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Cell_Biology_Alberts. a strong electric current transfers the protein onto the membrane. The membrane is then soaked in a solution of labeled antibody to reveal the protein of interest. This method of detecting proteins is called Western blotting, or immunoblotting (Figure 8–17). Sensitive Western-blotting methods can detect very small amounts of a specific protein (1 nanogram or less) in a total cell extract or some other heterogeneous protein mixture. The method can be very useful when assessing the amounts of a specific protein in the cell or when measuring changes in those amounts under various conditions.


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Cell_Biology_Alberts. Hydrodynamic Measurements Reveal the Size and Shape of a Protein Complex


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Cell_Biology_Alberts. Most proteins in a cell act as part of larger complexes, and knowledge of the size and shape of these complexes often leads to insights regarding their function. This information can be obtained in several important ways. Sometimes, a complex can be directly visualized using electron microscopy, as described in Chapter 9. A complementary approach relies on the hydrodynamic properties of a complex; that is, its behavior as it moves through a liquid medium. Usually, two separate measurements are made. One measure is the velocity of a complex as it moves under the influence of a centrifugal field produced by an ultracentrifuge (see Figure 8–7A). The sedimentation coefficient (or S value) obtained depends on both the size and the shape of the complex and does not, by itself, convey especially useful information. However, once a second hydrodynamic measurement is performed—by charting the migration of a complex through a gel-filtration chromatography column (see Figure 8–9B)—both the


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Cell_Biology_Alberts. useful information. However, once a second hydrodynamic measurement is performed—by charting the migration of a complex through a gel-filtration chromatography column (see Figure 8–9B)—both the approximate shape of a complex and its molecular weight can be calculated.


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Cell_Biology_Alberts. Molecular weight can also be determined more directly by using an analytical ultracentrifuge, a complex device that allows protein absorbance measurements to be made on a sample while it is subjected to centrifugal forces. In this approach, the sample is centrifuged until it reaches equilibrium, where the centrifugal force on a protein complex exactly balances its tendency to diffuse away. Because this balancing point is dependent on a complex’s molecular weight but not on its particular shape, the molecular weight can be directly calculated.


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Cell_Biology_Alberts. A frequent problem in cell biology and biochemistry is the identification of a protein or collection of proteins that has been obtained by one of the purification procedures discussed in the preceding pages. Because the genome sequences of most experimental organisms are now known, catalogs of all the proteins produced in those organisms are available. The task of identifying an unknown protein (or collection of unknown proteins) thus reduces to matching some of the amino acid


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Cell_Biology_Alberts. Figure 8–17 Western blotting. All the proteins from dividing tobacco cells in culture were first separated by two-dimensional polyacrylamide-gel electrophoresis. In (A), the positions of the proteins are revealed by a sensitive protein stain. In (B), the separated proteins on an identical gel were then transferred to a sheet of nitrocellulose and exposed to an antibody that recognizes only those proteins that are phosphorylated on threonine residues during mitosis. The positions of the few proteins that are recognized by this antibody are revealed by an enzyme-linked second antibody. (From J.A. Traas et al., Plant J. 2:723–732, 1992. With permission from Blackwell Publishing.)


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Cell_Biology_Alberts. Figure 8–18 The mass spectrometer. (A) Mass spectrometers used in biology contain an ion source that generates gaseous peptides or other molecules under conditions that render most molecules positively charged. The two major types of ion source are MALDI and electrospray, as described in the text. Ions are accelerated into a mass analyzer, which separates the ions on the basis of their mass and charge by one of three major methods: 1. Time-of-flight (TOF) analyzers determine the massto-charge ratio of each ion in the mixture from the rate at which it travels from the ion source to the detector. 2. Quadropole mass filters contain a long chamber lined by four electrodes that produce oscillating electric fields that govern the trajectory of ions; by varying the properties of the electric field over a wide range, a spectrum of ions with specific mass-to-charge ratios is allowed to pass through the chamber to the detector, while other ions are discarded. 3. Ion traps contain


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Cell_Biology_Alberts. electric field over a wide range, a spectrum of ions with specific mass-to-charge ratios is allowed to pass through the chamber to the detector, while other ions are discarded. 3. Ion traps contain doughnut-shaped electrodes producing a three-dimensional electric field that traps all ions in a circular chamber; the properties of the electric field can be varied over a wide range to eject a spectrum of specific ions to a detector. (B) Tandem mass spectrometry typically involves two mass analyzers separated by a collision chamber containing an inert, high-energy gas. The electric field of the first mass analyzer is adjusted to select a specific peptide ion, called a precursor ion, which is then directed to the collision chamber. Collision of the peptide with gas molecules causes random peptide fragmentation, primarily at peptide bonds, resulting in a highly complex mixture of fragments containing one or more amino acids from throughout the original peptide. The second mass analyzer is


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Cell_Biology_Alberts. fragmentation, primarily at peptide bonds, resulting in a highly complex mixture of fragments containing one or more amino acids from throughout the original peptide. The second mass analyzer is then used to measure the masses of the fragments (called product or daughter ions). With computer assistance, the pattern of fragments can be used to deduce the amino acid sequence of the original peptide.


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Cell_Biology_Alberts. sequences present in the unknown sample with known cataloged genes. This task is now performed almost exclusively by using mass spectrometry in conjunction with computer searches of databases. Charged particles have very precise dynamics when subjected to electrical and magnetic fields in a vacuum. Mass spectrometry exploits this principle to separate ions according to their mass-to-charge (m/z) ratio. It is an enormously sensitive technique. It requires very little material and is capable of determining the precise mass of intact proteins and of peptides derived from them by enzymatic or chemical cleavage. Masses can be obtained with great accuracy, often with an error of less than one part in a million.


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Cell_Biology_Alberts. Mass spectrometry is performed using complex instruments with three major components (Figure 8–18A). The first is the ion source, which transforms tiny amounts of a peptide sample into a gas containing individual charged peptide molecules. These ions are accelerated by an electric field into the second component, the mass analyzer, where electric or magnetic fields are used to separate the ions on the basis of their mass-to-charge ratios. Finally, the separated ions collide with a detector, which generates a mass spectrum containing a series of peaks representing the masses of the molecules in the sample.


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Cell_Biology_Alberts. There are many different types of mass spectrometer, varying mainly in the nature of their ion sources and mass analyzers. One of the most common ion sources depends on a technique called matrix-assisted laser desorption ionization (MALDI). In this approach, the proteins in the sample are first cleaved into short peptides by a protease such as trypsin. These peptides are mixed with an organic acid and then dried onto a metal or ceramic slide. A brief laser burst is directed toward the sample, producing a gaseous puff of ionized peptides, each carrying one or more positive charges. In many cases, the MALDI ion source is coupled to a mass analyzer called a time-of-flight (TOF) analyzer, which is a long chamber through which the ionized peptides are accelerated by an electric field toward a detector. Their mass and charge determine the time it takes them to reach the detector: large peptides move more slowly, and more highly charged molecules move more quickly. By analyzing those ionized


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Cell_Biology_Alberts. Their mass and charge determine the time it takes them to reach the detector: large peptides move more slowly, and more highly charged molecules move more quickly. By analyzing those ionized peptides that bear a single charge, the precise masses of peptides present in the original sample can be determined. This information is then used to search genomic databases, in which the masses of all proteins and of all their predicted peptide fragments have been tabulated from the genomic sequences of the organism. An unambiguous match to a particular open reading frame can often be made by knowing the mass of only a few peptides derived from a given protein.


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Cell_Biology_Alberts. By employing two mass analyzers in tandem (an arrangement known as MS/ MS; Figure 8–18B), it is possible to directly determine the amino acid sequences of individual peptides in a complex mixture. The MALDI-TOF instrument described above is not ideal for this method. Instead, MS/MS typically involves an electro-spray ion source, which produces a continuous thin stream of peptides that are ionized and accelerated into the first mass analyzer. The mass analyzer is typically either a quadropole or ion trap, which employs large electrodes to produce oscillating electric fields inside the chamber containing the ions. These instruments act as mass filters: the electric field is adjusted over a broad range to select a single peptide ion and discard all the others in the peptide mixture. In tandem mass spectrometry, this single ion is then exposed to an inert, high-energy gas, which collides with the peptide, resulting in fragmentation, primarily at peptide bonds. The second mass analyzer


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Cell_Biology_Alberts. mass spectrometry, this single ion is then exposed to an inert, high-energy gas, which collides with the peptide, resulting in fragmentation, primarily at peptide bonds. The second mass analyzer then determines the masses of the peptide fragments, which can be used by computational methods to determine the amino acid sequence of the original peptide and thereby identify the protein from which it came.


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Cell_Biology_Alberts. Tandem mass spectrometry is also useful for detecting and precisely mapping post-translational modifications of proteins, such as phosphorylations or acetylations. Because these modifications impart a characteristic mass increase to an amino acid, they are easily detected during the analysis of peptide fragments in the second mass analyzer, and the precise site of the modification can often be deduced from the spectrum of peptide fragments.


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Cell_Biology_Alberts. A powerful, “two-dimensional” mass spectrometry technique can be used to determine all of the proteins present in an organelle or another complex mixture of proteins. First, the mixture of proteins present is digested with trypsin to produce short peptides. Next, these peptides are separated by automated high-performance liquid chromatography (LC). Every peptide fraction from the chromatographic column is injected directly into an electrospray ion source on a tandem mass spectrometer (MS/MS), providing the amino acid sequence and post-translational modifications for every peptide in the mixture. This method, often called LC-MS/MS, is used to identify hundreds or thousands of proteins in complex protein mixtures from specific organelles or from whole cells. It can also be used to map all of the phosphorylation sites in the cell, or all of the proteins targeted by other post-translational modifications such as acetylation or ubiquitylation.


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Cell_Biology_Alberts. Sets of Interacting Proteins Can Be Identified by Biochemical Methods Because most proteins in the cell function as part of complexes with other proteins, an important way to begin to characterize the biological role of an unknown protein is to identify all of the other proteins to which it specifically binds. A key method for identifying proteins that bind to one another tightly is coimmunoprecipitation. A specific target protein is immunoprecipitated from a cell lysate using specific antibodies coupled to beads, as described earlier. If the target protein is associated tightly enough with another protein when it is captured by the antibody, the partner precipitates as well and can be identified by mass spectrometry. This method is useful for identifying proteins that are part of a complex inside cells, including those that interact only transiently—for example, when extracellular signal molecules stimulate cells (discussed in Chapter 15).


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Cell_Biology_Alberts. In addition to capturing protein complexes on columns or in test tubes, researchers are developing high-density protein arrays to investigate protein interactions. These arrays, which contain thousands of different proteins or antibodies spotted onto glass slides or immobilized in tiny wells, allow one to examine the biochemical activities and binding profiles of a large number of proteins at once. For example, if one incubates a fluorescently labeled protein with arrays containing thousands of immobilized proteins, the spots that remain fluorescent after extensive washing each contain a protein that specifically binds the labeled protein.


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Cell_Biology_Alberts. Once two proteins—or a protein and a small molecule—are known to associate, it becomes important to characterize their interaction in more detail. Proteins can associate with each other more or less permanently (like the subunits of RNA polymerase or the proteasome), or engage in transient encounters that may last only a few milliseconds (like a protein kinase and its substrate). To understand how a protein functions inside a cell, we need to determine how tightly it binds to other proteins, how rapidly it dissociates from them, and how covalent modifications, small molecules, or other proteins influence these interactions.


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Cell_Biology_Alberts. As we discussed in Chapter 3 (see Figure 3–44), the extent to which two proteins interact is determined by the rates at which they associate and dissociate. These rates depend, respectively, on the association rate constant (kon) and dissociation rate constant (koff). The kinetic rate constant koff is a particularly useful number because it provides valuable information about how long two proteins remain bound to one another. The ratio of the two kinetic constants (kon/koff) yields another very useful number called the equilibrium constant (K, also known as Keq or Ka), the inverse of which is the more commonly used dissociation constant Kd. The equilibrium constant is useful as a general indicator of the affinity of the interaction, and it can be used to estimate the amount of bound complex at different concentrations of the two protein partners—thereby providing insights into the importance of the interaction at the protein concentrations found inside the cell.


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Cell_Biology_Alberts. A wide range of methods can be used to determine binding constants for a two-protein complex. In a simple equilibrium binding experiment, two proteins are mixed at a range of concentrations, allowed to reach equilibrium, and the amount of bound complex is measured; half of the protein complex will be bound at a concentration that is equal to Kd. Equilibrium experiments often involve the use of radioactive or fluorescent tags on one of the protein partners, coupled with biochemical or optical methods for measuring the amount of bound protein. In a more complex kinetic binding experiment, the kinetic rate constants are determined using rapid methods that allow real-time measurement of the formation of a bound complex over time (to determine kon) or the dissociation of a bound complex over time (to determine koff).


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Cell_Biology_Alberts. Optical techniques provide particularly rapid, convenient, and accurate binding measurements, and in some cases the proteins do not even need to be labeled. Certain amino acids (tryptophan, for example) exhibit weak fluorescence that can be detected with sensitive fluorimeters. In many cases, the fluorescence intensity, or the emission spectrum of fluorescent amino acids located in a protein–protein interface, will change when two proteins associate. When this change can be detected by fluorimetry, it provides a simple and sensitive measure of protein binding that is useful in both equilibrium and kinetic binding experiments. A related but more widely useful optical binding technique is based on fluorescence anisotropy, a change in the polarized light that is emitted by a fluorescently tagged protein in the bound and free states (Figure 8–19).


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Cell_Biology_Alberts. Figure 8–19 Measurement of binding with fluorescence anisotropy. This method depends on a fluorescently tagged protein that is illuminated with polarized light at the appropriate wavelength for excitation; a fluorimeter is used to measure the intensity and polarization of the emitted light. If the fluorescent protein is fixed in position and therefore does not rotate during the brief period between excitation and emission, then the emitted light will be polarized at the same angle as the excitation light. This directional effect is called fluorescence anisotropy. Protein molecules in solution rotate or tumble rapidly, however, so that there is a decrease in the amount of anisotropic fluorescence. Larger molecules tumble at a slower rate and therefore have higher fluorescence anisotropy. (A) To measure the binding between a small molecule and a large receptor protein, the smaller molecule is labeled with a fluorophore. In the absence of its binding partner, the molecule tumbles


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Cell_Biology_Alberts. (A) To measure the binding between a small molecule and a large receptor protein, the smaller molecule is labeled with a fluorophore. In the absence of its binding partner, the molecule tumbles rapidly, resulting in low fluorescence anisotropy (top). When the small molecule binds to its larger partner, however, it tumbles less rapidly, resulting in an increase in fluorescence anisotropy (bottom). (B) In the equilibrium binding experiment shown here, a small, fluorescent peptide ligand was present at a low concentration, and the amount of fluorescence anisotropy (in millipolarization units, mP) was measured after incubation with various concentrations of a larger protein receptor for the ligand. From the hyperbolic curve that fits the data, it can be seen that 50% binding occurred at about 10 μM, which is equal to the dissociation constant Kd for the binding interaction.


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Cell_Biology_Alberts. Another optical method for probing protein interactions uses green fluorescent protein (discussed in detail below) and its derivatives of different colors. In this application, two proteins of interest are each labeled with a different fluorescent protein, such that the emission spectrum of one fluorescent protein overlaps the absorption spectrum of the second. If the two proteins come very close to each other (within about 1–5 nm), the energy of the absorbed light is transferred from one fluorescent protein to the other. The energy transfer, called fluorescence resonance energy transfer (FRET), is determined by illuminating the first fluorescent protein and measuring emission from the second (see Figure 9–26). When combined with fluorescence microscopy, this method can be used to characterize protein–protein interactions at specific locations inside living cells (discussed in Chapter 9).


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Cell_Biology_Alberts. Small chemical inhibitors of specific proteins have contributed a great deal to the development of cell biology. For example, the microtubule inhibitor colchicine is routinely used to test whether microtubules are required for a given biological process; it also led to the first purification of tubulin several decades ago. In the past, these small molecules were usually natural products; that is, they were synthesized by living creatures. Although natural products have been extraordinarily useful in science and medicine (see, for example, Table 6–4, p. 352), they act on a limited number of biological processes. However, the recent development of methods to synthesize hundreds of thousands of small molecules and to carry out large-scale automated screens holds the promise of identifying chemical inhibitors for virtually any biological process. In such approaches, large collections of small chemical compounds are simultaneously tested, either on living cells or in cell-free assays. Once


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Cell_Biology_Alberts. inhibitors for virtually any biological process. In such approaches, large collections of small chemical compounds are simultaneously tested, either on living cells or in cell-free assays. Once an inhibitor is identified, it can be used as a probe to identify, through affinity chromatography or other means, the protein to which the inhibitor binds. This general strategy, sometimes called chemical biology, has successfully identified inhibitors of many proteins that carry out key processes in cell biology. An inhibitor of a kinesin protein that functions in mitosis, for


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Cell_Biology_Alberts. Figure 8–20 Small-molecule inhibitors for manipulating living cells. (A) Chemical structure of monastrol, a kinesin inhibitor identified in a large-scale screen for small molecules that disrupt mitosis. (B) Normal mitotic spindle seen in an untreated cell. The microtubules are stained green and chromosomes blue. (C) Monopolar spindle that forms in cells treated with monastrol, which inhibits a kinesin protein required for separation of the spindle poles in early mitosis. (B and C, from T.U. Mayer et al., Science 286:971–974, 1999. With permission from AAAS.) example, was identified by this method (Figure 8–20). Chemical inhibitors give the cell biologist great control over the timing of inhibition, as drugs can be rapidly added to or removed from cells, allowing protein function to be switched on or off quickly.


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Cell_Biology_Alberts. The main technique that has been used to discover the three-dimensional structure of molecules, including proteins, at atomic resolution is x-ray crystallography. X-rays, like light, are a form of electromagnetic radiation, but they have a much shorter wavelength, typically around 0.1 nm (the diameter of a hydrogen 5 µm atom). If a narrow beam of parallel x-rays is directed at a sample of a pure protein, most of the x-rays pass straight through it. A small fraction, however, are scattered by the atoms in the sample. If the sample is a well-ordered crystal, the scattered waves reinforce one another at certain points and appear as diffraction spots when recorded by a suitable detector (Figure 8–21).


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Cell_Biology_Alberts. The position and intensity of each spot in the x-ray diffraction pattern contain information about the locations of the atoms in the crystal that gave rise to it. Deducing the three-dimensional structure of a large molecule from the diffraction pattern of its crystal is a complex task and was not achieved for a protein molecule until 1960. But in recent years x-ray diffraction analysis has become increasingly automated, and now the slowest step is likely to be the generation of suitable protein crystals. This step requires large amounts of very pure protein and often involves years of trial and error to discover the proper crystallization conditions; the pace has greatly accelerated with the use of recombinant DNA techniques to produce pure proteins and robotic techniques to test large numbers of crystallization conditions.


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Cell_Biology_Alberts. Analysis of the resulting diffraction pattern produces a complex three-dimensional electron-density map. Interpreting this map—translating its contours into a three-dimensional structure—is a complicated procedure that requires knowledge of the amino acid sequence of the protein. Largely by trial and error, the sequence and the electron-density map are correlated by computer to give the best possible fit. The reliability of the final atomic model depends on the resolution of the original crystallographic data: 0.5 nm resolution might produce a low-resolution map of the polypeptide backbone, whereas a resolution of 0.15 nm allows all of the non-hydrogen atoms in the molecule to be reliably positioned.


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Cell_Biology_Alberts. A complete atomic model is often too complex to appreciate directly, but simplified versions that show a protein’s essential structural features can be readily derived from it (see Panel 3–2, pp. 142–143). The three-dimensional structures of tens of thousands of different proteins have been determined by x-ray crystallography or by NMR spectroscopy (see page 461)—enough to allow the grouping of common structures into families (Movie 8.1). These structures or protein folds often seem to be more conserved in evolution than are the amino acid sequences that form them (see Figure 3–13).


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Cell_Biology_Alberts. X-ray crystallographic techniques can also be applied to the study of macromolecular complexes. The method was used, for example, to determine the structure of the ribosome, a large and complex machine made of several RNAs and more than 50 proteins (see Figure 6–62). The determination required the use of a synchrotron, a radiation source that generates x-rays with the intensity needed to analyze the crystals of such large macromolecular complexes.


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Cell_Biology_Alberts. beam of x-rays x-ray source protein crystal diffracted beams x-ray diffraction pattern obtained from the protein crystal beam stop (A) (B) (C)Figure 8–21 X-ray crystallography. (A) A narrow beam of x-rays is directed at a well-ordered crystal (B). Shown here is a protein crystal of ribulose bisphosphate carboxylase, an enzyme with a central role in CO2 fixation during photosynthesis. The atoms in the crystal scatter some of the beam, and the scattered waves reinforce one another at certain points and appear as a pattern of diffraction spots (C). This diffraction pattern, together with the amino acid sequence of the protein, can be used to produce an atomic model (D). The complete atomic model is hard to interpret, but this simplified version, derived from the x-ray diffraction data, shows the protein’s structural features clearly (α helices, green; β strands, red). The components pictured in A to D are not shown to scale. (B, courtesy of C. Branden; C, courtesy of J. Hajdu and I.


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Cell_Biology_Alberts. the protein’s structural features clearly (α helices, green; β strands, red). The components pictured in A to D are not shown to scale. (B, courtesy of C. Branden; C, courtesy of J. Hajdu and I. Andersson; D, adapted from original provided by B. Furugren.)


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Cell_Biology_Alberts. NMR Can Be Used to Determine Protein Structure in Solution Nuclear magnetic resonance (NMR) spectroscopy has been widely used for many years to analyze the structure of small molecules, small proteins, or protein domains. Unlike x-ray crystallography, NMR does not depend on having a crystalline sample. It simply requires a small volume of concentrated protein solution that is placed in a strong magnetic field; indeed, it is the main technique that yields detailed evidence about the three-dimensional structure of molecules in solution.


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Cell_Biology_Alberts. Certain atomic nuclei, particularly hydrogen nuclei, have a magnetic moment or spin: that is, they have an intrinsic magnetization, like a bar magnet. The spin aligns along the strong magnetic field, but it can be changed to a misaligned, excited state in response to applied radiofrequency (RF) pulses of electromagnetic radiation. When the excited hydrogen nuclei return to their aligned state, they emit RF radiation, which can be measured and displayed as a spectrum. The nature of the emitted radiation depends on the environment of each hydrogen nucleus, and if one nucleus is excited, it influences the absorption and emission of radiation by other nuclei that lie close to it. It is consequently possible, by an ingenious elaboration of the basic NMR technique known as two-dimensional NMR, to distinguish the signals from hydrogen nuclei in different amino acid residues, and to identify and measure the small shifts in these signals that occur when these hydrogen nuclei lie close enough


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Cell_Biology_Alberts. to distinguish the signals from hydrogen nuclei in different amino acid residues, and to identify and measure the small shifts in these signals that occur when these hydrogen nuclei lie close enough together to interact. Because the size of such a shift reveals the distance between the interacting pair of hydrogen atoms, NMR can provide information about the distances between the parts of the protein molecule. By combining this information with a knowledge of the amino acid


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Cell_Biology_Alberts. Figure 8–22 NMR spectroscopy. An example of the data from an NMR machine. This two-dimensional NMR spectrum is derived from the C-terminal domain of the enzyme cellulase. The spots represent interactions between hydrogen atoms that are near neighbors in the protein and hence reflect the distance that separates them. Complex computing methods, in conjunction with the known amino acid sequence, enable possible compatible structures to be derived. Ten structures of the enzyme, which all satisfy the distance constraints equally well, are shown superimposed on one another, giving a good indication of the probable three-dimensional structure. (Courtesy of P. Kraulis.) sequence, it is possible in principle to compute the three-dimensional structure of the protein (Figure 8–22).


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Cell_Biology_Alberts. P. Kraulis.) sequence, it is possible in principle to compute the three-dimensional structure of the protein (Figure 8–22). For technical reasons, the structure of small proteins of about 20,000 daltons or less can be most readily determined by NMR spectroscopy. Resolution decreases as the size of a macromolecule increases. But recent technical advances have now pushed the limit to about 100,000 daltons, thereby making the majority of proteins accessible for structural analysis by NMR. Because NMR studies are performed in solution, this method also offers a convenient means of monitoring changes in protein structure, for example during protein folding or when the protein binds to another molecule. NMR is also used widely to investigate molecules other than proteins and is valuable, for example, as a method to determine the three-dimensional structures of RNA molecules and the complex carbohydrate side chains of glycoproteins.


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Cell_Biology_Alberts. A third major method for the determination of protein structure, and particularly the structure of large protein complexes, is single-particle analysis by electron microscopy. We discuss this approach in Chapter 9. Having discussed methods for purifying and analyzing proteins, we now turn to a common situation in cell and molecular biology: an investigator has identified a gene important for a biological process but has no direct knowledge of the biochemical properties of its protein product.


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Cell_Biology_Alberts. Thanks to the proliferation of protein and nucleic acid sequences that are cataloged in genome databases, the function of a gene—and its encoded protein— can often be predicted by simply comparing its sequence with those of previously characterized genes. Because amino acid sequence determines protein structure, and structure dictates biochemical function, proteins that share a similar amino acid sequence usually have the same structure and usually perform similar biochemical functions, even when they are found in distantly related organisms. In modern cell biology, the study of a newly discovered protein usually begins with a search for previously characterized proteins that are similar in their amino acid sequences.


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Cell_Biology_Alberts. Searching a collection of known sequences for similar genes or proteins is typically done over the Internet, and it simply involves selecting a database and entering the desired sequence. A sequence-alignment program—the most popular is BLAST—scans the database for similar sequences by sliding the submitted sequence along the archived sequences until a cluster of residues falls into full or partial alignment (Figure 8–23). Such comparisons can predict the functions of individual proteins, families of proteins, or even most of the protein complement of a newly sequenced organism.


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Cell_Biology_Alberts. As was explained in Chapter 3, many proteins that adopt the same conformation and have related functions are too distantly related to be identified from a comparison of their amino acid sequences alone (see Figure 3–13). Thus, an ability to reliably predict the three-dimensional structure of a protein from its amino acid sequence would improve our ability to infer protein function from the sequence information in genomic databases. In recent years, major progress has been made in predicting the precise structure of a protein. These predictions are based, in part, on our knowledge of the thousands of protein structures that have already been determined by x-ray crystallography and NMR spectroscopy and, in part, on computations using our knowledge of the physical forces acting on the atoms. However, it remains a substantial and important challenge to predict the structures of proteins that are large or have multiple domains, or to predict structures at the very high levels of resolution


Cell_Biology_Alberts

Cell_Biology_Alberts. However, it remains a substantial and important challenge to predict the structures of proteins that are large or have multiple domains, or to predict structures at the very high levels of resolution needed to assist in computer-based drug discovery.


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Cell_Biology_Alberts. While finding related sequences and structures for a new protein will provide many clues about its function, it is usually necessary to test these insights through direct experimentation. However, the clues generated from sequence comparisons typically point the investigator in the correct experimental direction, and their use has therefore become one of the most important strategies in modern cell biology. Many methods exist for identifying proteins and analyzing their biochemical properties, structures, and interactions with other proteins. Small-molecule inhibitors allow the functions of proteins they act upon to be studied in living cells. Because proteins with similar structures often have similar functions, the biochemical activity of a protein can often be predicted by searching databases for previously characterized proteins that are similar in their amino acid sequences.


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Cell_Biology_Alberts. Until the early 1970s, DNA was the most difficult biological molecule for the biochemist to analyze. Enormously long and chemically monotonous, the string of nucleotides that forms the genetic material of an organism could be examined only indirectly, by protein sequencing or by genetic analysis. Today, the situation has changed entirely. From being the most difficult macromolecule of the cell to Figure 8–23 Results of a bLAST search.


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Cell_Biology_Alberts. Sequence databases can be searched to find similar amino acid or nucleic acid sequences. Here, a search for proteins similar to the human cell-cycle regulatory protein Cdc2 (Query) locates maize Cdc2 (Sbjct), which is 68% identical to human Cdc2 in its amino acid sequence. The alignment begins at residue 57 of the Query protein, suggesting that the human protein has an N-terminal region that is absent from the maize protein. The green blocks indicate differences in sequence, and the yellow bar summarizes the similarities: when the two amino acid sequences are identical, the residue is shown; similar amino acid substitutions are indicated by a plus sign (+). Only one small gap has been introduced—indicated by the red arrow at position 194 in the Query sequence—to align the two sequences maximally. The alignment score (Score), which is expressed in two different types of units, takes into account penalties for substitutions and gaps; the higher the alignment score, the better the match.


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Cell_Biology_Alberts. The alignment score (Score), which is expressed in two different types of units, takes into account penalties for substitutions and gaps; the higher the alignment score, the better the match. The significance of the alignment is reflected in the Expectation (E) value, which specifies how often a match this good would be expected to occur by chance. The lower the E value, the more significant the match; the extremely low value here (e–111) indicates certain significance. E values much higher than 0.1 are unlikely to reflect true relatedness. For example, an E value of 0.1 means there is a 1 in 10 likelihood that such a match would arise solely by chance.


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Cell_Biology_Alberts. analyze, DNA has become the easiest. It is now possible to determine the entire nucleotide sequence of a bacterial or fungal genome in a matter of hours, and the sequence of an individual human genome in less than a day. Once the nucleotide sequence of a genome is known, any individual gene can be easily isolated, and large quantities of the gene product (be it RNA or protein) can be made either by introducing the gene into bacteria or animal cells and coaxing these cells to over-express the foreign gene or by synthesizing the gene product in vitro. In this way, proteins and RNA molecules that might be present in only tiny amounts in living cells can be produced in large quantities for biochemical and structural analyses. And this approach can also be used to produce large quantities of human proteins (such as insulin, or interferon, or blood-clotting proteins) for use as human pharmaceuticals. As we will see later in this chapter, it is also possible for scientists to alter an


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Cell_Biology_Alberts. of human proteins (such as insulin, or interferon, or blood-clotting proteins) for use as human pharmaceuticals. As we will see later in this chapter, it is also possible for scientists to alter an isolated gene and transfer it back into the germ line of an animal or plant, so as to become a functional and heritable part of the organism’s genome. In this way, the biological roles of any gene can be assessed by observing—in the whole organism—the results of modifying it.


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Cell_Biology_Alberts. The ability to manipulate DNA with precision in a test tube or an organism, known as recombinant DNA technology has had a dramatic impact on all aspects of cell and molecular biology, allowing us to routinely study cells and their macromolecules in ways that were unimaginable even twenty years ago. Central to the technology are the following manipulations: 1. Cleavage of DNA at specific sites by restriction nucleases, which greatly facilitates the isolation and manipulation of individual pieces of a genome. 2. DNA ligation, which makes it possible to seamlessly join together DNA molecules from widely different sources. 3. DNA cloning (through the use of either cloning vectors or the polymerase chain reaction) in which a portion of a genome (often an individual gene) is “purified” away from the remainder of the genome by repeatedly copying it to generate many billions of identical molecules. 4.


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Cell_Biology_Alberts. 4. Nucleic acid hybridization, which makes it possible to identify any specific sequence of DNA or RNA with great accuracy and sensitivity based on its ability to selectively bind a complementary nucleic acid sequence. 5. DNA synthesis, which makes it possible to chemically synthesize DNA molecules with any sequence of nucleotides, whether or not the sequence occurs in nature. 6. Rapid determination of the sequence of nucleotides of any DNA or RNA molecule. In the following sections, we describe each of these basic techniques which, together, have revolutionized the study of cell and molecular biology.


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Cell_Biology_Alberts. Unlike a protein, a gene does not exist as a discrete entity in cells, but rather as a small region of a much longer DNA molecule. Although the DNA molecules in a cell can be randomly broken into small pieces by mechanical force, a fragment containing a single gene in a mammalian genome would still be only one among a hundred thousand or more DNA fragments, indistinguishable in their average size. How could such a gene be separated from all the others? Because all DNA molecules consist of an approximately equal mixture of the same four nucleotides, they cannot be readily separated, as proteins can, on the basis of their different charges and biochemical properties. The solution to this problem began to emerge with the discovery of restriction nucleases. These enzymes, which are purified from bacteria, cut the DNA double helix at specific sites defined by the local nucleotide sequence, thereby cleaving a long, double-stranded DNA molecule into fragments of strictly defined sizes.


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Cell_Biology_Alberts. Like many of the tools of recombinant DNA technology, restriction nucleases were discovered by researchers trying to understand an intriguing biological phenomenon. It had been observed that certain bacteria always degraded “foreign” DNA that was introduced into them experimentally. A search for the mechanism responsible revealed a then unanticipated class of bacterial nucleases that cleave DNA at specific nucleotide sequences. The bacterium’s own DNA is protected from cleavage by methylation of these same sequences, thereby protecting a bacterium’s own genome from being overrun by foreign DNA. Because these enzymes restrict the transfer of DNA into bacteria, they were called restriction nucleases. The pursuit of this seemingly arcane biological puzzle set off the development of technologies that have forever changed the way cell and molecular biologists study living things.


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Cell_Biology_Alberts. Different bacterial species produce different restriction nucleases, each cutting at a different, specific nucleotide sequence (Figure 8–24). Because these target sequences are short—generally four to eight nucleotide pairs—many sites of cleavage will occur, purely by chance, in any long DNA molecule. The reason restriction nucleases are so useful in the laboratory is that each enzyme will always cut a particular DNA molecule at the same sites. Thus for a given sample of DNA (which contains many identical molecules), a particular restriction nuclease will reliably generate the same set of DNA fragments.


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Cell_Biology_Alberts. The size of the resulting fragments depends on the length of the target sequences of the restriction nucleases. As shown in Figure 8–24, the enzyme HaeIII cuts at a sequence of four nucleotide pairs; a sequence this long would be expected to occur purely by chance approximately once every 256 nucleotide pairs (1 in 44). In comparison, a restriction nuclease with a target sequence that is eight nucleotides long would be expected to cleave DNA on average once every 65,536 nucleotide pairs (1 in 48). This difference in sequence selectivity makes it possible to cleave a long DNA molecule into the fragment sizes that are most suitable for a given application. Gel Electrophoresis Separates DNA Molecules of Different Sizes


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Cell_Biology_Alberts. The same types of gel-electrophoresis methods that have proved so useful in the analysis of proteins (see Figure 8–13) can be applied to DNA molecules. The procedure is actually simpler than for proteins: because each nucleotide in a nucleic acid molecule already carries a single negative charge (on the phosphate group), there is no need to add the negatively charged detergent SDS that is required to make protein molecules move uniformly toward the positive electrode. Larger DNA fragments will migrate more slowly because their progress is impeded to a greater extent by the gel matrix. Over several hours, the DNA fragments become spread out across the gel according to size, forming a ladder of discrete bands, each composed of a collection of DNA molecules of identical length (Figure 8–25A and B). To separate DNA molecules longer than 500 nucleotide pairs, the gel is made of a diluted solution of agarose (a polysaccharide isolated from seaweed). For DNA fragments less than 500


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Cell_Biology_Alberts. 8–25A and B). To separate DNA molecules longer than 500 nucleotide pairs, the gel is made of a diluted solution of agarose (a polysaccharide isolated from seaweed). For DNA fragments less than 500 nucleotides long, specially designed polyacrylamide gels allow the separation of molecules that differ in length by as little as a single nucleotide (see Figure 8–25C).


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Cell_Biology_Alberts. Figure 8–24 Restriction nucleases cleave DNA at specific nucleotide sequences.


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Cell_Biology_Alberts. Like the sequence-specific DNA-binding proteins we encountered in Chapter 7 (see Figure 7–8), restriction enzymes often work as dimers, and the DNA sequence they recognize and cleave is often symmetrical around a central point. Here, both strands of the DNA double helix are cut at specific points within the target sequence (orange). Some enzymes, such as HaeIII, cut straight across the double helix and leave two blunt-ended DNA molecules; with others, such as EcoRI and HindIII, the cuts on each strand are staggered. These staggered cuts generate “sticky ends”—short, single-stranded overhangs that help the cut DNA molecules join back together through complementary base-pairing. This rejoining of DNA molecules becomes important for DNA cloning, as we discuss below. Restriction nucleases are usually obtained from bacteria, and their names reflect their origins: for example, the enzyme EcoRI comes from Escherichia coli. There are currently hundreds of different restriction enzymes


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Cell_Biology_Alberts. are usually obtained from bacteria, and their names reflect their origins: for example, the enzyme EcoRI comes from Escherichia coli. There are currently hundreds of different restriction enzymes available; they can be ordered from companies that commercially produce them.


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Cell_Biology_Alberts. A variation of agarose-gel electrophoresis, called pulsed-field gel electrophoresis, makes it possible to separate extremely long DNA molecules, even those found in whole chromosomes. Ordinary gel electrophoresis fails to separate very large DNA molecules because the steady electric field stretches them out so that they travel end-first through the gel in snakelike configurations at a rate that is independent of their length. In pulsed-field gel electrophoresis, by contrast, the direction of the electric field changes periodically, which forces the molecules to reorient before continuing to move snakelike through the gel. This re-orientation takes much more time for larger molecules, so that longer molecules move more slowly than shorter ones. As a consequence, entire bacterial or yeast chromosomes separate into discrete bands in pulsed-field gels and so can be sorted and identified on the basis of their size (Figure 8–25D). Although a typical mammalian chromosome of 108 nucleotide


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Cell_Biology_Alberts. chromosomes separate into discrete bands in pulsed-field gels and so can be sorted and identified on the basis of their size (Figure 8–25D). Although a typical mammalian chromosome of 108 nucleotide pairs is still too long to be sorted even in this way, large segments of these chromosomes are readily separated and identified if the chromosomal DNA is first cut with a restriction nuclease selected to recognize sequences that occur only rarely.


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Cell_Biology_Alberts. The DNA bands on agarose or polyacrylamide gels are invisible unless the DNA is labeled or stained in some way. A particularly sensitive method of staining DNA is to soak the gel in the dye ethidium bromide, which fluoresces under ultraviolet light when it is bound to DNA (see Figure 8–25B and D). Even more sensitive detection methods incorporate a radioisotope or chemical marker into the DNA molecules before electrophoresis, as we next describe.


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Cell_Biology_Alberts. Figure 8–25 DNA molecules can be separated by size using gel electrophoresis. (A) Schematic illustration comparing the results of cutting the same DNA molecule (in this case, the genome of a virus that infects wasps) with two different restriction nucleases, EcoRI (middle) and HindIII (right). The fragments are then separated by gel electrophoresis using a gel matrix of agarose. Because larger fragments migrate more slowly than smaller ones, the lowermost bands on the gel contain the smallest DNA fragments. The sizes of the fragments can be estimated by comparing them to a set of DNA fragments of known sizes (left). (B) Photograph of an actual agarose gel showing DNA “bands” that have been stained with ethidium bromide.


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Cell_Biology_Alberts. A polyacrylamide gel with small pores was used to separate short DNA molecules that differ by only a single nucleotide. Shown here are the results of a dideoxy sequencing reaction, explained later in this chapter. From left to right, the bands in the four lanes were produced by adding G, A, T, and C chain-terminating nucleotides (see Panel 8–1). The DNA molecules were labeled with 32P, and the image shown was produced by laying a piece of photographic film over the gel and allowing the 32P to expose the film, producing the dark bands observed when the film was developed.


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Cell_Biology_Alberts. The technique of pulsed-field agarose-gel electrophoresis was used to separate the 16 different chromosomes of the yeast species Saccharomyces cerevisiae, which range in size from 220,000 to 2.5 million nucleotide pairs. The DNA was stained as in (B). DNA molecules as large as 107 nucleotide pairs can be separated in this way. (B, from U. Albrecht et al., J. Gen. Virol. 75:3353-3363, 1994; C, courtesy of Leander Lauffer and Peter Walter; D, from D. Vollrath and R.W. Davis, Nucleic Acids Res. 15:7865–7876, 1987. With permission from Oxford University Press.) 2.5 4 million 15 7 950,000 6.5 4.3 14 10 11 610,000 2.3 2 3 6 1 220,000 (A) slab of agarose gel (B) (C) (D) Purified DNA Molecules Can Be Specifically Labeled with Radioisotopes or Chemical Markers in vitro


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Cell_Biology_Alberts. The DNA polymerases that synthesize and repair DNA (discussed in Chapter 5) have become important tools in experimentally manipulating DNA. Because they synthesize sequences complementary to an existing DNA molecule, they are often used in the test tube to create exact copies of existing DNA molecules. The copies can include specially modified nucleotides (Figure 8–26). To synthesize DNA in this way, the DNA polymerase is presented with a template and a pool of nucleotide precursors that contain the modification. As long as the polymerase can use these precursors, it automatically makes new, modified molecules that match the sequence of the template. Modified DNA molecules have many uses. DNA labeled with the radioisotope 32P can be detected following gel electrophoresis by placing the gel next to a piece of photographic film (see Figure 8–25C). The 32P atoms emit β particles which expose the film, producing a visible record of every band on the gel. Alternatively, the gel can be


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Cell_Biology_Alberts. gel next to a piece of photographic film (see Figure 8–25C). The 32P atoms emit β particles which expose the film, producing a visible record of every band on the gel. Alternatively, the gel can be scanned by a detector that measures the β emissions directly. Other types of modified DNA, such as that labeled by digoxigenin (see Figure 8–26B), are useful for visualizing DNA molecules in whole cells, a topic we discuss later in this chapter.


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Cell_Biology_Alberts. Any DNA fragment can be cloned. In molecular biology, the term DNA cloning is used in two senses. It literally refers to the act of making many identical copies (typically billions) of a DNA molecule—the amplification of a particular DNA sequence. However, the term also describes the isolation of a particular stretch of DNA (often a particular gene) from the rest of the cell’s genome; the same term purifed fragment of duplex DNA DNA polymerase incorporates 32P nucleotides, resulting in a population of radiolabeled DNA molecules that contain sequences from both strands


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Cell_Biology_Alberts. Figure 8–26 Methods for labeling DNA molecules in vitro. (A) A purified DNA polymerase enzyme can incorporate radiolabeled nucleotides as it synthesizes new DNA molecules. In this way, radiolabeled versions of any DNA sequence can be prepared in the laboratory. (B) The method in (A) is also used to produce nonradioactive DNA molecules that carry a specific chemical marker that can be detected with an appropriate antibody. The base on the nucleoside triphosphate shown is an analog of thymine, in which the methyl group on T has been replaced by a spacer arm linked to the plant steroid digoxigenin. An anti-digoxigenin antibody coupled to a visible marker such as a fluorescent dye is then used to visualize the DNA. Other chemical labels, such as biotin, can be attached to nucleotides and used in the same way. The only requirements are that the modified nucleotides properly base-pair and appear “normal” to the DNA polymerase.


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Cell_Biology_Alberts. circular, is used because this isolation is usually accomplished by making many identical copies of only the DNA of interest. We note that elsewhere in the book, cloning, particularly when used in the context of developmental biology, can also refer to the generation of many genetically identical cells starting from a single cell or even to the generation of genetically identical organisms (see, for example, Figure 7–2). In all cases, cloning refers to the act of making many identical copies, and in this section, we use the term to refer to methods designed to generate many identical copies of a defined segment of nucleic acid.


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Cell_Biology_Alberts. DNA cloning can be accomplished in several ways. One of the simplest involves inserting a particular fragment of DNA into the purified DNA genome of a self-replicating genetic element—usually a plasmid. The plasmid vectors most widely used for gene cloning are small, circular molecules of double-stranded DNA derived from plasmids that occur naturally in bacterial cells. They generally account for only a minor fraction of the total host bacterial cell DNA, but owing to their small size, they can easily be separated from the much larger chromosomal DNA molecules, which precipitate as a pellet upon centrifugation. For use as cloning vectors, the purified plasmid DNA circles are first cut with a restriction nuclease to create linear DNA molecules. The DNA to be cloned is added to the cut plasmid and then covalently joined using the enzyme DNA ligase (Figure 8–27 and Figure 8–28). As discussed in Chapter 5, this enzyme is used by the cell to stitch together the Okazaki fragments produced


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Cell_Biology_Alberts. and then covalently joined using the enzyme DNA ligase (Figure 8–27 and Figure 8–28). As discussed in Chapter 5, this enzyme is used by the cell to stitch together the Okazaki fragments produced during DNA replication. The recombinant DNA circle is introduced back into bacterial cells that have been made transiently permeable to DNA. As the cells grow and divide, doubling in number every 30 minutes, the recombinant plasmids also replicate to produce an


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Cell_Biology_Alberts. Figure 8–27 The insertion of a DNA fragment into a bacterial plasmid with the enzyme DNA ligase. The plasmid is cut open with a restriction nuclease (in this case, one that produces staggered ends) and is mixed with the DNA fragment to be cloned (which has been prepared with the same restriction nuclease). DNA ligase and ATP are added. The staggered ends base-pair, and DNA ligase seals the nicks in the DNA backbone, producing a complete recombinant DNA molecule. In the accompanying micrographs, the inserted DNA is colored red. (Micrographs courtesy of Huntington Potter and David Dressler.) Figure 8–28 DNA ligase can join together any two DNA fragments in vitro to produce recombinant DNA molecules. ATP provides the energy necessary to reseal the sugar-phosphate backbone of DNA (see Figure 5–12).


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Cell_Biology_Alberts. DNA ligase can readily join two DNA fragments produced by the same restriction nuclease, in this case EcoRI. Note that the staggered ends produced by this enzyme enable the ends of the two fragments to base-pair correctly with each other, greatly facilitating their rejoining. DNA ligase can also be used to join DNA fragments produced by different restriction nucleases—for example, EcoRI and HaeIII. In this case, before the fragments undergo ligation, DNA polymerase plus a mixture of deoxyribonucleoside triphosphates (dNTPs) are used to fill in the staggered cut produced by EcoRI. Each DNA fragment shown in the figure is oriented so that its 5ʹ ends are at the left end of the upper strand and the right end of the lower strand, as indicated.


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Cell_Biology_Alberts. cell culture produces hundreds of millions of new bacteria enormous number of copies of DNA circles containing the foreign DNA (Figure 8–29). Once the cells are lysed and the plasmid DNA isolated, the cloned DNA fragment can be readily recovered by cutting it out of the plasmid DNA with the same restriction nuclease that was used to insert it, and then separating it from the plasmid DNA by gel electrophoresis. Together, these steps allow the amplification and purification of any segment of DNA from the genome of any organism.


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Cell_Biology_Alberts. A particularly useful plasmid vector is based on the naturally occurring F plasmid of E. coli. Unlike smaller bacterial plasmids, the F plasmid—and its engineered derivative, the bacterial artificial chromosome (BAC)—is present in only one or two copies per E. coli cell. The fact that BACs are kept in such low numbers means that they can stably maintain very long DNA sequences, up to 1 million nucleotide pairs in length. With only a few BACs present per bacterium, it is less likely that the cloned DNA fragments will become scrambled by recombination with sequences carried on other copies of the plasmid. Because of their stability, ability to accept large DNA inserts, and ease of handling, BACs are now the preferred vector for handling large fragments of foreign DNA. As we will see below, BACs were instrumental in determining the complete nucleotide sequence of the human genome.


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Cell_Biology_Alberts. Often it is useful to break up a genome into much smaller fragments and clone every fragment, separately, using a plasmid vector. This approach is useful because it allows scientists to work with easily managed, discrete pieces of a genome instead of whole, unwieldy chromosomes.


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Cell_Biology_Alberts. This strategy involves cleaving genomic DNA into small pieces using a restriction nuclease (or, in some cases, by mechanically shearing the DNA) and ligating the entire collection of DNA fragments into plasmid vectors, using conditions that favor the insertion of a single DNA fragment into each plasmid molecule. These recombinant plasmids are then introduced into E. coli at a concentration that ensures that no more than one plasmid molecule is taken up by each bacterium. The collection of cloned plasmid molecules is known as a DNA library. Because the DNA fragments were derived directly from the chromosomal DNA of the organism of interest, the resulting collection—called a genomic library—will represent the entire genome of that organism (Figure 8–30), spread out over tens of thousands of individual bacterial colonies.


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Cell_Biology_Alberts. An alternative strategy, one that enriches for protein-coding genes, is to begin the cloning process by selecting only those DNA sequences that are transcribed into mRNA and thus correspond to protein-encoding genes. This is done by extracting the mRNA from cells and then making a DNA copy of each mRNA Figure 8–30 Human genomic libraries containing DNA fragments that represent the whole human genome can be constructed using restriction nucleases and DNA ligase. Such a genomic library consists of a set of bacteria, each carrying a different fragment of human DNA. For simplicity, only the colored DNA fragments are shown in the library; in reality, all of the different gray fragments will also be represented.


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Cell_Biology_Alberts. Figure 8–29 A DNA fragment can be replicated inside a bacterial cell. To clone a particular fragment of DNA, it is first inserted into a plasmid vector, as shown in Figure 8–27. The resulting recombinant plasmid DNA is then introduced into a bacterium, where it is replicated many millions of times as the bacterium multiplies. For simplicity, the genome of the bacterial cell is not shown.


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Cell_Biology_Alberts. millions of genomic DNA fragments molecule present—a so-called complementary DNA, or cDNA. The copying reaction is catalyzed by the reverse transcriptase enzyme of retroviruses, which synthesizes a complementary DNA chain on an RNA template. The single-stranded cDNA molecules synthesized by the reverse transcriptase are converted by DNA polymerase into double-stranded cDNA molecules, and these molecules are inserted into a plasmid or virus vector and cloned (Figure 8–31). Each clone obtained in this way is called a cDNA clone, and the entire collection of clones derived from one mRNA preparation constitutes a cDNA library.


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Cell_Biology_Alberts. Figure 8–32 illustrates some important differences between genomic DNA clones and cDNA clones. Genomic clones represent a random sample of all of the DNA sequences in an organism—both coding and noncoding—and, with very rare exceptions, are the same regardless of the cell type used to prepare them. By contrast, cDNA clones contain only those regions of the genome that have been transcribed into mRNA. Because the cells of different tissues produce distinct sets of mRNA molecules, a distinct cDNA library is obtained for each type of cell used to prepare the library. tissue (e.g., brain) double-stranded cDNA copy of original mRNA


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Cell_Biology_Alberts. Figure 8–31 The synthesis of cDNA. Total mRNA is extracted from a particular tissue, and the enzyme reverse transcriptase (see Figure 5–62) is used to produce DNA copies (cDNA) of the mRNA molecules. For simplicity, the copying of just one of these mRNAs into cDNA is illustrated. A short oligonucleotide complementary to the poly-A tail at the 3ʹ end of the mRNA (discussed in Chapter 6) is first hybridized to the RNA to act as a primer for the reverse transcriptase, which then copies the RNA into a complementary DNA chain, thereby forming a DNA–RNA hybrid helix. Treating the DNA–RNA hybrid with a specialized nuclease (RNAse H) that attacks only the RNA produces nicks and gaps in the RNA strand. DNA polymerase then copies the remaining single-stranded cDNA into double-stranded cDNA. Because DNA polymerase can synthesize through the bound RNA molecules, the RNA fragment that is base-paired to the 3ʹ end of the first DNA strand usually acts as the primer for the second strand synthesis,


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Cell_Biology_Alberts. DNA polymerase can synthesize through the bound RNA molecules, the RNA fragment that is base-paired to the 3ʹ end of the first DNA strand usually acts as the primer for the second strand synthesis, as shown. Any remaining RNA is eventually degraded during subsequent cloning steps. As a result, the nucleotide sequences at the extreme 5ʹ ends of the original mRNA molecules are often absent from cDNA libraries.


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Cell_Biology_Alberts. Genomic libraries are especially useful in determining the nucleotide sequences of a whole genome. For example, to determine the nucleotide sequence of the human genome, it was broken up into roughly 100,000-nucleotide-pair pieces, each of which was inserted into a BAC plasmid and amplified in E. coli. The resulting genomic library consisted of tens of thousands of bacterial colonies, each containing a different human DNA insert. The nucleotide sequence of each insert was determined separately and the sequence of the entire genome was stitched together from the pieces. The most important advantage of cDNA clones, over genomic clones, is that they contain the uninterrupted coding sequence of a gene. When the aim of the cloning, for example, is to produce the protein in large quantities by expressing the cloned gene in a bacterial or yeast cell, it is more preferable to start with cDNA.


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Cell_Biology_Alberts. Genomic and cDNA libraries are inexhaustible resources, which are widely shared among investigators. Today, many such libraries are also available from commercial sources. Because the identity of each insert in a library is often known (through sequencing the insert), it is often possible to order a particular region of a chromosome (or, in the case of cDNA, a complete, intron-less protein-coding gene) and have it delivered by mail. Cloning DNA by using bacteria revolutionized the study of genomes and is still in wide use today. However, there is an even simpler way to clone DNA, one that can be carried out entirely in vitro. We discuss this approach, called the polymerase chain reaction, below. However, first we need to review a fundamental, far-reaching property of DNA and RNA called hybridization. Figure 8–32 The differences between cDNA clones and genomic DNA clones derived from the same region of DNA.


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Cell_Biology_Alberts. Figure 8–32 The differences between cDNA clones and genomic DNA clones derived from the same region of DNA. In this example, gene A is infrequently transcribed, whereas gene B is frequently transcribed, and both genes contain introns (orange). In the genomic DNA library, both the introns and the nontranscribed DNA (gray) are included in the clones, and most clones contain, at most, only part of the coding sequence of a gene (red). In the cDNA clones, the intron sequences (yellow) have been removed by RNA splicing during the formation of the mRNA (blue), and a continuous coding sequence is therefore present in each clone. Because gene B is transcribed more frequently than gene A in the cells from which the cDNA library was made, it is represented much more frequently than A in the cDNA library. In contrast, A and B are represented equally in the genomic DNA library. Hybridization Provides a Powerful, But Simple Way to Detect Specific Nucleotide Sequences


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Cell_Biology_Alberts. Hybridization Provides a Powerful, But Simple Way to Detect Specific Nucleotide Sequences Under normal conditions, the two strands of a DNA double helix are held together by hydrogen bonds between the complementary base pairs (see Figure 4–3). But these relatively weak, noncovalent bonds can be fairly easily broken. Such DNA denaturation will release the two strands from each other, but does not break the covalent bonds that link together the nucleotides within each strand. Perhaps the simplest way to achieve this separation involves heating the DNA to around 90°C. When the conditions are reversed—by slowly lowering the temperature— the complementary strands will readily come back together to re-form a double helix. This hybridization, or DNA renaturation, is driven by the re-formation of the hydrogen bonds between complementary base pairs (Figure 8–33). We saw in Chapter 5 that DNA hybridization underlies the crucial process of homologous recombination (see Figure 5–47).


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Cell_Biology_Alberts. This fundamental capacity of a single-stranded nucleic acid molecule, either DNA or RNA, to form a double helix with a single-stranded molecule of a complementary sequence provides a powerful and sensitive technique for detecting specific nucleotide sequences. Today, one simply designs a short, single-stranded DNA molecule (often called a DNA probe) that is complementary to the nucleotide sequence of interest. Because the nucleotide sequences of so many genomes are known—and are stored in publicly accessible databases—designing a probe to hybridize anywhere in a genome is straightforward. Probes are single-stranded, typically 30 nucleotides in length, and are usually synthesized chemically by a commercial service for pennies per nucleotide. A DNA sequence of 30 nucleotides will occur by chance only once every 1 × 1018 nucleotides (430); so, even in the human genome of 3 × 109 nucleotide pairs, a DNA probe designed to match a particular 30-nucleotide sequence will be highly unlikely to


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Cell_Biology_Alberts. only once every 1 × 1018 nucleotides (430); so, even in the human genome of 3 × 109 nucleotide pairs, a DNA probe designed to match a particular 30-nucleotide sequence will be highly unlikely to hybridize—by chance— anywhere else on the genome. This, of course, presumes that the sequence complementary to the probe does not occur multiple times in the genome, a condition that can be checked beforehand by scanning the genomic sequence in silico (using a computer) and designing probes that match only one spot. The hybridization conditions can be set so that even a single mismatch will prevent hybridization to “near-miss” sequences. The exquisite specificity of nucleic acid hybridization can be easily appreciated by the in situ (Latin for “in place”) hybridization experiment shown in Figure 8–34. As we will see throughout this chapter, nucleic acid


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Cell_Biology_Alberts. Figure 8–34 In situ hybridization can be used to locate genes on isolated chromosomes. Here, six different DNA probes have been used to mark the locations of their complementary nucleotide sequences on human Chromosome 5, isolated from a mitotic cell in metaphase (see Figure 4–59 and Panel 17–1, pp. 980–981). The DNA probes have been labeled with different chemical groups (see Figure 8–26B) and are detected using fluorescent antibodies specific for those groups. The chromosomal DNA has been partially denatured to allow the probes to base-pair with their complementary sequences. Both the maternal and paternal copies of Chromosome 5 are shown, aligned side by side. Each probe produces two dots on each chromosome because chromosomes undergoing mitosis have already replicated their DNA; therefore, each chromosome contains two identical DNA helices. The technique employed here is nicknamed FISH, for fluorescence in situ hybridization. (Courtesy of David C. Ward.)


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Cell_Biology_Alberts. Figure 8–33 A molecule of DNA can undergo denaturation and renaturation (hybridization). For two single-stranded molecules to hybridize, they must have complementary nucleotide sequences that allow base-pairing. In this example, the red and orange strands are complementary to each other, and the blue and green strands are complementary to each other. Although denaturation by heating is shown, DNA can also be renatured after being denatured by alkali treatment. hybridization has many uses in modern cell and molecular biology; one of the most powerful is in the cloning of DNA by the polymerase chain reaction, as we next discuss.


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Cell_Biology_Alberts. hybridization has many uses in modern cell and molecular biology; one of the most powerful is in the cloning of DNA by the polymerase chain reaction, as we next discuss. Genomic and cDNA libraries were once the only route to cloning genes and they are still used for cloning very large genes and for sequencing whole genomes. However, a powerful and versatile method for amplifying DNA, known as the polymerase chain reaction (PCR), provides a more rapid and straightforward approach to DNA cloning, particularly in organisms whose complete genome sequence is known. Today, since genome sequences are abundant, most cloning is carried out by PCR.


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Cell_Biology_Alberts. Invented in the 1980s, PCR revolutionized the way that DNA and RNA are analyzed. The technique can amplify any nucleotide sequence selectively and is performed entirely in a test tube. Eliminating the need for bacteria makes PCR convenient and rapid—billions of copies of a nucleotide can be generated in a matter of hours. Starting with an entire genome, PCR allows DNA from a specified region—selected by the experimenter—to be greatly amplified, effectively “purifying” this DNA away from the remainder of the genome, which remains unamplified. Because of its power to greatly amplify nucleic acids, PCR is remarkably sensitive: the method can be used to detect the trace amounts of DNA in a drop of blood left at a crime scene or in a few copies of a viral genome in a patient’s blood sample.


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Cell_Biology_Alberts. The success of PCR depends both on the selectivity of DNA hybridization and on the ability of DNA polymerase to copy a DNA template faithfully through repeated rounds of replication in vitro. As discussed in Chapter 5, this enzyme adds nucleotides to the 3ʹ end of a growing strand of DNA (see Figure 5–4). To copy DNA, the polymerase requires a primer—a short nucleotide sequence that provides a 3ʹ end from which synthesis can begin. For PCR, the primers are designed by the experimenter, synthesized chemically, and, by hybridizing to genomic DNA, “tell” the polymerase which part of the genome to copy. As discussed in the previous section, DNA primers (in essence, the same type of molecules as DNA probes but without a radioactive or fluorescent label) can be designed to uniquely locate any position on a genome.


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Cell_Biology_Alberts. PCR is an iterative process in which the cycle of amplification is repeated dozens of times. At the start of each cycle, the two strands of the double-stranded DNA template are separated and a different primer is annealed to each. These primers mark the right and left boundaries of the DNA to be amplified. DNA polymerase is then allowed to replicate each strand independently (Figure 8–35). In


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Cell_Biology_Alberts. Figure 8–35 A pair of primers directs the synthesis of a desired segment of DNA in a test tube. Each cycle of PCR includes three steps: (1) The double-stranded DNA is heated briefly to separate the two strands. (2) The DNA is exposed to a large excess of a pair of specific primers— designed to bracket the region of DNA to be amplified—and the sample is cooled to allow the primers to hybridize to complementary sequences in the two DNA strands. (3) This mixture is incubated with DNA polymerase and the four deoxyribonucleoside triphosphates so that DNA can be synthesized, starting from the two primers. To amplify the DNA, the cycle is repeated many times by reheating the sample to separate the newly synthesized DNA strands (see Figure 8–36).


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Cell_Biology_Alberts. The technique depends on the use of a special DNA polymerase isolated from a thermophilic bacterium; this polymerase is stable at much higher temperatures than eukaryotic DNA polymerases, so it is not denatured by the heat treatment shown in step 1. The enzyme therefore does not have to be added again after each cycle. products of frst cycle


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Cell_Biology_Alberts. Figure 8–36 PCR uses repeated rounds of strand separation, hybridization, and synthesis to amplify DNA. As the procedure outlined in Figure 8–35 is repeated, all the newly synthesized fragments serve as templates in their turn. Because the polymerase and the primers remain in the sample after the first cycle, PCR involves simply heating and then cooling the same sample, in the same test tube, again and again. Each cycle doubles the amount of DNA synthesized in the previous cycle, so that within a few cycles, the predominant DNA is identical to the sequence bracketed by and including the two primers in the original template. In the example illustrated here, three cycles of reaction produce 16 DNA chains, 8 of which (boxed in yellow) correspond exactly to one or the other strand of the original bracketed sequence. After four more cycles, 240 of the 256 DNA chains will correspond exactly to the original sequence, and after several more cycles, essentially all of the DNA strands will be


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Cell_Biology_Alberts. bracketed sequence. After four more cycles, 240 of the 256 DNA chains will correspond exactly to the original sequence, and after several more cycles, essentially all of the DNA strands will be this length. Typically, 20–30 cycles are carried out to effectively clone a region of DNA starting from genomic DNA; the rest of the genome remains unamplified, and its concentration is therefore negligible compared with that of the amplified region (Movie 8.2).


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Cell_Biology_Alberts. subsequent cycles, all the newly synthesized DNA molecules produced by the polymerase serve as templates for the next round of replication (Figure 8–36). Through this iterative amplification process, many copies of the original sequence can be made—billions after about 20 to 30 cycles. PCR is now the method of choice for cloning relatively short DNA fragments (say, under 10,000 nucleotide pairs). Each cycle takes only about five minutes, and automation of the whole procedure enables cell-free cloning of a DNA fragment in a few hours. The original template for PCR can be either DNA or RNA, so this method can be used to obtain either a genomic clone (complete with introns and exons) or a cDNA copy of an mRNA (Figure 8–37).


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Cell_Biology_Alberts. The PCR method is extraordinarily sensitive; it can detect a single DNA molecule in a sample if at least part of the sequence of that molecule is known. Trace amounts of RNA can be analyzed in the same way by first transcribing them into DNA with reverse transcriptase. For these reasons, PCR is frequently employed for uses that go beyond simple cloning. For example, it can be used to detect invading pathogens at very early stages of infection. In this case, short sequences complementary to a segment of the infectious agent’s genome are used as primers and following many cycles of amplification, even a few copies of an invading bacterial or viral genome in a patient’s sample can be detected (Figure 8–38). For many infections, PCR has replaced the use of antibodies against microbial molecules to isolate total DNA isolate total mRNA DNA segment to be cloned mRNA sequence to be cloned ADD FIRST PRIMER, REVERSE TRANSCRIPTASE, AND DEOXYRIBONUCLEOSIDE TRIPHOSPHATES SEPARATE STRANDS AND ADD


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Cell_Biology_Alberts. to isolate total DNA isolate total mRNA DNA segment to be cloned mRNA sequence to be cloned ADD FIRST PRIMER, REVERSE TRANSCRIPTASE, AND DEOXYRIBONUCLEOSIDE TRIPHOSPHATES SEPARATE STRANDS AND ADD SECOND PRIMER PCR AMPLIFICATION WITH BOTH PRIMERS PRESENTPCR AMPLIFICATION mRNA DNA SEPARATE STRANDS AND ADD PRIMERS genomic clones (A) cDNA clones (B) chromosomal DNA detect the presence of the invader. It is also used to verify the authenticity of a food source—for example, whether a sample of beef actually came from a cow.


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Cell_Biology_Alberts. Finally, PCR is now widely used in forensics. The method’s extreme sensitivity allows forensic investigators to isolate DNA from minute traces of human blood or other tissue to obtain a DNA fingerprint of the person who left the sample behind. rare HIV particle control, usingin plasma of blood frominfected person Figure 8–38 PCR can be used to detect the presence of a viral genome in a sample of blood. Because of its ability to amplify enormously the signal from a single molecule of nucleic acid, PCR is an extraordinarily sensitive method for detecting trace amounts of virus in a sample of blood or tissue, without the need to purify the virus. For HIV, the virus that causes AIDS, the genome is a single-stranded molecule of RNA, as illustrated here. In addition to HIV, many other viruses that infect humans are now detected in this way. Figure 8–37 PCR can be used to obtain either genomic or cDNA clones.


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Cell_Biology_Alberts. Figure 8–37 PCR can be used to obtain either genomic or cDNA clones. (A) To use PCR to clone a segment of chromosomal DNA, total genomic DNA is first purified from cells. PCR primers that flank the stretch of DNA to be cloned are added, and many cycles of PCR are completed (see Figure 8–36). Because only the DNA between (and including) the primers is amplified, PCR provides a way to obtain selectively any short stretch of chromosomal DNA in an effectively pure form. (B) To use PCR to obtain a cDNA clone of a gene, total mRNA is first purified from cells. The first primer is added to the population of mRNAs, and reverse transcriptase is used to make a DNA strand complementary to the specific RNA sequence of interest. The second primer is then added, and the DNA molecule is amplified through many cycles of PCR.


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Cell_Biology_Alberts. Figure 8–39 PCR is used in forensic science to distinguish one individual from another. The DNA sequences analyzed are short tandem repeats (STRs) composed of sequences such as CACACA… or GTGTGT… STRs are found in various positions (loci) in the human genome. The number of repeats in each STR locus is highly variable in the population, ranging from 4 to 40 in different individuals. Because of the variability in these sequences, individuals will usually inherit a different number of repeats at each STR locus from their mother and from their father; two unrelated individuals, therefore, rarely contain the same pair of sequences at a given STR locus. (A) PCR using primers that recognize unique sequences on either side of one particular STR locus produces a pair of bands of amplified DNA from each individual, one band representing the maternal STR variant and the other representing the paternal STR variant. The length of the amplified DNA, and thus its position after gel electrophoresis,


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Cell_Biology_Alberts. each individual, one band representing the maternal STR variant and the other representing the paternal STR variant. The length of the amplified DNA, and thus its position after gel electrophoresis, will depend on the exact number of repeats at the locus. (B) In the schematic example shown here, the same three STR loci are analyzed in samples from three suspects (individuals A, B, and C), producing six bands for each individual. Although different people can have several bands in common, the overall pattern is quite distinctive for each person. The band pattern can therefore serve as a DNA fingerprint to identify an individual nearly uniquely. The fourth lane (F) contains the products of the same PCR amplifications carried out on a hypothetical forensic DNA sample, which could have been obtained from a single hair or a tiny spot of blood left at a crime scene.


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Cell_Biology_Alberts. The more loci that are examined, the more confident one can be about the results. When examining the variability at 5–10 different STR loci, the odds that two random individuals would share the same fingerprint by chance are approximately one in 10 billion. In the case shown here, individuals A and C can be eliminated from inquiries, while B is a clear suspect. A similar approach is used routinely in paternity testing. With the possible exception of identical twins, the genome of each human differs in DNA sequence from that of every other person on Earth. Using primer pairs targeted at genome sequences that are known to be highly variable in the human population, PCR makes it possible to generate a distinctive DNA fingerprint for any individual (Figure 8–39). Such forensic analyses can be used not only to help identify those who have done wrong, but also—equally important—to exonerate those who have been wrongfully accused.


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Cell_Biology_Alberts. Most current methods of manipulating DNA, RNA, and proteins rely on prior knowledge of the nucleotide sequence of the genome of interest. But how were these sequences determined in the first place? And how are new DNA and RNA molecules sequenced today? In the late 1970s, researchers developed several strategies for determining, simply and quickly, the nucleotide sequence of any purified DNA fragment. The one that became the most widely used is called dideoxy sequencing or Sanger sequencing (Panel 8–1). This method was used to determine the nucleotide sequence of many genomes, including those of E. coli, fruit flies, nematode worms, mice, and humans. Today, cheaper and faster methods are routinely used to sequence DNA, and even more efficient strategies are being developed (see Panel 8–1). The original “reference” sequence of the human genome, completed in 2003, cost over $1 billion and required many scientists from around the world working together for 13 years. The enormous progress


Cell_Biology_Alberts

Cell_Biology_Alberts. The original “reference” sequence of the human genome, completed in 2003, cost over $1 billion and required many scientists from around the world working together for 13 years. The enormous progress made in the past decade makes it possible for a single person to complete the sequence of an individual human genome in less than a day.


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Cell_Biology_Alberts. The methods summarized in Panel 8–1 for rapidly sequencing DNA can also be applied to RNA. Although methods are being developed to sequence RNA directly, it is most commonly carried out by converting the RNA to complementary DNA (using reverse transcriptase) and using one of the methods described for DNA sequencing. It is important to keep in mind that although genomes remain the same from cell to cell and from tissue to tissue, the RNA produced from the genome can vary enormously. We will see later in this chapter that sequencing the entire repertoire of RNA from a cell or tissue (known as deep RNA sequencing, or RNA-seq) is a powerful way to understand how the information present in the genome is used by different cells under different circumstances. In the next section, we shall see how RNA-seq has also become a valuable tool for annotating genomes. To Be Useful, Genome Sequences Must Be Annotated


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Cell_Biology_Alberts. To Be Useful, Genome Sequences Must Be Annotated Long strings of nucleotides, at first glance, reveal nothing about how this genetic information directs the development of a living organism—or even what types of DNA, protein, and RNA molecules are produced by a genome. The process of genome annotation attempts to mark out all the genes (both protein-coding and noncoding) in a genome and ascribe a role to each. It also seeks to understand more subtle types of genome information, such as the cis-regulatory sequences that specify the time and place that a given gene is expressed and whether its mRNA undergoes alternative splicing to produce different protein isotypes. Clearly, this is a daunting task, and we are far short of completing it for any form of life, even the simplest bacterium. For many organisms, we know the approximate number of genes, and, for very simple organisms, we understand the functions of about half their genes.


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Cell_Biology_Alberts. In this section, we discuss broadly how genes are identified in genome sequences and what clues we can discern about their roles from simply inspecting their sequences. Later in the chapter, we turn to the more difficult problem of experimentally determining gene function. How does one begin to make sense of a genome sequence? The first step is usually to translate in silico the entire genome into protein. There are six different reading frames for any piece of double-stranded DNA (three on each strand). We saw in Chapter 6 that a random sequence of nucleotides, read in frame, will PANeL 8–1: DNA Sequencing Methods


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Cell_Biology_Alberts. OOCH25˜OH3˜3˜3˜OH allows strand extension at 3˜ end normal deoxyribonucleoside triphosphate (dNTP) OOCH25˜3˜ H prevents strand extension at 3˜end chain-terminating dideoxyribonucleoside triphosphate (ddNTP) basebase PPP PPP ATGTCAGTCCAG 3˜RESULT TACAGTCAGGTC 5˜GCAT ATG GCAT ATGTCAG GCAT ATGTCAGTCCAG G sequence of DNA primer sequence read from gel 3˜ CGTATACAGTCAGGTC 5˜3˜ CGTATACAGTCAGGTC 5˜5˜ GCAT 3˜GCAT A GCAT ATGTCA GCAT ATGTCAGTCCA GCAT AT GCAT ATGT GCAT ATGTCAGT GCAT ATGTC GCAT ATGTCAGTC GCAT ATGTCAGTCC A T C G A C C T G A C T G T A A T C G 3˜5˜ADD LABELED DNA PRIMER single-stranded DNA fragment to be sequenced T T T TTT T T T T A A A A AA AA GG GGG GG CC C C C CC C CADD DNA POLYMERASE AND DIVIDE INTO 4 SEPARATE TUBES ADD EXCESS AMOUNTS OF NORMAL dNTPs sequence of original DNA strand ADD SMALL AMOUNT OF ONE CHAIN-TERMINATING ddNTP TO EACH TUBE TT C T A T AG T G T C A CC T AAATA G C TT GG C G T AATC A T GG T(B) (A) GCATATG GCATA GCATATGT GCATAT GCATATGTC mixture of DNA


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Cell_Biology_Alberts. DNA strand ADD SMALL AMOUNT OF ONE CHAIN-TERMINATING ddNTP TO EACH TUBE TT C T A T AG T G T C A CC T AAATA G C TT GG C G T AATC A T GG T(B) (A) GCATATG GCATA GCATATGT GCATAT GCATATGTC mixture of DNA products, each containing a chain-terminating ddNTP labeled with a different fuorescent marker PRODUCTS LOADED ONTO CAPILLARY GEL electrophoresis size-separated products are read in sequence Dideoxy sequencing, or Sanger sequencing (named after the scientist who invented it), uses DNA polymerase, along with special chain-terminating nucleotides called dideoxyribonucleoside triphosphates (left), to make partial copies of the DNA fragment to be sequenced. These ddNTPs are derivatives of the normal deoxyribonucleoside triphosphates that lack the 3˜ hydroxyl group. When incorporated into a growing DNA strand, they block further elongation of that strand. DNA SEQUENCING MANUAL DIDEOXY SEQUENCING AUTOMATED DIDEOXY SEQUENCING To determine the complete sequence of a single-stranded fragment of DNA


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Cell_Biology_Alberts. DNA strand, they block further elongation of that strand. DNA SEQUENCING MANUAL DIDEOXY SEQUENCING AUTOMATED DIDEOXY SEQUENCING To determine the complete sequence of a single-stranded fragment of DNA (gray), the DNA is frst hybridized with a short DNA primer (orange) that is labeled with a fuorescent dye or radioisotope. DNA polymerase and an excess of all four normal deoxyribonucleoside triphosphates (blue A, C, G, or T) are added to the primed DNA, which is then divided into four reaction tubes. Each of these tubes receives a small amount of a single chain-terminating dideoxyribonucleoside triphosphate (red A, C, G, or T). Because these will be incorporated only occasionally, each reaction produces a set of DNA copies that terminate at different points in the sequence. The products of these four reactions are separated by electrophoresis in four parallel lanes of a polyacrylamide gel (labeled here A, T, C, and G). In each lane, the bands represent fragments that have terminated at a


Cell_Biology_Alberts

Cell_Biology_Alberts. four reactions are separated by electrophoresis in four parallel lanes of a polyacrylamide gel (labeled here A, T, C, and G). In each lane, the bands represent fragments that have terminated at a given nucleotide but at different positions in the DNA. By reading off the bands in order, starting at the bottom of the gel and reading across all lanes, the DNA sequence of the newly synthesized strand can be determined (see Figure 8–25C). The sequence, which is given in the green arrow to the right of the gel, is complementary to the sequence of the original gray single-stranded DNA. Fully automated machines can run dideoxy sequencing reactions. (A) The automated method uses an excess amount of normal dNTPs plus a mixture of four different chain-terminating ddNTPs, each of which is labeled with a fuorescent tag of a different color. The reaction products are loaded onto a long, thin capillary gel and separated by electrophoresis. A camera (not shown) reads the color of each band as it


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Cell_Biology_Alberts. with a fuorescent tag of a different color. The reaction products are loaded onto a long, thin capillary gel and separated by electrophoresis. A camera (not shown) reads the color of each band as it moves through the gel and feeds the data to a computer that assembles the sequence. (B) A tiny part of the data from such an automated sequencing run. Each colored peak represents a nucleotide in the DNA sequence. 5˜ GCAT 3˜ CGTA 479SEQUENCING WHOLE GENOMES Shotgun sequencing: To determine the nucleotide sequence of a whole genome, the genomic DNA is frst fragmented into small pieces and a genomic library is constructed, typically using plasmids and bacteria (see Figure 8–30). In shotgun sequencing, the nucleotide sequence of tens of thousands of individual clones is determined; the full genome sequence is then reconstructed by stitching together (in silico) the nucleotide sequence of each clone, using the overlaps between clones as a guide. The shotgun method works well for small


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Cell_Biology_Alberts. genome sequence is then reconstructed by stitching together (in silico) the nucleotide sequence of each clone, using the overlaps between clones as a guide. The shotgun method works well for small genomes (such as those of viruses and bacteria) that lack repetitive DNA. SECOND-GENERATION SEQUENCING TECHNOLOGIES The dideoxy method made it possible to sequence the genomes of humans and most of the other organisms discussed in this book. But newer methods, developed since 2005, have made genome sequencing even more rapid—and very much cheaper. With these so-called second-generation sequencing methods, the cost of sequencing DNA has decreased dramatically. Not surprisingly, the number of genomes that have been sequenced has increased enormously. These rapid methods allow multiple genomes to be sequenced in parallel in a matter of weeks, enabling investigators to examine thousands of individual human genomes, catalog the variation in nucleotide sequences from people around the world, and


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Cell_Biology_Alberts. be sequenced in parallel in a matter of weeks, enabling investigators to examine thousands of individual human genomes, catalog the variation in nucleotide sequences from people around the world, and uncover the mutations that increase the risk of various diseases, from cancer to autism. These methods have also made it possible to determine the genome sequence of extinct species, including Neanderthal man and the wooly mammoth. By sequencing genomes from many closely related species, they have also made it possible to understand the molecular basis of key evolutionary events in the tree of life, such as the “inventions” of multicellularity, vision, and language. The ability to rapidly sequence DNA has had major impacts on all branches of biology and medicine; it is almost impossible to imagine where we would be without it. BAC clones: Most plant and animal genomes are large (often over 109 nucleotide pairs) and contain extensive amounts of repetitive DNA spread throughout the genome.


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Cell_Biology_Alberts. where we would be without it. BAC clones: Most plant and animal genomes are large (often over 109 nucleotide pairs) and contain extensive amounts of repetitive DNA spread throughout the genome. Because a nucleotide sequence of a fragment of repetitive DNA will “overlap” every instance of the repeated DNA, it is diffcult, if not impossible, to assemble the fragments into a unique order solely by the shotgun method. To circumvent this problem, the human genome was frst broken down into very large DNA fragments (each approximately 100,000 nucleotide pairs) and cloned into BACs (see p. 469). The order of the BACs along a chromosome was determined by comparing the pattern of restriction enzyme cleavage sites in a given BAC clone with that of the whole genome. In this way, a given BAC clone can be mapped, say, to the left arm of human Chromosome 3. Once a collection of BAC clones was Thousands of genomes from individual humans have now been sequenced and it is not necessary to painstakingly


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Cell_Biology_Alberts. be mapped, say, to the left arm of human Chromosome 3. Once a collection of BAC clones was Thousands of genomes from individual humans have now been sequenced and it is not necessary to painstakingly reconstruct the order of DNA sequence “reads” each time; they are simply assembled using the order determined from the original human genome sequencing project. For this reason, resequencing, the term applied when the genome of a species is sequenced again (even though it may be from a different individual), is far easier than the original sequencing. sequence one strand of fragments GTTCAGCATTG------GCCATTAGTTCA---GCCATTAGTTCAGCATTG---original sequence reconstructed based on sequence overlap sequences of two fragments AA D B BA B C E C restriction pattern for individual BAC clones restriction map of one segment of human genome cleavage sites for restriction nucleases A, B, C, D, and E obtained that spanned the entire genome, each individual BAC was sequenced by the shotgun method. At the