Overview of Receptor Cloning - Wiley Online Library

Overview of Receptor Cloning Receptors are key elements in signal transduction pathways (see UNITS 1.1, 1.2, 1.3, & 2.1). They respond to discrete molecules that exist in the cellular environment by altering their inactive configurations to active forms. Initial attempts to isolate and clone receptors began in the 1970s, when investigators used painstaking protein purification techniques to solubilize and purify receptors from kilogram quantities of whole tissue. These techniques were often ineffective, because receptors are associated with membranes, which makes them difficult to purify and reconstitute. Nevertheless, these methods were successfully used to isolate the insulin receptor from rat liver and adipocyte membranes (Cuatrecasas, 1972), the epidermal growth factor receptor from A431 cells (Ullrich et al., 1984), and the nicotinic acetylcholine receptor from the electric organ of the ray Torpedo californica (Giraudat et al., 1982). The importance of these first cloning efforts can not be overemphasized. Thus, this unit presents cloning by protein purification as the first cloning strategy. The early, more tedious procedures have broadened knowledge about receptors and established a strong foundation that has enabled the utilization of the newer approaches discussed here, including: homology cloning by screening with oligonucleotides, DNA fragments or the products of direct polymerase chain reaction (PCR); expression cloning using function, ligand binding, and antibody recognition; and differential display techniques. Historically, receptors have been grouped according to their effector systems; more recently, they have been categorized by their structural and genetic similarities. According to the original classifications, receptors fall into four main families: steroid receptors, growth factor receptors, ionotropic receptors (i.e., ligand-gated ion channels), and metabotropic receptors (i.e., G protein–coupled receptors). Because of the large numbers of receptors, including receptor subtypes (which respond to the same agonist yet differ in their signal transduction) and isoforms (receptor subtypes that arise from differential mRNA processing of a single gene), these groups are now more commonly referred to as “superfamilies.” Steroid receptors exert their effects after they interact with specific steroid hormones. Once activated, the receptors migrate into the nucleus and bind to chromatin. Receptor affin-

UNIT 6.1

ity and target specificity for DNA are greatly enhanced by agonist binding. However, even the “inactive” receptor appears to shuttle from the cell surface to the nucleus, where it binds to random DNA sequences. Molecular biological techniques have allowed examination of the interaction of these receptors with their target DNA sequences. Footprinting analysis, a technique used to identify DNA domains that interact with specific proteins, has revealed that these receptor-binding regions are regulatory sequences that affect transcription of the gene(s) (Cato et al., 1988). Growth factor receptors are responsive to proteins and glycoproteins. As the name implies, these receptors stimulate cell division and differentiation, and increase hormone production (Fisher and Lakshmanan, 1990). Growth factor receptors have intrinsic tyrosine kinase activity, which permits them to catalyze the phosphorylation of distinct tyrosine residues in cellular target proteins (Feige and Chambaz, 1987). Depending on the protein targeted, phosphorylation can lead to an up- or downregulation of additional genes. Ionotropic receptors establish a unique configuration as they traverse the cell membrane. They are selectively activated by a variety of neurotransmitters including γ-amino butyric acid (GABA), glutamic acid, acetylcholine, ATP, and serotonin. On ligand binding, ionotropic receptors undergo a structural transformation that opens a channel through the membrane. This ligand-gated ion channel selectively allows the rapid flow of a particular ion through the cell membrane, an event that alters the polarity of the cell. In the case of neurons, this ion flux can cause excitation or inhibition of the cell. Cloning of these receptors has shown that they typically possess four membrane-spanning domains, a long extracellular N-terminal domain that contains multiple glycosylation sites, and a large hydrophilic loop between the third and fourth transmembrane domains (Fig. 6.1.1A; reviewed in Numa, 1989; see UNIT 1.1). The ion channel formed by these receptors is typically composed of five subunits and is probably not homomeric (Cooper et al., 1991; Brose et al., 1993). In other words, a combination of different subunits is normally required to produce a functional receptor in vivo. However some ionotropic receptors form functional Molecular Biology

Contributed by Norman Nash Current Protocols in Pharmacology (1998) 6.1.1-6.1.12 Copyright © 1998 by John Wiley & Sons, Inc.

6.1.1

A +

NH3

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Figure 6.1.1 (A) Schematic drawing of an ionotropic receptor subunit shows the extracellular Nand C-terminal domains and a large extracellular loop between the third and fourth transmembrane domains. Transmembrane domains are represented by stacked discs. (B) Schematic drawing of a G protein–coupled receptor shows the extracellular N-terminal domain, the intracellular C-terminal domain, and a large intracellular loop between the fifth and sixth transmembrane domains.

Overview of Receptor Cloning

homomeric receptors in certain expression systems. For example, both the α4 subunit of the neuronal nicotinic receptor and the R1 subunit of the NMDA receptor behave as functional receptors when expressed alone in Xenopus oocytes, but neither possesses all of the properties exhibited by native receptors in tissue preparations (Boulter et al., 1987; Moriyoshi et al., 1991). The final, and perhaps largest, group of receptors is the metabotropic receptors. Members of this superfamily are coupled to G proteins that activate a second messenger system within the cell. This class of receptors may be activated by either small molecules (e.g., acetylcholine) or peptides (e.g., bradykinin). The cellular response to activation of metabotropic receptors is determined by the G protein and can include inhibition or activation of adenylate cyclase, breakdown of phosphoinositides, and modulation of potassium channels (Gilman, 1984). Structurally, members of this superfamily of receptors contain seven transmembrane domains, an extracellular N-terminal domain, and a large intracellular loop between the fifth and sixth transmembrane domains (Fig. 6.1.1B).

Mutagenesis studies of various members of this family have shown that the C-terminal tail and the loop region interact with the G protein. It is probably the loop domain that specifies which type of G protein couples to a particular receptor (Wess et al., 1990).

CLONING STRATEGIES Cloning by Protein Purification Although this method is now less frequently employed, it is still effective and has laid the cornerstone for more contemporary techniques. To purify and clone receptors from whole tissue, it is first important to obtain either starting material rich in the desired receptor or a large quantity of the tissue. Traditionally, researchers acquired kilogram quantities of material from local slaughterhouses or produced liters of cultured cell lines expressing high levels of the target protein. After obtaining the starting material, several rounds of purification are required to isolate a pure sample of receptor. The receptor must first be solubilized from the tissue, typically with detergents (either ionic or nonionic), and then purified using various chromatographic separation techniques, including

6.1.2 Current Protocols in Pharmacology

ion-exchange, molecular size exclusion, and affinity (antibody- or ligand-binding) chromatographies. Depending on the protein, any or all of these methods might be required. The aim is to obtain microgram quantities of the desired protein, although with modern techniques, even nanogram quantities may be sufficient. The purified protein is then subjected to N-terminal amino acid sequencing, which provides information that is used to design degenerate oligonucleotide probes and peptides that can be used to generate specific antibodies. If the Nterminus is blocked (i.e., proteins expressed in E. coli and yeast are formylated at their initiating methionine and many eukaryotic proteins are acetylated at their amino termini), proteolytic digestion of the protein will be required. This treatment yields several smaller peptides that can be separated by HPLC or gel electrophoresis and subsequently sequenced. A degenerate oligonucleotide probe is designed from the known amino acid sequence of the peptide and is used for screening a cDNA or genomic library that can be prepared by the investigator or obtained commercially. Detailed protocols for preparing cDNA and genomic libraries can be found in Current Protocols in Molecular Biology Chapters 5 and 6 (Ausubel et al., 1998). Because the exact codon for each amino acid may vary, a degenerate oligonucleotide makes allowances for these differences (e.g., proline can be encoded by the codons CCC, CCA, CCG, and CCT). Therefore, a degenerate oligonucleotide would contain the sequence CCN, where N is a mixture of all four bases. Inosine, a metabolite in the purine synthesis pathway, may also be substituted for N, as it weakly binds to all four bases. Several companies, including Stratagene and Clontech, offer DNA libraries from various species and tissues that have been used to clone receptors from each of the four families. These libraries have been prepared by isolating mRNA (in the case of cDNA libraries) or genomic DNA from cells and subcloning this material into bacteriophage, plasmids, or cosmids. Because the subcloned material represents all of the information contained within the cell, it is called a library. Genomic libraries are useful because they retain all of the information encoded in a gene, including both exons and introns (protein coding and noncoding sequences, respectively). The cDNA library is more useful for obtaining only the information expressed in a particular cell type or tissue; that is, the information that usually results in the

translation of a protein product, in this case a functional receptor. A detailed explanation of library screening is presented in Current Protocols in Molecular Biology Chapter 6 (Ausubel et al., 1998). Briefly, the library is plated onto 150-mm dishes at a density of 40,000 to 50,000 plaque forming units (pfu) per plate for a bacteriophage library or 20,000 to 25,000 colonies per plate for a cosmid or plasmid library. Sufficient numbers of plates are used to ensure that at least one representation of each cDNA in the library is achieved. After allowing adequate time for growth, the plaques or colonies are directly transferred to a nylon or nitrocellulose membrane. A nylon membrane has the advantage of being more durable and more easily manipulated than a nitrocellulose membrane. The membranes are then treated with a solution containing NaOH to liberate and denature the DNA from the phage or colonies, and then neutralized with a Tris-buffered solution. The DNA can then be permanently fixed to the membrane by baking or cross-linking with UV irradiation. Hybridization of the oligonucleotide probe is accomplished by incubating the membranes in a solution containing blocking agents and the labeled oligonucleotide probe. Oligonucleotides can be labeled with radioactivity, digoxigenin, fluorescein, or biotin conjugated to the nucleotides (Fig. 6.1.2). The choice of label must be decided by each individual investigator, as there are advantages and disadvantages associated with each. All are comparably expensive when the cost of labeling, detection, and disposal are taken into consideration. However, 32P end-labeled oligonucleotide probes have been most successfully and consistently used for hybridization methodologies. Several companies market nonradioactive labeling and detection systems that are described as being as sensitive as radioactive detection; however, these kits are typically more technically challenging and can yield a higher background with more false positives. Thus, they require optimization by the individual user. The incubation is continued overnight to allow maximal binding of the oligonucleotide probe, after which the membrane is washed to remove all non-specifically bound probe. The membranes are then apposed to X-ray film in the case of radioactively labeled probes and exposed overnight at −80°C. In the case of fluorescein-, digoxigenin-, or biotin-labeled probes, the membranes are developed using antibodies and chemiluminescence, with sub-

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3′ 5′ multiple fragments generated; multiple labeled nucleotides incorporated

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Figure 6.1.2 Methods for labeling DNA. (A) A radiolabled phosphate group is added to the 5′ end of each DNA strand by T4 polynucleotide kinase. (B) Random primers, 6-10 bp in length, are annealed to denatured DNA. Label is incorporated by extending the annealed oligonucleotide primers with DNA polymerase in the presence of labeled and unlabeled dNTPs. (C) Multiple labeled dNTPs are added to the 3′ end of a DNA strand by incubation with terminal deoxynucleotidyltransferase.


sequent apposition to X-ray film. After the potential positive signals have been detected, the films are aligned with the original plates and plaques or colonies that contain the original DNA sequence of interest are recovered. The plaque or colony of interest is transferred to a tube containing a Tris⋅Cl buffer (for plaques) or LB medium (for colonies). Two or three rounds of plating using diluted plaques or colonies (1:100 to 1:10,000 is a good range to try) are required to obtain a pure clonal isolate. DNA is then isolated from the plaque or colony and sequenced to verify that the original sequence used for the oligonucleotide probe is contained in the clone. This allows determina-

tion of the correct reading frame for the entire protein coding sequence. It is essential to verify that the sequence truly represents the original protein sought. Beginning the process with multiple peptide sequences over the entire length of the clone will help to ensure that the sequence obtained is correct, as codons representative of the various peptides should be identifiable within the DNA sequence. It is wise to make several different oligonucleotides in order to obtain sequencing information over the entire length of the clone. These oligonucleotides should generate sequences that overlap by 25 to 50 base pairs to ensure that no sequence information is lost. It is also


good practice to sequence in both directions (i.e., both strands of DNA), especially when sequencing a novel clone. This will minimize any errors that may result from poor sequencing and will help ensure that the deduced amino acid sequence is correct, particularly for long clones, which can be difficult to sequence. Compressions in DNA sequence readout can often occur, making it difficult to be certain of the exact number of base pairs (i.e., groups of three or more Gs and Cs can be difficult to resolve). It is also important to verify that the cDNA isolated encodes the correct target protein. This can be accomplished by expressing the clones in Xenopus oocytes or mammalian cells and demonstrating the expected functional properties of the receptor. To express the clone in oocytes, DNA is first isolated from the plaque or colony and transcribed in vitro (several suppliers market kits for this purpose). Approximately 50 ng of the resulting RNA is then injected into an oocyte. After allowing sufficient time for expression (typically 48 hr), the oocyte is tested for response to an agonist of the receptor. It is critical to include control oocytes that have only been injected with water or a control RNA to ensure that the response detected is solely due to expression of the injected test RNA. Expression can also be examined in mammalian cells transfected with the cloned receptor DNA. To express the clone in mammalian cells, it is first necessary to subclone the receptor cDNA into an appropriate mammalian expression vector (e.g., pcDNA3 from Invitrogen is commonly used) and then transfect the new construct into a cell line that normally does not express the particular receptor. Agonist, antagonist, or binding assays are then performed to determine whether the cloned DNA corresponds to the receptor of interest. Again, control transfections of vector alone are important to ensure that the responses detected are not a result of the transfection. High-efficiency transfection kits are available from numerous suppliers; kits are chosen based on which cell line is being transfected.

Cloning by Homology Now that many receptors have been cloned, it is practical to take advantage of the similarities between members of a receptor family for designing oligonucleotides to screen libraries. Most researchers now deposit the sequences of novel clones into one of several databases (e.g.,

GenBank, EMBL (European Molecular Biological Laboratories), and the DNA Database of Japan). Internet searches can easily be performed utilizing programs maintained by the National Institutes of Health (http://www.ncbi. nlm.nih.gov). Searching these databases, it is evident that members of the various receptor superfamilies not only have functional similarities, but also share structural and sequence similarities. For example, the various opioid receptor subtypes are ∼60% identical in their amino acid sequences. When their transmembrane domains are examined, the identity increases to 73% to 76%. The homology with other members of the superfamily is also significant, as the amino acid sequence of the somatostatin receptor is 34% to 42% identical to that of the opioid receptors (Minami and Satoh, 1995). With this information, it is possible to design oligonucleotides that will allow cloning of any family (or superfamily) member by selecting amino acid sequences that are conserved between the most closely related receptors known and making degenerate oligonucleotides to these sequences, which are most often in transmembrane regions. To ensure that the maximum number of clones are identified in the library screening, it is wise to reduce the stringency of the wash conditions by lowering the temperature and increasing the salt concentration. This makes it possible to later incrementally raise the stringency conditions so that the hybridization clones can be grouped into pools of increasing complementarity to each of the oligonucleotide probes. Clones containing sequences that are exactly complementary to a probe will remain hybridized even under very high stringency conditions (65°C, 15 mM NaCl). Again, all clones must be sequenced and tested for function to determine which receptor is encoded by a particular cDNA. This strategy is most effective when attempting to isolate cDNAs encoding receptor homologs from different species. For example, if the clone of a receptor from rat is published, this information can quickly be exploited to design and synthesize probes for screening human libraries to isolate the human homolog. This technology has enabled the rapid expansion of clones in several receptor families, revealing that similarities between families also exist. Therefore, the broader term superfamily is now used to describe a group of structurally or functionally related families. Molecular Biology


PCR-Based Homology Cloning Polymerase chain reaction (PCR) is a technique used for amplifying small quantities of DNA. The reaction involves cycles of heat denaturing double-stranded template DNA, annealing sequence-specific primers to the template, and elongating the oligonucleotide primer with a DNA polymerase to produce a new complementary strand of DNA (Fig. 6.1.3; Elion, 1993; Kramer and Coen, 1995). When this technique was first employed, researchers manually transferred their reaction tubes between a water bath or block heater set to a temperature suitable for denaturing DNA, a second bath set to the annealing temperature, and a third bath set to a temperature that allowed optimal elongation by the DNA polymerase utilized. Originally, the Klenow fragment of E. coli DNA polymerase I was used because it lacked any 5′ to 3′ exonuclease activity but retained 3′ to 5′ exonuclease activity, which

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ensured high fidelity of nucleotide incorporation. The major problem that arose with early attempts to use the Klenow fragment was that significant numbers of nonspecifically primed sequences were also amplified (i.e., sequences that were not exactly complementary to the oligonucleotides, but contained some mismatches). This nonspecific amplification was a result of the relatively low temperature (37°C) required for elongation with Klenow fragment. The manual transfer of samples between baths of varying temperatures was also very awkward and resulted in extreme variability in the time it took to heat and cool samples to the desired temperatures. Two major developments in PCR technology have made it the powerful molecular biology tool it is today. The first was the substitution of heat-stable DNA polymerases (such as Taq, Pwo, or Pfu) for Klenow fragment. These enzymes allow the reaction temperatures needed for elongation to remain

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Figure 6.1.3 Amplification of DNA by PCR. (A) DNA template is denatured at high temperatures and then allowed to cool to an annealing temperature in the presence of excess specific primers and dNTPs. The temperature is then raised to the optimal temperature for DNA polymerase activity, allowing primer extension. (B) Subsequent rounds of PCR amplify the sequence between the two specific primers.


high (optimal temperature for Taq polymerase activity is 72°C), thereby reducing nonspecific amplification. The second was the invention of block heaters (thermal cyclers) that are able to heat and cool multiple samples in only a few seconds rather than several minutes. Both of these innovations have allowed PCR to become a very useful method for amplifying femtogram quantities of DNA into more workable nanogram and microgram amounts. Cloning receptors via PCR is the fastest and simplest method if the receptor sequence is already known. The time required to obtain a functional and sequence-verified clone can be as little as 2 weeks, but realistically is more like 1 to 2 months. This estimate includes the time needed to synthesize oligonucleotide primers, run the PCR reactions, subclone the PCR product into an appropriate vector for both expression and sequencing (this can be the same vector), verify the sequence to ensure that no mutations were introduced during PCR, and express the receptor in a eukaryotic expression system for functional testing. The most effective design for the PCR primers incorporates sequence (∼18 to 21 bases) specific for the 5′ or 3′ untranslated region and a restriction enzyme site that is not found within the receptor gene to facilitate subcloning into the sequencing vector. The choice of template for PCR can be a cDNA library, genomic DNA (if the receptor is known to lack introns), or RNA (either poly(A)+ mRNA or total RNA is suitable). If RNA is used, it is first necessary to perform a reverse transcription reaction to generate a cDNA strand suitable for use as a template, because the polymerases used in PCR require DNA as a template. The reverse transcription reaction is very simple to perform when following the protocols provided with most commercially available reverse transcriptases. (The Superscript II enzyme from Life Technologies is an excellent product and is highly recommended.) With both the template and primers in hand, it is very simple to set up the PCR reaction to amplify the sequence encoding the desired receptor. Typical reaction conditions include the following: 250 ng of each primer, 2.5 mM MgCl2, Tris/KCl buffer, pH 8.3 (usually supplied by manufacturer), DNA template (10 to 20 ng for cDNA, 100 ng for genomic DNA, or the volume of library equivalent to one to two representations, typically 1 to 2 µl), and 1 unit (U) of heat-stable polymerase. This reaction can then be subjected to 25 to 40 cycles of varying temperature conditions, with one cycle being: 30 sec at 94°C,

15 sec at 55° to 65°C (depending on the melting temperature of the oligonucleotide, which can be approximated by adding 4°C for every G or C and 2°C for every A or T and subtracting 5°C for every mismatch that occurs within the annealing sequence), and 1 min at 72°C for every 1000 base pairs of sequence being amplified (e.g., for a 3000-base pair sequence, 3 min at 72°C). The resulting product can then be subcloned and verified as the receptor of interest by sequence confirmation and functional analysis. PCR can also be used to isolate novel receptors. In much the same way that oligonucleotides can be used as probes, PCR products can be used for screening. If primers are designed to be complementary to conserved transmembrane domains, PCR can be utilized to amplify a fragment from a cDNA or genomic library and this fragment can be used as a probe for subsequent screening. The advantage of using a larger DNA fragment (typically several hundred bases) rather than an oligonucleotide as a probe is that the larger probe can be made much “hotter” (i.e., more radioactivity incorporated; Fig 6.1.2). Therefore, rare clones that may not be detectable with an oligonucleotide probe may be evident with a larger fragment probe. Similarly, if the sequence homology in the regions initially selected for the oligonucleotides is not sufficient to hybridize to the desired clone effectively, increasing the size of the probe may provide more regions of homology and allow recovery of the receptor of interest. Again, it is possible to use PCR to recover a fragment from a known receptor sequence and use this fragment as a probe for isolating a related receptor subunit or a homolog from a different species. It is also possible to use PCR to obtain the remaining 5′ and 3′ sequences flanking a fragment sequence presumed to be from a novel receptor. This rapid amplification of cDNA ends (RACE) is perhaps the most challenging PCR technique for isolating full-length cDNA clones (Schaefer, 1995). The technique employs the use of one specific oligonucleotide designed from the known fragment and a second, less specific oligonucleotide which can be of several different types. To amplify the 3′ end of a clone, it is possible to use a poly(dT) primer as a nonspecific 3′ primer. This oligonucleotide will bind to all poly(dA) sequences present in full-length cDNAs, including the one desired. This competition for the primer will result in a significant depletion of the poly(dT) primer and hence, a reduction in the amplification of the

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used as the original template and the reverse transcription reaction is primed with an oligonucleotide specific for the original fragment. The single-stranded cDNA produced from this reaction can then be “tailed” with a string of dCs using the enzyme terminal deoxynucleotidyl transferase (Fig. 6.1.2). This tailed cDNA is then used as the template for PCR using the second nested 3′ oligonucleotide and a poly(dG) 5′ oligonucleotide as primers. A slight variation to this second technique involves using a cDNA library as the template and performing a technique known as anchored PCR in which the mRNA has already been reverse transcribed and subcloned into a vector (either plasmid or bacteriophage). The same nested 3′ oligonucleotides can be employed, but this time a sequence specific to the vector which flanks the cDNA cloning site is used for the 5′ oligonucleotide. If the library was not made directionally, two separate reactions must be run; the only difference is the 5′ oligonucleotide used (Fig. 6.1.4). PCR can also be used to isolate genomic clones utilizing known cDNA sequences. This is very easily accomplished by designing oligonucleotide primers to sequences within the cDNA and using genomic DNA as the template. Initially, primers made 5′ and 3′ to the coding sequence can be used to quickly determine whether the gene contains introns, as the prod-

desired product. To help overcome this problem, a second round of PCR may be performed utilizing a small amount (1/10 to 1/100) of the initial reaction product as template and using a new “nested” oligonucleotide as the 5′ primer (Fig. 6.1.4). This nested primer should be a sequence unique to the fragment originally presumed to be present in the receptor of interest and situated 3′ of the original 5′ PCR primer. In this way, an added degree of specificity ensures that only those templates that were amplified in the first round of PCR and contain the known sequence of the original fragment will be amplified in this subsequent round of PCR. Amplification of the sequence toward the 5′ end of the clone is not quite as straightforward as there is no common sequence similar to the poly(A) to facilitate primer design. One of two approaches can be utilized for the 5′ amplification reaction. In both cases, the nested primer technique is employed, but this time for the 3′ specific oligonucleotide. In the first approach, the 5′ oligonucleotide can be a random decamer (an oligonucleotide that is ten nucleotides long) which will nonspecifically prime sequences within every 300 to 400 nucleotides. It is also important to use a reduced temperature for annealing of the primers (37° to 45°C) to allow the decamer an opportunity to anneal to the template. In the second approach, mRNA is

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Figure 6.1.4 Anchored PCR. Specific primers (striped) are unique to the cDNA of interest. The nested primer (stippled) is contained within the PCR product generated by the first amplification reaction. Specific anchored primers (gray) lie within the vector sequence and outside the inserted cDNAs. If the cDNA insert might have been subcloned into the vector in either orientation (i.e., a nondirectional subcloning), two separate amplifications will be required.


uct size will reflect both coding and noncoding sequences. If introns are present, oligonucleotide primers spaced throughout the cDNA sequence will generate genomic fragments that also span intron sequences. Multiple amplification reactions may be required to isolate the entire gene sequence, which can then be subcloned for subsequent examination.

Expression Cloning As the name implies, this technique screens for cloned receptors based on their ability to elicit a functional response. The method has been used very successfully for cloning G protein–coupled receptors (Masu et al., 1987; Tanaka et al., 1990) and the rat NMDAR1 subunit (Moriyoshi et al., 1991), part of an ionotropic receptor. For G protein–coupled receptors, the clone must be expressed in a cell capable of providing an appropriate G protein for coupling to the receptor of interest. Surprisingly, the NMDA receptor, which is comprised of multiple subunits, was cloned by expression in frog oocytes, even though the glutamate-induced response in this system was much less than the response in rat tissues (Moriyoshi et al., 1991). It was also subsequently shown that not all of the subunits of the NMDA receptor can elicit a functional response in their homomeric forms, which demonstrates that success sometimes requires a little good fortune along with hard work. The expression cloning technique requires introduction of either mRNA or cDNA into a cell that does not normally express the target receptor. After allowing sufficient time for transcription and translation, the injected or transfected cell is tested for a property or function characteristic of the receptor. Functional analysis can include ligand binding, electrophysiological recording, or antibody binding. After determining that introduction of the RNA or cDNA imparts the desired function, the clone is obtained and the sequence determined. Success in cloning by expression often takes a great deal of time and effort. Initially, good quality poly(A)+ RNA (the fraction of total RNA containing mRNA, usually only 3% to 5% of the total material; the remaining 95% is rRNA and tRNA) must be isolated from cells known to contain the functional receptor. This material is then subdivided into pools or reverse transcribed into cDNA. In either case, each subpool of material is tested for expression of the desired receptor. Pools that are positive for a functional response are then subdivided and retested. The process is repeated until a unique

clone is obtained. The benefit to first reverse transcribing the initial poly(A)+ RNA is that this material can then be subcloned into a vector with a T7, T3, or Sp6 promoter, amplified, and then transfected into mammalian cells or used to generate cRNA suitable for injection into cells to test for functional responses. The runoff transcription reaction that generates the cRNA involves linearizing the subcloned cDNA construct downstream of the original cloning site and then utilizing either T3, T7, or Sp6 RNA polymerase (depending on the vector used for subcloning; e.g., pBluescript from Stratagene carries both T3 and T7 promoter sequences for in vitro transcription). In vitro transcription kits are also available from several different manufacturers and provide the researcher with all of the components necessary to generate microgram quantities of cRNA suitable for expression testing. Another benefit to generating plasmid pools is that they are more stable and more easily manipulated in subsequent rounds of purification than poly(A)+ RNAs and are already in a suitable condition for final sequence confirmation. Functional studies can be performed in either Xenopus oocytes (Moriyoshi et al., 1991) or mammalian cell lines (Nash et al., 1997), or an additional technique can be performed in bacterial cells. If the original poly(A)+ RNA is reverse transcribed in a directional manner (i.e., with differing restriction sites introduced at the 5′ and 3′ ends of the molecules), the resulting cDNA can be cloned into a plasmid vector or bacteriophage in a specific orientation. For example, the λgt11 bacteriophage is designed to allow expression of cDNA inserted into its cloning site under the regulation of the inducible lac promoter. In a system like this, cDNAs can be induced to express their respective protein products. Then, utilizing a technique similar to plaque hybridization, the proteins can be detected with a specific antibody or ligand instead of an oligonucleotide probe. The drawback to this method is that there is often a very high level of nonspecific antibody binding. Thus, picking a truly positive clone requires experience as well as luck. A more detailed description of screening libraries with antibodies can be found in Current Protocols in Molecular Biology Chapter 6 (Ausubel et al., 1998).

Differential Display PCR A combination of expression cloning and PCR has been employed to recover genes that are expressed specifically in one cell type and

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not another, or expressed only after stimulation with an inducing agent. This has been called differential display PCR (Liang and Pardee, 1992). Differential display PCR cloning relies on the fact that most eukaryotic mRNA molecules contain a poly(A)+ tail preceded by twelve different pairs of nucleotides XY, where X = A/C/G/T and Y = C/G/T. Therefore, the sequence XY-poly(A) would describe the last nucleotides of the vast majority of messenger RNA present in a cell. The length of the poly(A)+ sequence may vary significantly between RNA molecules, but in every case the poly(A)+ sequence must be preceded by two nucleotides of the order XY (note: an A in position Y would simply extend the poly(A)+ tail by an additional nucleotide). This means that poly(A)+ RNA can be subdivided into twelve distinct pools, each with a different XY combination preceding the poly(A)+ tail. Therefore, after reverse transcribing the poly(A)+ RNA isolated from a particular cell type with a poly(dT) primer, twelve separate PCR amplifications can be run utilizing a 3′ primer with the sequence dT12-MN (where dT12 = dTTTTTTTTTTTT; M = dA/C/G and N = dA/C/G/T) and a random nonamer or decamer as the 5′ primer. Amplification conditions are adjusted so that the annealing temperature is only 42°C, the nucleotide concentration is reduced to 2 µM for each dNTP, and a radioactive nucleotide ([α-35S]dATP or [α33P]dATP) is included in the reaction mix to allow visualization of the PCR products. This assay typically yields 50 to 100 small amplification products (