Tuesday, April 6, 2021
Monday, April 5, 2021
Monday, November 2, 2020
Covalent bonds, the strongest interactions between atoms, are responsible for holding atoms together to form molecules. They form when two atoms come together and share a pair of electrons (Figure 2.1). For example, methane (CH4) is formed when four hydrogen atoms share electrons with one carbon atom (Figure 2.1A). The number of covalent bonds that an atom can form is determined by the number of unpaired electrons in its outer electron shell (its valence). Carbon has four unpaired electrons whereas hydrogen has one, so carbon can form covalent bonds with four hydrogen atoms. The other principal atoms of living organisms, oxygen and nitrogen, have two and three unpaired electrons, respectively.
The carbohydrates include simple sugars as well as polysaccharides. These simple sugars, such as glucose, are the major nutrients of cells. As discussed in Chapter 3, their breakdown provides both a source of cellular energy and the starting material for the synthesis of other cell constituents. Polysaccharides are storage forms of sugars and form structural components of the cell. In addition, polysaccharides and shorter polymers of sugars act as markers for a variety of cell recognition processes, including the adhesion of cells to their neighbors and the transport of proteins to appropriate intracellular destinations.
Lipids have three major roles in cells. First, they provide an important form of energy storage. Second, and of great importance in cell biology, lipids are the major components of cell membranes. Third, lipids play important roles in cell signaling, both as steroid hormones (e.g., estrogen and testosterone) and as messenger molecules that convey signals from cell surface receptors to targets within the cell. The simplest lipids are fatty acids, which consist of long hydrocarbon chains, most frequently containing 16 or 18 carbon atoms, with a carboxyl group (COO–) at one end (Figure 2.8). Unsaturated fatty acids contain one or more double bonds between carbon atoms; in saturated fatty acids all of the carbon atoms are bonded to the maximum number of hydrogen atoms. The long hydrocarbon chains of fatty acids contain only nonpolar C—H bonds, which are unable to interact with water. The hydrophobic nature of these fatty acid chains is responsible for much of the behavior of complex lipids, particularly in the formation of biological membranes.
Wednesday, October 28, 2020
Although the electron microscope has allowed detailed visualization of cell structure, microscopy alone is not sufficient to define the functions of the various components of eukaryotic cells. To address many questions concerning the function of subcellular organelles, it is necessary to isolate the organelles of eukaryotic cells in a form that can be used for biochemical studies. This is usually accomplished by differential centrifugation a method developed largely by Albert Claude, Christian de Duve, and their colleagues in the 1940s and 1950s to separate the components of cells on the basis of their size and density. The first step in subcellular fractionation is the disruption of the plasma membrane under conditions that do not destroy the internal components of the cell. Several methods are used, including sonication (exposure to high-frequency sound), grinding in a mechanical homogenizer, or treatment with a high-speed blender. All these procedures break the plasma membrane and the endoplasmic reticulum into small fragments, while leaving other components of the cell (such as nuclei, lysosomes, peroxisomes and mitochondria) intact. The suspension of broken cells (called a lysate or homogenate) is then fractionated into its components by a series of centrifugations, with an ultracentrifuge used to rotate samples at very high speeds (over 100,000 rpm), producing forces up to 500,000 times greater than gravity. This force causes cell components to move toward the bottom of the centrifuge tube and form a pellet (a process called sedimentation) at a rate that depends on their size and density, with the largest and heaviest structures sedimenting most rapidly (Figure 1.40). Usually the cell homogenate is first centrifuged at a low speed, which sediments only unbroken cells and the largest sub- cellular structures the nuclei. Thus, an enriched fraction of nuclei can be recovered from the pellet of such a low-speed centrifugation while the other cell components remain suspended in the supernatant (the remaining solution). The supernatant is then centrifuged at a higher speed to sediment mitochondria, chloroplasts, lysosomes, and peroxisomes. Recentrifugation of the supernatant at an even higher speed sediments fragments of the plasma membrane and the endoplasmic reticulum. A fourth centrifugation at a still higher speed sediments ribosomes, leaving only the soluble portion of the cytoplasm (the cytosol) in the supernatant.
Because of the limited resolution of the light microscope, analysis of the details of cell structure has required the use of more powerful microscopic techniques namely electron microscopy, which was developed in the 1930s and first applied to biological specimens by Albert Claude, Keith Porter, and George Palade in the 1940s and 1950s. The electron microscope can achieve a much greater resolution than that obtained with the light microscope because the wavelength of electrons is shorter than that of light. The wavelength of electrons in an electron microscope can be as short as 0.004 nm about 100,000 times shorter than the wavelength of visible light. Theoretically, this wavelength could yield a resolution of 0.002 nm, but such a resolution cannot be obtained in practice, because resolution is determined not only by wave-length, but also by the properties of the microscope lens and the specimen being examined. Consequently, for biological samples the practical limit of resolution of the electron microscope is 1–2 nm. Although this resolution is much less than that predicted simply from the wavelength of electrons, it represents more than a hundredfold improvement over the resolving power of the light microscope (see Figure 1.25).
Monday, October 5, 2020
SUPER-RESOLUTION MICROSCOPY: BREAKING THE DIFFRACTION BARRIER
An exciting advance in recent years has been the development of super-resolution microscopy techniques that break the diffraction barrier and increase the resolution of fluorescence microscopy to the range of 10-100 nm, about tenfold less than the theoretical limit of resolution of the light microscope (see Figure 1.25). Several methods of super-resolution microscopy use fluorescent probes that shift the limit of resolution from the wavelength of visible light to the molecular level.
SHARPENING THE FOCUS AND SEEING CELLS IN THREE DIMENSIONS
The images obtained by conventional fluorescence microscopy are blurred as a result of out-of-focus fluorescence. These images can be improved by a computational approach called image deconvolution, in which a computer analyzes images obtained from different depths of focus and generates a sharper image than would have been expected from a single focal point. Alternatively, confocal microscopy allows images of increased contrast and detail to be obtained by analyzing fluorescence from only a single point in the specimen. A small point of light, usually supplied by a laser, is focused on the specimen at a particular depth. The emitted fluorescent light is then collected using a detector, such as a video camera. Before the emitted light reaches the detector, however, it must pass through a pinhole aperture (called a confocal aperture) placed at precisely the point where light emitted from the chosen depth of the specimen comes to a focus (Figure the plane of focus is able to reach the detector. Scanning across the specimen generates a two-dimensional image of the plane of focus, a much sharper image than that obtained with standard fluorescence microscopy (Figure 1.34). Moreover, a series of images obtained at different depths can be used to reconstruct a three-dimensional image of the sample.
FOLLOWING PROTEIN MOVEMENTS AND INTERACTIONS
A variety of methods have been developed to follow the movement and interactions of GFP-labeled proteins within living cells. One widely used method for studying the movements of GFP-labeled proteins is fluorescence recovery after photobleaching (FRAP) (Figure 1.31). In this technique, a region of interest in a cell expressing a GFP-labeled protein is bleached by exposure to high-intensity light. Fluorescence recovers over time due to the movement of unbleached GFP-labeled molecules into the bleached region, allowing the rate at which the protein moves within the cell to be determined.