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Chapter 4 Molecular Cloning Methods. Introduction The significance of gene cloning To elucidate the structure and function of genes. i.e.: investigating.

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Presentation on theme: "Chapter 4 Molecular Cloning Methods. Introduction The significance of gene cloning To elucidate the structure and function of genes. i.e.: investigating."— Presentation transcript:

1 Chapter 4 Molecular Cloning Methods

2 Introduction The significance of gene cloning To elucidate the structure and function of genes. i.e.: investigating hGH gene hGH gene: <10 -6 of human genome Problem 1: need kilograms of human genome DNA for 1μg hGH gene genome DNA for 1μg hGH gene Problem 2: how to separate the gene from the rest DNA rest DNA

3 4.1 Gene Cloning The procedure in a gene cloning experiment is 1. To place a foreign gene into a bacterial cell; 2. To grow a clone of those modified bacteria. The principle factors for gene cloning experiment:  Restriction endonucleases  Vectors  Specific probe

4   The Role of Restriction Endonucleases   Vectors Plasmids as Vectors Phages as Vectors λ Phage Vectors Cosmids M13 phage vectors Phagemids Eukaryotic Vectors   Identifying a Specific Clone with a Specific Probe Polynucleotide Probes

5 4.1.1 The Role of Restriction Endonucleases Restriction : restrict the host range of the virus Restriction : restrict the host range of the virus Endonucleases : cut at sites within the foreign DNA Endonucleases : cut at sites within the foreign DNA How to name: the first 3 letters of the Latin name of the microorganism + the strain designation + Roman numeral How to name: the first 3 letters of the Latin name of the microorganism + the strain designation + Roman numeral

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7 Recognition sequence Recognition frequency 4 bp 4 4 =254 bp 6 bp 4 6 =4096 bp 8 bp 4 8 =65000bp Rere cutters

8 Many restriction enzymes make staggered cut in the two DNA strands, leaving a sticky ends, that can base-pair together briefly. Enzymes that recognize identical sequences are called isoschizomers. The main advantage of restriction enzyme is there ability to cut a DNA reproducibly in the same place; this is the basis of many techniques used to analyze genes. Many restriction enzymes make staggered cut in the two DNA strands, leaving a sticky ends, that can base-pair together briefly. Enzymes that recognize identical sequences are called isoschizomers.

9 Restriction- modification system R-M system Almost all restriction nucleases are paired with methylases that recognize and methylate the same DNA sites

10 Figure 4.1 Maintaining restriction endonuclease resistance after DNA replication We begin with an EcoRI site that is methylated (red) on both strands. After replication, the parental strand of each daughter DNA duplex remains methylated, but the newly made strand of each duplex has not been methylated yet. The one methylated strand in these hemimethylated DNAs is enough to protect both strands against cleavage by EcoRI. Soon, the methylase recognizes the unmethylated strand in each EcoRI site and methylates it, regenerating the fully methylated DNA. We begin with an EcoRI site that is methylated (red) on both strands. After replication, the parental strand of each daughter DNA duplex remains methylated, but the newly made strand of each duplex has not been methylated yet. The one methylated strand in these hemimethylated DNAs is enough to protect both strands against cleavage by EcoRI. Soon, the methylase recognizes the unmethylated strand in each EcoRI site and methylates it, regenerating the fully methylated DNA.

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12 Figure 4.2 The first cloning experiment involving a recombinant DNA assembled in vitro. Boyer and Cohen cut two plasmids, pSC101 and RSF1010, with the same restriction endonuclease, EcoRI. This gave the twolinear DNAs the same stickyends, which were then linked in vitro using DNA ligase. The investigators reintroduced the recombinant DNA into E. coli cells by transformation and selected clones that were resistant to both tetracycline and streptomycin. These clones were therefore harboring the recombinant plasmid. Boyer and Cohen cut two plasmids, pSC101 and RSF1010, with the same restriction endonuclease, EcoRI. This gave the twolinear DNAs the same stickyends, which were then linked in vitro using DNA ligase. The investigators reintroduced the recombinant DNA into E. coli cells by transformation and selected clones that were resistant to both tetracycline and streptomycin. These clones were therefore harboring the recombinant plasmid.

13 Restriction endonucleases recognize specific sequences in DNA molecules and make cuts in both strands. This allows very specific cutting of DNAs. Also, because the cuts in the two strands are frequently staggered, restriction enzymes can create sticky ends that help link together two DNAs to form a recombinant DNA in vitro. Summary:

14 4.1.2 Vectors Vectors serve as carriers to allow replication of recombinant DNAs. Origin of replication Origin of replication Multiple cloning site(MCS) Multiple cloning site(MCS) Selection gene Selection gene Plasmids pBR322 pUC Plasmids pBR322 pUC Phages λphage cosmids M13 Phagemids

15 Plasmids as Vectors

16 Summary: The first generations of plasmid cloning vectors were pBR322 and the pUC plasmids. The former has two antibiotic resistance genes and a variety of unique restriction sites into which one can introduce foreign DNA. Most of these sites interrupt one of the antibiotic resistance genes, making screening straightforward. Screening is even easier with the pUC plasmids. These have an ampicillin resistance gene and a multiple cloning site that interrupts a partial β-galactosidase gene. One screens for ampicillin-resistant clones that do not make active β-galactosidase and therefore do not turn the indicator, X-gal, blue. The multiple cloning site also makes it convenient to carry out directional cloning into two different restriction sites. The first generations of plasmid cloning vectors were pBR322 and the pUC plasmids. The former has two antibiotic resistance genes and a variety of unique restriction sites into which one can introduce foreign DNA. Most of these sites interrupt one of the antibiotic resistance genes, making screening straightforward. Screening is even easier with the pUC plasmids. These have an ampicillin resistance gene and a multiple cloning site that interrupts a partial β-galactosidase gene. One screens for ampicillin-resistant clones that do not make active β-galactosidase and therefore do not turn the indicator, X-gal, blue. The multiple cloning site also makes it convenient to carry out directional cloning into two different restriction sites.

17 Figure 4.3 The plasmid pBR322, showing the locations of 11 unique restriction sites that can be used to insert foreign DNA The locations of the two antibiotic resistance genes (Ampr =ampicillin resistance; Tetr =tetracycline resistance) and the origin of replication (ori ) are also shown. Numbers refer to kilobase pairs (kb) from the EcoRI site. The locations of the two antibiotic resistance genes (Ampr =ampicillin resistance; Tetr =tetracycline resistance) and the origin of replication (ori ) are also shown. Numbers refer to kilobase pairs (kb) from the EcoRI site.

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19 Figure 4.4 Cloning foreign DNA using the PstI site of pBR322. We cut both the plasmid and the insert (yellow) with PstI, then join them through these sticky ends with DNA ligase. Next, we transform bacteria with the recombinant DNA and screen for tetracycline-resistant, ampicillin- sensitive cells. The recombinant plasmid no longer confers ampicillin resistance because the foreign DNA interrupts that resistance gene (blue). We cut both the plasmid and the insert (yellow) with PstI, then join them through these sticky ends with DNA ligase. Next, we transform bacteria with the recombinant DNA and screen for tetracycline-resistant, ampicillin- sensitive cells. The recombinant plasmid no longer confers ampicillin resistance because the foreign DNA interrupts that resistance gene (blue).

20 Figure 4.5 Screening bacteria by replica plating. (a) The replica plating process. We touch a velvet-covered circular tool to the surface of the first dish containing colonies of bacteria. Cells from each of these colonies stick to the velvet and can be transferred to the replica plate in the same positions relative to each other. (b) Screening for inserts in the pBR322 ampicillin resistance gene by replica plating. The original plate contains tetracycline, so all colonies containing pBR322 will grow. The replica plate contains ampicillin, so colonies bearing pBR322 with inserts in the ampicillin resistance gene will not grow (these colonies are depicted by dotted circles). The corresponding colonies from the original plate can then be picked.

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22 pUC

23 lacZ’ : coding for the amino terminalportion of the enzyme β – galactosidease. Host E.coli strain carry a gene fragment that codes the carboxyl potion of β – galactosidease; When X-gal cleaved by β –galactosidease, it releases galactose plus an indigo dye that stains the bacterial colony blue.

24 Figure 4.7 Joining of vector to insert. (a) Mechanism of DNA ligase. Step 1: DNA ligase reacts with an AMP donor—either ATP or NAD(nicotinamide adenine dinucleotide), depending on the type of ligase. This produces an activated enzyme (ligase-AMP). Step 2: The activated enzyme donates a phosphate to the free 5’-phosphate at the nick in the lower strand of the DNA duplex, creating a high- energy diphosphate group on one side of the nick. Step 3: With energy provided by cleavage of the diphosphate, a new phosphodiester bond is created, sealing the nick in the DNA. This reaction can also occur in both DNA strands at once, so two independent DNAs can be joined together by DNA ligase. Step 1: DNA ligase reacts with an AMP donor—either ATP or NAD(nicotinamide adenine dinucleotide), depending on the type of ligase. This produces an activated enzyme (ligase-AMP). Step 2: The activated enzyme donates a phosphate to the free 5’-phosphate at the nick in the lower strand of the DNA duplex, creating a high- energy diphosphate group on one side of the nick. Step 3: With energy provided by cleavage of the diphosphate, a new phosphodiester bond is created, sealing the nick in the DNA. This reaction can also occur in both DNA strands at once, so two independent DNAs can be joined together by DNA ligase.

25 Figure 4.7 Joining of vector to insert. (b)Alkaline phosphatase prevents vector re-ligation. Step 1: We cut the vector(blue, top left) with BamHI. This produces sticky ends with 5’-phosphates(red). Step 2: We remove the phosphates with alkaline phosphatase, making it impossible for the vector to re-ligate with itself. Step 3: We also cut the insert(yellow, upper right) with BamHI, producing sticky ends with phosphates that we do not remove. Step 4: Finally, we ligate the vector and insert together. The phosphates on the insert allow two phosphodiester bonds to form(red), but leave two unformed bonds, or nicks, These will be completed once the DNA is in the transformed bacterial cell. Step 1: We cut the vector(blue, top left) with BamHI. This produces sticky ends with 5’-phosphates(red). Step 2: We remove the phosphates with alkaline phosphatase, making it impossible for the vector to re-ligate with itself. Step 3: We also cut the insert(yellow, upper right) with BamHI, producing sticky ends with phosphates that we do not remove. Step 4: Finally, we ligate the vector and insert together. The phosphates on the insert allow two phosphodiester bonds to form(red), but leave two unformed bonds, or nicks, These will be completed once the DNA is in the transformed bacterial cell.

26 Phages as vectors Natural advantages over plasmid: They infect cells much more efficiently than plasmids transform cells, so the yield of clones with phage vectors is usually higher.

27 Summary: Two kinds of phages have been especially popular as cloning vectors. The first of these is λ, from which certain nonessential genes have been removed to make room for inserts. Some of these engineered phages can accommodate inserts up to 20 kb, which makes them useful for building genomic libraries, in which it is important to have large pieces of genomic DNA in each clone. Cosmids can accept even larger inserts—up to 50 kb—making them a favorite choice for genomic libraries. The second major class of phage vector is composed of the M13 phages. These vector have the convenience of a multiple cloning site and the further advantage of producing single-stranded recombinant DNA, which can be used for DNA sequencing and for site-direct mutagenesis. Plasmids called phagemids have also been engineered to produce single-stranded DNA in the presence of helper phages. Two kinds of phages have been especially popular as cloning vectors. The first of these is λ, from which certain nonessential genes have been removed to make room for inserts. Some of these engineered phages can accommodate inserts up to 20 kb, which makes them useful for building genomic libraries, in which it is important to have large pieces of genomic DNA in each clone. Cosmids can accept even larger inserts—up to 50 kb—making them a favorite choice for genomic libraries. The second major class of phage vector is composed of the M13 phages. These vector have the convenience of a multiple cloning site and the further advantage of producing single-stranded recombinant DNA, which can be used for DNA sequencing and for site-direct mutagenesis. Plasmids called phagemids have also been engineered to produce single-stranded DNA in the presence of helper phages.

28 Figure 4.8 Cloning in Charon 4. (a) Forming the recombinant DNA. We cut the vector (yellow) with EcoRI to remove the stuffer fragment and save the arms. Next, we ligate partially digested insert DNA (red) to the arms. (b) Packaging and cloning the recombinant DNA. We mix the recombinant DNA from (a) with an in vitro packaging extract that contains λ phage head and tail components and all other factors needed to package the recombinant DNA into functional phage particles. Finally, we plate these particles on E.coli and collect the plaques that form. (b) Packaging and cloning the recombinant DNA. We mix the recombinant DNA from (a) with an in vitro packaging extract that contains λ phage head and tail components and all other factors needed to package the recombinant DNA into functional phage particles. Finally, we plate these particles on E.coli and collect the plaques that form.

29 Figure 4.9 Selection of positive genomic clones by plaque hybridization. First, we touch a nitrocellulose ot similar filter to the surface of the dish containing the Charon 4 plaques from Figure 4.8. Phage DNA released naturally from each plaque will stick to the filter. Next, we denature the DNA with alkali and hybridize the filter to a labeled probe for the gene we are studying, then use X-ray film to reveal the position of the label. Cloned DNA from one plaque near the center of the filter has hybridized, as shown by the dark spot on the film.

30 Cosmids Behave both as plasmids and as phages; Contain the cos sites of λ and plasmid origin of replication; Have room for kb inserts.

31 M13 phage vectors β –galactosidease gene fragment pUC family MCS Single stranded DNA genome

32 Figure 4.10 Obtaining single- stranded DNA by cloning in M13 phage. Foreign DNA (red), cut with HindIII, is inserted into the HindIII site of the double- stranded phage DNA. The resulting recombinant DNA is used to transform E.coli cells, whereupon the DNA replicates by a rolling circle mechanism, producing many single-stranded product DNAs. The product DNAs are called positive (+) strands, by convention. The template DNA is therefore the negative (-) strand.

33 PhagemidesSingle-stranded; Both phage and plasmid characteristics; Help phage Two RNA polymerase promoters (T7and T3)

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35 Summary Two kinds of phages have been especially popular as cloning vectors. The first of these is λ, from which certain nonessential genes have been removed to make room for inserts. Some of these engineered phages can accommodate inserts up to 20 kb, which makes them useful for building genomic libraries, in which it is important to have large pieces of genomic DNA in each clone. Cosmids can accept even larger inserts—up to 50 kb—making them a favorite choice for genomic libraries. The second major class of phage vector is composed of the M13 phages. These vector have the convenience of a multiple cloning site and the further advantage of producing single-stranded recombinant DNA, which can be used for DNA sequencing and for site-direct mutagenesis. Plasmids called phagemids have also been engineered to produce single-stranded DNA in the presence of helper phages.

36 4.1.3 Identifying a Specific Clone with a Specific Probe Polynucleotide Probes High stringency High stringency Low stringency Low stringency

37 Summary Specific clones can be identified using polynucleotide probes that bind to the gene itself. Knowing the amino acid sequence of a gene product, one can design a set of oligonucleotides that encode part of this amino acid sequence. This can be one of the quickest and most accurate means of identifying a particular clone. Specific clones can be identified using polynucleotide probes that bind to the gene itself. Knowing the amino acid sequence of a gene product, one can design a set of oligonucleotides that encode part of this amino acid sequence. This can be one of the quickest and most accurate means of identifying a particular clone.

38 4.2 The Polymerase Chain Reaction (PCR) PCR amplifies a region of DNA between two predetermined sites. Oligo- nucleotides complementary to these sites serve as primers for synthesis of copies of the DNA between the sites. Each cycle of PCR double the number of copies of the amplified DNA until a large quantity has been made. PCR amplifies a region of DNA between two predetermined sites. Oligo- nucleotides complementary to these sites serve as primers for synthesis of copies of the DNA between the sites. Each cycle of PCR double the number of copies of the amplified DNA until a large quantity has been made.

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41 Figure 4.12 Amplifying DNA by the polymerase chain reaction. Cycle 1: Start with a DNA duplex (top) and heat it to separate its two strands (red and blue). Then add short, single-stranded DNA primers (purple and yellow) complementary to sequences on either side of the region (X) to be amplified. The primers hybridize to the appropriate sites on the separated DNA strands; now a special heat-stable DNA polymerase uses these primers to start synthesis of complementary DNA strands. The arrows represent newly made DNA, in which replication has stopped at the tip of the arrowhead. At the end of cycle 1, two DNA duplexes are present, including the region to be amplified, whereas we started with only one. The 5’→3’ polarities of all DNA strands and primers are indicated throughout cycle 1. The same principles apply in cycle 2. Cycle 2: Repeat the process, heating to separate DNA strands, cooling to allow annealing with primers, and letting the heat-stable DNA polymerase make more DNA. Now each of the four DNA strands, including the two newly made ones, can serve as templates for complementary DNA synthesis. The result is four DNA duplexes that have the region to be amplified. Notice that each cycle doubles the number of molecules of DNA because the products of each cycle join the parental molecules in serving as templates for next cycle. This exponential increase yields 8 molecules in the next cycle and 16 in the cycle after that. This process obviously leads to very high numbers in only a short time. Cycle 1: Start with a DNA duplex (top) and heat it to separate its two strands (red and blue). Then add short, single-stranded DNA primers (purple and yellow) complementary to sequences on either side of the region (X) to be amplified. The primers hybridize to the appropriate sites on the separated DNA strands; now a special heat-stable DNA polymerase uses these primers to start synthesis of complementary DNA strands. The arrows represent newly made DNA, in which replication has stopped at the tip of the arrowhead. At the end of cycle 1, two DNA duplexes are present, including the region to be amplified, whereas we started with only one. The 5’→3’ polarities of all DNA strands and primers are indicated throughout cycle 1. The same principles apply in cycle 2. Cycle 2: Repeat the process, heating to separate DNA strands, cooling to allow annealing with primers, and letting the heat-stable DNA polymerase make more DNA. Now each of the four DNA strands, including the two newly made ones, can serve as templates for complementary DNA synthesis. The result is four DNA duplexes that have the region to be amplified. Notice that each cycle doubles the number of molecules of DNA because the products of each cycle join the parental molecules in serving as templates for next cycle. This exponential increase yields 8 molecules in the next cycle and 16 in the cycle after that. This process obviously leads to very high numbers in only a short time.

42 4.2.1 cDNA Cloning Nick translation Nick translation Reverse transcriptase Reverse transcriptase RNase H RNase H Terminal transferase Terminal transferase

43 Figure 4.13 Making a cDNA library. This figure focuses on cloning a single cDNA, but the method can be applied to a mixture of mRNAs and produce a library of corresponding cDNAs. (a) Use oligo(dT) as a primer and reverse transcriptase tocopy the mRNA (blue), producing a cDNA (red) that is hybridized to the mRNA template. (b) Use RNase H to partially digest the mRNA, yielding a set of RNA primers base-paired to the first-strand cDNA. (c) Use E.coli DNA polymerase I under nick translation conditions to build second-strand cDNAs on the RNA primers. (d) The second-strand cDNA growing from the leftmost primer (blue) has been extended all the way to the 3’-end of the oligo(dA) corresponding to the oligo(dT) primer on the first-strand cDNA. (e) To give the double- stranded cDNA sticky ends, add oligo(dC) with terminal transferase. (f) Anneal the oligo(dC) ends of the cDNA to complementary oligo(dG) ends of a suitable vector (black). The recombinant DNA can then be used to transform bacterial cells. Enzymes in these cells remove remaining nicks and replace any remaining RNA with DNA. This figure focuses on cloning a single cDNA, but the method can be applied to a mixture of mRNAs and produce a library of corresponding cDNAs. (a) Use oligo(dT) as a primer and reverse transcriptase tocopy the mRNA (blue), producing a cDNA (red) that is hybridized to the mRNA template. (b) Use RNase H to partially digest the mRNA, yielding a set of RNA primers base-paired to the first-strand cDNA. (c) Use E.coli DNA polymerase I under nick translation conditions to build second-strand cDNAs on the RNA primers. (d) The second-strand cDNA growing from the leftmost primer (blue) has been extended all the way to the 3’-end of the oligo(dA) corresponding to the oligo(dT) primer on the first-strand cDNA. (e) To give the double- stranded cDNA sticky ends, add oligo(dC) with terminal transferase. (f) Anneal the oligo(dC) ends of the cDNA to complementary oligo(dG) ends of a suitable vector (black). The recombinant DNA can then be used to transform bacterial cells. Enzymes in these cells remove remaining nicks and replace any remaining RNA with DNA.

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45 Figure 4.15 Using RT-PCR to clone a single cDNA. (a) Use a reverse primer (red) with a HindIII site (yellow) at its 5’-end to start first-strand cDNA synthesis, with reverse transcriptase to catalyze the reaction. (b) Denature the mRNA-cDNA hybrid and anneal a forward primer (red) with a BamHI site (green) at its 5’-end. (c) This forward primer initiates second-strand cDNA synthesis, with DNA polymerase catalyzing the reaction. (d) Continue PCR with the same two primers to amplify the double-stranded cDNA. (e) Cut the cDNA with BamHI and HindIII to generate sticky ends. (f) Ligate the cDNA to the BamHI and HindIII sites of a suitable vector (purple). Finally, transform cells with the recombinant cDNA to produce a clone.

46 Figure 4.16 RACE procedure to fill in the 5’-end of a cDNA. (a) Hybridize an incomplete cDNA (red), or an oligonucleotide segment of a cDNA to mRNA (green), and use reverse transcriptase to extend the cDNA to the 5’-end of the mRNA. (b) Use terminal transferase and dCTP to add C residues to the 3’end of the extended cDNA; also, use RNase H to degrade the mRNA. (c) Use an oligo(dG) primer and DNA polymerase to synthesize a second strand of cDNA (blue). (d) Perform PCR with oligo(dG) as the forward primer and an oligonucleotide that hybridizes to the 3’-end of the cDNA as the reverse primer. (e)The product is a cDNA that has been extended to the 5’-end of the mRNA. A similar procedure (3’-RACE) can be used to extend the cDNA in the 3’-direction. In that case, there is no need to tail the 3’-end of the cDNA with terminal transferase because the mRNA already contains poly(A); thus, the reverse primer would be oligo(dT).

47 Summary To make a cDNA library, we can synthesize cDNAs one strand at a time, using mRNAs from a cell as templates for the first strands and these first strands as temletes for the second strands. Reverse trnscriptase generates the first strands and E.coli DNA polymerase I generates the second strands. We can endow the double stranded cDNAs with oligonucleotide tails that base-par with complementary tails on a cloning vector. We can then use these recombinant DNAs to transform bacteria. We can use RT-PCR to generate a cDNA from a single type of mRNA, but we must know the sequence of the mRNA in order to design the primers for the PCR step. If we put restriction sites on the PCR primers, we place these sites at the ends of the cDNA,so it is easy to ligate the cDNA into a vector. We can detect particular clones by colony hybridazation with redioactive DNA probes,or with antibodies if an expression vector such as λgt11 is used. To make a cDNA library, we can synthesize cDNAs one strand at a time, using mRNAs from a cell as templates for the first strands and these first strands as temletes for the second strands. Reverse trnscriptase generates the first strands and E.coli DNA polymerase I generates the second strands. We can endow the double stranded cDNAs with oligonucleotide tails that base-par with complementary tails on a cloning vector. We can then use these recombinant DNAs to transform bacteria. We can use RT-PCR to generate a cDNA from a single type of mRNA, but we must know the sequence of the mRNA in order to design the primers for the PCR step. If we put restriction sites on the PCR primers, we place these sites at the ends of the cDNA,so it is easy to ligate the cDNA into a vector. We can detect particular clones by colony hybridazation with redioactive DNA probes,or with antibodies if an expression vector such as λgt11 is used.

48 4.3 Methods of Expressing Cloned Genes

49 4.3.1 Expression Vectors Expression vectors with strong promoters Expression vectors with strong promoters Inducible Expression Vectors Inducible Expression Vectors Expression vectors produce fusion proteins Expression vectors produce fusion proteins Eukaryotic expression vectors Eukaryotic expression vectors

50 Figure 4.17 Figure 4.17 Producing a fusion protein by cloning in a pUC plasmid. Insert foreign DNA (yellow) into the multiple cloning site (MCS); transcription from the lac promoter (purple) gives a hybrid mRNA beginning with a few lacZ’ codons, changing to insert sequence, then back to lacZ’ (red). This mRNA is translated to a fusion protein containing a few β- galactosidase amino acids for the remainder ofthe protein. Because the insert contains a translation stop codon, the remaining lacZ’ codons are not translated. Insert foreign DNA (yellow) into the multiple cloning site (MCS); transcription from the lac promoter (purple) gives a hybrid mRNA beginning with a few lacZ’ codons, changing to insert sequence, then back to lacZ’ (red). This mRNA is translated to a fusion protein containing a few β- galactosidase amino acids for the remainder ofthe protein. Because the insert contains a translation stop codon, the remaining lacZ’ codons are not translated.

51 Figure 4.18 Using a P BAD vector. The green fluorescent protein (GFP) gene was cloned into a vector under control of the P BAD promoter and promoter activity was induced with increasing concentrations of arabinose. GFP production was monitored by electrophoresing extracts from cells induced with the arabinose concentrations given at top, blotting the proteins to a membrane, and detecting GFP with an anti-GFP antibody. The green fluorescent protein (GFP) gene was cloned into a vector under control of the P BAD promoter and promoter activity was induced with increasing concentrations of arabinose. GFP production was monitored by electrophoresing extracts from cells induced with the arabinose concentrations given at top, blotting the proteins to a membrane, and detecting GFP with an anti-GFP antibody.

52 Summary: Expression vectors are designed to yield the protein product of a cloned gene, usually in the greatest amount possible. To optimize expression, these vectors provide strong bacterial promoters and bacterial ribosome binding sites that would be missing on cloned eukaryotic genes. Most cloning vectors are inducible, to avoid premature overproduction of a foreign product that could poison the bacterial host cells. Expression vectors are designed to yield the protein product of a cloned gene, usually in the greatest amount possible. To optimize expression, these vectors provide strong bacterial promoters and bacterial ribosome binding sites that would be missing on cloned eukaryotic genes. Most cloning vectors are inducible, to avoid premature overproduction of a foreign product that could poison the bacterial host cells.

53 Figure 4.19 Using an oligohistidine expression vector. (a) Map of a generic oligohistidine vector. Just after the ATG initiation codon (green) lies a coding region (red) encoding six histidine in a row [(His)6]. This is followed by a region (orange) encoding a recognition site for the proteolytic enzyme enterokinase (EK). Finally, the vector has a multiple cloning site (MCS, blue). Usually, the vector comes in three forms with the MCS sites in each of the three reading frames. One can select the vector that puts the gene in the right reading frame relative to the oligohistidine. Just after the ATG initiation codon (green) lies a coding region (red) encoding six histidine in a row [(His)6]. This is followed by a region (orange) encoding a recognition site for the proteolytic enzyme enterokinase (EK). Finally, the vector has a multiple cloning site (MCS, blue). Usually, the vector comes in three forms with the MCS sites in each of the three reading frames. One can select the vector that puts the gene in the right reading frame relative to the oligohistidine.

54 Figure 4.19 Using an oligohistidine expression vector. (b) Using the vector. 1. Insert the gene of interest (yellow) into the vector in frame with the oligohistidine coding region (red) and transform bacterial cells with the recombinant vector. The cells produce the fusion protein (red and yellow), along with other, bacterial proteins (green). 2. Lyse the cells, releasing the mixture of proteins. 3. Pour the cell lysate through a nickel affinity chromatography column, which binds the fusion protein but not the other proteins. 4. Release the fusion protein from the column with histidine or with imidazole, a histidine analogue, which competes with the oligohistidine for binding to the nickel. 5. Cleave the fusion protein with enterokinase. 6. Pass the cleaved protein through the nickel column once more to separate the oligehistidine from the desired protein.

55 Summary: Expression vectors frequently produce fusion proteins, with one part of the protein coming from coding sequences in the vector and the other part from sequences in the cloned gene itself. Many fusion proteins have the great advantage of being simple to isolate by affinity chromatography. The λgt11 vector produces fusion proteins that can be detected in plaques with a specific antiserum. Expression vectors frequently produce fusion proteins, with one part of the protein coming from coding sequences in the vector and the other part from sequences in the cloned gene itself. Many fusion proteins have the great advantage of being simple to isolate by affinity chromatography. The λgt11 vector produces fusion proteins that can be detected in plaques with a specific antiserum.

56 Figure 4.20 Forming a fusion protein in λgt11. The gene to be expressed (green) is inserted into the EcoRI site near the end of the lacZ coding region (red) just upstream of the transcription terminator. Thus, upon induction of the lacZ gene by IPTG, a fused mRNA results, containing the inserted coding region just downstream of that of β-galactosidase. This mRNA is translated by the host cell to yield a fusion protein. The gene to be expressed (green) is inserted into the EcoRI site near the end of the lacZ coding region (red) just upstream of the transcription terminator. Thus, upon induction of the lacZ gene by IPTG, a fused mRNA results, containing the inserted coding region just downstream of that of β-galactosidase. This mRNA is translated by the host cell to yield a fusion protein.

57 Figure 4.21 Detecting positiveλgt11 clones by antibody screening. A filter is used to blot proteins from phage plaques on a Petri dish. One of the clones (red) has produced a plaque containing a fusion protein including β- galactosidase and a part of the protein of interest. The filter with its blotted proteins is incubated with an antibody directed against our protein of interest, then with labeled Staphylococcus protein A, which binds specifically to antibodies. It will therefore bind only to the antibody-antigen complexes at the spot corresponding to our positive clone. A dark spot on the film placed in contact with the filter reveals the location of our positive clone. A filter is used to blot proteins from phage plaques on a Petri dish. One of the clones (red) has produced a plaque containing a fusion protein including β- galactosidase and a part of the protein of interest. The filter with its blotted proteins is incubated with an antibody directed against our protein of interest, then with labeled Staphylococcus protein A, which binds specifically to antibodies. It will therefore bind only to the antibody-antigen complexes at the spot corresponding to our positive clone. A dark spot on the film placed in contact with the filter reveals the location of our positive clone.

58 Figure 4.22 Expressing a gene in a baculovirus.

59 First, insert the gene to be expressed (red), into a baculovirus transfer vector. In this case, the vector contains the powerful polyhedrin promoter (Polh), flanked bythe DNA sequences (yellow) that normally surround the polyhedrin gene, including a gene (green) that is essential for virus replication, the polyhedrin coding region itself is missing from this transfer vector. Just downstream of the promoter is a BamHI restriction site, which can be used to open up the vector (step a) so it can accept the foreign gene (red) by ligation (step b). In step c, mix the recombinant transfer vector with linear viral DNA that has been cut so as to remove the essential gene. Transfect insect cells with the two DNAs together. This process is known as co-transfection. The two DNAs are not drawn to scale, the viral DNA is actually almost 15 times the size of the vector. Inside the cell, the two DNAs recombine by a double crossover that inserts the gene to be expressed, along with the essential gene, into the viral DNA. The result is a recombinant virus DNA that has the gene of interest under the control of the polyhedrin promoter. Next, infect cells with the recombinant virus. Finally, in step d and e, infect cells with the recombinant virus and collect the protein product these cells make. Notice that the original viral DNA is linear and it is missing the essential gene, so it cannot infect cells (f). This lack of infectivity selects automatically for recombinant viruses; they are the only ones that can infect cells. First, insert the gene to be expressed (red), into a baculovirus transfer vector. In this case, the vector contains the powerful polyhedrin promoter (Polh), flanked bythe DNA sequences (yellow) that normally surround the polyhedrin gene, including a gene (green) that is essential for virus replication, the polyhedrin coding region itself is missing from this transfer vector. Just downstream of the promoter is a BamHI restriction site, which can be used to open up the vector (step a) so it can accept the foreign gene (red) by ligation (step b). In step c, mix the recombinant transfer vector with linear viral DNA that has been cut so as to remove the essential gene. Transfect insect cells with the two DNAs together. This process is known as co-transfection. The two DNAs are not drawn to scale, the viral DNA is actually almost 15 times the size of the vector. Inside the cell, the two DNAs recombine by a double crossover that inserts the gene to be expressed, along with the essential gene, into the viral DNA. The result is a recombinant virus DNA that has the gene of interest under the control of the polyhedrin promoter. Next, infect cells with the recombinant virus. Finally, in step d and e, infect cells with the recombinant virus and collect the protein product these cells make. Notice that the original viral DNA is linear and it is missing the essential gene, so it cannot infect cells (f). This lack of infectivity selects automatically for recombinant viruses; they are the only ones that can infect cells.

60 Summary: Foreign genes can be expressed in eukaryotic cells, and these eukaryotic systems have some advantages over their prokaryotic counterparts for producing eukaryotic proteins. Foreign genes can be expressed in eukaryotic cells, and these eukaryotic systems have some advantages over their prokaryotic counterparts for producing eukaryotic proteins. Two of the most important advantages are (1) Eukaryotic proteins made in eukaryotic cells tend to be folded properly, so they are soluble, rather than aggregated into insoluble inclusion bodies. (2) Eukaryotic proteins made in eukaryotic cells are modified (phosphorylated, glycosylated, etc.) in a eukaryotic manner. Two of the most important advantages are (1) Eukaryotic proteins made in eukaryotic cells tend to be folded properly, so they are soluble, rather than aggregated into insoluble inclusion bodies. (2) Eukaryotic proteins made in eukaryotic cells are modified (phosphorylated, glycosylated, etc.) in a eukaryotic manner.

61 4.3.2 Other Eukaryotic Vectors Yeast Artificial chromosomes (YACs) Yeast Artificial chromosomes (YACs) Using the Ti plasmid to transfer genes to plants Using the Ti plasmid to transfer genes to plants

62 Essential components of YAC vectors Centromers (CEN), telomeres (TEL) and autonomous replicating sequence (ARS) for proliferation in the host cell. amp r for selective amplification and markers such as TRP1 and URA3 for identifying cells containing the YAC vector. Recognition sites of restriction enzymes (e.g., EcoRI and BamHI)

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64 BAC vectors Bacterial artificial chromosomes are based on the F factor of E. coli and can be used to clone up to 350 kb of genomic DNA in a conveniently handled E. coli host. They are a morre stable and easier to use alternative to YAC. Bacterial artificial chromosomes are based on the F factor of E. coli and can be used to clone up to 350 kb of genomic DNA in a conveniently handled E. coli host. They are a morre stable and easier to use alternative to YAC.

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66 Using the Ti Plasmid to Transfer Gees to Plants

67 Nopaline and octopine Ti plasmids carry a variety of genes, including T-regions that have overlapping functions

68 T-DNA has almost identical repeats of 25 bp at each end in the Ti plasmid. The right repeat is necessary for transfer and integration to a plant genome. T-DNA that is integrated in a plant genome has a precise junction that retains 1-2 bp of the right repeat, but the left junction varies and may be up to 100 bp short of the left repeat.

69 Figure 4.24 Crown gall tumors. (a) Formation of a crown gall. 1. Agrobacterium cells enter a wound in the plant, usually at the crown, or the junction of root an stem. 2. The Agrobacterium contains a Ti plasmid in addition to the much larger bacterial chromosome. The Ti plasmid has a segment (the T-DNA, red) that promotes tumor formation in infected plants. 3. The bacterium contributes its Ti plasmid to the plant cell, and the T-DNA from the Ti plasmid into grates into the plant’s chromosomal DNA. 4. The genes in the T-DNA direct the formation of a crown gall, which nourishes the invading bacteria.

70 Figure 4.24 Crown gall tumors. (b) Photograph of a crown gall tumor genetated by cutting off the top of a tobacco plant and inoculating with Agrobacterium. This crown gall tumor is a teratoma, which generates normal as well as tumorous tissues springing from the tumor.

71 Figure 4.25 Using a T-DNA plasmid to introduce a gene into tobacco plant. (a) A plasmid is formed with a foreign gene (red) under the control of the mannopine synthetase promoter (blue). This plasmid is used to transform Agrobacterium cells. (a) A plasmid is formed with a foreign gene (red) under the control of the mannopine synthetase promoter (blue). This plasmid is used to transform Agrobacterium cells. (b) The transformed bacterial cells divide repeatedly. (c) A disk of tobacco leaf tissue is removed and incubated in nutrient medium, along with the transformed Agrobacterium cells. These cells infect the tobacco tissue, transferring the plasmid bearing the cloned foreign gene. (d) The disk of tobacco tissue sends out roots into the surrounding medium. (e) One of these roots is transplanted to another kind of medium, where it forms a shoot. This plantlet grows into a transgenic tobacco plant that can be tested for expression of the transplanted gene.

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73 Summary: Molecular biologists can clone hundreds of thousands of base pairs of DNA at a time in yeast artificial chromosomes (YACs). If they wish to transfer cloned genes to plants, creating transgenic organisms with altered characteristics, they use a plant vector such as the Ti plasmid Molecular biologists can clone hundreds of thousands of base pairs of DNA at a time in yeast artificial chromosomes (YACs). If they wish to transfer cloned genes to plants, creating transgenic organisms with altered characteristics, they use a plant vector such as the Ti plasmid.


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