></td></tr><tr><td>Centrifugation tube 2 mL</td><td>EPPENDORF</td><td>30120094</td><td></td></tr><tr><td>Chloroquine diphosphate salt</td><td>Sigma Aldrich</td><td>C6628</td><td></td></tr><tr><td>Conical tube 15 mL</td><td>Falcon</td><td>352097</td><td></td></tr><tr><td>cover slips 12 mm #1</td><td>epredia</td><td>CB00120RA120MNZ0</td><td></td></tr><tr><td>cover slips 32 mm #1</td><td>epredia</td><td>CB00320RA140MNZ0</td><td></td></tr><tr><td>CuSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma Aldrich</td><td>209198</td><td></td></tr><tr><td>DMEM high glucose medium&amp;nbsp;</td><td>Dutscher</td><td>L0104-500</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Saline (PBS)&amp;nbsp;</td><td>Dutscher</td><td>L0615-500</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>biowest</td><td>S1810-500</td><td></td></tr><tr><td>Fibroblast 533T</td><td>-</td><td>-</td><td>Collected from healthy individual</td></tr><tr><td>FIJI ImageJ 2.9.0</td><td>-</td><td>-</td><td></td></tr><tr><td>Gelatin</td><td>Bio-RAD</td><td>170-6537</td><td></td></tr><tr><td>Guanidine HCl</td><td>Sigma Aldrich</td><td>50950</td><td></td></tr><tr><td>HeLa cells</td><td>ATCC</td><td>CCL-2</td><td></td></tr><tr><td>Hoechst 33342</td><td>Sigma Aldrich</td><td>14533</td><td></td></tr><tr><td>Imaris 10.2</td><td>-</td><td>-</td><td></td></tr><tr><td>K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Euromedex</td><td>PB0447-B</td><td>Anhydrous</td></tr><tr><td>LSM 780 Confocal Microscopy&amp;nbsp;</td><td>Zeiss</td><td>-</td><td></td></tr><tr><td>MCF7</td><td>ATCC</td><td>HTB-22</td><td></td></tr><tr><td>N-acetylmannosamine (ManNAc)</td><td>BIOSYNTH</td><td>MA05269</td><td></td></tr><tr><td>NaCl</td><td>Carlo Erba</td><td>479687</td><td></td></tr><tr><td>N-azidoacetylmannosamine (ManNAz)</td><td>BIOSYNTH</td><td>MA46002</td><td></td></tr><tr><td>Objectif &quot;Plan-Apochromat&quot; 63x/1,4 Oil DIC M27</td><td>Zeiss</td><td>420782-9900-799&amp;nbsp;</td><td></td></tr><tr><td>Phosphate Buffered Saline (PBS) 10x</td><td>Euromedex</td><td>ET330</td><td></td></tr><tr><td>Proteinase K</td><td>Sigma Aldrich</td><td>P2308</td><td>from&amp;nbsp;Tritirachium album</td></tr><tr><td>purified mouse GM130 antibody</td><td>BD Bioscience</td><td>610822</td><td>50 &amp;micro;g</td></tr><tr><td>Sodium acrylate&amp;nbsp;</td><td>Sigma Aldrich</td><td>408220</td><td></td></tr><tr><td>T75 Flask</td><td>Corning</td><td>430641</td><td></td></tr><tr><td>TEMED</td><td>Sigma Aldrich</td><td>T9281</td><td></td></tr><tr><td>tris Acetate EDTA (TAE) 10x</td><td>Euromedex</td><td>EU0202-B</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>X-100</td><td></td></tr><tr><td>Trypan Blue</td><td>Dutscher</td><td>702630</td><td></td></tr><tr><td>Trypsine-EDTA 1x</td><td>Dutscher</td><td>L0930-100</td><td></td></tr></tbody></table>

visualizing intracellular sialylation with click chemistry and expansion microscopy

Metabolic labeling techniques allow the incorporation of bioorthogonal reporters into glycans, enabling the targeted bioconjugation of molecular dyes within cells through click and bioorthogonal chemistry. Metabolic oligosaccharide engineering (MOE) has attracted considerable interest due to the essential role of glycosylation in numerous biological processes that involve molecular recognition and its impact on pathologies ranging from cancer to genetic disorders to viral and bacterial infections.
Although MOE is better known for the detection of cell surface glycoconjugates, it is also a very important methodology for the study of intracellular glycans in physiological and pathological contexts. Such studies greatly benefit from high spatial resolution. However, super-resolution microscopy is not readily available in most laboratories and poses challenges for daily implementation. Expansion microscopy is a recent alternative that enhances the resolution of microscopy by physically enlarging biological specimens labeled with fluorescent markers. By embedding the sample in a swellable gel and causing it to expand isotropically through chemical treatment, subcellular structures can be visualized with enhanced precision and resolution without the need for super-resolution techniques.
In this work, we illustrate the capacity of expansion microscopy to visualize intracellular sialylated glycans through the combined use of MOE and click chemistry. Specifically, we propose a procedure for bioorthogonal labeling and expansion microscopy that employs a reporter targeting sialylation, which may be associated with immunofluorescence for co-localization studies. This protocol enables localization studies of sialoconjugate biosynthesis, intracellular trafficking, and recycling.

Shown below is the application of the protocol to visualize sialylated glycoproteins in HeLa cells (Figure 3A) and MCF7 cells (Figure 3B), omitting CQ treatment (protocol section 2) and immunofluorescence co-localization staining (protocol step 5.3).
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67479/67479fig03.jpg" /> 
Figure 3: Comparison of the sialylation labeling pattern before (left) and after (right) the ExM protocol. (A) Hela cells and (B) MCF7 cells were incubated for 24 h with 500 &#181;M ManNAz. Labeling was carried out through CuAAC with Alexa Fluor 488 alkyne (depicted in green). Nuclei were stained with Hoechst 33342 (depicted in blue). Scale bars = 20 &#181;m. Abbreviations: ExM = Expansion Microscopy; ManNAz = N-azidoacetylmannosamine; CuAAC = Copper-catalyzed Azide-Alkyne Cycloaddition. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67479/67479fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Typically, sialylation labeling consists of three main fluorescence pools: a Golgi pool indicating nascent glycoconjugates being synthesized, a vesicular pool (usually less intense) representing glycoconjugates being exported to or recycled from the cell membrane, and a membrane pool showing glycoconjugates that have been expressed13. It is important to note that only biopolymer glycoconjugates tagged with the metabolically generated Neu5Az are detected, while any unreacted excess reporter or probes are eliminated during the washing steps. Images post ExM were acquired on a confocal microscope and reveal much finer details and lesser background, more particularly in the Golgi apparatus. The protocol can be applied to any type of adherent cells that express sialylated glycoconjugates.
Different cell or tissue types may be impacted to varying degrees by the partial loss of signal intrinsic to ExM, which is due to the volumetric dilution of fluorophores. This may also be worsened by a poor retention of fluorophores during the radical polymerization, or of biomolecules due to suboptimal anchoring or digestion, which can lead to a drastic loss of signal (Figure 4). This point is the main topic of the discussion below. It is, therefore, advised to adjust the parameters of the digestion steps to the cell or tissue type under scrutiny if needed. Digestion can be optimized through various means, depending on the specific structures or biomolecules being observed. We recommend starting with a standardized digestion protocol and then empirically adjusting parameters such as digestion time, detergent concentration, and/or enzyme concentration to achieve the desired results. It is important to note that more efficient digestion leads to a better isotropic expansion avoiding distortion, but this can also increase the risk of signal loss. Therefore, a balance must be found between achieving high-quality isotropic expansion and retaining signal integrity. Signal retention can be further enhanced by using specifically designed molecular anchors. For studies focused on protein observation, protocols using only Sodium Dodecyl Sulfate (SDS) at elevated temperatures have been shown to effectively digest the embedded sample while better preserving protein signals26,27.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/67479/67479fig04.jpg" /> 
Figure 4: Example of suboptimal result. This figure illustrates the importance of adjusting conditions to each type of cell, tissue, and monitored biomolecule, especially for the digestion step. HeLa cells were incubated 24 h with 500 &#181;M ManNAz and labeled with Alexa Fluor 488 alkyne (green, sialylated N-glycoproteins) and Hoechst 33342 (blue, nuclei). (A) Labeling pattern pre expansion. (B) labeling pattern post expansion. (Top) Poorly optimized digestion parameters in terms of composition, concentration, and/or length may lead to drastic loss of signal due to a lack of grafted biomolecule retention in the hydrogel, preventing bioimaging data acquisition with sufficient sensitivity. In this particular case, the digestion buffer was similar to that used in protocol step 7.3. However, the digestion time was 90 min, compared to the 180 min digestion time used in the subsequent image. (Bottom) Image obtained after optimization of the digestion parameters (see protocol step 7.3). Scale bars = 20 &#181;m. Abbreviation: ManNAz = N-azidoacetylmannosamine. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67479/67479fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
To ensure the reproducibility of the experiment, it is crucial to preserve the samples in solution and protect them from light, even after embedding them in the hydrogel. The hydrogel must be maintained in a humid chamber to prevent it from drying out. It is strongly recommended to perform the three main steps-gelation (protocol step 7.2), digestion (protocol step 7.3), and expansion (protocol step 7.4)-on the same day. Once expanded, the sample is stable for a few days if kept at 4 &#176;C in deionized water and protected from light, allowing time for bioimaging data acquisition.
During the analysis of microscopy images, it is recommended to evaluate the expansion factor (EF) by measuring the average ratio of the nucleus Feret diameter before (preExM) and after expansion (postExM). Here, in this protocol, nuclei are stained with Hoechst 33342, but the more conventional DAPI is also compatible with expansion microscopy. The Feret diameter corresponds to the maximum distance between two points situated at the boundary of the nucleus. Typically, successful experiments provide an EF value of 4 to 5 (Figure 5), corresponding to a maximum lateral resolution of ~60-70 nm, depending on the utilized wavelength and objective. The average Feret diameter of nuclei is determined by measuring at least 60 to 100 cells across several images for a statistical approach. For greater accuracy, z-stacks enabling 3D reconstruction can be systematically acquired and processed to ensure that the measured distance corresponds to the actual Feret diameter for all nuclei. However, this approach may be significantly more time-consuming, both in terms of acquisition depending on the microscope and objectives used, and particularly in terms of data treatment. This may be cross-checked by macroscale measurement of the EF by comparing the size of the hydrogel before and after expansion, which, in our experience, leads to similar results.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/67479/67479fig05.jpg" /> 
Figure 5: Determination of the Expansion Factor. The average size of nuclei before and after expansion is determined through the measurement of their Feret diameter (i.e., the longest distance between two points at the edge of the nucleus) in the ImageJ software. Calculation of the ratio between the average Feret diameter post ExM and pre ExM provides the linear expansion factor. Human primary fibroblasts 533T were submitted to the expansion protocol. Nuclei were stained with Hoechst 33342 (depicted in blue). Scale bars = 20 &#181;m. Abbreviation: ExM = Expansion Microscopy. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67479/67479fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
This protocol is designed to address biological questions requiring specific cell treatments or comparisons between various physiological and pathological conditions. This is exemplified here with its application on human fibroblasts treated with the antimalarial drug chloroquine (CQ). CQ, a diprotic weak base with pKa values of 8.4 and 10.8, preferentially accumulates in lysosomes via pH trapping, resulting in increased lysosomal pH and subsequent loss of function.
In this experiment, ManNAz is incorporated into sialylated glycans, which are then labeled with Alexa Fluor 488 through CuAAC, while the Golgi apparatus is stained with the mouse anti-GM130 antibody. Notably, distinct differences in relative intensities are observed between control cells (Figure 6A) and CQ-treated cells (Figure 6B). Under physiological conditions in control cells, the primary pool of sialoconjugate labeling is localized within the Golgi apparatus. In contrast, CQ-treated cells exhibit sialic acid accumulation in lysosomes, evidencing the trapping of recycled glycoconjugates. This is particularly relevant in the context of studying congenital disorders of glycosylation or lysosomal storage diseases13,14,15.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/67479/67479fig06.jpg" /> 
Figure 6: PostExM visualization of the effect of chloroquine on the localization of sialylated glycoproteins postExM. Human primary fibroblasts 533T were either (A) not treated with CQ or (B) treated with 100 &#181;M CQ for 8 h prior to incubation with 500 &#181;M ManNAz for 24 h and CuAAC labeling with Alexa Fluor 488 alkyne (green channel). Co-localization staining of the Golgi apparatus (red channel) was carried out with a mouse anti-GM130 primary antibody and an anti-mouse Alexa Fluor 546 labeled secondary antibody. The cells were subjected to anchoring, gelation, digestion, and expansion as described. Scale bars = 20 &#181;m. Abbreviations: ExM = Expansion Microscopy; CQ = chloroquine; ManNAz = N-azidoacetylmannosamine; CuAAC = Copper-catalyzed Azide-Alkyne Cycloaddition. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67479/67479fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Visualizing Intracellular Sialylation with Click Chemistry and Expansion Microscopy

Here, we propose a simple protocol combining metabolic oligosaccharide engineering, click chemistry, and expansion microscopy that allows bioimaging of intracellular sialylated N-glycoproteins with improved resolution using routine microscopy equipment.

Metabolic labeling techniques allow the incorporation of bioorthogonal reporters into glycans, enabling the targeted bioconjugation of molecular dyes ...

visualizing-intracellular-sialylation-with-click-chemistry-expansion

Nanyang Technological University

Research

JoVE Journal

Biology

378 Views.  Univ. Lille, CNRS, UMR 8576 - UGSF - Unité de Glycobiologie Structurale et Fonctionnelle. We presenteren een eenvoudig protocol voor het visualiseren van intracellulaire gesialyleerde glycoconjugaten met een hoge ruimtelijke resolutie, waardoor er geen gespecialiseerde apparatuur nodig is. Deze methode vergemakkelijkt de studie van de biosynthese, handel en recycling in verschillende pathologische contexten. Om dit te bereiken, combineren we metabole labeling, een klikchemie en expansiemicroscopie.Superresolutiemicroscopietechnieken zoals S...

Video: Visualisatie van intracellulaire sialylering met klikchemie en expansiemicroscopie

A Label-free Technique for the Spatio-temporal Imaging of Single Cell Secretions

Inter-cellular communication is an integral part of a complex system that helps in maintaining basic cellular activities. As a result, the malfunctioning of such signaling can lead to many disorders. To understand cell-to-cell signaling, it is essential to study the spatial and temporal nature of the secreted molecules from the cell without disturbing the local environment. Various assays have been developed to study protein secretion, however, these methods are typically based on fluorescent probes which disrupt the relevant signaling pathways. To overcome this limitation, a label-free technique is required.
In this paper, we describe the fabrication and application of a label-free localized surface plasmon resonance imaging (LSPRi) technology capable of detecting protein secretions from a single cell. The plasmonic nanostructures are lithographically patterned onto a standard glass coverslip and can be excited using visible light on commercially available light microscopes. Only a small fraction of the coverslip is covered by the nanostructures and hence this technique is well suited for combining common techniques such as fluorescence and bright-field imaging.
A multidisciplinary approach is used in this protocol which incorporates sensor nanofabrication and subsequent biofunctionalization, binding kinetics characterization of ligand and analyte, the integration of the chip and live cells, and the analysis of the measured signal. As a whole, this technology enables a general label-free approach towards mapping cellular secretions and correlating them with the responses of nearby cells.

Inter-cellular communication is critical for controlling various physiological activities within and outside the cell. This paper describes a protocol for measuring the spatio-temporal nature of single cell secretions. To achieve this, a multidisciplinary approach is used which integrates label-free nanoplasmonic sensing with live cell imaging.

Inter-cellular communication is an integral part of a complex system that helps in maintaining basic cellular activities. As a result, the malfunctioning ...

Bioengineering

Optimized Incorporation of Alkynyl Fatty Acid Analogs for the Detection of Fatty Acylated Proteins using Click Chemistry

Fatty acylation, the covalent addition of saturated fatty acids to protein substrates, is important in regulating a myriad of cellular functions in addition to its implications in cancer and neurodegenerative diseases. Recent developments in fatty acylation detection methods have enabled efficient and non-hazardous detection of fatty acylated proteins, particularly through the use of click chemistry with bio-orthogonal labeling. However, click chemistry detection can be limited by the poor solubility and potential toxic effects of adding long chain fatty acids to cell culture. Described here is a labeling approach with optimized delivery using saponified fatty acids in combination with fatty-acid free BSA, as well as delipidated media, which can improve detection of hard to detect fatty acylated proteins. This effect was most pronounced with the alkynyl-stearate analog, 17-ODYA, which has been the most commonly used fatty acid analog in click chemistry detection of acylated proteins. This modification will improve cellular incorporation and increase sensitivity to acylated protein detection. In addition, this approach can be applied in a variety of cell types and combined with other assays such as pulse-chase analysis, stable isotope labeling with amino acids in cell culture, and mass spectrometry for quantitative profiling of fatty acylated proteins.

The study of fatty acylation has important implications in cellular protein interactions and diseases. Presented here is a modified protocol to improve click chemistry detection of fatty acylated proteins, which can be applied in various cell types and combined with other assays, including pulse-chase and mass spectrometry.

Fatty acylation, the covalent addition of saturated fatty acids to protein substrates, is important in regulating a myriad of cellular functions in ...

Biochemistry

An Endothelial Planar Cell Model for Imaging Immunological Synapse Dynamics

Adaptive immunity is regulated by dynamic interactions between T cells and antigen presenting cells (&#39;APCs&#39;) referred to as &#39;immunological synapses&#39;. Within these intimate cell-cell interfaces discrete sub-cellular clusters of MHC/Ag-TCR, F-actin, adhesion and signaling molecules form and remodel rapidly. These dynamics are thought to be critical determinants of both the efficiency and quality of the immune responses that develop and therefore of protective versus pathologic immunity. Current understanding of immunological synapses with physiologic APCs is limited by the inadequacy of the obtainable imaging resolution. Though artificial substrate models (e.g., planar lipid bilayers) offer excellent resolution and have been extremely valuable tools, they are inherently non-physiologic and oversimplified. Vascular and lymphatic endothelial cells have emerged as an important peripheral tissue (or stromal) compartment of &#39;semi-professional APCs&#39;. These APCs (which express most of the molecular machinery of professional APCs) have the unique feature of forming virtually planar cell surface and are readily transfectable (e.g., with fluorescent protein reporters). Herein a basic approach to implement endothelial cells as a novel and physiologic &#39;planar cellular APC model&#39; for improved imaging and interrogation of fundamental antigenic signaling processes will be described.

Adaptive immunity is controlled by dynamic 'immunological synapses' formed between T cells and antigen presenting cells. This protocol describes methods for investigating endothelial cells both as understudied physiologic APCs and as a novel type of 'planar cellular APC model'.

Adaptive immunity is regulated by dynamic interactions between T cells and antigen presenting cells (&#39;APCs&#39;) referred to as &#39;immunological ...

Immunology and Infection

Label-free Super-resolution Imaging Enabled by Vibrational Imaging of Swelled Tissue and Analysis

The universal utilization of fluorescence microscopy, especially super-resolution microscopy, has greatly advanced knowledge about modern biology. Conversely, the requirement of fluorophore labeling in fluorescent techniques poses significant challenges, such as photobleaching and non-uniform labeling of fluorescent probes and prolonged sample processing. In this protocol, the detailed working procedures of vibrational imaging of swelled tissue and analysis (VISTA) are presented. VISTA circumvents obstacles associated with fluorophores and achieves label-free super-resolution volumetric imaging in biological samples with spatial resolution down to 78 nm. The procedure is established by embedding cells and tissues in hydrogel, isotropically expanding the hydrogel sample hybrid, and visualizing endogenous protein distributions by vibrational imaging with stimulated Raman scattering microscopy. The method is demonstrated on both cells and mouse brain tissues. Highly correlative VISTA and immunofluorescence images were observed, validating the protein origin of imaging specificities. Exploiting such correlation, a machine learning-based image-segmentation algorithm was trained to achieve multi-component prediction of nuclei, blood vessels, neuronal cells, and dendrites from label-free mouse brain images. The procedure was further adapted to investigate pathological poly-glutamine (polyQ) aggregates in cells and amyloid-beta (A&#946;) plaques in brain tissues with high throughput, justifying its potential for large-scale clinical samples.

By combining sample-expansion hydrogel chemistry with label-free chemical-specific stimulated Raman scattering microscopy, the protocol describes how to achieve label-free super-resolution volumetric imaging in biological samples. With an additional machine learning image segmentation algorithm, protein-specific multi-component images in tissues without antibody labeling were obtained.

The universal utilization of fluorescence microscopy, especially super-resolution microscopy, has greatly advanced knowledge about modern biology. ...

Chemistry

Metabolic Glycoengineering of Sialic Acid Using N-acyl-modified Mannosamines

Sialic acid (Sia) is a highly important constituent of glycoconjugates, such as N- and O-glycans or glycolipids. Due to its position at the non-reducing termini of oligo- and polysaccharides, as well as its unique chemical characteristics, sialic acid is involved in a multitude of different receptor-ligand interactions. By modifying the expression of sialic acid on the cell surface, sialic acid-dependent interactions will consequently be influenced. This can be helpful to investigate sialic acid-dependent interactions and has the potential to influence certain diseases in a beneficial way. Via metabolic glycoengineering (MGE), the expression of sialic acid on the cell surface can be modulated. Herein, cells, tissues, or even entire animals are treated with C2-modified derivatives of N-acetylmannosamine (ManNAc). These amino sugars act as sialic acid precursor molecules and therefore are metabolized to the corresponding sialic acid species and expressed on glycoconjugates. Applying this method produces intriguing effects on various biological processes. For example, it can drastically reduce&#160;the expression of polysialic acid (polySia) in treated neuronal cells and thus affects neuronal growth and differentiation. Here, we show the chemical synthesis of two of the most common C2-modified N-acylmannosamine derivatives, N-propionylmannosamine (ManNProp) as well as N-butanoylmannosamine (ManNBut), and further show how these non-natural amino sugars can be applied in cell culture experiments. The expression of modified sialic acid species is quantified by high performance liquid chromatography (HPLC) and further analyzed via mass spectrometry. The effects on polysialic acid expression are elucidated via Western blot using a commercially available polysialic acid antibody.

Metabolic Glycoengineering of Sialic Acid Using N-acyl-modified Mannosamines

Sialic acid is a typical monosaccharide-unit found in glycoconjugates. It is involved in a plethora of molecular and cellular interactions. Here we present a method to modify cell surface sialic acid expression using metabolic glycoengineering with N-acetylmannosamine derivatives.

Sialic acid (Sia) is a highly important constituent of glycoconjugates, such as N- and O-glycans or glycolipids. Due to its position at the non-reducing ...

Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze membrane fusion within eukaryotic cells. The SNARE protein family consists of about 36 different members. Specific intracellular transport routes are catalyzed by specific sets of 3 or 4 SNARE proteins that thereby contribute to the specificity and fidelity of membrane trafficking. However, studying the precise function of SNARE proteins is technically challenging, because SNAREs are highly abundant and functionally redundant, with most SNAREs having multiple and overlapping functions. In this protocol, a new method for the visualization of SNARE complex formation in live cells is described. This method is based on expressing SNARE proteins C-terminally fused to fluorescent proteins and measuring their interaction by F&#246;rster resonance energy transfer (FRET) employing fluorescence lifetime imaging microscopy (FLIM). By fitting the fluorescence lifetime histograms with a multicomponent decay model, FRET-FLIM allows (semi-)quantitative estimation of the fraction of the SNARE complex formation at different vesicles. This protocol has been successfully applied to visualize SNARE complex formation at the plasma membrane and at endosomal compartments in mammalian cell lines and primary immune cells, and can be readily extended to study SNARE functions at other organelles in animal, plant, and fungal cells.

This protocol describes a new method allowing for the quantitative visualization of complex formation of SNARE proteins, based on F&#246;rster resonance energy transfer, and fluorescence lifetime imaging microscopy.

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze ...

Imaging the Human Immunological Synapse

The purpose of the method is to generate an immunological synapse (IS), an example of cell-to-cell conjugation formed by an antigen-presenting cell (APC) and an effector helper T lymphocyte (Th) cell, and to record the images corresponding to the first stages of the IS formation and the subsequent trafficking events (occurring both in the APC and in the Th cell). These events will eventually lead to polarized secretion at the IS. In this protocol, Jurkat cells challenged with Staphylococcus enterotoxin E (SEE)-pulsed Raji cells as a cell synapse model was used, because of the closeness of this experimental system to the biological reality (Th cell-APC synaptic conjugates). The approach presented here involves cell-to-cell conjugation, time-lapse acquisition, wide-field fluorescence microscopy (WFFM) followed by image processing (post-acquisition deconvolution). This improves the signal-to-noise ratio (SNR) of the images, enhances the temporal resolution, allows the synchronized acquisition of several fluorochromes in emerging synaptic conjugates and decreases fluorescence bleaching. In addition, the protocol is well matched with the end point cell fixation protocols (paraformaldehyde, acetone or methanol), which would allow further immunofluorescence staining and analyses. This protocol is also compatible with laser scanning confocal microscopy (LSCM) and other state-of-the-art microscopy techniques. As a main caveat, only those T cell-APC boundaries (called IS interfaces) that were at the right 90&#176; angle to the focus plane along the Z-axis could be properly imaged and analyzed. Other experimental models exist that simplify imaging in the Z dimension and the following image analyses, but these approaches do not emulate the complex, irregular surface of an APC, and may promote non-physiological interactions in the IS. Thus, the experimental approach used here is suitable to reproduce and to confront some biological complexities occurring at the IS.

This protocol images both the immunological synapse formation and the subsequent polarized secretory traffic towards the immunological synapse. Cellular conjugates were formed between a superantigen-pulsed Raji cell (acting as an antigen-presenting cell) and a Jurkat clone (acting as an effector helper T lymphocyte).

The purpose of the method is to generate an immunological synapse (IS), an example of cell-to-cell conjugation formed by an antigen-presenting cell (APC) ...

Identifying Cell Surface Markers of Primary Neural Stem and Progenitor Cells by Metabolic Labeling of Sialoglycan

Neural stem and progenitor cells (NSPCs) are the cellular basis for the complex structures and functions of the brain. They are located in specialized niches in vivo and can be isolated and expanded in vitro, serving as an important resource for cell transplantation to repair brain damage. However, NSPCs are heterogeneous and not clearly defined at the molecular level or purified due to a lack of specific cell surface markers. The protocol presented, which has been previously reported, combines a neural-endothelial co-culture system with a metabolic glycan labeling method to identify the surface sialoglycoproteome of primary NSPCs. The NSPC-endothelial co-culture system allows self-renewal and expansion of primary NSPCs in vitro, generating a sufficient number of NSPCs. Sialoglycans in cultured NSPCs are labeled using an unnatural sialic acid metabolic reporter with bioorthogonal functional groups. By comparing the sialoglycoproteome from self-renewing NSPCs expanded in an endothelial co-culture with differentiating neural culture, we identify a list of membrane proteins that are enriched in NSPCs. In detail, the protocol involves: 1) set-up of an NSPC-endothelial co-culture and NSPC differentiating culture; 2) labeling with azidosugar per-O-acetylated N-azidoacetylmannosamine (Ac4ManNAz); and 3) biotin conjugation to modified sialoglycan for imaging after fixation of neural culture or protein extraction from neural culture for mass spectrometry analysis. Then, the NSPC-enriched surface marker candidates are selected by comparative analysis of mass spectrometry data from both the expanded NSPC and differentiated neural cultures. This protocol is highly sensitive for identifying membrane proteins of low abundance in the starting materials, and it can be applied to marker discovery in other systems with appropriate modifications

Presented here is a protocol that combines an in vitro neural-endothelial co-culture system and metabolic incorporation of sialoglycan with bioorthogonal functional groups to expand primary neural stem and progenitor cells and label their surface sialoglycoproteins for imaging or mass-spectrometry analysis of cell surface markers.

Neural stem and progenitor cells (NSPCs) are the cellular basis for the complex structures and functions of the brain. They are located in specialized ...

Neuroscience

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

Reversed-Phase High-Performance Liquid Chromatography: A Robust Method for Quantitation of Fluorescently Labeled and Derivatized Sialic Acids Isolated from Mouse Liver

Source: Cao, C. et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/v/56030">Determination of Sialic Acids in Liver and Milk Samples of Wild-type and CMAH Knock-out Mice.</a>&#160;J. Vis. Exp.&#160;(2017)
In this video, we demonstrate the detection of sialic acids, N-acetylneuraminic acid and N-glycolylneuraminic acid, using high-performance liquid chromatography, HPLC, following derivatization with a suitable fluorescent labeling reagent to aid in fluorescence detection.

Source: Cao, C. et al. Determination of Sialic Acids in Liver and Milk Samples of Wild-type and CMAH Knock-out Mice.&#160;J. Vis. Exp.&#160;(2017)
In this ...

Visualizing Intracellular Sialylation with Click Chemistry and Expansion Microscopy

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