BD Pathway bioimaging systems

When should I use confocal?

The system is supplied with an Olympus Plan fluo 0.75 NA, 20x objective and the following optional objectives can be used:

How does the Nipkow spinning disk system work?

The BD Pathway™ high-content bioimager uses a multi-pinhole Nipkow spinning disk, so named after its inventor Paul Nipkow. The disk contains multiple sets of spirally arranged pinholes placed in the image plane of the objective lens. A select region of the disk is illuminated from the rear by a column of light. The highly reduced images of the pinholes are focused by the objective lens onto the specimen. When spinning at a rapid rate, the pinhole coverage of the sample is completed several hundred times per second, effectively creating a full image of the focal plane in real time. Emitted light is collected and imaged using a high resolution and high quantum efficiency CCD camera.

What are the principles of confocal microscopy?

The principle of confocal microscopy is the elimination of out-of-focus light, thus producing a high z-resolution image. Confocal fluorescence microscopes achieve this via two principal mechanisms. First, incident light is focused to a particular point within the specimen by passing it through a very small pinhole. The focusing helps to limit the excitation of fluorophores above and below the plane of focus. Second, any emission that is above or below the plane of focus is blocked from reaching the detector by passing it through the same pinhole. The specimen is placed in the light path at a conjugate focal plane such that movement in the vertical (z) direction keeps the focus at a fixed distance from the objective, which effectively scans in layers through the specimen.

Passing light through a pinhole limits the illumination of the sample to one particular spot, a spot much smaller than the typical field of view. To fully develop an image, light must be delivered to every point of the specimen within the focal plane. Two general strategies accomplish this. The first involves scanning the sample with light in a raster pattern over a period of time such that a complete image is made of the focus plane within the sample. Emitted light is then detected using a photomultiplier tube (PMT), and the image is reconstructed by mapping the light output at each point. This is how the conventional laser scanning microscope works.

What are the main advantages of spinning disk systems over laser scanning confocal systems?

The most significant advantage of a spinning disk confocal system is its ability to simultaneously monitor the entire imaging area. Therefore rapidly occurring events within living cells can be observed confocally without compromising resolution. These events would be missed during the time required to perform a laser scan. The rapid scanning (1000 scans per second) also enables the user to observe the confocal image through the eyepiece. The spinning disk can also be moved out of the light path allowing the user to quickly go from confocal imaging to wide field imaging. High scanning rates also allow 3D and 4D recordings to be made at much higher speeds than attainable by laser scanning confocals.

The high-speed scanning of a high quantum efficiency CCD camera can capture images unlike a PMT used in the laser scanner. The use of a white light source on the spinning disk system allows full spectrum (36–600 nm) confocal. The flexibility afforded by the 100–200W halide/mercury arc lamps allows users to select fluorescent probes based on experimental design and not on the lasers available to them on their confocal system. In addition, a combination of multipoint illumination and high QE EM CCD cameras significantly reduces photobleaching and phototoxicity of the specimen, allowing longer imaging times without loss of image quality.

What are the main applications of spinning disk systems?

The fast scanning speeds of the spinning disk system are conducive to high z-resolution live cell imaging. The system also can be used to improve the signal/noise ratio in fixed cell imaging by excluding out-of-focus light.

What is the depth of penetration of halide light compared to laser scanning?

Neither Hg/halide light or laser scanning impact depth of penetration. Both have a penetration of about 30–100 µm. To penetrate deeper, use multiphoton confocal imaging, which uses high wavelength (700–1200 nm) lasers that can penetrate to about 100–200 nm. Note that wavelength of light and opacity of tissue impact depth of penetration. The higher the wavelength, the deeper the penetration.

Is the Z resolution of the BD Pathway high-content bioimager lower than the conventional laser scanner?

No. For a given pinhole size, there should be no difference between the two systems. The actual Z resolution depends on the pinhole diameter, the NA (numerical aperture), and the magnification of the objective. With conventional laser scanners, the Z resolution for a given objective can be changed by altering the pinhole size. In the spinning disk system used by the BD Pathway™ Bioimager, the pinhole diameter is fixed and matched for the high-NA, high-magnification objectives.

What is the Z resolution for my objectives?

The measured Z resolutions for the BD Pathway are:

• For 100x PlanApo 1.4NA, PSF = 0.5 µm
• For 60x PlanApo 1.4NA, PSF = 0.8 µm
• For 40x PlanApo 0.9 NA, PSF = 2.0 µm
• For 20x PlanApo 0.75NA, PSF = 2.8 µm

How do I calculate an optimal number of z-sections to be imaged for a given objective (or what interval size should I use when collecting a confocal z-stack)?

To capture all possible axial detail, the z-step spacing should be at least half the z-resolution of the system. For instance, when a 100x 1.4 NA objective is used, the z-resolution (PSF) is 0.5 µm. Therefore sampling should be done at 0.25 µm steps at minimum.

For 60x 1.4 NA, the z-resolution is 0.8 µm; therefore, sampling should be done at 0.4 µm steps at minimum.

During 3D rendering, an aspect ratio of the imaged sample is required. The aspect ratio is calculated by dividing the z-step size by the XY resolution. The XY resolution is usually measured; however, it can be calculated by dividing the pixel size by the magnification. For example, an ORCA ER camera has a pixel size of 6.4x6.4 µm. When a 100x objective is used, the XY resolution is 6.4/100= 0.064 µm. Therefore, the aspect ratio for a 100x objective at 0.25- µm steps would be 0.25/0.064= 3.9.

When a 60x objective is used, the XY resolution is 6.4/60= 0.106 µm. The aspect ratio at 0.4 µm steps would be 0.4/0.106= 3.7. If the camera was binned 2x2, then the pixel size would be 6.4 x 2= 12.8. The XY resolution for the 60x would be 12.8/60= 0.21 µm. The aspect ratio at 0.4 µm steps would be 0.4/0.21= 1.9.

Do you need a laser to get true confocal?

No. Lasers are bright (many photons per second can be focused into a single diffraction-limited spot), but the photons they produce are just like those produced by Hg arc lamps or Hg/metal halide lamps or other sources, except lasers produce thousands of times more photons in a diffraction-limited spot. The laser beam does not penetrate more or less. Minski's first functioning confocal microscope used an Hg source and a vacuum photodiode detector. It was confocal because of the two small apertures he inserted. One aperture was used to limit the effective size of the source and the second aperture was used to limit the effective size of the detector.

Confocals often use lasers to get one mW of light into a single spot. However, this causes the dye at the focus spot to saturate. If you have many pinholes, like in a spinning disk confocal, you can get enough photons to form images fairly fast using arc sources. The BD Pathway™ instrument uses a large number of pinholes that allow thousands of times more light than a single round aperture. Thus, it compensates for the reduced light of an Hg light source.