S

BamHI site

E. coli cells. With certain refinements of the PCR, DNA segments >10 kb in length can be amplified and cloned in this way.

Note that this method does not involve cloning of large numbers of restriction fragments derived from genomic DNA and their subsequent screening to identify the specific fragment of interest. In effect, the PCR method inverts this traditional approach and thus avoids its most tedious aspects. The PCR method is useful for isolating gene sequences to be manipulated in a variety of useful ways described later. In addition the PCR method can be used to isolate gene sequences from mutant organisms to determine how they differ from the wild-type.

Preparation of Probes Earlier we discussed how oligonu-cleotide probes for hybridization assays can be chemically synthesized. Preparation of such probes by PCR amplification requires chemical synthesis of only two relatively short primers corresponding to the two ends of the target sequence. The starting sample for PCR amplification of the target sequence can be a preparation of genomic DNA. Alternatively, if the target sequence corresponds to a mature mRNA sequence, a complete set of cellular cDNAs synthesized from the total cellular mRNA using reverse transcrip-tase or obtained by pooling cDNA from all the clones in a X cDNA library can be used as a source of template DNA. To generate a radiolabeled product from PCR, 32P-labeled dNTPs are included during the last several amplification cycles. Because probes prepared by PCR are relatively long and have many radioactive 32P atoms incorporated into them, these probes usually give a stronger and more specific signal than chemically synthesized probes.

Tagging of Genes by Insertion Mutations Another useful application of the PCR is to amplify a "tagged" gene from the genomic DNA of a mutant strain. This approach is a sim

▲ EXPERIMENTAL FIGURE 9-26 Southern blot technique can detect a specific DNA fragment in a complex mixture of restriction fragments. The diagram depicts three different restriction fragments in the gel, but the procedure can be applied to a mixture of millions of DNA fragments. Only fragments that pler method for identifying genes associated with a particular mutant phenotype than screening of a library by functional complementation (see Figure 9-20).

The key to this use of PCR is the ability to produce mutations by insertion of a known DNA sequence into the genome of an experimental organism. Such insertion mutations can be generated by use of mobile DNA elements, which can move (or transpose) from one chromosomal site to another. As discussed in more detail in Chapter 10, these DNA sequences occur naturally in the genomes of most organisms and may give rise to loss-of-function mutations if they transpose into a protein-coding region.

For example, researchers have modified a Drosophila mobile DNA element, known as the P element, to optimize its use in the experimental generation of insertion mutations. Once it has been demonstrated that insertion of a P element causes a mutation with an interesting phenotype, the genomic sequences adjacent to the insertion site can be amplified by a variation of the standard PCR protocol that uses synthetic primers complementary to the known P-element sequence but that allows unknown neighboring sequences to be amplified. Again, this approach avoids the cloning of large numbers of DNA fragments and their screening to detect a cloned DNA corresponding to a mutated gene of interest.

Similar methods have been applied to other organisms for which insertion mutations can be generated using either mobile DNA elements or viruses with sequenced genomes that can insert randomly into the genome.

Blotting Techniques Permit Detection of Specific DNA Fragments and mRNAs with DNA Probes

Two very sensitive methods for detecting a particular DNA or RNA sequence within a complex mixture combine separation by gel electrophoresis and hybridization with a complementary radiolabeled DNA probe. We will encounter hybridize to a labeled probe will give a signal on an autoradiogram. A similar technique called Northern blotting detects specific mRNAs within a mixture. [See E. M. Southern, 1975, J. Mol. Biol. 98:508.]

references to both these techniques, which have numerous applications, in other chapters.

Southern Blotting The first blotting technique to be devised is known as Southern blotting after its originator E. M. Southern. This technique is capable of detecting a single specific restriction fragment in the highly complex mixture of fragments produced by cleavage of the entire human genome with a restriction enzyme. In such a complex mixture, many fragments will have the same or nearly the same length and thus migrate together during electrophoresis. Even though all the fragments are not separated completely by gel elec-trophoresis, an individual fragment within one of the bands can be identified by hybridization to a specific DNA probe. To accomplish this, the restriction fragments present in the gel are denatured with alkali and transferred onto a nitrocellulose filter or nylon membrane by blotting (Figure 9-26). This procedure preserves the distribution of the fragments in the gel, creating a replica of the gel on the filter, much like the replica filter produced from clones in a \ library. (The blot is used because probes do not readily diffuse into the original gel.) The filter then is incubated under hybridization conditions with a specific radiolabeled DNA probe, which usually is generated from a cloned restriction frag-

ment. The DNA restriction fragment that is complementary to the probe hybridizes, and its location on the filter can be revealed by autoradiography.

Northern Blotting One of the most basic ways to characterize a cloned gene is to determine when and where in an organism the gene is expressed. Expression of a particular gene can be followed by assaying for the corresponding mRNA by Northern blotting, named, in a play on words, after the related method of Southern blotting. An RNA sample, often the total cellular RNA, is denatured by treatment with an agent such as formaldehyde that disrupts the hydrogen bonds between base pairs, ensuring that all the RNA molecules have an unfolded, linear conformation. The individual RNAs are separated according to size by gel elec-trophoresis and transferred to a nitrocellulose filter to which the extended denatured RNAs adhere. As in Southern blotting, the filter then is exposed to a labeled DNA probe that is complementary to the gene of interest; finally, the labeled filter is subjected to autoradiography. Because the amount of a specific RNA in a sample can be estimated from a Northern blot, the procedure is widely used to compare the amounts of a particular mRNA in cells under different conditions (Figure 9-27).

UN 48 h

96 h

UN 48 h

96 h

/8-globin mRNA

▲ EXPERIMENTAL FIGURE 9-27 Northern blot analysis reveals increased expression of p-globin mRNA in differentiated erythroleukemia cells. The total mRNA in extracts of erythroleukemia cells that were growing but uninduced and in cells induced to stop growing and allowed to differentiate for 48 hours or 96 hours was analyzed by Northern blotting for (3-globin mRNA. The density of a band is proportional to the amount of mRNA present. The p-globin mRNA is barely detectable in uninduced cells (UN lane) but increases more than 1000-fold by 96 hours after differentiation is induced. [Courtesy of L. Kole.]

E. coli Expression Systems Can Produce Large Quantities of Proteins from Cloned Genes a Many protein hormones and other signaling or regulatory proteins are normally expressed at very low concentrations, precluding their isolation and purification in large quantities by standard biochemical techniques. Widespread therapeutic use of such proteins, as well as basic research on their structure and functions, depends on efficient procedures for producing them in large amounts at reasonable cost. Recombinant DNA techniques that turn E. coli cells into factories for synthesizing low-abundance proteins now are used to commercially produce factor VIII (a blood-clotting factor), granulocyte colony-stimulating factor (G-CSF), insulin, growth hormone, and other human proteins with therapeutic uses. For example, G-CSF stimulates the production of granulocytes, the phagocytic white blood cells critical to defense against bacterial infections. Administration of G-CSF to cancer patients helps offset the reduction in gran-ulocyte production caused by chemotherapeutic agents, thereby protecting patients against serious infection while they are receiving chemotherapy. I

The first step in producing large amounts of a low-abundance protein is to obtain a cDNA clone encoding the full-length protein by methods discussed previously. The second step is to engineer plasmid vectors that will express large amounts of the encoded protein when it is inserted into E. coli cells. The key to designing such expression vectors is

▲ EXPERIMENTAL FIGURE 9-28 Some eukaryotic proteins can be produced in E. coli cells from plasmid vectors containing the lac promoter. (a) The plasmid expression vector contains a fragment of the E. coli chromosome containing the lac promoter and the neighboring lacZ gene. In the presence of the lactose analog IPTG, RNA polymerase normally transcribes the lacZ gene, producing lacZ mRNA, which is translated into the encoded protein, p-galactosidase. (b) The lacZ gene can be cut out of the expression vector with restriction enzymes and replaced by a cloned cDNA, in this case one encoding granulocyte colony-stimulating factor (G-CSF). When the resulting plasmid is transformed into E. coli cells, addition of IPTG and subsequent transcription from the lac promoter produce G-CSF mRNA, which is translated into G-CSF protein.

To aid in purification of a eukaryotic protein produced in an E. coli expression system, researchers often modify the cDNA encoding the recombinant protein to facilitate its separation from endogenous E. coli proteins. A commonly used modification of this type is to add a short nucleotide sequence to the end of the cDNA, so that the expressed protein will have six histidine residues at the C-terminus. Proteins modified in this way bind tightly to an affinity matrix that contains chelated nickel atoms, whereas most E. coli proteins will not bind to such a matrix. The bound proteins can be released from the nickel atoms by decreasing the pH of the surrounding medium. In most cases, this procedure yields a pure recombinant protein that is functional, since addition of short amino acid sequences to either the C-terminus or the N-terminus of a protein usually does not interfere with the protein's biochemical activity.

Plasmid Expression Vectors Can Be Designed for Use in Animal Cells

One disadvantage of bacterial expression systems is that many eukaryotic proteins undergo various modifications (e.g., glycosylation, hydroxylation) after their synthesis on ribosomes (Chapter 3). These post-translational modifications generally are required for a protein's normal cellular function, but they cannot be introduced by E. coli cells, which lack the necessary enzymes. To get around this limitation, cloned genes are introduced into cultured animal cells, a process called transfection. Two common methods for transfecting animal cells differ in whether the recombinant vector DNA is or is not integrated into the host-cell genomic DNA.

In both methods, cultured animal cells must be treated to facilitate their initial uptake of a recombinant plasmid vector. This can be done by exposing cells to a preparation of lipids that penetrate the plasma membrane, increasing its permeability to DNA. Alternatively, subjecting cells to a brief electric shock of several thousand volts, a technique known as electroporation, makes them transiently permeable to DNA. Usually the plasmid DNA is added in sufficient concentration to ensure that a large proportion of the cultured cells will receive at least one copy of the plasmid DNA.

inclusion of a promoter, a DNA sequence from which transcription of the cDNA can begin. Consider, for example, the relatively simple system for expressing G-CSF shown in Figure 9-28. In this case, G-CSF is expressed in E. coli transformed with plasmid vectors that contain the lac promoter adjacent to the cloned cDNA encoding G-CSF. Transcription from the lac promoter occurs at high rates only when lactose, or a lactose analog such as isopropylthiogalactoside (IPTG), is added to the culture medium. Even larger quantities of a desired protein can be produced in more complicated E. coli expression systems.

Transient Transfection The simplest of the two expression methods, called transient transfection, employs a vector similar to the yeast shuttle vectors described previously. For use in mammalian cells, plasmid vectors are engineered also to carry an origin of replication derived from a virus that infects mammalian cells, a strong promoter recognized by mammalian RNA polymerase, and the cloned cDNA encoding the protein to be expressed adjacent to the promoter (Figure 9-29a). Once such a plasmid vector enters a mammalian cell, the viral origin of replication allows it to replicate efficiently, generating numerous plasmids from which the protein is ex-

(a) Transient transfection

Promoter cDNA

cDNA

Promoter

Viral origin of replication

Viral origin of replication

Transfect cultured cells by lipid treatment or electroporation

Transfect cultured cells by lipid treatment or electroporation

Protein is expressed from cDNA in plasmid DNA

Protein is expressed from cDNA in plasmid DNA

(b) Stable transfection (transformation) cDNA

Promoter

Promoter

Transfect cultured cells by lipid treatment or electroporation

Select for G-418 resistance

G-418-resistant clones

Transfect cultured cells by lipid treatment or electroporation

Select for G-418 resistance

G-418-resistant clones

Protein is expressed from cDNA integrated into host chromosome

pressed. However, during cell division such plasmids are not faithfully segregated into both daughter cells and in time a substantial fraction of the cells in a culture will not contain a plasmid, hence the name transient transfection.

Stable Transfection (Transformation) If an introduced vector integrates into the genome of the host cell, the genome is permanently altered and the cell is said to be transformed. Integration most likely is accomplished by mammalian enzymes that function normally in DNA repair and recombination. Because integration is a rare event, plasmid expression vectors designed to transform animal cells must carry a se

M EXPERIMENTAL FIGURE 9-29 Transient and stable transfection with specially designed plasmid vectors permit expression of cloned genes in cultured animal cells. Both methods employ plasmid vectors that contain the usual elements—ORI, selectable marker (e.g., amp), and polylinker— that permit propagation in E. coli and insertion of a cloned cDNA with an adjacent animal promoter. For simplicity, these elements are not depicted. (a) In transient transfection, the plasmid vector contains an origin of replication for a virus that can replicate in the cultured animal cells. Since the vector is not incorporated into the genome of the cultured cells, production of the cDNA-encoded protein continues only for a limited time. (b) In stable transfection, the vector carries a selectable marker such as ned, which confers resistance to G-418. The relatively few transfected animal cells that integrate the exogenous DNA into their genomes are selected on medium containing G-418. These stably transfected, or transformed, cells will continue to produce the cDNA-encoded protein as long as the culture is maintained. See the text for discussion.

lectable marker in order to identify the small fraction of cells that integrate the plasmid DNA. A commonly used selectable marker is the gene for neomycin phosphotransferase (designated neor), which confers resistance to a toxic compound chemically related to neomycin known as G-418. The basic procedure for expressing a cloned cDNA by stable transfec-tion is outlined in Figure 9-29b. Only those cells that have integrated the expression vector into the host chromosome will survive and give rise to a clone in the presence of a high concentration of G-418. Because integration occurs at random sites in the genome, individual transformed clones resistant to G-418 will differ in their rates of transcribing the inserted cDNA. Therefore, the stable transfectants usually are screened to identify those that produce the protein of interest at the highest levels.

Epitope Tagging In addition to their use in producing proteins that are modified after translation, eukaryotic expression vectors provide an easy way to study the intracellular localization of eukaryotic proteins. In this method, a cloned cDNA is modified by fusing it to a short DNA sequence encoding an amino acid sequence recognized by a known monoclonal antibody. Such a short peptide that is bound by an antibody is called an epitope; hence this method is known as epitope tagging. After transfection with a plasmid expression vector containing the fused cDNA, the expressed epitope-tagged form of the protein can be detected by immunofluorescence labeling of the cells with the monoclonal antibody specific for the epitope. Figure 9-30 illustrates the use of this method to localize AP1 adapter proteins, which participate in formation of clathrin-coated vesicles involved in intracellular protein trafficking (Chapter 17). Epitope tagging of a protein so it is detectable with an available monoclonal antibody obviates the time-consuming task of producing a new monoclonal antibody specific for the natural protein.

AP1 Furin Merged

▲ EXPERIMENTAL FIGURE 9-30 Epitope tagging facilitates cellular localization of proteins expressed from cloned genes. In this experiment, the cloned cDNA encoding one subunit of the API adapter protein was modified by addition of a sequence encoding an epitope for a known monoclonal antibody. Plasmid expression vectors, similar to those shown in Figure 9-29, were constructed to contain the epitope-tagged API cDNA. After cells were transfected and allowed to express the epitope-tagged version of the API protein, they were fixed and labeled with monoclonal antibody to the epitope and with antibody to furin, a marker protein for the late Golgi and endosomal membranes. Addition of a green fluorescently labeled secondary antibody specific for the anti-epitope antibody visualized the API protein (left). Another secondary antibody with a different (red) fluorescent signal was used to visualize furin (center). The colocalization of epitope-tagged API and furin to the same intracellular compartment is evident when the two fluorescent signals are merged (right). [Courtesy of Ira Mellman, Yale University School of Medicine.]

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