CELL BIO WEEK 11

NIH 3T3 cells

An immortal mouse cell line that spontaneously immortalized and has a stable growth rate.

Immortal (immortalized) cell line

A population of cells from a multicellular organism that, because of mutation, keeps going through the cell cycle and dividing, when normally such cells would not proliferate indefinitely. The mutations can occur naturally or be induced by experimenters.

Immortal cell lines vs. stem cells

Stem cells divide indefinitely as part of the organism’s normal developmental program; immortal cell lines divide indefinitely because of mutations. NIH 3T3 cells are not stem cells.

Adherent cells

Cells that grow attached to the surface of the culture vessel, so they must be detached (here with trypsin/EDTA) to be passaged. [definition not in manual]

Confluent / percent confluence

How much of the culture surface is covered by cells; a confluent culture covers the whole surface. You record percent confluence before passaging. [definition not in manual]

Ethanol fixation

Ethanol is a non-additive precipitant fixative: it fixes cell components by dehydration and precipitation, without forming covalent bonds. How well it fixes depends on how much water is present.

What happens to lipids in ethanol fixation, and why does it matter for PI?

Lipids are not preserved, and some dissolve in the ethanol, which makes the plasma membrane permeable to PI. PI can then stain all the double-stranded nucleic acids in the cells.

Can PI enter living cells?

No. An intact cell membrane is not permeable to PI; a permeable membrane is a sign of cell death or apoptosis, so in a growing culture PI stains only dead cells.

Propidium iodide (PI)

A fluorescent dye that binds double-stranded nucleic acids, both DNA and RNA. It is carcinogenic, so wear gloves.

Structure of PI

It resembles ethidium bromide: three joined, conjugated six-membered rings. The molecule is flat.

How does PI bind DNA and dsRNA?

By intercalation: the flat molecule inserts itself between the stacked base pairs of dsDNA and dsRNA.

RNase

A nuclease that degrades both double-stranded and single-stranded RNA into small fragments; small dsRNA pieces under 10 base pairs become single-stranded.

Why add RNase while staining with PI?

It removes all double-stranded RNA, so PI binds DNA specifically. The PI fluorescence of each cell then measures its DNA content.

Why doesn’t the free, unbound PI around the cells fluoresce much?

Free PI absorbs the excitation light, but solvent water quenches most of its excited-state energy. PI intercalated in DNA sits in a hydrophobic, water-free environment, so it emits a photon as fluorescence.

How is DNA content measured in this lab?

PI-stained, RNase-treated cells are photographed by fluorescence microscopy, and FIJI measures the total fluorescence (integrated density) of each nucleus.

DNA content through the cell cycle

G1: 2N (diploid). S: between 2N and 4N as DNA is replicated. G2 and M: 4N. [not in manual]

How does integrated density tell you a cell’s cell-cycle stage?

Integrated density is the total PI fluorescence of the nucleus, which reflects its DNA content: nuclei with about twice the G1 value are in G2/M, and those in between are in S. [the G1/S/G2 values are not in manual]

Why must every fluorescence photo use the same settings?

So the data are consistent between images: same camera zoom and same camera illumination setting, and on the AxioLab A.1 the same microscope fluorescence intensity.

Why must the cells be in sharp focus?

FIJI can’t threshold out-of-focus cells, so blurry images can’t be analyzed.

Photos needed for the FIJI analysis

Several fields of PI-stained cells under fluorescence at 200X with at least 40 sharply focused cells, plus one brightfield photo of the scale bar at the same magnification and phone zoom.

Microscopes and camera mount for PI imaging

Zeiss AxioLab A.1 or Zeiss Primostar fluorescence microscopes, with your phone in a Gosky mount.

PI stain: first spin of the ethanol-fixed cells

Centrifuge at 800 x g (1,000 rpm in the IEC centrifuge) for 3 minutes. Mark the tube at 0.5 and 0.1 mL, remove ethanol down to 500 µL, and gently resuspend by flicking.

PI stain: washing in PBS + BSA

While vortexing at low speed, add 4.5 mL PBS + BSA dropwise (final volume 5.0 mL), mix by inversion, spin at 800 x g for 3 minutes, and withdraw the supernatant down to 0.1 mL.

PBS + BSA

Phosphate-buffered saline plus 0.5 mg/mL bovine serum albumin.

PI stain: counting the cells

Resuspend by pipetting 50 µL up and down at least 10 times without bubbles, load 10 µL into a hemocytometer, and count under phase contrast at 100X (Axiostar set up in Koehler illumination during the spin).

Target cell concentration for PI staining

1.0 × 10^6 cells/mL. Dilute with PBS + BSA; if you have fewer, re-centrifuge and remove 50 µL of supernatant.

How much PI staining solution (with RNase)?

11 µL per 100 µL of cell suspension (5.5 µL if you have 50 µL), in an Eppendorf tube.

PI staining time and conditions

At least 20 minutes at room temperature, protected from light.

Making the PI slide

Resuspend by gentle flicking, put 5 µL on a clean slide, add a coverslip, seal the edges with nail polish, and let it dry in the fume hood with the hood lights off before going near a microscope.

What is Fiji?

“Fiji Is Just ImageJ”: free ImageJ image-analysis software (fiji.sc). The toolbar’s title bar reads “(Fiji Is Just) ImageJ.”

What if Fiji’s main menu bar doesn’t show up?

Select the Fiji toolbar window and the main menu bar should appear.

Lookup table (LUT) button

A button on the Fiji toolbar that changes the colors used to display an image (also under Image > Lookup Tables). Hovering over any toolbar button shows what it does in the bottom-left of the toolbar.

.czi files and metadata

Carl Zeiss Image files from a professional Zeiss microscope. Proprietary formats like this contain extra information; “Display metadata” shows details such as the lasers and filters used.

When does the Bio-Formats Import Options dialog appear?

When you open a proprietary microscope file such as a .czi, not an ordinary image like a JPEG.

Two Bio-Formats import options you must turn on

Split channels (splits the image into its individual colors) and View stack with: Hyperstack (imports it in a useful format).

The three channels in the FIJI video’s example image

Green: glucocorticoid receptor, a transcription factor, labeled with an antibody. Blue: DAPI, a DNA dye that marks the nucleus. Red: an endosomal protein that binds the green protein.

DAPI

A DNA dye that marks the nucleus (the blue channel in the FIJI video).

Can split channels be put back together?

Yes, they can be merged back into one image (Image > Color > Merge Channels).

What does Invert LUT do?

Swaps the display so dark colors become light and light colors become dark, giving every color scheme a second look. The underlying image is the same.

How do you save your image for the workbook?

File > Save As > Jpeg.

Why open the scale bar image along with the cell image?

To set the scale: the image starts out measured in pixels (shown at the top left of the window), and it must be changed to microns to get real area measurements of the nuclei.

Two toolbar tools used for the measurements

The straight line tool (to measure the scale bar) and the wand (tracing) tool (to select individual nuclei).

First step before thresholding

Convert the image to grayscale: Image > Type > 8-bit.

Why convert to 8-bit grayscale before thresholding?

The computer can only use the intensity information properly in grayscale. Once the image is a single dimension of intensity, it can threshold against it to find the nuclei.

How do you open the threshold tool?

Image > Adjust > Threshold.

Threshold methods compared in the Thresholding video

Default (IsoData) didn’t select the nuclei well at first; Huang looked similar to Default; Intermodes worked well for this image. You can also drag the threshold sliders to adjust any method.

Display options in the Threshold dialog

You can change how the threshold is shown (for example Red or Over/Under) and flip the background with the Dark background box. The video uses Red, with the background shown red and the nuclei left white.

What do you save after thresholding?

Save at least your first thresholded image (File > Save As > Jpeg) to show you did it.

Setting the scale, step 1

Pick the straight line tool, drag a line across the whole scale bar, and measure it (Analyze > Measure, or Cmd+M / Ctrl+M) to get the scale bar’s length in pixels.

Setting the scale, step 2

Analyze > Set Scale: “Distance in pixels” is already filled in from your line. Enter the known distance, which you calculate yourself from the magnification as described in the workbook, and change the unit of length from pixel to micron. [the video’s example uses 400X; this lab’s photos are taken at 200X]

How can you tell the scale has been set?

The top-left of the image window shows microns instead of pixels.

Which three measurements do you record for each nucleus?

Area, mean gray value and integrated density (chosen in Analyze > Set Measurements).

How do you measure one nucleus?

Click it with the wand (tracing) tool to select it, then Analyze > Measure (Cmd+M); the values appear in the Results table.

Why must you avoid clusters of nuclei?

A cluster gives a falsely high integrated density and area. In the video, a cluster measured 848 area and 159,907 integrated density, versus about 440–478 and 78,000–85,000 for single nuclei.

How many cells do you measure, and what goes in the lab report?

At least 40 individual cells, avoiding clusters. Include the table of area, mean gray value and integrated density.

Mean gray value vs. integrated density

Mean gray value: the average pixel brightness in the selection. Integrated density: area × mean gray value, the object’s total signal. [not in manual]

Hood rules for passaging

Work only on the solid surface; spray 70% ethanol on everything that goes in, including gloved hands; have all materials inside before you start; don’t open media or cells until ready; keep your hands in the hood.

Materials for passaging

Pipets, tips, pipettors, two waste beakers (solids and liquids), a 15 mL conical tube, cells in a 25 cm² flask, DMEM+, trypsin/EDTA, sterile PBS and BCS, with a 70% alcohol spray outside the hood.

Passaging: before removing the medium

Scan the whole flask on the inverted phase contrast microscope and record percent confluence, then pour the expired medium into the waste beaker without disturbing the cells.

Passaging: PBS rinse

Pipette 4 mL sterile PBS into the flask at the neck, tilt to rinse the cells, and pour it out without disturbing them.

Passaging: trypsin/EDTA step

Add 1 mL trypsin/EDTA and tilt until it covers the whole surface. Check every 2 minutes on the inverted microscope; when the cells round up and detach, tilt until no attached cells remain and they float.

Passaging: after the cells detach

Add 4 mL PBS and 0.500 mL BCS, pipette it over the surface 3–4 times to rinse off remaining cells, and put the ~5 mL suspension into a 15 mL conical tube.

Passaging: spin and resuspend

Centrifuge at 1,200 rpm for 5 min (1,000 x g in the IEC). In the hood, pour off the supernatant, add 1 mL PBS, finger-vortex until there are no clumps, and keep on ice.

Why finger-vortex before pipetting cells into the wells?

Cells left undisturbed for several minutes sink to the bottom of the 15 mL tube.

6-well plate setup for the apoptosis experiment

A cohort of three groups shares a plate: 1X, 2X, 3X (experimental) on top and 1C, 2C, 3C (controls) below. Each well gets 1.4 × 10^4 cells in 2 mL DMEM+.

Labeling the 6-well plate

Write your bench number on one side of the plate cover; don’t label the tops of the wells, because you look through them.

What happens to the 6-well plate next week?

Apoptosis will be induced and observed directly by 200X phase contrast on the inverted microscopes.

Is a new 25 cm² flask started this week?

No.