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Experiments were carried out by measuring cell fluorescence at 25°C (λEx = 488nm, λEm = 516nm) before and after the addition of compounds.
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In an experiment reported by Newman et. al. [ 24], the abundance of yeast proteins was compared between two states: YEPD (rich), and SD (minimal) by measuring cells' fluorescence due to GFP-tagging of individual proteins.
Samples were withdrawn at 0, 8, 24, 48, 72 and 96 h to measure cell concentration, fluorescence (GFP) and luminescence (Gluc).
We used flow cytometry to measure cell number and individual cells' fluorescence.
To measure total cell fluorescence intensity, cells were fixed with paraformaldehyde (3.7%, RT, 15 min) and permeabilized with 0.1% Triton X-100.
Data were obtained before the addition of the probe to measure background cell fluorescence and after 10 min equilibration with TMA-DPH (240 nM final).
To quantify these differences, we measured cell size with a fluorescence-activated cell sorter (FACS); the forward-angle light scatter (FSC) parameter correlates with cell diameter [7].
This was found to be more reliable than measuring whole-cell fluorescence.
Flow cytometry was utilized for lipid analysis by measuring the single cell fluorescence of cells stained with Nile Red, a lipophilic dye, as a relative lipid indicator.
However, when cell mortality starts to be important, the model cannot be quantitatively compared to the experimental measures of cell fluorescence, since in the simulations dead cells are immediately removed, while in the culture cells that do not divide seem to accumulate fluorescence before lysing, thus increasing the weight of high fluorescence classes.
Calcium release from intracellular stores and store-operated calcium influx after treatment with either bFGF or calcium ionophore A 23187 were measured by single cell fluorescence technique.
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