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Cell_Biology_Alberts. 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 end, the sequences of all the BAC inserts were stitched together using the knowledge of the position of each BAC insert in the human genome. In all, approximately 30,000 BAC clones were sequenced to complete the human genome. random fragmentation multiple copies of genome 479


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Cell_Biology_Alberts. ILLUMINA® SEQUENCING Several second-generation sequencing methods are now in wide use, and we will discuss two of the most common. Both rely on the construction of libraries of DNA fragments that represent—in toto—the DNA of the genome. Instead of using bacterial cells to generate these libraries, as we saw in Figure 8–30), they are made using PCR amplifcation of billions of DNA fragments, each attached to a solid support. The amplifcation is carried out so that the PCR-generated copies, instead of foating away in solution, remain bound in proximity to the original DNA fragment. This process generates clusters of DNA fragments, where each cluster contains about 1000 identical copies of a small bit of the genome. These clusters—a billion of which can ft in a single slide or plate—are then sequenced at the same time; that is, in parallel. One method, known as Illumina sequencing, is based on the dideoxy method described above, but it incorporates several innovations. Here, each


Cell_Biology_Alberts

Cell_Biology_Alberts. then sequenced at the same time; that is, in parallel. One method, known as Illumina sequencing, is based on the dideoxy method described above, but it incorporates several innovations. Here, each nucleotide is attached to a removable fuorescent molecule (a different color for each of the four bases) as well as a special chain-terminating chemical adduct: instead of a 3’-OH group, as in conventional dideoxy sequencing, the nucleotides carry a chemical group that blocks elongation by DNA polymerase but which can be removed chemically. Sequencing is then carried out as follows: the four fuorescently labeled nucleotides along with DNA polymerase are added to billions of DNA clusters immobilized on a slide. Only the appropriate nucleotide (that is complementary to the next nucleotide in the template) is covalently incorporated at each cluster; the unincorporated nucleotides are washed away, and a high-resolution digital camera takes an image that registers which of the four nucleotides


Cell_Biology_Alberts

Cell_Biology_Alberts. template) is covalently incorporated at each cluster; the unincorporated nucleotides are washed away, and a high-resolution digital camera takes an image that registers which of the four nucleotides was added to the chain at each cluster. The fuorescent label and the 3˜-OH blocking group are then removed enzymatically, washed away, and the process is repeated many times. In this way, billions of sequencing reactions are carried out simultaneously. By keeping track of the color changes occuring at each cluster, the DNA sequence represented by each spot can be read. Although each individual sequence read is relatively short (approximately 200 nucleotides), the billions that are carried out simultaneously can produce several human genomes worth of sequence in about a day. Principle behind Illumina sequencing. This reaction is carried out stepwise, on billions of DNA clusters at once. The method relies on a color digital camera that rapidly scans all the DNA clusters after each round of


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Cell_Biology_Alberts. sequencing. This reaction is carried out stepwise, on billions of DNA clusters at once. The method relies on a color digital camera that rapidly scans all the DNA clusters after each round of modifed nucleotide incorporation. The DNA sequence of each cluster is then determined by the sequence of color changes it undergoes as the elongation reaction proceeds stepwise. Each round of modifed nucleotide incorporation, image acquisition, and removal of the 3˜block and the fuorescent group takes less than an hour. Each cluster on the slide contains many copies of different, random bits of a genome; in preparing the clusters, a DNA sequence (specifed by the experimenter) is joined to each copy in every cluster, and a primer complementary to this sequence is used to begin the elongation reaction by DNA polymerase. 100 µm A slide showing individual clusters of PCR-generated DNA molecules. Each cluster carries about 1000 identical DNA molecules; the four colors are produced by incorporation of


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Cell_Biology_Alberts. by DNA polymerase. 100 µm A slide showing individual clusters of PCR-generated DNA molecules. Each cluster carries about 1000 identical DNA molecules; the four colors are produced by incorporation of C, G, A, or T, each of which has a different color fuorophore. The image has been taken just after a fuorescent nucleotide has been incorporated into each growing DNA chain. (From Illumina Sequencing Overview, 2013.) PPP PP fuor 3˜ block 5˜3˜ block incorporation into growing chain by DNA polymerase P 5˜free 3˜ end P DNA template OH fuor removed block removed next cycle no fuor photo taken 480


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Cell_Biology_Alberts. ION TORRENT™ SEQUENCING THE FUTURE OF DNA SEQUENCING Another widely used strategy for rapid DNA sequencing is called the ion torrent method. Here, a genome is fragmented, and the individual fragments are attached to microscopic beads. Using PCR, each DNA fragment is then amplifed so that copies of it eventually coat the bead to which it was initially attached. This process produces a library of billions of individual beads, each covered with identical copies of a particular DNA fragment. Like eggs in a carton, the beads are placed into individual wells on an array that can hold a billion beads in a square inch. Beginning with a primer, DNA synthesis is then initiated on each bead. A hydrogen ion (H+) is released (along with pyrophosphate) each time a nucleotide is incorporated into a growing DNA chain (see Figure 5–3), and the ion torrent method is based on this simple fact. Each of the four nucleotides is washed in, one at a time, over the array of beads; when a nucleotide is


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Cell_Biology_Alberts. a growing DNA chain (see Figure 5–3), and the ion torrent method is based on this simple fact. Each of the four nucleotides is washed in, one at a time, over the array of beads; when a nucleotide is incorporated in the DNA of a given bead, the release of an H+ ion changes the pH, which is registered by a semiconductor chip placed beneath the array of wells. In this way, the DNA sequence on a given bead can be read from the pattern of pH changes observed as nucleotides are washed over them. Like a high-resolution sensor in a digital camera, the ion torrent semiconductor chip can register enormous amounts of information and can thus keep track of billions of parallel sequencing reactions. Using this technology it is currently possible, using a single chip, to determine the nucleotide sequences of several human genomes in just a few hours. PPP PP 5˜DNA template A OH OH H+ pH change detected volts 1 µm voltage-sensing chip well bead coated with DNA template A DNA sequencing by the ion


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Cell_Biology_Alberts. of several human genomes in just a few hours. PPP PP 5˜DNA template A OH OH H+ pH change detected volts 1 µm voltage-sensing chip well bead coated with DNA template A DNA sequencing by the ion torrent method. Beads, each coated with a DNA molecule that has been amplifed many times, are placed in wells along with primers and DNA polymerase. As nucleotides are sequentially washed over the beads, those incorporated by the polymerase cause a pH change. In the example shown, an A is incorporated; thus, the template must have a T in this position. As the four nucleotides are sequentially washed over the beads, the sequence of the DNA on each bead can be “read” by the pattern of pH fuctuations. Billions of beads are monitored at once by a voltage-sensitive semiconductor chip placed below the array of beads. Even newer, potentially faster, methods of sequencing DNA are being developed. Some of these “third-generation” technologies bypass the DNA amplifcation steps altogether and determine the


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Cell_Biology_Alberts. of beads. Even newer, potentially faster, methods of sequencing DNA are being developed. Some of these “third-generation” technologies bypass the DNA amplifcation steps altogether and determine the sequence of single molecules of DNA. In one technique, a DNA molecule is pushed through a tiny channel, like a thread through the eye of a needle. As the DNA molecule moves through the pore, it generates electrical currents that depend on its sequence of nucleotides; the pattern of currents can then be used to deduce the nucleotide sequence. Other methods visualize single DNA molecules using electron or atomic force microscopy; the nucleotide sequence is read from the small differences in the “appearance” of the DNA as it is scanned. Finally, another method is based on immobilizing a single DNA polymerase molecule (with a template) and measuring the “dwell” time of each of the four nucleotides, which are labeled with different removable fuorescent dyes. Nucleotides that reside longer on the


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Cell_Biology_Alberts. polymerase molecule (with a template) and measuring the “dwell” time of each of the four nucleotides, which are labeled with different removable fuorescent dyes. Nucleotides that reside longer on the polymerase (before their dye is removed) are those incorporated by the polymerase. Although the two methods we have described in detail (Illumina and ion torrent) are now used extensively, it is likely that faster and cheaper methods will continue to be developed. 2001 2003 2005 2007 2009 2011 2013 2015 100,000,000 10,000,000 1,000,000 100,000 10,000 1000 100 cost in dollars years Shown here are the costs of sequencing a human genome, which was $100 million in 2001 and about a thousand dollars by the end of 2014. (Data from the National Human Genome Research Initiative.) 481 (A) reading direction for sequence of top DNA strand frames –3 reading direction for sequence of bottom DNA strand 500 base pairs contain a stop codon about every 20 amino acids; protein-coding regions will, in


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Cell_Biology_Alberts. direction for sequence of top DNA strand frames –3 reading direction for sequence of bottom DNA strand 500 base pairs contain a stop codon about every 20 amino acids; protein-coding regions will, in contrast, usually contain much longer stretches without stop codons (Figure 8–40). Known as open reading frames (ORFs), these usually signify bona fide protein-coding genes. This assignment is often “double-checked” by comparing the ORF amino acid sequence to the many databases of documented proteins from other species. If a match is found, even as imperfect one, it is very likely that the ORF will code for a functional protein (see Figure 8–23).


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Cell_Biology_Alberts. This strategy works very well for compact genomes, where intron sequences are rare and ORFs often extend for many hundreds of amino acids. However, in many animals and plants, the average exon size is 150–200 nucleotide pairs (see Figure 6–31) and additional information is usually required to unambiguously locate all the exons of a gene. Although it is possible to search genomes for splicing signals and other features that help to identify exons (codon bias, for example), one of the most powerful methods is simply to sequence the total RNA produced from the genome in living cells. As can be seen in Figure 7–3, this RNA-seq information, when mapped onto the genome sequence, can be used to accurately locate all the introns and exons of even complex genes. By sequencing total RNA from different cell types, it is also possible to identify cases of alternative splicing (see Figure 6–26).


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Cell_Biology_Alberts. RNA-seq also identifies noncoding RNAs produced by a genome. Although the function of some of them can be readily recognized (tRNAs or snoRNAs, for example), many have unknown functions and still others probably have no function at all (discussed in Chapter 7, pp. 429–436). The existence of the many noncoding RNAs and our relative ignorance of their function is the main reason that we know only the approximate number of genes in the human genome.


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Cell_Biology_Alberts. But even for protein-coding genes that have been unambiguously identified, we still have much to learn. Thousands of genomes have been sequenced, and we know from comparative genomics that many organisms share the same basic set of proteins. However, the functions of a very large number of identified proteins remain unknown. Depending on the organism, approximately one-third of the proteins encoded by a sequenced genome do not clearly resemble any protein that has been studied biochemically. This observation underscores a limitation of the emerging field of genomics: although comparative analysis of genomes reveals a great deal of information about the relationships between genes and organisms, it often does not provide immediate information about how these genes function, or what roles they have in the physiology of an organism. Comparison of the full Figure 8–40 Finding the regions in a DNA sequence that encode a protein.


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Cell_Biology_Alberts. (A) Any region of the DNA sequence can, in principle, code for six different amino acid sequences, because any one of three different reading frames can be used to interpret the nucleotide sequence on each strand. Note that a nucleotide sequence is always read in the 5ʹ-to-3ʹ direction and encodes a polypeptide from the N-terminus to the C-terminus. For a random nucleotide sequence read in a particular frame, a stop signal for protein synthesis is encountered, on average, about once every 20 amino acids. In this sample sequence of 48 base pairs, each such signal (stop codon) is colored blue, and only reading frame 2 lacks a stop signal. (B) Search of a 1700-base-pair DNA sequence for a possible protein-encoding sequence. The information is displayed as in (A), with each stop signal for protein synthesis denoted by a blue line. In addition, all of the regions between possible start and stop signals for protein synthesis (see pp. 347–349) are displayed as red bars. Only reading frame 1


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Cell_Biology_Alberts. synthesis denoted by a blue line. In addition, all of the regions between possible start and stop signals for protein synthesis (see pp. 347–349) are displayed as red bars. Only reading frame 1 actually encodes a protein, which is 475 amino acid residues long.


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Cell_Biology_Alberts. gene complement of several thermophilic bacteria, for example, does not reveal why these bacteria thrive at temperatures exceeding 70°C. And examination of the genome of the incredibly radioresistant bacterium Deinococcus radiodurans does not explain how this organism can survive a blast of radiation that can shatter glass. Further biochemical and genetic studies, like those described in the other sections of this chapter, are required to determine how genes, and the proteins they produce, function in the context of living organisms. DNA Cloning Allows Any Protein to be Produced in Large Amounts


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Cell_Biology_Alberts. In the last section, we saw how protein-coding genes can be identified in genome sequences. Using the genetic code (and provided the intron and exon boundaries are known), the amino acid sequence of any protein coded in a genome can be deduced. As was discussed earlier, this sequence can often provide an important clue to the protein’s function if found to be similar to the amino acid sequence of a protein that has already been studied (see Figure 8–23). Although this strategy is often successful, it typically provides only the likely biochemical function of the protein; for example, whether the protein resembles a kinase or a protease. It usually remains for the experimenter to verify (or refute) this assignment and, most importantly, to discover the protein’s biological function in the whole organism; that is, to what attributes of the organism does the kinase or the protease contribute and in what molecular pathways does it function? Nowadays, most new proteins are “discovered”


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Cell_Biology_Alberts. whole organism; that is, to what attributes of the organism does the kinase or the protease contribute and in what molecular pathways does it function? Nowadays, most new proteins are “discovered” through genome sequencing, and it often remains a great challenge to ascertain their functions.


Cell_Biology_Alberts

Cell_Biology_Alberts. An important approach in determining gene function is to alter the gene (or in some cases, its expression pattern), to put the altered copy back into the germ line of the organism, and to deduce the function of the normal gene by the changes caused by its alteration. Various techniques to implement this strategy are discussed in the next section of this chapter. But it is equally important to study the biochemical and structural properties of a gene product, as outlined in the first part of this chapter. One of the most important contributions of DNA cloning to cell and molecular biology is the ability to produce any protein, even the rare ones, in nearly unlimited amounts—as long as the gene coding for it is known. Such high-level production is usually carried out in living cells using expression vectors (Figure 8–41). These are generally plasmids that have been designed to produce a large amount of stable mRNA that can be efficiently translated into protein when the plasmid is


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Cell_Biology_Alberts. expression vectors (Figure 8–41). These are generally plasmids that have been designed to produce a large amount of stable mRNA that can be efficiently translated into protein when the plasmid is introduced into bacterial, yeast, insect, or mammalian cells. To prevent the high level of the foreign protein from interfering with the cell’s growth, the expression vector is often designed to delay the synthesis of the foreign mRNA and protein until shortly before the cells are harvested and lysed (Figure 8–42).


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Cell_Biology_Alberts. Because the desired protein made from an expression vector is produced inside a cell, it must be purified away from the host-cell proteins by chromatography following cell lysis; but because it is such a plentiful species in the cell (often 1–10% of the total cell protein), the purification is usually easy to accomplish in only a few steps. As we saw in the first part of this chapter, many expression


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Cell_Biology_Alberts. Figure 8–41 Production of large amounts of a protein from a protein-coding DNA sequence cloned into an expression vector and introduced into cells. A plasmid vector has been engineered to contain a highly active promoter, which causes unusually large amounts of mRNA to be produced from an adjacent protein-coding gene inserted into the plasmid vector. Depending on the characteristics of the cloning vector, the plasmid is introduced into bacterial, yeast, insect, or mammalian cells, where the inserted gene is efficiently transcribed and translated into protein. If the gene to be overexpressed has no introns (typical for genes from bacteria, archaea, and simple eukaryotes), it can simply be cloned from genomic DNA by PCR. For cloned animal and plant genes, it is often more convenient to obtain the gene as cDNA, either from a cDNA library (see Figure 8–32) or cloned directly by PCR from RNA isolated from the organism (see Figure 8–37). Alternatively, the DNA coding for the protein can be


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Cell_Biology_Alberts. the gene as cDNA, either from a cDNA library (see Figure 8–32) or cloned directly by PCR from RNA isolated from the organism (see Figure 8–37). Alternatively, the DNA coding for the protein can be made by chemical synthesis (see p. 472).


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Cell_Biology_Alberts. vectors have been designed to add a molecular tag—a cluster of histidine residues or a small marker protein—to the expressed protein to facilitate easy purification by affinity chromatography (see Figure 8–11). A variety of expression vectors is available, each engineered to function in the type of cell in which the protein is to be made. This technology is also used to make large amounts of many medically useful proteins, including hormones (such as insulin and growth factors) used as human pharmaceuticals, and viral coat proteins for use in vaccines. Expression vectors also allow scientists to produce many proteins of biological interest in large enough amounts for detailed structural studies. Nearly all three-dimensional protein structures depicted in this book are of proteins produced in this way. Recombinant DNA techniques thus allow scientists to move with ease from protein to gene, and vice versa, so that the functions of both can be explored on multiple fronts (Figure 8–43).


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Cell_Biology_Alberts. DNA cloning allows a copy of any specific part of a DNA or RNA sequence to be selected from the millions of other sequences in a cell and produced in unlimited amounts in pure form. DNA sequences can be amplified after breaking up chromosomal DNA and inserting the resulting DNA fragments into the chromosome of a self-replicating genetic element such as a plasmid. The resulting “genomic DNA library” is housed in millions of bacterial cells, each carrying a different cloned DNA fragment. Individual cells from this library that are allowed to proliferate produce large amounts of a single cloned DNA fragment. Bypassing cloning vectors and bacterial cells altogether, the polymerase chain reaction (PCR) allows DNA cloning to be performed directly with a DNA polymerase and DNA primers—provided that the DNA sequence of interest is already known.


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Cell_Biology_Alberts. The procedures used to obtain DNA clones that correspond in sequence to mRNA molecules are the same, except that a DNA copy of the mRNA sequence, called cDNA, is first made. Unlike genomic DNA clones, cDNA clones lack intron sequences, making them the clones of choice for analyzing the protein product of a gene. Nucleic acid hybridization reactions provide a sensitive means of detecting any nucleotide sequence of interest. The enormous specificity of this hybridization reaction allows any single-stranded sequence of nucleotides to be labeled with a radioisotope or chemical and used as a probe to find a complementary partner strand, even in a cell or cell extract that contains millions of different DNA and RNA sequences. DNA hybridization also makes it possible to use PCR to amplify any section of any genome once its sequence is known. direction of electrophoresis


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Cell_Biology_Alberts. direction of electrophoresis Figure 8–42 Production of large amounts of a protein by using a plasmid expression vector. In this example, an expression vector that overproduces a DNA helicase has been introduced into bacteria. In this expression vector, transcription from this coding sequence is under the control of a viral promoter that becomes active only at a temperature of 37°C or higher. The total cell protein, either from bacteria grown at 25°C (no helicase protein made) or after a shift of the same bacteria to 42°C for up to 2 hours (helicase protein has become the most abundant protein species in the lysate), has been analyzed by SDS polyacrylamide-gel electrophoresis. (Courtesy of Jack Barry.) sequence of a peptide fragment E. coli or otherPROTEIN host cell to produce protein DNA primers for PCR insert protein-coding region of gene into expression vector (from cDNA clone)


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Cell_Biology_Alberts. E. coli or otherPROTEIN host cell to produce protein DNA primers for PCR insert protein-coding region of gene into expression vector (from cDNA clone) Figure 8–43 Recombinant DNA techniques make it possible to move experimentally from gene to protein and from protein to gene. If a gene has been identified (right), its protein-coding sequence can be inserted into an expression vector to produce large quantities of the protein (see Figure 8–41), which can then be studied biochemically or structurally. If a protein has been purified based on its biochemical properties, mass spectrometry (see Figure 8–18) can be used to obtain a partial amino acid sequence, which is used to search a genome sequence for the corresponding nucleotide sequence. The complete gene can then be cloned by PCR from a sequenced genome (see Figure 8–37). The gene can also be manipulated and introduced into cells or organisms to study its function, a topic covered in the next section of this chapter.


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Cell_Biology_Alberts. The nucleotide sequence of any genome can be determined rapidly and simply by using highly automated techniques based on several different strategies. Comparison of the genome sequences of different organisms allows us to trace the evolutionary relationships among genes and organisms, and it has proved valuable for discovering new genes and predicting their functions. Taken together, these techniques for analyzing and manipulating DNA have made it possible to sequence, identify, and isolate genes from any organism of interest. Related technologies allow scientists to produce the protein products of these genes in the large quantities needed for detailed analyses of their structure and function, as well as for medical purposes.


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Cell_Biology_Alberts. Ultimately, one wishes to determine how genes—and the proteins they encode— function in the intact organism. Although it may seem counterintuitive, one of the most direct ways to find out what a gene does is to see what happens to the organism when that gene is missing. Studying mutant organisms that have acquired changes or deletions in their nucleotide sequences is a time-honored practice in biology and forms the basis of the important field of genetics. Because mutations can disrupt cell processes, mutants often hold the key to understanding gene function. In the classical genetic approach, one begins by isolating mutants that have an interesting or unusual appearance: fruit flies with white eyes or curly wings, for example. Working backward from the phenotype—the appearance or behavior of the individual—one then determines the organism’s genotype, the form of the gene responsible for that characteristic (Panel 8–2).


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Cell_Biology_Alberts. Today, with numerous genome sequences available, the exploration of gene function often begins with a DNA sequence. Here, the challenge is to translate sequence into function. One approach, discussed earlier in the chapter, is to search databases for well-characterized proteins that have similar amino acid sequences to the protein encoded by a new gene. From there, the protein (or for noncoding genes, the RNA molecule) can be overexpressed and purified and the methods described in the first part of this chapter can be employed to study its three-dimensional structure and its biochemical properties. But to determine directly a gene’s function in a cell or organism, the most effective approach involves studying mutants that either lack the gene or express an altered version of it. Determining which cell processes have been disrupted or compromised in such mutants will usually shed light on a gene’s biological role.


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Cell_Biology_Alberts. In this section, we describe several approaches to determining a gene’s function, starting either from an individual with an interesting phenotype or from a DNA sequence. We begin with the classical genetic approach, which starts with a genetic screen for isolating mutants of interest and then proceeds toward identification of the gene or genes responsible for the observed phenotype. We then describe the set of techniques that are collectively called reverse genetics, in which one begins with a gene or gene sequence and attempts to determine its function. This approach often involves some intelligent guesswork—searching for similar sequences in other organisms or determining when and where a gene is expressed—as well as generating mutant organisms and characterizing their phenotype.


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Cell_Biology_Alberts. Before the advent of gene cloning technology, most genes were identified by the abnormalities produced when the gene was mutated. Indeed, the very concept of the gene was deduced from the heritability of such abnormalities. This classical genetic approach—identifying the genes responsible for mutant phenotypes—is most easily performed in organisms that reproduce rapidly and are amenable to genetic manipulation, such as bacteria, yeasts, nematode worms, and fruit flies. Although spontaneous mutants can sometimes be found by examining extremely PANeL 8–2: Review of Classical Genetics GENES AND PHENOTYPES Gene: a functional unit of inheritance, usually corresponding to the segment of DNA coding for a single protein. Genome: all of an organism’s DNA sequences. locus:the site of the gene in the genome Wild-type: the normal, Mutant: differing from the alleles:alternative forms of a gene naturally occurring type wild-type because of a genetic change (a mutation)


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Cell_Biology_Alberts. Wild-type: the normal, Mutant: differing from the alleles:alternative forms of a gene naturally occurring type wild-type because of a genetic change (a mutation) GENOTYPE: the specifc set of alleles forming the genome of an individual PHENOTYPE: the visible character of the individual allele A is dominant (relative to a); allele a is recessive (relative to A) In the example above, the phenotype of the heterozygote is the same as that of one of the homozygotes; in cases where it is different from both, the two alleles are said to be co-dominant. a chromosome at the beginning of the cell cycle, in G1 phase; the single long bar centromere represents one long double helix of DNA a chromosome near the end of the cell cycle, in metaphase; it is duplicated and condensed, consisting of mother father two identical sister chromatids (each containing one DNA double helix) joined at the centromere.


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Cell_Biology_Alberts. “p” arm “q” arm A normal diploid chromosome set, as seen in a metaphase spread, prepared pair of autosomes by bursting open a cell at metaphase and staining the scattered maternal 1 chromosomes. In the example shown schematically here, there are three pairs of autosomes (chromosomes both parents, regardless of sex) and two sex chromosomes—an X from the mother and a Y from the father. The maternal 2 numbers and types of sex determination are variable from one class of organisms to another, as is the X number of pairs of autosomes. For simplicity, the cycle is shown for only sex chromosomes one chromosome/chromosome pair. MEIOSIS AND GENETIC RECOMBINATION The greater the distance chromosome, the greater is the chance that they will be


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Cell_Biology_Alberts. MEIOSIS AND GENETIC RECOMBINATION The greater the distance chromosome, the greater is the chance that they will be RECOMBINATION site of crossing-over occurring at a site between them. If two genes are thus reassorted in x% of gametes, they are said to be separated on a chromosome by a genetic map AB distance of x map units (or genotype ab haploid gametes (eggs or sperm) x centimorgans). POINT MUTATION: maps to a single site in the genome, corresponding to a single nucleotide pair or a very small part of a single gene INVERSION: inverts a segment of a chromosome lethal mutation: causes the developing organism to die prematurely. conditional mutation: produces its phenotypic effect only under certain conditions, called the restrictive conditions. Under other conditions—the permissive conditions—the effect is not seen. For a temperature-sensitive mutation, the restrictive condition typically is high temperature, while the permissive condition is low temperature.


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Cell_Biology_Alberts. loss-of-function mutation: either reduces or abolishes the activity of the gene. These are the most common class of mutations. Loss-of-function mutations are usually recessive—the organism can usually function normally as long as it retains at least one normal copy of the affected gene. null mutation: a loss-of-function mutation that completely abolishes the activity of the gene. TWO GENES OR ONE? Given two mutations that produce the same phenotype, how can we tell whether they are mutations in the same gene? If the mutations are recessive (as they most often are), the answer can be found by a complementation test. COMPLEMENTATION: MUTATIONS IN TWO DIFFERENT GENES DELETION: deletes a segment of a chromosome


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Cell_Biology_Alberts. COMPLEMENTATION: MUTATIONS IN TWO DIFFERENT GENES DELETION: deletes a segment of a chromosome TRANSLOCATION: breaks off a segment from one chromosome and attaches it to another gain-of-function mutation: increases the activity of the gene or makes it active in inappropriate circumstances; these mutations are usually dominant. dominant-negative mutation: dominant-acting mutation that blocks gene activity, causing a loss-of-function phenotype even in the presence of a normal copy of the gene. This phenomenon occurs when the mutant gene product interferes with the function of the normal gene product. suppressor mutation: suppresses the phenotypic effect of another mutation, so that the double mutant seems normal. An intragenic suppressor mutation lies within the gene affected by the frst mutation; an extragenic suppressor mutation lies in a second gene—often one whose product interacts directly with the product of the frst.


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Cell_Biology_Alberts. In the simplest type of complementation test, an individual who is homozygous for one mutation is mated with an individual who is homozygous for the other. The phenotype of the offspring gives the answer to the question. NONCOMPLEMENTATION: TWO INDEPENDENT MUTATIONS IN THE SAME GENE hybrid offspring shows normal phenotype: hybrid offspring shows mutant phenotype: one normal copy of each gene is present no normal copies of the mutated gene are present large populations—thousands or tens of thousands of individual organisms—isolating mutant individuals is much more efficient if one generates mutations with chemicals or radiation that damage DNA. By treating organisms with such mutagens, very large numbers of mutant individuals can be created quickly and then screened for a particular defect of interest, as we discuss shortly.


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Cell_Biology_Alberts. An alternative approach to chemical or radiation mutagenesis is called insertional mutagenesis. This method relies on the fact that exogenous DNA inserted randomly into the genome can produce mutations if the inserted fragment interrupts a gene or its regulatory sequences. The inserted DNA, whose sequence is known, then serves as a molecular tag that aids in the subsequent identification and cloning of the disrupted gene (Figure 8–44). In Drosophila, the use of the transposable P element to inactivate genes has revolutionized the study of gene function in the fly. Transposable elements (see Table 5–4, p. 288) have also been used to generate mutations in bacteria, yeast, mice, and the flowering plant Arabidopsis. Genetic Screens Identify Mutants with Specific Abnormalities


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Cell_Biology_Alberts. Genetic Screens Identify Mutants with Specific Abnormalities Once a collection of mutants in a model organism such as yeast or fly has been produced, one generally must examine thousands of individuals to find the altered phenotype of interest. Such a search is called a genetic screen, and the larger the genome, the less likely it is that any particular gene will be mutated. Therefore, the larger the genome of an organism, the bigger the screening task becomes. The phenotype being screened for can be simple or complex. Simple phenotypes are easiest to detect: one can screen many organisms rapidly, for example, for mutations that make it impossible for the organism to survive in the absence of a particular amino acid or nutrient.


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Cell_Biology_Alberts. More complex phenotypes, such as defects in learning or behavior, may require more elaborate screens (Figure 8–45). But even genetic screens that are used to dissect complex physiological systems can be simple in design, which permits the simultaneous examination of large numbers of mutants. As an example, one particularly elegant screen was designed to search for genes involved in visual processing in zebrafish. The basis of this screen, which monitors the fishes’ response to motion, is a change in behavior. Wild-type fish tend to swim in the direction of a perceived motion, whereas mutants with defects in their visual processing systems swim in random directions—a behavior that is easily detected. One mutant discovered in this screen is called lakritz, which is missing 80% of the retinal ganglion cells that help to relay visual signals from the eye to the brain. As the cellular organization of the zebrafish retina is similar to that of all vertebrates, the study of such mutants


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Cell_Biology_Alberts. ganglion cells that help to relay visual signals from the eye to the brain. As the cellular organization of the zebrafish retina is similar to that of all vertebrates, the study of such mutants should also provide insights into visual processing in humans.


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Cell_Biology_Alberts. Because defects in genes that are required for fundamental cell processes— RNA synthesis and processing or cell-cycle control, for example—are usually lethal, the functions of these genes are often studied in individuals with Figure 8–44 Insertional mutant of the snapdragon, Antirrhinum. A mutation in a single gene coding for a regulatory protein causes leafy shoots (left) to develop in place of flowers, which occur in the normal plant (right). The mutation causes cells to adopt a character that would be appropriate to a different part of the normal plant, so instead of a flower, the cells produce a leafy shoot. (Courtesy of Enrico Coen and Rosemary Carpenter.)


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Cell_Biology_Alberts. Figure 8–45 A behavioral phenotype detected in a genetic screen. (A) Wild-type C. elegans engage in social feeding. The worms migrate around until they encounter their neighbors and commence feeding on bacteria. (B) Mutant animals feed by themselves. (Courtesy of Cornelia Bargmann, Cell 94: cover, 1998. With permission from Elsevier.) mutant cells proliferate and form a colony at the permissive temperature mutant cells fail to proliferate and form a colony at the nonpermissive conditional mutations. The mutant individuals function normally as long as “permissive” conditions prevail, but demonstrate abnormal gene function when subjected to “nonpermissive” (restrictive) conditions. In organisms with temperature-sensitive mutations, for example, the abnormality can be switched on and off experimentally simply by changing the ambient temperature; thus, a cell containing a temperature-sensitive mutation in a gene essential for survival will die at a nonpermissive temperature but


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Cell_Biology_Alberts. off experimentally simply by changing the ambient temperature; thus, a cell containing a temperature-sensitive mutation in a gene essential for survival will die at a nonpermissive temperature but proliferate normally at the permissive temperature (Figure 8–46). The temperature-sensitive gene in such a mutant usually contains a point mutation that causes a subtle change in its protein product; for example, the mutant protein may function normally at low temperatures but unfold at higher temperatures.


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Cell_Biology_Alberts. Temperature-sensitive mutations were crucial to find the bacterial genes that encode the proteins required for DNA replication. The mutants were identified by screening populations of mutagen-treated bacteria for cells that stop making DNA when they are warmed from 30°C to 42°C. These mutants were later used to identify and characterize the corresponding DNA replication proteins (discussed in Chapter 5). Similarly, screens for temperature-sensitive mutations led to the identification of many proteins involved in regulating the cell cycle, as well as many proteins involved in moving proteins through the secretory pathway in yeast. Related screening approaches demonstrated the function of enzymes involved in the principal metabolic pathways of bacteria and yeast (discussed in Chapter 2) and identified many of the gene products responsible for the orderly development of the Drosophila embryo (discussed in Chapter 21). Mutations Can Cause Loss or Gain of Protein Function


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Cell_Biology_Alberts. Mutations Can Cause Loss or Gain of Protein Function Gene mutations are generally classed as “loss of function” or “gain of function.” A loss-of-function mutation results in a gene product that either does not work or works too little; thus, it can reveal the normal function of the gene. A gain-of-function mutation results in a gene product that works too much, works at the wrong time or place, or works in a new way (Figure 8–47).


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Cell_Biology_Alberts. An important early step in the genetic analysis of any mutant cell or organism is to determine whether the mutation causes a loss or a gain of function. A standard test is to determine whether the mutation is dominant or recessive. A dominant mutation is one that still causes the mutant phenotype in the presence of a single copy of the wild-type gene. A recessive mutation is one that is no longer able to cause the mutant phenotype in the presence of a single wild-type copy of the gene. Although cases have been described in which a loss-of-function mutation is dominant or a gain-of-function mutation is recessive, in the vast majority of cases, recessive mutations are loss of function and dominant mutations are gain Figure 8–46 Screening for temperature-sensitive bacterial or yeast mutants.


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Cell_Biology_Alberts. Figure 8–46 Screening for temperature-sensitive bacterial or yeast mutants. Mutagenized cells are plated out at the permissive temperature. They divide and form colonies, which are transferred to two identical Petri dishes by replica plating. One of these plates is incubated at the permissive temperature, the other at the nonpermissive temperature. Cells containing a temperature-sensitive mutation in a gene essential for proliferation can divide at the normal, permissive temperature but fail to divide at the elevated, nonpermissive temperature. Temperature-sensitive mutations of this type were especially useful for identifying genes needed for DNA replication, an essential process. Figure 8–47 Gene mutations that affect their protein product in different ways.


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Cell_Biology_Alberts. Figure 8–47 Gene mutations that affect their protein product in different ways. In this example, the wild-type protein has a specific cell function denoted by the red rays. Mutations that eliminate this function or inactivate it at higher temperatures are shown. The conditional mutant protein carries an amino acid substitution (red) that prevents its proper folding at 37ºC, but allows the protein to fold and function normally at 25ºC. Such temperature-sensitive conditional mutations are especially useful for studying essential genes; the organism can be grown under the permissive condition and then be moved to the nonpermissive condition to study the consequences of losing the gene product.


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Cell_Biology_Alberts. of function. It is easy to determine if a mutation is dominant or recessive. One simply mates a mutant with a wild type to obtain diploid cells or organisms. The progeny from the mating will be heterozygous for the mutation. If the mutant phenotype is no longer observed, one can conclude that the mutation is recessive and is very likely to be a loss-of-function mutation (see Panel 8–2). Complementation Tests Reveal Whether Two Mutations Are in the Same Gene or Different Genes


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Cell_Biology_Alberts. A large-scale genetic screen can turn up many different mutations that show the same phenotype. These defects might lie in different genes that function in the same process, or they might represent different mutations in the same gene. Alternative forms of the same gene are known as alleles. The most common difference between alleles is a substitution of a single nucleotide pair, but different alleles can also bear deletions, substitutions, and duplications. How can we tell, then, whether two mutations that produce the same phenotype occur in the same gene or in different genes? If the mutations are recessive—if, for example, they represent a loss of function of a particular gene—a complementation test can be used to ascertain whether the mutations fall in the same gene or in different genes. To test complementation in a diploid organism, an individual that is homozygous for one mutation—that is, it possesses two identical alleles of the mutant gene in question—is mated with an


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Cell_Biology_Alberts. genes. To test complementation in a diploid organism, an individual that is homozygous for one mutation—that is, it possesses two identical alleles of the mutant gene in question—is mated with an individual that is homozygous for the other mutation. If the two mutations are in the same gene, the offspring show the mutant phenotype, because they still will have no normal copies of the gene in question (Figure 8–48). If, in contrast, the mutations fall in different genes, the resulting offspring show a normal phenotype, because they retain one normal copy (and one mutant copy) of each gene; the mutations thereby complement one another and restore a normal phenotype. Complementation testing of mutants identified during genetic screens has revealed, for example, that 5 different genes are required for yeast to digest the sugar galactose, 20 genes are needed for E. coli to build a functional flagellum, 48 genes are involved in assembling bacteriophage T4 viral particles, and hundreds of


Cell_Biology_Alberts

Cell_Biology_Alberts. for yeast to digest the sugar galactose, 20 genes are needed for E. coli to build a functional flagellum, 48 genes are involved in assembling bacteriophage T4 viral particles, and hundreds of genes are involved in the development of an adult nematode worm from a fertilized egg.


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Cell_Biology_Alberts. Once a set of genes involved in a particular biological process has been identified, the next step is often to determine in which order the genes function. Gene order is perhaps easiest to explain for metabolic pathways, where, for example, enzyme A is necessary to produce the substrate for enzyme B. In this case, we would say that the gene encoding enzyme A acts before (upstream of) the gene encoding enzyme B in the pathway. Similarly, where one protein regulates the activity of another protein, we would say that the former gene acts before the latter. Gene order can, in many cases, be determined purely by genetic analysis without any knowledge of the mechanism of action of the gene products involved. Suppose we have a biosynthetic process consisting of a sequence of steps, such that performance of step B is conditional on completion of the preceding step A; and suppose gene A is required for step A, and gene B is required for step


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Cell_Biology_Alberts. B. Then a null mutation (a mutation that abolishes function) in gene A will arrest the process at step A, regardless of whether gene B is functional or not, whereas a null mutation in gene B will cause arrest at step B only if gene A is still active. In such a case, gene A is said to be epistatic to gene B. By comparing the phenotypes of the different combinations of mutations, we can therefore discover the order in which the genes act. This type of analysis is called epistasis analysis. As an example, the pathway of protein secretion in yeast has been analyzed in this way. Different mutations in this pathway cause proteins to accumulate aberrantly in the endoplasmic reticulum (ER) or in the Golgi apparatus. When a yeast cell is engineered to carry both a mutation that blocks protein processing in the ER and a mutation that blocks processing in the Golgi apparatus, proteins accumulate in


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Cell_Biology_Alberts. Figure 8–48 A complementation test can reveal that mutations in two different genes are responsible for the same abnormal phenotype. When an albino (white) bird from one strain is bred with an albino from a different strain, the resulting offspring (bottom) have normal coloration. This restoration of the wild-type plumage indicates that the two white breeds lack color because of recessive mutations in different genes. (From W. Bateson, Mendel’s Principles of Heredity, 1st ed. Cambridge, UK: Cambridge University Press, 1913.) the ER. This indicates that proteins must pass through the ER before being sent to the Golgi before secretion (Figure 8–49). Strictly speaking, an epistasis analysis can only provide information about gene order in a pathway when both mutations are null alleles. When the mutations retain partial function, their epistasis interactions can be difficult to interpret.


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Cell_Biology_Alberts. Sometimes, a double mutant will show a new or more severe phenotype than either single mutant alone. This type of genetic interaction is called a synthetic phenotype, and if the phenotype is death of the organism, it is called synthetic lethality. In most cases, a synthetic phenotype indicates that the two genes act in two different parallel pathways, either of which is capable of mediating the same cell process. Thus, when both pathways are disrupted in the double mutant, the process fails altogether, and the synthetic phenotype is observed.


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Cell_Biology_Alberts. Once a collection of mutant organisms with interesting phenotypes has been obtained, the next task is to identify the gene or genes responsible for the altered phenotype. If the phenotype has been produced by insertional mutagenesis, locating the disrupted gene is fairly simple. DNA fragments containing the insertion (a transposon or a retrovirus, for example) are amplified by PCR, and the nucleotide sequence of the flanking DNA is determined. The gene affected by the insertion can then be identified by a computer-aided search of the complete genome sequence of the organism.


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Cell_Biology_Alberts. If a DNA-damaging chemical was used to generate the mutations, identifying the inactivated gene is often more laborious, but there are several powerful strategies available. If the genome size of the organism is small (for example, for bacteria or simple eukaryotes), it is possible to simply determine the genome sequence of the mutant organism and identify the affected gene by comparison with the wild-type sequence. Because of the continuous accumulation of neutral mutations, there will probably be differences between the two genome sequences in addition to the mutation responsible for the phenotype. One way of proving that a mutation is causative is to introduce the putative mutation back into a normal organism and determine whether or not it causes the mutant phenotype. We will discuss how this is accomplished later in the chapter.


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Cell_Biology_Alberts. Genetic screens in model experimental organisms have been spectacularly successful in identifying genes and relating them to various phenotypes, including many that are conserved between these organisms and humans. But how can we study humans directly? They do not reproduce rapidly, cannot be treated with mutagens, and, if they have a defect in an essential process such as DNA replication, would die long before birth. Despite their limitations compared to model organisms, humans are becoming increasingly attractive subjects for genetic studies. Because the human


Cell_Biology_Alberts

Cell_Biology_Alberts. Despite their limitations compared to model organisms, humans are becoming increasingly attractive subjects for genetic studies. Because the human Figure 8–49 Using genetics to determine the order of function of genes. In normal cells, secretory proteins are loaded into vesicles, which fuse with the plasma membrane to secrete their contents into the extracellular medium. Two mutants, A and B, fail to secrete proteins. In mutant A, secretory proteins accumulate in the ER. In mutant B, secretory proteins accumulate in the Golgi. In the double mutant AB, proteins accumulate in the ER; this indicates that the gene defective in mutant A acts before the gene defective in mutant B in the secretory pathway.


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Cell_Biology_Alberts. population is so large, spontaneous, nonlethal mutations have arisen in all human genes—many times over. A substantial proportion of these remain in the genomes of present-day humans. The most deleterious of these mutations are discovered when the mutant individuals call attention to themselves by seeking medical help. With the recent advances that have enabled the sequencing of entire human genomes cheaply and quickly, we can now identify such mutations and study their evolution and inheritance in ways that were impossible even a few years ago. By comparing the sequences of thousands of human genomes from all around the world, we can begin to identify directly the DNA differences that distinguish one individual from another. These differences hold clues to our evolutionary origins and can be used to explore the roots of disease. Linked Blocks of Polymorphisms Have Been Passed Down from Our Ancestors


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Cell_Biology_Alberts. Linked Blocks of Polymorphisms Have Been Passed Down from Our Ancestors When we compare the sequences of multiple human genomes, we find that any two individuals will differ in roughly 1 nucleotide pair in 1000. Most of these variations are common and relatively harmless. When two sequence variants coexist in the population and both are common, the variants are called polymorphisms. The majority of polymorphisms are due to the substitution of a single nucleotide, called single-nucleotide polymorphisms or SNPs (Figure 8–50). The rest are due largely to insertions or deletions—called indels when the change is small, or copy number variations (CNVs) when it is large. Although these common variants can be found throughout the genome, they are not scattered randomly—or even independently. Instead, they tend to travel in groups called haplotype blocks—combinations of polymorphisms that are inherited as a unit.


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Cell_Biology_Alberts. To understand why such haplotype blocks exist, we need to consider our evolutionary history. It is thought that modern humans expanded from a relatively small population—perhaps around 10,000 individuals—that existed in Africa about 60,000 years ago. Among that small group of our ancestors, some individuals will have carried one set of genetic variants, others a different set. The chromosomes of a present-day human represent a shuffled combination of chromosome segments from different members of this small ancestral group of people. Because only about two thousand generations separate us from them, large segments of these ancestral chromosomes have passed from parent to child, unbroken by the crossover events that occur during meiosis. As described in Chapter 5, only a few crossovers occur between each set of homologous chromosomes during each meiosis (see Figure 5–53).


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Cell_Biology_Alberts. As a result, certain sets of DNA sequences—and their associated polymorphisms—have been inherited in linked groups, with little genetic rearrangement across the generations. These are the haplotype blocks. Like genes that exist in different allelic forms, haplotype blocks also come in a limited number of variants that are common in the human population, each representing a combination of DNA polymorphisms passed down from a particular ancestor long ago. Figure 8–50 Single-nucleotide polymorphisms (SNPs) are sites in the genome where two or more alternative choices of a nucleotide are common in the population. Most such variations in the human genome occur at locations where they do not significantly affect a gene’s function. Polymorphisms Can Aid the Search for Mutations Associated with Disease


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Cell_Biology_Alberts. Mutations that give rise, in a reproducible way, to rare but clearly defined abnormalities, such as albinism, hemophilia, or congenital deafness, can often be identified by studies of affected families. Such single-gene, or monogenic, disorders are often referred to as Mendelian because their pattern of inheritance is easy to track. Moreover, individuals who inherit the causative mutation will exhibit the abnormality irrespective of environmental factors such as diet or exercise. But for many common diseases, the genetic roots are more complex. Instead of a single allele of a single gene, such disorders stem from a combination of contributions from multiple genes. And often, environmental factors have strong influences on the severity of the disorder. For these multigenic conditions, such as diabetes or arthritis, population studies are often helpful in tracking down the genes that increase the risk of getting the disease.


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Cell_Biology_Alberts. In population studies, investigators collect DNA samples from a large number of people who have the disease and compare them to samples from a group of people who do not have the disease. They look for variants—SNPs, for example— that are more common among the people who have the disease. Because DNA sequences that are close together on a chromosome tend to be inherited together, the presence of such SNPs could indicate that an allele that increases the risk of the disease might lie nearby (Figure 8–51). Although, in principle, the disease could be caused by the SNP itself, the culprit is much more likely to be a change that is merely linked to the SNP as part of a haplotype block.


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Cell_Biology_Alberts. Such genome-wide association studies have been used to search for genes that predispose individuals to common diseases, including diabetes, coronary artery disease, rheumatoid arthritis, and even depression. For many of these conditions, the DNA polymorphisms identified increase the risk of disease only slightly. Moreover, environmental factors (diet, exercise, for example) play an important role in the onset and severity of the disease. Nonetheless, the identification of genes affected by these polymorphisms is leading to a mechanistic understanding of some of our most common disorders. Genomics Is Accelerating the Discovery of Rare Mutations That Predispose Us to Serious Disease


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Cell_Biology_Alberts. Genomics Is Accelerating the Discovery of Rare Mutations That Predispose Us to Serious Disease The genetic variants that have thus far allowed us to identify some of the genes that increase our risk of disease are common ones. They arose long ago in our evolutionary past and are now present, in one form or another, in a substantial portion (1% or more) of the population. Such polymorphisms are thought to account


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Cell_Biology_Alberts. Figure 8–51 Genes that affect the risk of developing a common disease can often be tracked down through linkage to SNPs. Here, the patterns of SNPs are compared between two sets of individuals—a set of healthy controls and a set affected by a particular common disease. A segment of a typical chromosome is shown. For most polymorphic sites in this segment, it is a random matter whether an individual has one SNP variant (red vertical bars) or another (blue vertical bars); this same randomness is seen both for the control group and for the affected individuals. However, in the part of the chromosome that is shaded in darker gray, a bias is seen: most normal individuals have the blue SNP variants, whereas most affected individuals have the red SNP variants. This suggests that this region contains, or is close to, a gene that is genetically linked to these red SNP variants and that predisposes individuals to the disease. Using carefully selected controls and thousands of affected


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Cell_Biology_Alberts. contains, or is close to, a gene that is genetically linked to these red SNP variants and that predisposes individuals to the disease. Using carefully selected controls and thousands of affected individuals, this approach can help track down disease-related genes, even when they confer only a slight increase in the risk of developing the disease.


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Cell_Biology_Alberts. for about 90% of the differences between one person’s genome and another’s. But when we try to tie these common variants to differences in disease susceptibility or other heritable traits, such as height, we find that they do not have as much predictive power as we had anticipated: thus, for example, most confer relatively small increases—less than twofold—in the risk of developing a common disease.


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Cell_Biology_Alberts. In contrast to polymorphisms, rare DNA variants—those much less frequent in humans than SNPs—can have large effects on the risk of developing some common diseases. For example, a number of different loss-of-function mutations, each individually rare, have been found to increase greatly the predisposition to autism and schizophrenia. Many of these are de novo mutations, which arose spontaneously in the germ-line cells of one or the other parent. The fact that these mutations arise spontaneously with some frequency could help explain why these common disorders—each observed in about 1% of the population—remain with us, even though the affected individuals leave few or no descendants. These rare mutations may arise in any one of hundreds of different genes, which could explain much of the clinical variability of autism and schizophrenia. Because they are kept rare by natural selection, most such variants with a large effect on risk would be missed by genome-wide association studies.


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Cell_Biology_Alberts. Now that DNA sequencing has become fast and inexpensive, the most efficient and cost-effective way to identify these rare, large-effect mutations is by sequencing the genomes of affected individuals, along with those of their parents and siblings as controls. Reverse Genetics Begins with a Known Gene and Determines Which Cell Processes Require Its Function


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Cell_Biology_Alberts. As we have seen, classical genetics starts with a mutant phenotype (or, in the case of humans, a range of characteristics) and identifies the mutations, and consequently the genes, responsible for it. Recombinant DNA technology has made possible a different type of genetic approach, one that is used widely in a variety of genetically tractable species. Instead of beginning with a mutant organism and using it to identify a gene and its protein, an investigator can start with a particular gene and proceed to make mutations in it, creating mutant cells or organisms so as to analyze the gene’s function. Because this approach reverses the traditional direction of genetic discovery—proceeding from genes to mutations, rather than vice versa—it is commonly referred to as reverse genetics. And because the genome of the organism is deliberately altered in a particular way, this approach is also called genome engineering or genome editing. We shall see in this chapter that this approach can be


Cell_Biology_Alberts

Cell_Biology_Alberts. the genome of the organism is deliberately altered in a particular way, this approach is also called genome engineering or genome editing. We shall see in this chapter that this approach can be scaled upward so that whole collections of organisms can be created, each of which has a different gene altered.


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Cell_Biology_Alberts. There are several ways a gene of interest can be altered. In the simplest, the gene can simply be deleted from the genome, although in a diploid organism, this requires that both copies—one on each chromosome homolog—be deleted. Although somewhat counterintuitive, one of the best ways to discover the function of a gene is by observing the effects of not having it. Such “gene knockouts” are especially useful if the gene is not essential. Through reverse genetics, the gene in question (even if it is essential) can also be replaced by one that is expressed in the wrong tissue or at the wrong time in development; this type of manipulation often provides important clues to the gene’s normal function. For example, a gene of interest can be modified to be expressed at will by the experimenter (Figure 8–52). Finally, genes can also be engineered so that they are expressed normally in most cell types and tissues but deleted in certain cell types or tissues selected by the experimenter (see


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Cell_Biology_Alberts. (Figure 8–52). Finally, genes can also be engineered so that they are expressed normally in most cell types and tissues but deleted in certain cell types or tissues selected by the experimenter (see Figure 5–66). This approach is especially useful when a gene has different roles in different tissues.


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Cell_Biology_Alberts. It is also possible to make subtler changes to a gene. It is sometimes useful to make slight changes in a protein’s structure so that one can begin to dissect which portions of a protein are important for its function. The activity of an enzyme, for example, can be studied by changing a single amino acid in its active site. It is also possible, through genome engineering, to create new types of proteins in an


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Cell_Biology_Alberts. Figure 8–52 engineered genes can be turned on and off with small molecules. Here, the DNA-binding portion of a bacterial protein (the tetracycline, Tet, repressor) has been fused to a portion of a mammalian transcriptional activator and expressed in cultured mammalian cells. The engineered gene X, present in place of the normal gene, has its usual gene control region replaced by cis-regulatory sequences recognized by the tetracycline repressor. In the absence of doxycycline (a particularly stable version of tetracycline), the engineered gene is expressed; in the presence of doxycycline, the gene is turned off because the drug causes the tetracycline repressor to dissociate from the DNA. This strategy can also be used in mice by incorporating the engineered genes into the germ line. In many tissues, the gene can be turned on and off simply by adding or removing doxycycline from the animal’s water. If the tetracycline repressor construct is placed under the control of a tissue-specific


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Cell_Biology_Alberts. tissues, the gene can be turned on and off simply by adding or removing doxycycline from the animal’s water. If the tetracycline repressor construct is placed under the control of a tissue-specific gene control region, the engineered gene will be turned on and off only in that tissue.


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Cell_Biology_Alberts. animal. For example, a gene can be fused to the gene for a fluorescent protein. When this altered gene is introduced into the genome, the protein can be tracked in the living organism by monitoring its fluorescence.


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Cell_Biology_Alberts. Altered genes can be created in several ways. Perhaps the simplest is to chemically synthesize the DNA that makes up the gene. In this way, the investigator can specify any type of variant of the normal gene. It is also possible to construct altered genes using recombinant DNA technology, as described earlier in this chapter. Once obtained, altered genes can be introduced into cells in a variety of ways. DNA can be microinjected into mammalian cells with a glass micropipette or introduced by a virus that has been engineered to carry foreign genes. In plant cells, genes are frequently introduced by a technique called particle bombardment: DNA samples are painted onto tiny gold beads and then literally shot through the cell wall with a specially modified gun. Electroporation is the method of choice for introducing DNA into bacteria and some other cells. In this technique, a brief electric shock renders the cell membrane temporarily permeable, allowing foreign DNA to enter the cytoplasm.


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Cell_Biology_Alberts. To be most useful to experimenters, the altered gene, once it is introduced into a cell, must recombine with the cell’s genome so that the normal gene is replaced. In simple organisms such as bacteria and yeasts, this process occurs with high frequency using the cell’s own homologous recombination machinery, as described in Chapter 5. In more complex organisms that have elaborate developmental programs, the procedure is more complicated because the altered gene must be introduced into the germ line, as we next describe.


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Cell_Biology_Alberts. Animals and plants that have been genetically engineered by gene deletion or gene replacement are called transgenic organisms, and any foreign or modified genes that are added are called transgenes. We discuss transgenic plants later in this chapter and, for now, concentrate our discussion on transgenic mice, as enormous progress has been made in this area. If a DNA molecule carrying a mutated mouse gene is transferred into a mouse cell, it often inserts into the chromosomes at random, but methods have been developed to direct the mutant gene to replace the normal gene by homologous recombination. By exploiting these “gene targeting” events, any specific gene can be altered or inactivated in a mouse cell by a direct gene replacement. In the case in which both copies of the gene of interest are completely inactivated or deleted, the resulting animal is called a “knockout” mouse. The technique is summarized in Figure 8–53.


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Cell_Biology_Alberts. TRANSGENIC MOUSE WITH ONE COPY OF TARGET GENE REPLACED BY ALTERED GENE IN GERM LINE SOMATIC CELLS OF OFFSPRING TESTED FOR PRESENCE OF ALTERED GENE, AND SELECTED MICE BRED TO TEST FOR GENE IN GERM-LINE CELLS BIRTH MATE AND HARVEST EARLY EMBRYOS HYBRID EARLY EMBRYO PARTLY FORMED FROM ES CELLS INTRODUCE HYBRID EARLY EMBRYO INTO PSEUDOPREGNANT MOUSE isolated early embryo in culture altered version of target gene constructed by genetic engineering INTRODUCE A DNA FRAGMENT CONTAINING ALTERED GENE INTO MANY CELLS TEST FOR THE RARE COLONY IN WHICH THE DNA FRAGMENT HAS REPLACED ONE COPY OF THE NORMAL GENE LET EACH CELL PROLIFERATE TO FORM A COLONY ES cells with one copy of target gene replaced by mutant gene INJECT ES CELLS INTO EARLY EMBRYO


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Cell_Biology_Alberts. The ability to prepare transgenic mice lacking a known normal gene has been a major advance, and the technique has been used to determine the functions of many mouse genes (Figure 8–54). If the gene functions in early development, a knockout mouse will usually die before it reaches adulthood. These lethal defects can be carefully analyzed to help determine the function of the missing gene. As described in Chapter 5, an especially useful type of transgenic animal takes advantage of a site-specific recombination system to excise—and thus disable— the target gene in a particular place or at a particular time (see Figure 5–66). In this case, the target gene in embryonic stem (ES) cells is replaced by a fully functional version of the gene that is flanked by a pair of the short DNA sequences, called lox sites, that are recognized by the Cre recombinase protein. The transgenic mice that result are phenotypically normal. They are then mated with transgenic mice that express the Cre


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Cell_Biology_Alberts. called lox sites, that are recognized by the Cre recombinase protein. The transgenic mice that result are phenotypically normal. They are then mated with transgenic mice that express the Cre recombinase gene under the control of an inducible promoter. In


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Cell_Biology_Alberts. Figure 8–53 Summary of the procedures used for making gene replacements in mice. In the first step (A), an altered version of the gene is introduced into cultured ES (embryonic stem) cells. These cells are described in detail in Chapter 22. Only a few ES cells will have their corresponding normal genes replaced by the altered gene through a homologous recombination event. These cells can be identified by PCR and cultured to produce many descendants, each of which carries an altered gene in place of one of its two normal corresponding genes. In the next step of the procedure (B), these altered ES cells are injected into a very early mouse embryo; the cells are incorporated into the growing embryo, and a mouse produced by such an embryo will contain some somatic cells (indicated by orange) that carry the altered gene. Some of these mice will also contain germ-line cells that contain the altered gene; when bred with a normal mouse, some of the progeny of these mice will contain one copy


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Cell_Biology_Alberts. that carry the altered gene. Some of these mice will also contain germ-line cells that contain the altered gene; when bred with a normal mouse, some of the progeny of these mice will contain one copy of the altered gene in all of their cells.


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Cell_Biology_Alberts. The mice with the transgene in their germ line are then bred to produce both a male and a female animal, each heterozygous for the gene replacement (that is, they have one normal and one mutant copy of the gene). When these two mice are mated (not shown), one-fourth of their progeny will be homozygous for the altered gene. the specific cells or tissues in which Cre is switched on, it catalyzes recombination between the lox sequences—excising a target gene and eliminating its activity (see Figure 22–5). The Bacterial CRISPR System Has Been Adapted to Edit Genomes in a Wide Variety of Species One of the difficulties in making transgenic mice by the procedure just described is that the introduced DNA molecule (bearing the experimentally altered gene) often inserts at random in the genome, and many ES cells must therefore be screened individually to find one that has the “correct” gene replacement.


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Cell_Biology_Alberts. Creative use of the CRISPR system, discovered in bacteria as a defense against viruses, has largely solved this problem. As described in Chapter 7, the CRISPR system uses a guide RNA sequence to target (through complementary base-pairing) double-stranded DNA, which it then cleaves (see Figure 7–78). The gene coding for the key component of this system, the bacterial Cas9 protein, has been transferred into a variety of organisms, where it greatly simplifies the process of making transgenic organisms (Figure 8–55A and B). The basic strategy is as follows: Cas9 protein is expressed in ES cells along with a guide RNA designed by the experimenter to target a particular location on the genome. The Cas9 and guide RNA associate, the complex is brought to the matching sequence on the genome, and the Cas9 protein makes a double-strand break. As we saw in Chapter 5, double-strand breaks are often repaired by homologous recombination; here, the template chosen by the cell to repair the damage is


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Cell_Biology_Alberts. Cas9 protein makes a double-strand break. As we saw in Chapter 5, double-strand breaks are often repaired by homologous recombination; here, the template chosen by the cell to repair the damage is often the altered gene, which is introduced to ES cells by the experimenter. In this way, the normal gene can be selectively damaged by the CRISPR system and replaced at high efficiency by the experimentally altered gene.


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Cell_Biology_Alberts. The CRISPR system has a variety of other uses. Its particular power lies with its ability to target Cas9 to thousands of different positions across a genome through the simple rules of complementary base-pairing. Thus, if a catalytically inactive Cas9 protein is fused to a transcription activator or repressor, it is possible, in principle, to turn any gene on or off (Figure 8–55C and D).


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Cell_Biology_Alberts. Figure 8–54 Transgenic mice engineered to express a mutant DNA helicase show premature aging. The helicase, encoded by the Xpd gene, is involved in both transcription and DNA repair. Compared with a wild-type mouse of the same age (A), a transgenic mouse that expresses a defective version of Xpd (B) exhibits many of the symptoms of premature aging, including osteoporosis, emaciation, early graying, infertility, and reduced life-span. The mutation in Xpd used here impairs the activity of the helicase and mimics a mutation that in humans causes trichothiodystrophy, a disorder characterized by brittle hair, skeletal abnormalities, and a very reduced life expectancy. These results indicate that an accumulation of DNA damage can contribute to the aging process in both humans and mice. (From J. de Boer et al., Science 296:1276–1279, 2002. With permission from AAAS.)


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Cell_Biology_Alberts. Figure 8–55 Use of CRISPR to study gene function in a wide variety of species. (A) The Cas9 protein (artificially expressed in the species of interest) binds to a guide RNA, designed by the experimenter and also expressed. The portion of RNA in light blue is needed for associations with Cas9; that in dark blue is specified by the experimenter to match a position on the genome. The only other requirement is that the adjacent genome sequence includes a short PAM (protospacer adjacent motif) that is needed for Cas9 to cleave. As described in Chapter 7, this sequence is how the CRISPR system in bacteria distinguishes its own genome from that of invading viruses. (B) When directed to make double-strand breaks, the CRISPR system greatly improves the ability to replace an endogenous gene with an experimentally altered gene since the altered gene is used to “repair” the double-strand break (C, D). By using a mutant form of Cas9 that can no longer cleave DNA, Cas9 can be used to activate a


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Cell_Biology_Alberts. an experimentally altered gene since the altered gene is used to “repair” the double-strand break (C, D). By using a mutant form of Cas9 that can no longer cleave DNA, Cas9 can be used to activate a normally dormant gene (C) or turn off an actively expressed gene (D). (Adapted from P. Mali et al., Nat. Methods 10:957– 963, 2013. With permission from Macmillan Publishers Ltd.)


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Cell_Biology_Alberts. The CRISPR system has several advantages over other strategies for experimentally manipulating gene expression. First, it is relatively easy for the experimenter to design the guide RNA: it simply follows standard base pairing convention. Second, the gene to be controlled does not have to be modified; the CRISPR strategy exploits DNA sequences already present in the genome. Third, numerous genes can be controlled simultaneously. Cas9 has to be expressed only once, but many guide RNAs can be expressed in the same cell; this strategy allows the experimenter to turn on or off a whole set of genes at once.


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Cell_Biology_Alberts. The export of the CRISPR system from bacteria to virtually all other experimental organisms (including mice, zebrafish, worms, flies, rice, and wheat) has revolutionized the study of gene function. Like the earlier discovery of restriction enzymes, this breakthrough came from scientists studying a fascinating phenomenon in bacteria without—at first—realizing the enormous impact these discoveries would have on all aspects of biology. Large Collections of Engineered Mutations Provide a Tool for Examining the Function of Every Gene in an Organism


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Cell_Biology_Alberts. Extensive collaborative efforts have produced comprehensive libraries of mutations in a variety of model organisms, including S. cerevisiae, C. elegans, Drosophila, Arabidopsis, and even the mouse. The ultimate aim in each case is to produce a collection of mutant strains in which every gene in the organism has been systematically deleted or altered in such a way that it can be conditionally disrupted. Collections of this type provide an invaluable resource for investigating gene function on a genomic scale. For example, a large collection of mutant organisms can be screened for a particular phenotype. Like the classic genetic approaches described earlier, this is one of the most powerful ways to identify the genes responsible for a particular phenotype. Unlike the classical genetic approach, however, the set of mutants is “pre-engineered,” so that there is no need to rely on chance events such as spontaneous mutations or transposon insertions. In addition, each of the individual


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Cell_Biology_Alberts. however, the set of mutants is “pre-engineered,” so that there is no need to rely on chance events such as spontaneous mutations or transposon insertions. In addition, each of the individual mutations within the collection is often engineered to contain a distinct molecular “barcode”—in the form of a unique DNA sequence—designed to make identification of the altered gene rapid and routine (Figure 8–56).


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Cell_Biology_Alberts. Figure 8–56 Making barcoded collections of mutant organisms. A deletion construct for use in yeast contains DNA sequences (red) homologous to each end of a target gene x, a selectable marker gene (blue), and a unique “barcode” sequence approximately 20 nucleotide pairs in length (green). This DNA is introduced into yeast cells, where it readily replaces the target gene by homologous recombination. Cells that carry a successful gene replacement are identified by expression of the selectable marker gene, typically a gene that provides resistance to a drug. By using a collection of such constructs, each specific for one gene, a library of yeast mutants was constructed containing a mutant for every gene. Essential genes cannot be studied this way, as their deletion from the genome causes the cells to die. In this case, the target gene is replaced by a version of the gene that can be regulated by the experimenter (see Figure 8–52). The gene can then be turned off and the effect of this can


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Cell_Biology_Alberts. to die. In this case, the target gene is replaced by a version of the gene that can be regulated by the experimenter (see Figure 8–52). The gene can then be turned off and the effect of this can be monitored before the cells die.


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Cell_Biology_Alberts. Figure 8–57 Genome-wide screens for fitness using a large pool of barcoded yeast deletion mutants. A large pool of yeast mutants, each with a different gene deleted and present in equal amounts, is grown under conditions selected by the experimenter. Some mutants (blue) grow normally, but others show reduced growth (orange and green) or no growth at all (red). The fitness of each mutant is experimentally determined in the following way. After the growth phase is completed, genomic DNA (isolated from the mixture of strains) is purified and the relative abundance of each mutant is determined by quantifying the level of the DNA barcode matched to each deletion. This can be done by sequencing the pooled genomic DNA or hybridizing it to microarrays (see Figure 8–64) that contain DNA oligonucleotides complementary to each barcode. In this way, the contribution of every gene to growth under the specified condition can be rapidly ascertained. This type of study has revealed that of the


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Cell_Biology_Alberts. complementary to each barcode. In this way, the contribution of every gene to growth under the specified condition can be rapidly ascertained. This type of study has revealed that of the approximately 6000 coding genes in yeast, only about 1000 are essential under standard growth conditions.


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Cell_Biology_Alberts. In S. cerevisiae, the task of generating a complete set of 6000 mutants, each missing only one gene, was accomplished several years ago. Because each mutant strain has an individual barcode sequence embedded in its genome, a large mixture of engineered strains can be grown under various selective test conditions— such as nutritional deprivation, a temperature shift, or the presence of various drugs—and the cells that survive can be rapidly identified by the unique sequence tags present in their genomes. By assessing how well each mutant in the mixture fares, one can begin to discern which genes are essential, useful, or irrelevant for growth under the various conditions (Figure 8–57).


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Cell_Biology_Alberts. The insights generated by examining mutant libraries can be considerable. For example, studies of an extensive collection of mutants in Mycoplasma genitalium—the organism with the smallest known genome—have identified the minimum complement of genes essential for cellular life. Growth under laboratory conditions requires about three-quarters of the 480 protein-coding genes in M. genitalium. Approximately 100 of these essential genes are of unknown function, which suggests that a surprising number of the basic molecular mechanisms that underlie life have yet to be discovered.


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Cell_Biology_Alberts. Collections of mutant organisms are also available for many animal and plant species. For example, it is possible to “order,” by phone or email from a consortium of investigators, a deletion or insertion mutant for almost all coding genes in Drosophila. Likewise, a nearly complete set of mutants exists for the “model” plant Arabidopsis. And the adaptation of the CRISPR system for use in mice means that, in the near future, we can expect to be able to turn on or off—at will—each gene in the mouse genome. Although we are still ignorant about the function of most genes in most organisms, these technologies allow an exploration of gene function on a scale that was unimaginable a decade ago. RNA Interference Is a Simple and Rapid Way to Test Gene Function


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Cell_Biology_Alberts. RNA Interference Is a Simple and Rapid Way to Test Gene Function Although knocking out (or conditionally expressing) a gene in an organism and studying the consequences is the most powerful approach for understanding the functions of the gene, RNA interference (RNAi, for short), is an alternative, particularly convenient approach. As discussed in Chapter 7, this method exploits a natural mechanism used in many plants, animals, and fungi to protect themselves against viruses and transposable elements. The technique introduces into a cell or organism a double-stranded RNA molecule whose nucleotide sequence matches that of part of the gene to be inactivated. After the RNA is processed, it hybridizes with the target-gene RNA (either mRNA or noncoding RNA) and reduces its expression by the mechanisms shown in Figure 7–75.


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Cell_Biology_Alberts. RNAi is frequently used to inactivate genes in Drosophila and mammalian cell culture lines. Indeed, a set of 15,000 Drosophila RNAi molecules (one for every coding gene) allows researchers, in several months, to test the role of every fly gene in any process that can be monitored using cultured cells. RNAi has also been widely used to study gene function in whole organisms, including the nematode pool of barcoded yeast mutants, each deleted for a different gene E. coli, expressing double-stranded RNA, eaten by worm


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Cell_Biology_Alberts. E. coli, expressing double-stranded RNA, eaten by worm Figure 8–58 Gene function can be tested by RNA interference. (A) Double-stranded RNA (dsRNA) can be introduced into C. elegans by (1) feeding the worms E. coli that express the dsRNA or (2) injecting the dsRNA directly into the animal’s gut. (B) In a wild-type worm embryo, the egg and sperm pronuclei (red arrowheads) come together in the posterior half of the embryo shortly after fertilization. (C) In an embryo in which a particular gene has been inactivated by RNAi, the pronuclei fail to migrate. This experiment revealed an important but previously unknown function of this gene in embryonic development. (B and C, from P. Gönczy et al., Nature 408:331–336, 2000. With permission from Macmillan Publishers Ltd.)


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Cell_Biology_Alberts. C. elegans. When working with worms, introducing the double-stranded RNA is quite simple: the RNA can be injected directly into the intestine of the animal, or the worm can be fed with E. coli engineered to produce the RNA (Figure 8–58). The RNA is amplified (see p. 431) and distributed throughout the body of the worm, where it inhibits expression of the target gene in different tissue types. RNAi is being used to help in assigning functions to the entire complement of worm genes (Figure 8–59).


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Cell_Biology_Alberts. A related technique has also been applied to mice. In this case, the RNAi molecules are not injected or fed to the mouse; rather, recombinant DNA techniques are used to make transgenic animals that express the RNAi under the control of an inducible promoter. Often this is a specially designed RNA that can fold back on itself and, through base-pairing, produce a double-stranded region that is recognized by the RNAi machinery. In the simplest cases, the process inactivates only the genes that exactly match the RNAi sequence. Depending on the inducible promoter used, the RNAi can be produced only in a specified tissue or only at a particular time in development, allowing the functions of the target genes to be analyzed in elaborate detail. RNAi has made reverse genetics simple and efficient in many organisms, but it has several potential limitations compared with true genetic knockouts. For C. elegans each well contains WORMS INGEST E. coli; RESULTING PHENOTYPES RECORDED AND ANALYZED


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Cell_Biology_Alberts. C. elegans each well contains WORMS INGEST E. coli; RESULTING PHENOTYPES RECORDED AND ANALYZED E. coli expressing a different dsRNA ADD TO WELLS IN PLATE 96-well plate


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Cell_Biology_Alberts. Figure 8–59 RNA interference provides a convenient method for conducting genome-wide genetic screens. In this experiment, each well in this 96-well plate is filled with E. coli that produce a different double-stranded RNA. Each interfering RNA matches the nucleotide sequence of a single C. elegans gene, thereby inactivating it. About 10 worms are added to each well, where they ingest the genetically modified bacteria. The plate is incubated for several days, which gives the RNAs time to inactivate their target genes—and the worms time to grow, mate, and produce offspring. The plate is then examined in a microscope, which can be controlled robotically, to screen for genes that affect the worms’ ability to survive, reproduce, develop, and behave. Shown here are normal worms alongside worms that show an impaired ability to reproduce due to inactivation of a particular “fertility” gene. (From B. Lehner et al., Nat. Genet. 38:896–903, 2006. With permission from Macmillan Publishers Ltd.)


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Cell_Biology_Alberts. that show an impaired ability to reproduce due to inactivation of a particular “fertility” gene. (From B. Lehner et al., Nat. Genet. 38:896–903, 2006. With permission from Macmillan Publishers Ltd.) unknown reasons, RNAi does not efficiently inactivate all genes. Moreover, within whole organisms, certain tissues may be resistant to the action of RNAi (for example, neurons in nematodes). Another problem arises because many organisms contain large gene families, the members of which exhibit sequence similarity. RNAi therefore sometimes produces “off-target” effects, inactivating related genes in addition to the targeted gene. One strategy to avoid such problems is to use multiple small RNA molecules matched to different regions of the same gene. Ultimately, the results of any RNAi experiment must be viewed as a strong clue to, but not necessarily a proof of, normal gene function.


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Cell_Biology_Alberts. In the preceding section, we discussed how genetic approaches can be used to assess a gene’s function in cultured cells or, even better, in the intact organism. Although this information is crucial to understanding gene function, it does not generally reveal the molecular mechanisms through which the gene product works in the cell. For example, genetics on its own rarely tells us all the places in the organism where the gene is expressed, or how its expression is controlled. It does not necessarily reveal whether the gene acts in the nucleus, the cytosol, on the cell surface, or in one of the numerous other compartments of the cell. And it does not reveal how a gene product might change its location or its expression pattern when the external environment of the cell changes. Key insights into gene function can be obtained by simply observing when and where a gene is expressed. A variety of approaches, most involving some form of genetic engineering, can easily provide this critical


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Cell_Biology_Alberts. into gene function can be obtained by simply observing when and where a gene is expressed. A variety of approaches, most involving some form of genetic engineering, can easily provide this critical information.


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Cell_Biology_Alberts. As discussed in detail in Chapter 7, cis-regulatory DNA sequences, located upstream or downstream of the coding region, control gene transcription. These regulatory sequences, which determine precisely when and where the gene is expressed, can be easily studied by placing a reporter gene under their control and introducing these recombinant DNA molecules into cells (Figure 8–60). In this way, the normal expression pattern of a gene can be determined, as well as 123 start site for RNA pattern of normal gene X DNA sequences that determine the expression of gene X cells 123 pattern of reporter gene Y expression CONCLUSIONS —cis-regulatory sequence 3 normally turns on gene X in cell B —cis-regulatory sequence 2 normally turns on gene X in cells D, E, and F —cis-regulatory sequence 1 normally turns off gene X in cell D


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Cell_Biology_Alberts. Figure 8–60 Using a reporter protein to determine the pattern of a gene’s expression. (A) In this example, the coding sequence for protein X is replaced by the coding sequence for reporter protein Y. The expression patterns for X and Y are the same. (B) Various fragments of DNA containing candidate cis-regulatory sequences are added in combinations to produce test DNA molecules encoding reporter gene Y. These recombinant DNA molecules are then tested for expression after introducing them into a variety of different types of mammalian cells. The results are summarized in (C). For experiments in eukaryotic cells, two commonly used reporter proteins are the enzyme β-galactosidase (β-gal) (see Figure 7–28) and green fluorescent protein (GFP) (see Figure 9–22). the contribution of individual cis-regulatory sequences in establishing this pattern (see also Figure 7–29).


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Cell_Biology_Alberts. the contribution of individual cis-regulatory sequences in establishing this pattern (see also Figure 7–29). Reporter genes also allow any protein to be tracked over time in living cells. Here, the reporter gene typically encodes a fluorescent protein, often green fluorescent protein (GFP), the molecule that gives luminescent jellyfish their greenish glow. The GFP is simply attached—in the coding frame—to the protein-coding gene of interest. The resulting GFP fusion protein often behaves in the same way the normal protein does and its location can be monitored by fluorescence microscopy, a topic that is discussed in the next chapter (see Figure 9–25). GFP fusion has become a standard strategy for tracking not only the location but also the movement of specific proteins in living cells. In addition, the use of multiple GFP variants that fluoresce at different wavelengths can provide insights into how different cells interact in a living tissue (Figure 8–61).


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Cell_Biology_Alberts. In situ Hybridization Can Reveal the Location of mRNAs and Noncoding RNAs


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Cell_Biology_Alberts. It is also possible to directly observe the time and place that an RNA product of a gene is expressed using in situ hybridization. For protein-coding genes, this strategy often provides the same general information as the reporter gene approaches described above; however, it is crucial for genes whose final product is RNA rather than protein. We encountered in situ hybridization earlier in the chapter (see Figure 8–34); it relies on the basic principles of nucleic acid hybridization. Typically, tissues are gently fixed so that their RNA is retained in an exposed form that can hybridize with a labeled complementary DNA or RNA probe. In this way, the patterns of differential gene expression can be observed in tissues, and the location of specific RNAs can be determined (Figure 8–62). An advantage of in situ hybridization over other approaches is that genetic engineering is not required. Thus, it is often simpler and faster and can be used for genetically intractable species.


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Cell_Biology_Alberts. Expression of Individual Genes Can Be Measured Using Quantitative RT-PCR Although reporter genes and in situ hybridization accurately reveal patterns of gene expression, they are not the most powerful methods for quantifying amounts of individual RNAs in cells. We have seen that RNA sequencing can provide information about the relative abundance of different RNA molecules (see Figure 7–3). Here, the number of “sequence reads” (short bits of nucleotide sequence) is proportional to the abundance of the RNA species. But this method is limited to RNAs