Formalin

Upon sacrifice, ulnae were dissected and fixed overnight at 4 °C. For histological analysis, specimens were fixed in 10% formalin (Sigma-Aldrich), decalcified, embedded in paraffin, sectioned longitudinally in 5μm increments, and stained with hematoxylin and eosin (H&E) (Sigma-Aldrich) for 10 min and 30 s, respectively. Micrographs were collected with a CKX41 inverted microscope (Olympus) at 20× magnification. To determine whether our ablation model was successful in inducing cell death, ulnae were fixed in 4% paraformaldehyde (Sigma-Aldrich), decalcified, and cryosectioned longitudinally in 5μm increments. GFP was visualized and micrographs were collected with a confocal microscope (Olympus) at 100× magnification.
To detect primary cilia in vitro, isolated and sorted primary periosteal OCPs were seeded on glass bottom dishes (MatTek, Ashland, MA) at 2.5 k per dish 24 h prior to experimentation. Following treatment, cells were fixed in 10% formalin, blocked with 15% goat serum, and incubated in a primary antibody for acetylated α-tubulin acquired from a C3B9 hybridoma line (Sigma-Aldrich), followed by a fluorescent secondary (1:500, Alexa-Fluor 488, Life Technologies). All incubations were conducted at room temperature for 1 h. Micrographs were collected with a confocal microscope (Olympus) at 100X magnification.

The lungs of each mouse were first inflated with 1 mL formalin (Sigma-Aldrich) via the trachea, then excised and fixed in 4% buffered formalin. Lung tissue was embedded in paraffin, cut into 5-mm sections, and stained with hematoxylin and eosin. Inflammatory cell infiltration and lung architecture were assessed by light microscopy. The mucus secretion level was detected by periodic acid-Schiff (PAS; Sigma-Aldrich) staining. Lung sections were deparaffinized and hydrated in water, and stained with periodic acid for 5 minutes. After rinsing with distilled H₂O, lung sections were stained with Schiff reagent for 15 minutes and washed in tap water. Finally, the sections were counterstained in Mayer's hematoxylin, dehydrated, and mucus secretion was assessed by light microscopy.

Marc-145 cells were incubated with the master seed virus inoculum in MEM with 5% FBS and 1% penicillin-streptomycin at 37°C with 5% CO₂. The AHSV infectivity titers were calculated before concentration as ≥1 × 10⁷ plaque-forming units/mL. After 7 days of infection, virus particles and cell suspension were harvested by freeze-thawing 3 times. After centrifugation, AHSV was inactivated by the addition of 0.1% formalin (Merck, Germany) and incubated for 2 weeks. The virus was concentrated 10 times and formalin removed using the Labscale™ Tangential Flow Filtration System (Millipore, USA) for prototype vaccine production. The inactivated virus solution was tested for residual viable virus by passaging in MARC-145 cells in T75 tissue culture flasks. The AHSV’s residual viral RNA was detected in the final passages by PCR. All inactivated viral suspensions were stored at 2–8°C until formulation with an adjuvant [14 , 15 ].

To produce experimental tonic pain in rats, 50 μL of 2.5% formalin (prepared by diluting 38% formalin (Merck, Cat. No.: 818708) with distilled water) was injected subcutaneously (SC) into both the hind paw and the right paw using a 1 mL (27‐gauge) syringe.
The formalin pain test was performed first, followed by the euthanasia of the animals with CO₂ gas. The raphe magnus nucleus, thalamus, and PAG nucleus were isolated. The samples were then immediately transferred to the freezer where they were kept at −70°C for the next steps (RT‐qPCR and ELISA).

One-hundred, 500 or 1000 keratinocytes were plated on a 3T3 feeder layer per 10 cm diameter dish or per well of a six-well dish. After 12 days, feeders were removed and keratinocyte colonies were fixed in 10% formalin (Sigma) for 10 min then stained with 1% Rhodanile Blue (1:1 mixture of Rhodamine B and Nile Blue A [Acros Organics]). Colonies were scored using ImageJ and clonogenicity was calculated as the percentage of plated cells that formed colonies.