The F. oxysporum f. sp. lycopersici race 2 strain 4287 (FGSC 9935) was used in all experiments. The F. oxysporum mutant constitutively expressing histone H1 fused to mCherry red fluorescent protein (H1-mCherry) was previously described (Ruiz-Roldan et al., 2010). To obtain a F. oxysporum strain simultaneously expressing both H1-mCherry and the Lifeact-sGFP fluorescent reporter for F-actin visualization (Fernández-Ábalos et al., 1998; Riedl et al., 2008), protoplasts of the previously obtained H1-mCherry strain were co-transformed with a hygromycin resistance cassette plus a P_gpdA::LifeAct-sGFP linear fragment (for details see Supplementary Text S1 and Supplementary Table S1), as previously described (Di Pietro et al., 2001; López-Berges et al., 2012). For microconidia production, cultures were grown in potato dextrose broth (PDB; Sigma P6685) at 28°C and 120 rpm for 4–7 days. Microconidia were filtered through a custom-made cotton filter system, harvested by centrifugation and washed twice with pure water.

The U. maydis strains expressing opsin1 fused to enhanced green fluorescent protein (eGFP) were described in detail before (Panzer et al., 2019) and derived from the wild type isolate FB1 (Banuett and Herskowitz, 1989). Either the strain FB1 ΔUmOps1 P_crg::UmOps1-eGFP K1 or FB1 P_crg::UmOps1-eGFP KA was used for ExM experiments. U. maydis sporidia were grown as described before (Panzer et al., 2019). Sporidia were grown in PDB for 15–24 h at 28°C and 100 rpm, harvested by centrifugation (4000 × g, 3 min), washed once in ddH₂O and resuspended in ddH₂O to a final density of 6.5 x 10⁵ sporidia/mL. If required, expression of UmOps1-eGFP was induced by treatment with arabinose-containing induction medium (Panzer et al., 2019).

The non-homologous end joining-deficient A. fumigatus strain AfS35 (Krappmann et al., 2006; Wagener et al., 2008) was transformed with plasmids pYZ011 and pYZ012. pYZ012 harbors a phleomycine resistance cassette and encodes a mitochondria-targeted red fluorescent protein (RFP) fusion protein that consists of the N-terminal part (52 amino acids) of Aspergillus niger citrate synthase followed by mRFP1 under control of the Aspergillus nidulans gpdA promoter. pYZ011 harbors a pyrithiamine resistance cassette and encodes A. fumigatus AFUA_1G10040 including 230 bp of its promoter region, followed without stop codon by the coding sequence of a GFP (S65T) (Heim et al., 1995), FPbase ID: B6J33 derivative with the following modifications: M1_S2insV, E235_K238delinsSCTSKISRPRETW. The strain was cultivated on solid Aspergillus minimal medium (AMM) (Hill and Kafer, 2001) in T75 tissue culture flasks (Sarstedt) in order to avoid uncontrolled spreading of the hydrophobic fungal spores. Spores were harvested by submerging them with PBS and resuspending them by means of glass beads.

Fungal spores and sporidia were seeded onto poly-D-lysine (PDL)-coated coverslips in 4-well tissue culture plates (800 μL/well). While sporidia were investigated after 30 min of sedimentation, conidia were allowed to germinate in the respective culture medium for 18 h (A. fumigatus) or 14–19 h (F. oxysporum). If indicated, fungal cells were incubated for 5–10 min in 0.5 μM mCling-Atto643 fluorescent dye dissolved in nutrient media to stain the membrane. After three washing steps with PBS the samples were either directly fixed in 4% formaldehyde and 0.25% glutaraldehyde for 15 min (standard procedure) or, as in case of microtubule detection in U. maydis sporidia, treated according to the protocol of Michie et al. (2017). Briefly, sporidia were prefixed and permeabilized for 1 min in prewarmed cytoskeleton buffer (10 mM MES buffer pH 6.1, 150 mM NaCl, 5 mM EGTA, 5 mM glucose and 5 mM MgCl₂) containing additionally 0.25% Triton X-100 and 0.3% glutaraldehyde and finally fixed for 10 min in cytoskeleton buffer supplied with 2% glutaraldehyde. For quenching of autofluorescence, fixation was followed by a 7 min incubation in 0.1% NaBH₄. After consecutive washing, cell walls were digested for 1 h at RT with a cell wall lytic enzyme solution, based on an enzymatic mix (0.1 g lysing enzyme of T. harzianum, 0.25 g driselase, and 0.5 mg chitinase dissolved in 10 ml 0.7 M NaCl) that was used for the generation of Fusarium fujikuroi protoplasts before (García-Martínez et al., 2015). The enzymatic solution was either directly used in the experiments or stored at −80°C for later use. For treatment of young germlings and U. maydis sporidia the enzyme solution was diluted in a ratio of 1:5 with 0.7 M NaCl. In case additional antibody-staining was required, the samples were blocked for 30 min in 5% BSA / 0.25% Triton X-100 and subsequently incubated with the primary anti-α-tubulin antibody (abcam, ab18251) for 1 h. After washing, the samples were incubated for another hour in the corresponding secondary antibody (Alexa 488-label, Thermo-fisher, A11008 or ATTO647N-label, Sigma, 40839) resolved in blocking solution and washed with PBS. All samples were instantly processed for ExM.

Immediately before gelation, as previously published (Chozinski et al., 2016; Kunz et al., 2019), the samples were incubated for 10 min with 0.25% glutaraldehyde and washed with PBS. Thereafter, a droplet of the monomer solution [8.625% sodium acrylate (Sigma, 408220), 2.5% acrylamide (Sigma, A9926), 0.15% N,N′-methylenbisacrylamide (Sigma, A9926), 2 M NaCl (Sigma, S5886), 1 × PBS and 0.2% freshly added ammonium persulfate (APS, Sigma, A3678) and tetramethylethylenediamine (TEMED, Sigma, T7024)] was prepared on parafilm in a humid Petri dish. Using tweezers, the coverslip with the attached fungi was then transferred upside-down on the gelation droplet. The sample was allowed to gelate for at least 1 h at RT in the closed dish. To ensure isotropic expansion, samples were homogenized (Gao et al., 2017) in digestion buffer [50 mM Tris pH 8.0, 1 mM EDTA (Sigma, ED2P), 0.5% Triton X-100 (Thermo Fisher, 28314), and 0.8 M guanidine HCl (Sigma, 50933)] supplied with 8 U/ml proteinase K (Thermo Fisher, AM2548) for 1 h to overnight. This step is required to reduce the cohesion of the fixed proteins while most cellular compounds are washed out the gel. At the same time the majority of the fluorophores remain attached to the polymer (Tillberg et al., 2016). Digested samples were expanded in ddH₂O for 3–4 h. The water was changed every hour until the size of the gel did not increase any more. Expanded gels were stored at 4°C until use. The expansion factor was determined by both the diameter of the fungi as well as by the gel size before and after expansion. Imaging was performed in PDL-coated chambers (Merck, 734-2055) to immobilize the gels.

Imaging was performed on a confocal inverted system (Zeiss LSM700) or on SIM system (Zeiss ELYRA S.1 SR-SIM) equipped with an 63x oil (used for unexpanded samples) and a 63x water-immersion objective (used for ExM samples; C-Apochromat, 63 x 1.2 NA, Zeiss, 441777-9970). The water objective was necessary to provide sufficient working distance to be able to image the expanded samples. Unexpanded CLSM images were captured using the ideal pixel sizes provided by the software (between 740 x 740 and 856 x 856 pixels) and a pixel dwell time between 1.58 μs and 4.24 μs using laser powers ranging between 1.5 and 10%. The expanded samples were then imaged again with the optimum pixel size and pixel dwell time ranging from 6.30 μs to 8.43 μs, using laser powers between 10 and 26% with lasers of 488 nm, 555 nm and 639 nm. The pinhole was adjusted to 1 airy unit and the photomultiplier was set to 700. SIM images were reconstructed with the ZEN image processing platform of the SIM module, with a fixed pixel size of 31 nm. Laser power ranged between 8 and 25% using laser of 488 nm, 561 nm, and 642 nm with integration times between 100 ms and 300 ms. For final image processing, Imaris 8.4.1 and FIJI 1.51 (Schindelin et al., 2012) were used.

Expansion microscopy relies on isotropic expansion of all cellular structures during gel swelling. Therefore, it is crucial that the fungal cell wall is completely digested to enable the uniform movement of labeled proteins and/or fluorophores during the swelling process.

The fungal cell wall contains rigid polymers to maintain turgor pressure and to avoid undesired cell swelling in fungi, and thus needs to be removed to enable uniform expansion of the fungal cell in the polyelectrolyte hydrogel. We successfully degraded the cell wall in F. oxysporum and A. fumigatus germlings and U. maydis sporidia using a mixture of glucanex, driselase and chitinase. Complete removal of cell wall material was confirmed by labeling with the chitin-specific dye calcofluor white (Supplementary Figure S1). The used expansion protocol was based on previously established protocols (Chozinski et al., 2016; Kunz et al., 2019). We noted that the time point of cell wall digestion within the protocol workflow significantly affected the morphology obtained after the expansion process. Cell wall lysis before fixation resulted in the generation of protoplasts with spherical shape, leading to impaired distribution of the subcellular structures as compared to untreated cells (Figure 1). Protoplasts might still be useful for imaging depending on the scientific question addressed. On the other hand, when cell wall digestion was performed after fixation, the original shape of the hyphae and sporidia was preserved (Figure 1).

In the three fungal species tested, cell wall removal was successfully accomplished after fixation with either 4% formaldehyde or 4% formaldehyde + 0.25% glutaraldehyde. As reported previously (Li et al., 2017; Wu and Chou, 2019), the germination time of F. oxysporum conidia had a significant impact on the outcome of cell wall digestion, with longer germination times resulting in incomplete cell wall digestion that impaired the expansion process (Supplementary Figure S2).

After digestion of the cell wall, fungal cells were expanded by a factor of 4.53 ± 0.08 (n = 9) as determined from the gel dimensions before and after expansion. In expanded samples, details of subcellular structures were visualized by CLSM and SIM as described in detail in the following sections.

Ustilago maydis is a dimorphic fungus, which undergoes a morphological transition from the yeast form to the filamentous form (Kahmann and Kämper, 2004; Vollmeister et al., 2012). The haploid sporidia represent the yeast form, which proliferates by budding. Untreated sporidia have a diameter of 2.4 ± 0.35 μm (n = 26) and exhibit subcellular structures that are below the diffraction limit of resolution and thus cannot be resolved by conventional microscopy (Figure 2A). Here we found that the cell wall of sporidia was easily removed by treatment with the lytic enzyme cocktail, which allowed expansion by a factor of 4.6 resulting in a diameter of 11 ± 1.08 μm (n = 9). Confocal images of expanded cells accurately visualized the cytoskeleton and plasma membrane (Figure 2B). Most importantly, membrane vesicles were clearly visible in the expanded samples (Figure 2B). These results demonstrate that ExM can isotropically expand intracellular structures of fungi.

Next, we investigated whether or not membrane proteins in U. maydis can be visualized in expanded samples. To test this, we used a U. maydis strain heterologously expressing UmOps1-eGFP, a microbial rhodopsin that was recently shown to act as a green-light driven proton pump (Panzer et al., 2019). Localization of UmOps1 in the plasma membrane could be observed before (Figure 2C) and after expansion (Figure 2D). In the latter case, the fluorescent membrane protein was isotropically expanded 4.5-fold, visualizing the shape of the expanded sporidium, and the fluorophore density decreased 91-fold. As a consequence, in some areas the fluorescence appeared weak or non-homogeneously labeled in the expanded images (Figure 2D).

The soil-inhabiting ascomycete F. oxysporum causes vascular wilt disease in more than a 100 different crop species and has been reported as an opportunistic human pathogen. Similar to U. maydis, the plasma membrane of F. oxysporum can be stained with mCling dye, which stably remains in membranes after fixation (Revelo and Rizzoli, 2016).

We used a F. oxysporum strain expressing histone H1 labeled with mCherry (Ruiz-Roldan et al., 2010). With a hyphal diameter of 2.89 ± 0.49 μm (n = 17), the standard SIM image provides only limited information on the intracellular structure (Figure 3A). In contrast, imaging of the expanded fungal hypha with an average diameter of 12.73 ± 0.71 μm (n = 4), clearly revealed different membrane embedded organelles and vesicles in addition to the histone-H1-labeled nuclei (Figure 3B). Depending on the time of staining, mCling tends to stain also the intracellular membranes as seen in the expanded F. oxysporum hypha that was incubated with mCling for 10 min (Figure 3B).

By using the F. oxysporum histone H1-mCherry strain as genetic background, we generated a strain expressing both histone H1-mCherry and the F-actin reporter Lifeact-sGFP (Riedl et al., 2008). Figures 3C,D show the distribution of filamentous actin together with the nuclei. The formation of actin filaments at the hyphal tip was visible both in the expanded and non-expanded sample, but the distinct actin cables (bundles of actin filaments) could not be resolved well in the non-expanded sample due to the small diameter of F. oxysporum hyphae, a situation similar to that reported in A. nidulans (Bergs et al., 2016). The superior resolution provided by ExM allowed observation of the actin cables, similar to N. crassa hyphae which naturally exhibit a much larger hyphal diameter (Berepiki et al., 2010). Within the nucleus, regions with higher and others with lower fluorescence intensity were visible, possibly reflecting differences in histone density as described in living cells of mammals (Nozaki et al., 2017) and plants (Rutowicz et al., 2018) PREPRINT. Such differences may appear more pronounced due to the diluted fluorescence intensity after expansion.

Aspergillus fumigatus, a mold with worldwide distribution, produces masses of highly hydrophobic conidia that are ubiquitous in the air and can cause life-threatening invasive aspergillosis when inhaled by immunocompromised patients. We used ExM to visualize an A. fumigatus strain expressing mRFP fused to the N-terminus of citrate synthase, thus conferring localization in the mitochondria. Cell wall digestion of A. fumigatus hyphae was obtained within the first 18 h of germination (Figure 4). The membrane dye mCling was successfully used to visualize the shape of hyphae. Typically, A. fumigatus hyphae exhibited mean diameters of 2.43 ± 0.27 μm (n = 9), which increased 4.4-fold to 10.72 ± 0.69 μm (n = 17) after digestion and expansion.

While the shape of single mitochondria could not be distinguished well before expansion (Figure 4A), it was clearly resolved in the expanded hyphae (Figure 4B). However, fluorescence intensity of mRFP1 was low, possibly due to the proteinase treatment. The use of different proteinases or antibodies as an alternative to fluorescent proteins could further enhance the fluorescence signal.