Seventy to One Hundred Then Three Days for the Medium

9:47 AM — Box labelled “Lichen—March 6” at back of bench. Eight thin sections floating in 70% ethanol, three months old. Originally sliced for the spot test work in June but never mounted. Fixative smells like solvent and time.

Pulled one section out with forceps. Translucent, gray-green cortex visible against white medulla layer. About 2mm thick—hand-sectioned with a razor blade, not microtomed. Rough edges.

First mistake: tried to mount it wet directly from the ethanol. Drop on slide, coverslip, check under scope at 40×. Blurry. Air bubbles trapped at the interface, specimen floating, coverslip skating around on surface tension. Also the ethanol evaporated in under two minutes, leaving the specimen stuck to the glass but dried and opaque.

Alcohol gradient. Should’ve remembered from the March microscopy session—prepared slides aren’t just “put thing on glass.” There’s a protocol.

10:23 AM — Dug out the paraffin embedding kit I bought for something else and never opened. Four small jars: 80% ethanol, 95% ethanol, 100% ethanol, xylene. Label says “dehydration series.” The lichen section has been sitting in 70% for three months. Need to walk it up to 100% before infiltrating with wax or mounting medium.

Transfer to 80%: five minutes. Transfer to 95%: five minutes. Transfer to 100%: ten minutes. During this I read about “clearing agents”—xylene is miscible with both alcohol and paraffin, acts as a bridge. Without it, the wax won’t penetrate because water and wax don’t mix, and ethanol and wax barely tolerate each other.

Into xylene: specimen turned transparent. Genuinely transparent—could see through the medulla to the far cortex. “Clearing” is literal. Chemical makes tissues optically uniform by replacing fluids with something that has a refractive index close to the cellular structure. Same principle as immersion oil in high-magnification objectives.

11:04 AM — Mounting medium is Permount. Comes in a brown glass bottle with a dropper, smells like industrial solvent, label warns about spontaneous polymerization if you leave the cap off. Refractive index 1.52, close to glass at 1.518, which means less light scatter at boundaries.

Drop of Permount on slide. Specimen transferred from xylene bath directly into the drop using bent forceps. Coverslip lowered at 45-degree angle to avoid trapping air. Pressed gently. Medium spread to edges, excess squeezed out. Set aside to cure.

Curing time: 24-48 hours depending on temperature and how much medium you used. Used too much. Pooled around the edges, will take three days minimum.

Second specimen while waiting. This time I want to stain it first.

11:38 AM — Safranin stain, leftover from something. Instructions say it binds to lignin and cellulose, turns plant cell walls red. Lichen thalli aren’t quite plants—they’re fungal hyphae wrapped around algal cells—but both components have cell walls, so maybe.

Dehydration series again: 70% → 80% → 95%, then into safranin solution for two minutes. Specimen turned dark pink immediately. Rinsed in 95% ethanol—some dye washed out, leaving differential staining. Cortex darker than medulla.

Then 100% ethanol, xylene, Permount, coverslip. This one I used less medium. Looks better already under the stereoscope at 20×.

12:46 PM — First slide won’t be ready for days, but I checked it under 100× anyway. Mostly just saw mounting medium and bubbles. One corner of the coverslip had settled enough to bring the specimen into the focal plane: fungal hyphae visible as thin threads, algal cells as dark green spheres clustered in the medulla layer. Could see individual cell walls in the cortex where the safranin stained.

Köhler illumination setup from March still holds. Adjusted condenser height slightly for the increased thickness—this slide is maybe 1.2mm total with the specimen, vs 1mm for a bare slide.

Coverslip thickness: #1.5, which is 0.17mm. Bought a box of 100 from the lab supply place. The cheap coverslips I tried first were unmarked thickness, probably #1 or thinner, caused spherical aberration above 40× magnification. Image looked soft, low contrast, like shooting a lens wide open with the wrong filter. Switched to certified #1.5 and the 100× objective snapped into focus. Thickness matters more than I expected.

1:52 PM — Third specimen, trying hematoxylin. This is the BSC-certified batch—Biological Stain Commission tested it, stamped the label, published the lot number in Biotechnic & Histochemistry. The non-certified stuff I almost bought would’ve been cheaper and possibly useless. Stain consistency varies between manufacturers, sometimes between batches. Certification is the only guarantee you’re getting actual hematoxylin and not mystery dye that happens to be purple.

Hematoxylin stains nuclei blue-purple. Lichen cells have nuclei—both fungal and algal—so this should differentiate them from the extracellular matrix.

Dehydration to 95%, into hematoxylin for ninety seconds, rinse in tap water (tap water, not distilled—pH matters for the colour shift, municipal water is slightly alkaline and “blues” the stain), quick rinse in 95%, then 100%, xylene, mount.

This one I got the medium thickness right. Thin layer, coverslip settled flat, no excess pooling. Under 40× magnification: nuclei showed up as dark purple dots throughout the medulla and cortex. Algal cells more densely packed, multiple nuclei per cluster. Fungal hyphae walls unstained except where nuclei interrupted them.

2:31 PM — Seven specimens left in the box. Five more sections to process, or I can leave them in 70% ethanol for another three months until I need them.

Permount is drying on my fingers. Smells like xylene and impatience. The first slide still won’t be ready until Thursday.

The second slide—safranin-stained, less medium—is already readable at 200×. Cell walls sharp, hyphae threading between algal clusters, cortex structure layered like shingles. Could see where I cut through during the March sectioning: torn cells at the edge, ragged boundary, not the clean microtomed sections you’d get in a real histology lab. But good enough to read the anatomy.

Slide labels are written in pencil on frosted glass at one end. “Lichen—March 6—Safranin—June 23.” Pencil is permanent, supposedly archival. Ink fades, especially under immersion oil. Labels are half the work—unlabeled slides are mysteries six months later.

Valap sealant recipe is equal parts vaseline, lanolin, paraffin wax. Melt together, paint around the coverslip edge to seal against evaporation and oxidation. Haven’t tried it yet. Permount polymerizes and seals itself, but for wet mounts—live specimens, aqueous media—you need Valap or the preparation dries out in twenty minutes. Filing that away.

The third slide is curing. Hematoxylin-stained, nuclei visible, ready for higher magnification once the medium hardens enough to stop the coverslip from shifting under pressure.

Five unstained sections still in the ethanol jar.