Protocol for the DeltaVision Microscope
First thing, it is kinda hot (since it is kept at 37°C, so don't be wearing a jumper under the lab coat.
First thing to do is to set up the deconvolution settings. Unfortunately I don't think I watched closely enough to be able to do it next time. For toxo slides prepared at botany, we used 10um distance from the cover slip to the cells, and we were using number 1.5 cover slips (which most labs use, I understand), and Fluorogel was approximated as vector shield (ie. refractive index 1.44), since we couldn't find the refractive index of fluorogel with a quick google. So we used the 1.522 oil, dabbing it on each slide (NOTE: not the lens).
Need to change lenses manually. We used 100×.
Use 512x512 image with 1x1 box. The 2x2 box has an average (median?) pixel for each 4 1x1 pixels, which seems like loss of information. We are pretty close to the resolution limit though.
For my samples, the easiest thing was to
- Put oil on cover slip
- Move the lens down (spin clockwise looking from the RHS of the scope) to make room for the slide
- Put the cover slip face down
- Align the lens with the oil by using the knob inside the chamber on top of the metal attached to the slide. This is faster than using the joystick.
- Move the lens back up until it just touches the oil (spin counter-clockwise)
- Put the light source above the slide down and close the chamber. The light source can be moved up and down with a scroller to the left and up a bit in the chamber. It should be ~1cm above the slide.
- switch to eye-mode, using the knob down and right of the eye piece
- Turn the fluorescence to a channel that will work, or the polarised light mode.
- To switch, using the ring near the eye piece. It changes on the computer screen - the bottom right of the square displays the relevant channel
- Turn on excitation or light, using either of the 2 buttons on the bottom left of the control pad. Make sure that it is working by looking into the chamber.
- Move lens up until the picture comes up into focus. Can turn up the %T on the computer if there is not enough light.
Then, to do things the lazy (easy) way:
- Once in focus, abandon the eye piece and go back to the computer (switch the knob below right of the eye piece).
- Take a picture using the relevant channel. It should be in reasonable focus because it was just focused on the eye piece.
- Adjust the settings (%T, exposure time) to make sure that the camera isn't being saturated.
- Use the spiral mode to the left of the map to search for parasites, using a channel that clearly delineates the parasites from the host cells (e.g. the TgTom40 antibody works well for this, where Hoechst also stains the host cell nuclei)
- Select points on the map (which should show as bright spots on the map), then adjust the focus so it is about in the middle, then save each of the points.
- When you are happy you are imaging enough parasites, setup the experiment. Make sure to:
- Refresh the channel-specific settings (they don't auto-carry over from what was just done).
- Set the post-process tasks up - do the deconvolution, do the maximal projection, export as TIFF (see below).
- Make sure thin enough sections are being taken (0.15μm seemed sensible, and with the thickness (10um?) 60 slices was ok).
- Use the lens tissue to soak up the excess oil from the lens (don't wipe, except maybe gently around in a circle).
- roll over the lens with a cotton bud tip, using the 1:1 ethanol:chloroform
- move the lens down as far as possible. On start the system doesn't know where it is, so can sometimes put the lens to push up on the slide, if one is there.
Exporting images as TIFFs
Unfortunately, the machine uses its own proprietry format when exporting There are 3 ways to do this, probably the first way is the way that I should be using
- Add a post-process filter when acquiring images (i.e. after deconvolution). That way it will just happen automatically, and probably requires the least effort in the long run.
- Do it singly. To do this, open up the relevant
- Do a whole bunch at once
You can use ImageJ afterwards instead on the DV files, using the loci tools, e.g. the ImageJ plugin, or using the cmdline e.g. https://github.com/wwood/bbbin/blob/master/microscopy_automate_dv.rb
To put images on a USB stick, the stick needs to be formatted using the FAT32 filesystem, not NTFS. To insert the USB stick, open the door in front of the computer, and the slot is on the right.