Confocal Fluorescence Microscopy: A Technique to Determine the Localization of Proteins in Mouse Fibroblasts

Source: Dominique R. Bollino¹, Eric A. Legenzov², Tonya J. Webb^1
1 Department of Microbiology and Immunology, University of Maryland School of Medicine and the Marlene and Stewart Greenebaum Comprehensive Cancer Center, Baltimore, Maryland 21201
² Center for Biomedical Engineering and Technology, University of Maryland School of Medicine, Baltimore, Maryland 21201

Confocal fluorescence microscopy is an imaging technique that enables increased optical resolution as compared to conventional "wide-field" epifluorescence microscopy. Confocal microscopes are able to achieve improved x-y optical resolution through "laser scanning"- typically a set of voltage-controlled mirrors (galvanometer or "galvo" mirrors) that direct laser-illumination to each pixel of the specimen at a time. More importantly, confocal microscopes achieve superior z-axial resolution by using a pinhole to remove out of focus light originating from locations that are not in the z-plane being scanned, thus enabling the detector to collect data from a specified z-plane. Because of the high z-resolution achievable in confocal microscopy, it is possible to collect images from a series of z-planes (also-called z-stack) and construct a 3D image through software.

Before discussing the mechanism of a confocal microscope, it is important to consider how a sample interacts with light. Light is composed of photons, packets of electromagnetic energy. A photon impinging on a biological sample can interact with the molecules comprising the sample in one of four ways: 1) the photon does not interact and passes through the sample; 2) the photon is reflected/scattered; 3) the photon is absorbed by a molecule and the absorbed energy is released as heat through processes collectively known as nonradiative decay; and 4) the photon is absorbed and the energy is then rapidly reemitted as a secondary photon through the process known as fluorescence. A molecule whose structure permits fluorescence emission is called a fluorophore. Most biological samples contain negligible endogenous fluorophores; therefore exogenous fluorophores must be used to highlight features of interest in the sample. During fluorescence microscopy, the sample is illuminated with light of the appropriate wavelength for absorption by the fluorophore. Upon absorbing a photon, a fluorophore is said to be "excited" and the process of absorption is referred to as "excitation". When a fluorophore gives up energy in the form of a photon, the process is known as "emission", and the emitted photon is called fluorescence.

The light beam used to excite a fluorophore is focused by the objective lens of a microscope and converges at a "focal point" where it is maximally focused. Beyond the focal point the light again diverges. The entering and exiting beams may be visualized as a pair of cones touching at the focal point (see Figure 1, left panel). The phenomenon of diffraction imposes a limit on how tightly a beam of light can be focused - the beam actually focuses to a spot of finite size. Two factors determine the size of the focal spot: 1) the wavelength of the light, and 2) the light-gathering ability of the objective lens, which is characterized by its numerical aperture (NA). The focal "spot" extends not only in the x-y plane, but also in the z direction, and is in reality a focal volume. The dimensions of this focal volume define the maximum resolution achievable by optical imaging. Although the number of photons is greatest within the focal volume, the conical light paths above and below the focus also contain a lower density of photons. Any fluorophore in the light path can thus be excited. In conventional (wide-field) epifluorescence microscopy, emission from fluorophores above and below the focal plane contribute out-of-focus fluorescence (a "hazy background"), which reduces image resolution and contrast, as demonstrated in Figure 1, with the red cube representing fluorophore emission above the focal plane (red star) that results in out-of-focus fluorescence (top right). This problem is ameliorated in confocal microscopy, due to the utilization of a pinhole. (Figure 2, bottom right). As depicted in Figure 3, the pinhole allows emissions originating from the focal place to reach the detector (left), while blocking the out-of-focus fluorescence (right) from reaching the detector, thus improving both resolution and contrast.

Figure 1. Optical resolution of epifluorescence versus confocal microscopy. Please click here to view a larger version of this figure.

The light beam used to excite a fluorophore is focused by the objective lens of a microscope and converges at a focal volume and then diverges (left). The red star represents the focal plane of a sample that is being imaged while the red square represents fluorophore emission above the focal plane. When capturing an image of this sample using an epifluorescent microscope, the emission from the out-of-focus red square will visible and contribute to a "hazy background" (top right). Confocal microscopes have a pinhole which prevents the detection of light emitted outside of the focal plane, eliminating the "hazy background" (bottom right).

Figure 2. Pinhole effect in confocal microscopy. Please click here to view a larger version of this figure.

Although the highest intensity of the excitation light is at the focal point of the lens (left, red oval), other parts of the sample not in the focal point (right, red star) will get light and fluoresce. To prevent light emitted from these out-of-focus regions to reach the detector, a screen with a pinhole is present in front of the detector. Only the in-focus light (left) emitting from the focal plane is able to travel through the pinhole and reach the detector. The out-of-focus light (right) is blocked with the pinhole and fails to reach the detector.

Figure 3. Principal components of a confocal laser scanning microscope. Please click here to view a larger version of this figure.

For the sake of simplicity, the mechanistic description of a confocal microscope will be limited to that of the Nikon Eclipse Ti A1R. Although there may be minor technical differences between different confocal microscopes, the A1R serves well as a good model for describing confocal microscope function. The excitation light beam, produced by an array of diode lasers, is reflected by the primary dichroic mirror into the objective, which focuses the light onto the specimen being imaged. The primary dichroic mirror selectively reflects the excitation light while allowing light at other wavelengths to pass through. The light then encounters the scanning mirrors that sweep the light beam across the specimen in an x-y manner, illuminating a single (x,y) pixel at a time. Fluorescence emitted by fluorophores at the illuminated pixel is collected by the objective lens and passes through the primary dichroic mirror to reach an array of photomultiplier tubes (PMTs). Secondary dichroic mirrors direct the emission light to the appropriate PMT. Excitation light scattered by the sample back into the objective is reflected by the primary dichroic mirror back towards the specimen, and thus prevented from entering the detection light path and reaching the PMTs (see Figure 3). This allows the relatively feeble fluorescence to be quantified without contamination by light scattered from the excitation light beam, which is typically orders of magnitude more intense than the fluorescence. Because the pinhole blocks light from outside of the focal volume, the light arriving at the detector comes from a narrow, selected z-plane. Therefore, images can be collected from a series of adjacent z-planes; this series of images is often referred to as a 'z-stack'. By using the appropriate software, a z-stack can be processed to generate a 3D image of the specimen. A particular advantage of confocal microscopy is the ability to distinguish the subcellular localization of staining. For example, the differentiation between membrane staining from intracellular staining, which is very challenging with conventional epifluorescence microscopy (1, 2, 3).

Sample preparation is an important facet of confocal imaging. A strength of optical microscopy techniques is the flexibility to image live or fixed cells. When attempting to produce 3D images, because of the number of images that must be acquired for a z-stack, the difficulty of maintaining cell health, and the movement of live cells and their organelles, the use of fixed cells is typical. The procedure for fixing and staining cells for confocal fluorescence is similar to that conventionally used in immunofluorescence. After culture in chamber slides or on coverslips, cells are fixed using paraformaldehyde to preserve cellular morphology. Non-specific antibody binding is blocked using bovine serum albumin, milk, or normal serum. In order to maintain the specificity of the secondary antibodies, the solution used should not originate from the same species in which the primary antibodies were generated. The cells are incubated with primary antibodies that bind the antigen of interest. When labeling several cellular targets, the primary antibodies must each be derived from a different species. Antibodies tagging an antigen are then bound by fluorophore-conjugated secondary antibodies. Fluorophore-conjugated secondary antibodies should be selected so that they are compatible with the wavelengths of laser excitation available in the confocal microscope. When visualizing multiple antigens, the excitation/emission spectra of the fluorophores should differ enough so that their signals can be discriminated by microscopic analysis. The stained specimen is then mounted on a slide for imaging. A mounting medium is used to prevent photobleaching and specimen dehydration. If desired, a mounting medium containing a nuclear counterstain (e.g. DAPI or Hoechst) can be used (4).

In the following protocol, mouse fibroblasts transfected to express CD1d (LCD1) were stained with antibodies recognizing CD1d and CD107a (LAMP-1). CD1d is a major histocompatibility complex 1 (MHC 1)-like receptor present on the surface of antigen presenting cells that presents lipid antigens. LAMP-1 (lysosomal associated membrane protein-1) is a transmembrane protein primarily present in lysosomal membranes. For proper antigen presentation, CD1d is trafficked through the low pH lysosomal compartment, so LAMP-1 is being used as a marker of the lysosomal compartment for this protocol. By probing the LCD1 cells with anti-CD1d and anti- LAMP-1 that were produced in different species, secondary antibodies with unique fluorophores can be used to determine the localization of each protein in the cell and whether CD1d is present in the LAMP-1 positive lysosomal compartments.