Application Note 09

Cell Culture and Fluorescence
Staining in the µ-Slide VI 0.4
Fluorescent actin stainings, particularly using phalloidin,
are a powerful approach for getting insight into cellularstructures and processes using microscopy. Thisapproach allows the specific visualization of actin
filaments, making fluorescent actin staining anindispensable tool for scientists to answer numerous
questions related to cell biology, including cytoskeletalorganization and dynamics.
In this Application Note, we present a simple protocol forthe cultivation and fluorescent actin staining of theadherent human fibrosarcoma cell line HT-1080 using
the µ-Slide VI 0.4.

For more insights on the benefits of using theµ-Slide VI 0.4, take a look at Application Note 03 “CellCulture in ibidi Channel Slides”.

Please note: before starting a fluorescent actin staining,several important parameters must be checked, such as
the optimal cell density, the ideal cell culture vesselgeometry and substrate/coating. In addition, positive
and negative controls are necessary to validate thestaining. Therefore, we strongly recommend a thoroughliterature research before the experiment.

Do you want to combine this protocol with an
immunofluorescence assay? Find more detailed
information about planning and conducting
immunofluorescence assays in our Application
Chapter “Immunofluorescence”.

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Application Note 09

1. Materials
1.1. Reagents and Buffers

Cell Culture••••

HT-1080 cells (85111505, Sigma Aldrich)
Cell culture medium: DMEM (D6546, Sigma Aldrich) with 10 % Fetal Bovine Serum(F1283, Sigma Aldrich)
D-PBS (14190144, Gibco)
Accutase (A1110501, Gibco)

Fluorescence Staining and Imaging••
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D-PBS (14190144, Gibco)
Formalin, 10 %, ready to use (HT5011, Sigma Aldrich)
Triton-X-100 (A16046, Thermo Fisher Scientific)
Permeabilization Buffer (0.1 % Triton-X-100 in D-PBS)
Bovine Serum Albumin (BSA) (A1470-10G, Sigma Aldrich)
Blocking Buffer (1 % BSA in D-PBS)
Phalloidin solution: 1 µl of Phalloidin-iFluor 488 Reagent (ab176753, Abcam) in 1 ml ofblocking buffer
ibidi Mounting Medium With DAPI (50011, ibidi)
ibidi Immersion Oil (50101, ibidi)

1.2. Equipment••••µ-Slide VI 0.4 ibiTreat (80606, ibidi)µ-Slide Rack (80003, ibidi)Standard cell culture equipment (pipettes, sterile working bench, cell culture incubator,culture flasks, cell culture medium, hemocytometer, etc.)
Inverted fluorescence microscope with appropriate filter sets

2. Methods
2.1. Cell Cultivation
Please read the Instructions before working with the µ-Slide VI 0.4. Perform all steps under sterile
conditions. It is recommended to place the µ-Slide VI 0.4 and the cell culture medium into the incubatorthe day before seeding the cells to avoid the formation of air bubbles during handling. Before starting
the experiment, prepare the HT-1080 cells in a standard cell culture flask (e.g., T75) with adherent
cells at the bottom. The cells should ideally be subconfluent and healthy on the day of the experiment.
It is essential to work swiftly during the whole procedure to prevent the cells from drying.
If not stated otherwise, all given volumes are per channel, and all incubation steps are at roomtemperature.
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Application Note 09

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Add 10 ml Accutase to the T75 flask for celldetachment; incubate for 5 min in the incubator(37 °C, 5 % CO2).
Harvest the cell suspension, centrifuge, and dilute it
in a low amount of cell culture medium for counting.
Count the cells and adjust to a final concentration of
3 × 105 cells/ml in cell culture medium.
Unpack an ibidi µ-Slide VI 0.4 and put it on a µ-Slide
Rack or an appropriate surface.
Pipet 30 µl of the HT-1080 cell suspension into eachchannel by pipetting directly into the channel. Quick
dispensing helps to avoid trapped air bubbles.
Remove trapped air bubbles from the channel by
inclining the µ-Slide and tapping on one edge.
Cover the reservoirs with the supplied lid.
Put the µ-Slide with the rack into the incubator (37 °C,5 % CO2) and let the cells attach for 1 h.
Fill 60 µl of cell-free cell culture medium into each of thereservoirs. Do not trap air bubbles.
Put the µ-Slide with the rack into the incubator (37 °C,
5 % CO2) and incubate the cells overnight.
For extended cell cultivation, we recommend acontinuous medium exchange every second day (seeChapter 2.2).

Filling a cell suspension into achannel of the µ-Slide VI 0.4.

Filling the Luer reservoirs withcell-free culture medium.

2.2. Continuous Medium Exchange••••

Carefully aspirate the medium from the reservoirs. Do not aspirate any liquid from thechannel itself.
Gently introduce 120 µl of cell-free culture medium into one reservoir, allowing the channel
to replenish.
Aspirate the old culture medium from the opposite reservoir. Be careful when using a cell
culture aspiration device, as this may flush away partially attached cells or clusters.
Refill the reservoirs using 60 µl cell-free culture medium per reservoir.

Continuous medium exchange with minimum three times the channel volume.

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2.3. Fixation, Permeabilization, and Blocking
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Perform all steps quickly to ensure that the channels do not dry out. Prepare enough
Permeabilization Buffer and Blocking Buffer for your experiment.
Replace the cell culture medium with D-PBS through continuous medium exchange forwashing.
Aspirate most of the D-PBS. Do not aspirate the entire volume.
Fix the cells with 100 µl formalin (10 %) for 20 min.
Wash the cells three times with 200 µl D-PBS through continuous medium exchange.
Aspirate most of the D-PBS. Do not aspirate the entire volume.
Incubate the cells in 100 µl Permeabilization Buffer for 5 min.
Wash the cells with 200 µl D-PBS through continuous medium exchange.
Aspirate most of the D-PBS. Do not aspirate the entire volume.
Block with 100 µl Blocking Buffer for 20 min.
Wash the cells with 200 µl D-PBS.

2.4. Staining•••

Aspirate all D-PBS from reservoirs and channel. The
use of a 5 ml syringe to fill the empty channels
minimizes the risk of introducing air bubbles.
Immediately apply 30 µl of the phalloidin solution to thechannel; incubate for 20 min in the dark. From this pointon, the samples should be kept in the dark wheneverpossible.
Wash the cells twice with 200 µl D-PBS throughcontinuous medium exchange.

2.5. Mounting•••

Aspirate all D-PBS and immediately add ibidi Mounting
Medium With DAPI for nuclear staining until thechannel is filled. If a different mounting medium isused, please note that it must be non-drying to avoid
damage of the µ-Slide.
Store at 4 °C in the dark until imaging.
The stained µ-Slide can be stored for up to 4 weeks.
Ideally, proceed immediately with imaging since longer
storage times can reduce image quality.

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2.6. Imaging••

Observe the cells under a fluorescence microscopewith appropriate filter sets and, if necessary, withibidi Immersion Oil.
Optionally, overlay images to create a merged image.

Which immersion oils are compatible with the ibidi labware
products? Find out at: ibidi.com/oil

3. Results

Widefield fluorescence microscopyof HT1080 cells in a µ-Slide VI 0.4.
The F-actin cytoskeleton wasvisualized using phalloidin (green).
Nuclei were stained with DAPI(blue). The imaging was performed
on a Zeiss Axiovert 135 microscopeusing a Plan-Neofluar 40x/0.75
objective lens.

Your fluorescence staining did not work out as expected? Please refer to ourimmunofluorescence troubleshooting guide!

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