Molecular and Cellular Imaging Basics
Light microscopes provide a highly magnified image of an object and allow us to visualize and measure fine details at the cellular and even subcellular level. The microscopic image is produced in 2 steps: the objective forms a real, inverted, magnified image (=intermediate image), which is then further enlarged to a virtual image by the oculars. Transmitted light microscopy employs various optical/physical techniques to produce an image: regular bright field, dark field (oblique illumination of the object), polarized light, phase contrast, and differential interference contrast (DIC). All these techniques are based on 6 principal interactions of light and matter: absorption (the most common interaction in biological microscopy), reflection (light hits a surface at a certain angle), refraction (light travels through different media), diffraction (light scattering), polarization (light is emitted in a single plane and the rays behave differently when they pass through crystalline or birefringent material), and fluorescence (see below). Subcellular and very thin objects are difficult to visualize in native samples, due to the lack of absorption and/or diffraction of the light. Histological or fluorescent stains, or physical contrasting techniques (e.g., Phase, DIC) are therefore often necessary to resolve these structures.
The limit of optical resolution in the light microscope, i.e., the minimal distance between 2 points which can still be distinguished as separate points is generally 200 nm, although newer techniques and systems can nowadays decrease this distance to less than 100 nm.
When a fluorescent molecule absorbs light of a specific, suitable wavelength, one part of the absorbed light (=energy) is converted into heat, the remainder is emitted as light of longer wavelength. In this process, called fluorescence, the absorbed light (photon) has sufficient energy to boost an electron from the ground-state energy level (G0) to an excited state energy level (S1). The electron remains in the excited state only for a short period of time and then ‘falls’ back to G0, during which the light emission occurs. The entire process is very fast and has a half-life of <10-8 seconds.
In the fluorescence microscope, the light source is a mercury or xenon arc lamp, which emit high-intensity light, concentrated at certain wavelengths. A specific filter set reflects only the desired range of wavelengths, which then excite the fluorescent molecules in the sample. The emitted light passes through the same filter set, this time only the desired emission wavelengths are transmitted.
Fluorescence microscopy has several advantages, but also disadvantages:
- Specific structures or molecules can be labeled
- Co-localization can be observed
- Very small objects are visible, which would not be visible in transmitted light
- Phototoxicity – many of the fluorescent dyes are phototoxic
- The fluorescent dyes bleach sooner or later, making this imaging method time-sensitive
- Crosstalk – two fluorescent dyes have an overlapping emission range
The principle of confocal imaging was advanced by Marvin Minsky and patented in 1957, and is employed in all modern confocal microscopes.
In a conventional widefield microscope, the entire specimen is bathed in light from a mercury or xenon source, and the image can be viewed directly by eye or an image can be taken. The significant amount of fluorescence emission that occurs at points above and below the objective focal plane is not confocal with the pinhole and forms extended Airy disks in the aperture plane, usually experienced as a blurry image.
In contrast, the illumination in a confocal laser scanning microscope (CLSM) is achieved by scanning one or more focused laser beams across the specimen. The emitted light only from the focal plane in the specimen is detected by a photomultiplier tube (PMT) through a detection pinhole, and transformed into electrical signals which are converted to images by the specific software and displayed on the computer screen. The detection pinhole suppresses signal from structures that are out of focus. In the spinning disk confocal micoscope (SD), the laser beam illuminates the entire sample (no scanning). Only one laser can be activated at the time. The emitted light of the sample passes a spinning disk before it is captured by a high-resolution monochrome camera. The spinniing disk has thousands of small pinholes in a specific arrangement, thus creating the confocal image of a thin optical section. The depth of the focal plane is determined by the excitation wavelength, the numerical aperture of the objective, and the diameter of the detection pinhole. Although unstained specimens can be viewed using light reflected back from the specimen, they usually are labeled with one or more fluorescent probes.
The electron microscope instead of using light from an incandescent bulb or a laser beam, as in light and confocal microscopy, generates a cloud of electrons by heating a filament, filters them using a high voltage field then forms them into a beam focusing them on a specimen using a number of electromagnetic field lenses. The electrons pass through the specimen in a Transmission Electron Microscope (TEM) or generate a new set of electrons from the sample as the beam is moved across its surface (raster), in the Scanning Electron Microscope, (SEM).
Those passing through the sample in the TEM can be collected on film or using a CCD camera. The electrons generated from the sample in an SEM are captured using a charged detector and imaged light to dark according to their energy, thus giving a false 3D effect.
Proper preparation of the samples is a challenge for all forms of microscopy. To preserve the fine structures in the living system from enzymatic degradation, the high vacuum and intense bombardment of electrons in the EM, the sample preparation for EM can be painstaking and laborious. Heavy metal dyes are used to add contrast and generate sufficient signal. Other, more sophisticated, methods and equipment are employed to produce a sample, which will adequately represent the living state for biology and the active state for materials.
In the EM, properly prepared samples yield accurate images of samples, their component molecules and their products and interactions far beyond the ability of any other tool to visualize.
Cryo-electron microscopy (cryo-EM) is a broad term that encompasses several electron microscopy techniques. In most cases, cryo-EM is based upon the principle of imaging radiation sensitive biological samples under cryogenic (frozen) conditions to minimize distortions and artifacts cased by conventional sample fixation procedures. Cryo-EM techniques include cryo-electron tomography, single-particle cryo-electron microscopy and electron crystallography. Electron tomography is a technique that uses a series of two-dimensional image of a specimen to reconstruct a magnified three-dimensional volume. When combine with cryogenic sample preparation methods, electron tomography is a powerful tool for probing cellular ultrastructure, sub-cellular structures, viral particles and large macromolecular complexes.