15.2: Make and Screen a cDNA Library
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To make a cDNA library, first isolate cellular mRNA. This extract should represent all of the transcripts in the cells at the time of isolation, or the cell’s transcriptome. This term is used in analogy to genome. While the genome is all of the genetic information of an organism, a transcriptome (usually eukaryotic) reflects all of the genes expressed in a given cell type at a moment in time. Reverse-transcribed cDNAs from an mRNA extract are also referred to as a transcriptome. A tube full of bacterial cells that were transformed with plasmids recombined with cDNAs is a cDNA library. cDNA libraries made from mRNAs from different cell types are in fact different transcriptomes; likewise, cDNA libraries from mRNAs taken from the same cells but grown under different conditions. A cDNA library reflects the mRNAs transcribed in cells at the moment of extraction. When cells in a cDNA library are spread out on a nutrient agar petri dish, each cell grows into a colony; each cell in the colony is a clone of the starting cell. cDNA libraries can be used to isolate and then to sequence the DNA encoding a polypeptide that you are studying.
Recall that the mature mRNA in eukaryotic cells has been spliced. Thus, cDNAs made from eukaryotic cells do not include introns. Introns, as well as sequences of enhancers and other regulatory elements in and surrounding a gene, must be studied in genomic libraries (to be discussed later). Here we look at how to make a cDNA library.
15.2.1 cDNA Construction
mRNA is only a few percent of eukaryotic cell RNA; most is rRNA. But that small amount of mRNA can be isolated from other RNAs because of their 3′ poly(A) tails. Simply pass a total RNA extract over an oligo(dT) column like the one illustrated in Figure 15.2.
The strings of thymidine (T) in the oligo(dT) column form H-bonds with the poly(A) tails of mRNAs, tethering them to the column. All RNAs without a 3′ poly(A) tail will flow through the column as waste. A second buffer is passed through the column to destabilize the H-bonds to allow elution of an mRNA fraction. If “free” oligo(dT)is then added to the eluted mRNA, it also forms H-bonds with the poly(A) tails of the mRNAs, where it can serve as a primer for the synthesis of cDNA copies of the poly(A) mRNAs originally in the cells. Adding four deoxynucleotide DNA precursors and reverse transcriptase (e.g., from chicken retrovirus– infected cells) will start reverse transcription. Figure 15.3 shows the synthesis of a cDNA strand complementary to an mRNA.
After being heated to separate the cDNAs from the mRNAs, the cDNA is replicated to produce double-stranded cDNA, or (ds)cDNA (Figure 15.4).
Replication of the second cDNA strand is also catalyzed by reverse transcriptase. Second-strand synthesis is primed by the 3′ end of a stem-loop structure that can form with most mRNAs. Reverse transcriptase is also a DNA polymerase, recognizing DNA as well as RNA templates, with the same 5′-to-3′ DNA polymerizing activity as all DNA and RNA polymerases. After second cDNA strand synthesis, an S1 nuclease (a specifically single stranded endonuclease originally isolated from an East Asian fungus!) is added. This nuclease will open the loop of the (ds)cDNA structure and trim the rest of the single-stranded DNA. What remains is the (ds)cDNA.
15.2.2 Cloning cDNAs into Plasmid Vectors
To understand cDNA cloning and other aspects of making recombinant DNA, let’s look again at what’s in the recombinant DNA tool kit. In addition to reverse transcriptase and S1 nuclease, other enzymes in the kit include restriction endonucleases (REs or restriction enzymes for short) and DNA ligase. The natural function of REs in bacteria is to recognize and hydrolyze specific restriction site sequences in phage DNA, destroying the phage DNA and avoiding infection.
At 4–6 bp long, typical RE sites occur often and by chance in DNA. How does a phage-infected bacterium keep its RE from attacking its own genome?
Some restriction enzymes cut through the two strands of the double helix to leave blunt ends. Others make a staggered cut on each strand at their restriction site, leaving behind complementary (“sticky”) ends for H-bond formation (Figure 15.5).
Typical RE sites are palindromes. Consider one strand of this EcoR1 RE site: 5′-GAATTC-3′. Fill in the opposite strand to see the palindrome and look up how EcoR1 digestion leaves staggered ends.
If two dsDNAs from different sources (e.g., different species) are digested with an RE that makes a staggered cut, the resulting fragments will have the same sticky ends. If you mix these DNAs, their complementary ends will form H-bonds linking the fragments. If the linked fragments are from different sources, we can add DNA ligase to covalently seal the fragments. We say then that the fragments have been recombined. Such a protocol makes it relatively easy to recombine any two different DNAs at will. Let’s look at how we recombine plasmid DNAs and cDNA—the first steps in cloning cDNAs.
15.2.3 Preparing Recombinant Plasmid Vectors Containing cDNA Inserts
Vectors (such as plasmids or phage DNA) are carriers engineered to recombine with foreign DNAs of interest. When a recombinant vector with its foreign DNA insert gets into a host cell, it can replicate many copies of itself, enough for easy isolation and study. cDNAs are typically inserted into plasmid vectors (usually “store-bought”). They can be cut with an RE at a suitable location, leaving those sticky ends. However, it would not do to digest (ds)cDNA with restriction endonucleases, since the goal is not to clone cDNA fragments but entire cDNA molecules. Therefore, it will be necessary to attach linkers to either end of the (ds)cDNAs. Plasmid DNAs and cDNA-linker constructs can then be digested with the same restriction enzyme to produce compatible sticky ends. Figure 15.6 shows the steps in the preparation of a vector and (ds)cDNA for recombination.
To prepare for recombination, a plasmid vector is digested with a restriction enzyme to open the DNA circle. The (ds)cDNAs to be inserted into the plasmid vector are mixed with linkers and DNA ligase to attach a linker DNA at both ends of the (ds)cDNA. The linkers are short, synthetic (ds)DNA oligomers, containing restriction sites recognized by the same restriction enzyme that linearized the plasmid. Linkers attached to the (ds)cDNAs ends are digested with the appropriate restriction enzyme to give them “sticky ends” complementary to the staggered, plasmid ends.
15.2.4 Recombining Plasmids and cDNA Inserts and Transforming Host Cells
The next step is to mix the digested plasmids with the digested linker-ended cDNAs in just the right proportions so that most of the cDNAs form H-bonds with most of the sticky plasmid ends (and not with each other!). Once the H-bonds have formed between the cDNAs and plasmids, the addition of DNA ligase forms phosphodiester linkages between plasmid and cDNA insert to complete the recombinant circle of DNA (Figure 15.7).
In early cloning experiments, an important consideration was how to generate plasmids with only one copy of a given cDNA insert (rather than how to make lots of re-ligated plasmids with no inserts or how to make lots of plasmids with multiple inserts). Improved, better-engineered vector and linker combinations made this issue less important.
Improvements include vectors with multiple restriction- enzyme sites. Vectors with two different restriction enzymes could then be ligated to cDNAs with different oligonucleotide linkers at each end. How might this fix solve some of the recombination issues noted here?
15.2.5 Transforming Host Cells with Recombinant Plasmids
The recombinant DNA molecules are now ready for the next step: the creation of a cDNA library. The recombinant DNAs are added to E. coli (or other) host cells, which have been made permeable so that they are easily transformed. Recall that transformation (as defined by Griffith) is the bacterial uptake of foreign DNA, leading to a genetic change. The transforming principle in cloning is the recombinant plasmid! The transformation step that results in a tube full of transformed cells (which is, in fact, the cDNA library) is shown in Figure 15.8.
15.2.6 Plating a cDNA Library on Antibiotic-Agar to Select Recombinant Plasmids
After all these treatments, not all plasmid molecules in the mix are recombinant; some cells in the mix haven’t even taken up a plasmid. So, when the recombinant cells are plated on agar, how do you tell which of the colonies that grow came from cells that took up a recombinant plasmid? Both the host strain of E. coli and plasmid vectors used these days were further engineered to solve this problem. Specifically, a plasmid vector was designed that carried an antibiotic-resistance gene. In the following example, ampicillin-sensitive cells are transformed with recombinant plasmids containing the ampicillin-resistance gene. The cells are then plated on media containing ampicillin (a form of penicillin); the results are shown in Figure 15.9.
Transformed cells—that is, those that that took up a recombinant plasmid carrying the ampicillin-resistance gene—can grow on the ampicillin-agar medium. Untransformed cells (cells that failed to take up a plasmid) lack the ampicillin-resistance gene and cannot grow on the ampicillin medium.
But there is still a question: How can you tell whether the cells that grew were really transformed by a recombinant plasmid containing a cDNA insert? It is possible that some of the transformants are nonrecombinant plasmids that still have the ampicillin-resistance gene! To address this issue, plasmids were further engineered to contain a streptomycin-resistance gene (as well as the ampicillin-resistance gene). But in this case, the restriction-enzyme sites in the plasmid that were to be used for recombination were placed in the middle of the gene. So inserting a cDNA into this plasmid would disrupt and inactivate the streptomycin resistance gene. This second bit of genetic engineering enabled harvesting only cells transformed with recombinant plasmids (i.e., those containing a cDNA insert).
We can tell transformants with recombinant plasmids apart from those with nonrecombinant plasmids by the technique of replica plating (Figure 15.10).
After colonies grow on the ampicillin-agar plate, lay a filter over the plate. The filter will pick up a few cells from each colony, in effect becoming a replica filter (i.e., a mirror image) of the colonies on the plate. Next, place the replica filter on a new agar plate containing streptomycin. Any colonies that grow on the filter must be streptomycin-resistant, containing only nonrecombinant plasmids. Colonies containing recombinant plasmids, those that did not grow in streptomycin, are easily identified on the original ampicillin-agar plate.
In practice, highly efficient recombination and transformation procedures typically reveal very few streptomycin-resistant cells (i.e., colonies) after replica plating. When this happens, the ampicillin-resistant cells constitute a good cDNA library, ready for screening
15.2.7 Identifying Colonies Containing Plasmids with Inserts of Interest
The next step is to screen colonies in a cDNA library for those containing a specific cDNA. Since cells typically make thousands of proteins at the same time, a cDNA library should contain thousands of cDNAs made from thousands of mRNAs. Finding a single cDNA of interest can require plating the cDNA library in a tube on more than a few agar plates.
Continuing with the example above, actual screening would be done using multiple replica filters of ampicillin-resistant cells. The number of replica filters to be screened can be calculated from assumptions and formulas for estimating the number of colonies to be screened in order to represent an entire transcriptome (i.e., the number of different mRNAs in the original cellular mRNA source (including in this example, globin mRNAs).
Once the requisite number of replica filters is made, they are subjected to in situ lysis to disrupt cell walls and membranes. The result is that the cell contents are released, and the DNA is denatured (unwound to become single-stranded). The DNA then adheres to the filter in place (in situ), where the colonies were. The result of in situ lysis is a filter with faint traces of the original colonies (Figure 15.11).
Next, a molecular probe is used to identify DNA containing the sequence of interest. The probe is often a synthetic oligonucleotide, the sequence of which was inferred from known amino acid sequences. These oligonucleotides are made radioactive and placed in a bag with the filter(s). DNA from cells that contained recombinant plasmids with a cDNA of interest will bind the complementary probe. The results of in situ lysis and hybridization of a radioactive probe to a replica filter are shown in Figure 15.12 (below).
264 Probing a Replica Plate Filter
The filters are rinsed to remove the unbound radioactive oligomer probe and then placed on X-ray film. After a period of exposure, the film is developed. Black spots will form on the film from radioactive exposure, creating an autoradiograph of the filter. The black spots in the autoradiograph in Figure 15.13 correspond to colonies on a filter that contain a recombinant plasmid with the target cDNA sequence.
From a positive on the film, recombinant colonies are located on the original plate and grown in liquid culture for plasmid DNA isolation. Next, cDNAs are sequenced and the amino acid sequences they encode are inferred from the genetic-code dictionary. Once verified, recombinant plasmids can be isolated, tagged (made radioactive or fluorescent), and used for various purposes:
- To probe for the genes from which they originated
- To identify and to quantitate the mRNA and even to locate the transcripts in the cells
- To quantitatively measure amounts of specific mRNAs
Isolated plasmid cDNAs can even be expressed in suitable cells to make the encoded protein. These days, diabetics no longer receive pig insulin; rather, they get synthetic human insulin made from expressed human cDNAs. Moreover, while the introduction of the polymerase chain reaction (PCR; see section 15.5) has superseded some uses of cDNAs, they still play a role in genome-level and transcriptome-level studies.