Ultra plus field emission scanning electron microscope

Transverse sections of all grass leaves were imaged for light microscopy under 10× and 40× objectives using a Nikon Eclipse 50i upright microscope (Nikon Instruments). SEM was performed using a Zeiss Ultra Plus field emission scanning electron microscope at 3 kV. To quantify pit field distribution, two z-stacks from two leaf tissues per species were obtained using a Leica SP8 multiphoton confocal microscope (Leica Microsystems). Details can be found in Danila et al. (2016) (link).

For scanning electron microscopy, leech specimens were treated with 16% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA, USA) or relaxation solution (4.8 mM NaCl₂, 1.2 mM KCl, 10 mM MgCl₂, 8% EtOH) while feeding or relaxing. After treatment, the head region containing the proboscis was cut and fixed in 4% PFA at room temperature overnight. The tissues were washed three times with PBT (1X PBS + 0.1% Tween-20) for 20 min at room temperature, and then fixed in 1% osmium tetroxide (Ted Pella Inc., Redding, CA, USA) in 1 M PBS for 1 h. Osmium tetroxide was removed by washing three times with PBT. Thereafter, the tissues were gradually dehydrated with ethanol (30%, 50%, 60%, 70%, 80%, 90%, 95%, 100% in 1X PBS) for 20 min per step. Dehydrated tissues were treated with stepwise concentrated isopentyl acetate (Alfa Aesar, Ward Hill, MA, USA) (isopentyl acetate: EtOH = 1:3, 1:1, and 3:1) for 15 min per step, and then transferred to 100% isopentyl acetate. After the solution was removed, the samples were dried for 3 days in a fume hood. Dried samples were coated with gold particles and examined with an UltraPlus field emission scanning electron microscope (Carl Zeiss, Oberkochen, BW, Germany).

Primary cultures of lung fibroblasts from Sec13^+/+ and Sec13^H/− mice were prepared for FESEM imaging as previously described47 (link)48 (link). Briefly, cells were detached by trypsinization, washed in PBS, and subjected to two separate rounds of hypotonic treatment (15 mM Tris-HCl, pH 7.4, 10 mM NaCl, 3 mM MgCl₂). The cells were then resuspended in PBS+10% glycerol, spun down onto poly-lysine-coated silicon chips and fixed in 3% glutaraldehyde in PBS. Further processing for FESEM included postfixation in 1% osmium tetroxide, dehydration through a graded ethanol series and critical-point drying on a CPD030 apparatus (Bal-Tec). Samples were sputter-coated with 1–2 nm chromium on an EMITECH K575X apparatus and imaged with an in-lens detector for secondary electrons on a Zeiss ULTRA plus field emission scanning electron microscope.

Mineralogical composition mapping of a single long-term P. onkodes skeleton was also carried out at ANU (Centre for Advanced Microscopy, Dr Frank Brink) using QEMSCAN. This mapping integrates scanning electron microscopy and energy-dispersive X-ray spectroscopy (SEM-EDS) hardware with software to generate micron-scale compositional maps of rocks and sediments and is widely used in mining and petroleum industries [58 ]. To apply this method of mineral analyses to the MgCO₃ coralline algae skeleton, the technique was modified to determine changes in the Mg intensity (i.e., high Mg-calcite, dolomite and magnesite), rather than changes between minerals. Images of these changes in Mg intensity were taken at a resolution of 5 μm. SEM-EDS, using a Zeiss Ultraplus field emission scanning electron microscope, operated at 15.0 kV, 10.9 mm working distance was then used to provide elemental composition using spot analysis within each of these bands. Samples were carbon coated and mounted using carbon tape. As outlined in Nash et al., [8 , 48 ], Mg-calcite was identified ranging from 8–25 mol% MgCO_3, dolomite as 38–62%, and magnesite as >80% (however measurements between 62–99% are thought to be magnesite with small amounts of the neighbouring Mg-calcite or dolomite).

Rab21CA transfomants were plated on different ECM coated surfaces and incubated for 4–6 hrs at 35°C and processed for SEM analysis.
Briefly, log phase trophozoites were harvested and washed with complete medium and then resuspended in warm complete medium containing 15% adult bovine serum. The trophozoites were plated on glass coverslips in a four well plate placed in a BD EZ Gas Pak and incubated at 37°C for an hour for attachment. Further the medium in the wells was replaced with medium containing 2 mg/ml of collagen type I and 1mg/ml of matrigel and trophozoites were incubated for additional 6–8hrs at 37°C in the GasPak. Thereafter, the medium was removed and the cells were briefly washed with warm 0.1M phosphate buffer (pH 7.4) and fixed using 2.5% EM grade gultaraldehyde in 0.1M phosphate buffer (pH 7.4) at 4°C for overnight. Following day, the cells were dehydrated in a graded series of alcohol (25%, 50%, 75%, 95%) for 15min each at RT followed by 100% for 15 min at RT for three times. The samples were then left for drying at RT for 48–56hrs covered with an aluminum foil. The dried samples were sputter coated with gold using Quorum Q150R ES and were examined and photographed with Zeiss ULTRA PLUS field emission scanning electron microscope operating at 5kV.

Bone marrow‐derived macrophages or mid‐logarithmic phase cultures of M. catarrhalis incubated either in the presence or absence of recombinant mGBP2 were washed in PBS, fixed with 2.5% glutaraldehyde in PBS for 3 h and further washed with PBS. Cells were fixed in 1% osmium tetroxide in distilled water for 1 h. Samples were subsequently dehydrated in a series of alcohol and subjected to liquid carbon dioxide critical point drying. Samples were then sputter‐coated with platinum (3 nm thickness) at 15 mA for 2 min using the EMI TECH K550 Sputter coater and visualised under a Zeiss UltraPlus Field emission scanning electron microscope at 2–5 kV.