The RNAs that are transcribed from a genome under a given set of conditions can be determined using DNA sequencing methods described in Chapter 8. The approach is called RNA-Seq (Fig. 9-15). RNA is first isolated from a tissue or a population of cells. The RNA is fragmented and converted to double-stranded DNA using reverse transcriptase (see Fig. 9-12a). This DNA is then subjected to deep DNA sequencing, which reveals both the RNAs that are present and the relative abundance of each (if more copies of one RNA are present, they will give rise to more DNA sequencing reads). This method is sensitive enough to apply to single cells, an approach called single cell RNA-Seq, or scRNA-Seq. It allows investigators to catalog the RNAs being transcribed in different parts of a tissue.

The transcriptional state of a human cell or tissue can be diagnostic of conditions ranging from diabetes to cancer. If a particular gene under study is expressed in a certain tissue or under particular metabolic conditions, the result provides a new functional clue. Detailed knowledge about what genes are expressed in a given tumor may eventually help guide treatment options. RNA-Seq can reveal patterns of gene regulation and expression. It has special importance in studying cancerous tumors, in which rapid evolution triggered by genome instability creates a range of cell types. The mRNAs that are present in tumor cells provide a clue to the proteins that may be present, although not all mRNAs are immediately translated into protein. RNA-Seq also reveals the presence of many types of noncoding RNAs (described in Chapter 26) that are now being defined.

A more direct way to establish protein presence or absence is to assess the cellular proteome. Mass spectrometry (Chapter 3) can accurately catalog and quantify the thousands of proteins present in a typical cell. This approach is a complement to RNA-Seq, as it provides a comprehensive list of the genes that are both transcribed and translated into protein. Mass spectrometry also provides information about how those proteins are modified, in turn allowing an assessment of their regulatory state.

Often, an important clue to the function of a gene product comes from determining its location within the cell. For example, a protein found exclusively in the nucleus could be involved in processes that are unique to that organelle, such as transcription, replication, or chromatin condensation. Researchers often engineer fusion proteins for the purpose of locating a protein in the cell or organism. Some of the most useful fusions are the attachment of marker proteins that signal the location by direct visualization or by immunofluorescence.

A particularly useful marker is the green fluorescent protein (GFP) (Fig. 9-16), discovered by Osamu Shimomura. As subsequently shown by Martin Chalfie, a target gene (encoding the protein of interest) fused to the GFP gene generates a fusion protein that is highly fluorescent — it literally lights up when exposed to blue light — and can be visualized directly in a living cell. GFP is a protein derived from the jellyfish Aequorea victoria (Fig. 9-16a). The protein has a β-barrel structure with a fluorophore (the fluorescent component of the protein) in the center (Fig. 9-16b). The fluorophore is derived from a rearrangement and oxidation of three amino acid residues (Fig. 9-16c). Because this reaction is autocatalytic and requires no proteins or cofactors other than molecular oxygen, GFP is readily cloned in an active form in almost any cell. Just a few molecules of this protein can be observed microscopically, allowing the study of its location and movements in a cell.

Careful protein engineering by Roger Tsien, coupled with the isolation of related fluorescent proteins from other marine coelenterates, has made variants of these proteins available in an array of colors (Fig. 9-16d) and other characteristics (brightness, stability). If fusion to GFP does not impair the function or properties of a protein one wishes to study, the fusion protein can be used to reveal the protein’s location in the cell under a range of conditions and to detect interactions with other labeled proteins. With this technology, for example, the protein GLR1 (a glutamate receptor of nervous tissue) has been visualized as a GLR1-GFP fusion protein in the nematode Caenorhabditis elegans (Fig. 9-16e).

In some cases, the GFP fusion protein may be inactive or may not be expressed at sufficient levels to allow visualization. Immunofluorescence is an alternative approach for visualizing the endogenous (unaltered) protein. This approach requires fixation (and thus death) of the cell. The protein of interest is sometimes expressed as a fusion protein with an epitope tag, a short protein sequence that is bound tightly by a well-characterized, commercially available antibody. Fluorescent molecules (fluorochromes) are attached to this antibody. More commonly, the target protein is unaltered and is bound by an antibody that is specific for the protein. Next, a second antibody is added that binds specifically to the first one, and it is the second antibody that has the attached fluorochrome(s) (Fig. 9-17). A variation of this indirect approach to visualization is to attach biotin molecules to the first antibody, then add streptavidin (a bacterial protein closely related to avidin, a protein that binds biotin; see Table 5-1) complexed with fluorochromes. The interaction between biotin and streptavidin is one of the strongest and most specific known, and the potential to add multiple fluorochromes to each target protein gives this method great sensitivity. In all of these cases, the end product is a microscopic view of a cell in which a spot of light (a focus) reveals the location of the protein.

By fusing the gene encoding a protein under study with the gene for an epitope tag, investigators can precipitate the protein product of the fusion gene by complexing it with the antibody that binds the epitope. This process is called immunoprecipitation (Fig. 9-18). If the tagged protein is expressed in cells, other proteins that bind to it precipitate with it. Identifying the associated proteins reveals some of the intracellular protein-protein interactions of the tagged protein. There are many variations of this process. For example, a crude extract of cells that express a tagged protein is added to a column containing immobilized antibody (see Fig. 3-17c for a description of affinity chromatography). The tagged protein binds to the antibody, and proteins that interact with the tagged protein are sometimes also retained on the column. The connection between the protein and the tag is cleaved with a specific protease. The protein complexes are eluted from the column, and the proteins in them are identified by mass spectrometry. Researchers can use these methods to define complex networks of interactions within a cell. In principle, the chromatographic approach to analyzing protein-protein interactions can be used with any type of protein tag (His tag, GST, etc.) that can be immobilized on a suitable chromatographic medium.

The selectivity of this approach can be enhanced with tandem affinity purification (TAP) tags. Two consecutive tags are fused to a target protein, and the fusion protein is expressed in a cell (Fig. 9-19). The first tag is protein A, a protein found at the surface of the bacterium Staphylococcus aureus that binds tightly to mammalian immunoglobulin G (IgG). The second tag is often a calmodulin-binding peptide. A crude extract containing the TAP-tagged fusion protein is passed through a column matrix with attached IgG antibodies that bind protein A. Most of the unbound cellular proteins are washed through the column, but proteins that normally interact with the target protein in the cell are retained. The first tag is then cleaved from the fusion protein with a highly specific protease, TEV protease, and the shortened fusion target protein and any proteins associated noncovalently with the target protein are eluted from the column. The eluate is then passed through a second column containing a matrix with attached calmodulin that binds the second tag. Loosely bound proteins are again washed from the column. After the second tag is cleaved, the target protein is eluted from the column with its associated proteins. The two consecutive purification steps eliminate most weakly bound contaminants. False positives are minimized, and protein interactions that persist through both steps are likely to be functionally significant.

A sophisticated genetic approach to defining protein-protein interactions is based on the properties of the Gal4 protein (Gal4p; see Fig. 28-32), which activates the transcription of GAL genes (encoding the enzymes of galactose metabolism) in yeast. Gal4p has two domains: one that binds a specific DNA sequence, and another that activates RNA polymerase to synthesize mRNA from an adjacent gene. The two domains of Gal4p are stable when separated, but activation of RNA polymerase requires interaction with the activation domain, which in turn requires positioning by the DNA-binding domain. Hence, the domains must be brought together to function correctly.

In yeast two-hybrid analysis, the protein-coding regions of the genes to be analyzed are fused to the yeast gene for either the DNA-binding domain or the activation domain of Gal4p, and the resulting genes express a series of fusion proteins (Fig. 9-20). If a protein fused to the DNA-binding domain interacts with a protein fused to the activation domain, transcription is activated. The reporter gene transcribed by this activation is generally one that yields a protein required for growth or an enzyme that catalyzes a reaction with a colored product. Thus, when grown on the proper medium, cells that contain a pair of interacting proteins are easily distinguished from those that do not.

A library can be set up with a particular yeast strain in which each cell in the library has a gene fused to the Gal4p DNA-binding domain gene, and many such genes are represented in the library. In a second yeast strain, a gene of interest is fused to the gene for the Gal4p activation domain. The yeast strains are mated, and individual diploid cells are grown into colonies. The only cells that grow on the selective medium, or that produce the appropriate color, are those in which the gene of interest is binding to a partner, allowing transcription of the reporter gene. This allows large-scale screening for cellular proteins that interact with the target protein. The interacting protein that is fused to the Gal4p DNA-binding domain present in a particular selected colony can be quickly identified by DNA sequencing of the fusion protein’s gene. Some false positive results occur, due to the formation of multiprotein complexes.

“CRISPR” stands for clustered, regularly interspaced short palindromic repeats; as the name suggests, these consist of a series of regularly spaced short repeats in the bacterial genome. A Cas (CRISPR-associated) protein is a nuclease. The CRISPR sequences and Cas protein are components of a kind of immune system that evolved to allow bacteria to survive infection by bacteriophages. CRISPR sequences are embedded in the bacterial genome, surrounding sequences derived from phage pathogens that previously infected the bacterium without killing it. The viral sequences are, in effect, spacer sequences separating the CRISPR sequences. When the same bacteriophage again attacks a bacterium that has the corresponding CRISPR/Cas system, the CRISPR sequence and Cas protein act together to destroy the viral DNA. First, the CRISPR sequences are transcribed to RNA, and individual viral spacer sequences are cleaved to form products called guide RNAs (gRNAs), which include some adjacent repeat RNA. A gRNA forms a complex with one or more Cas proteins and, in some cases, with another RNA called a trans-activating CRISPR RNA, or tracrRNA. The resulting complex binds specifically to the invading bacteriophage DNA, cleaving and destroying it through the nuclease activities associated with the Cas proteins.

The current technology was made possible by discovery of a relatively simple CRISPR/Cas system in Streptococcus pyogenes. This system requires only a single Cas protein, Cas9, to cleave DNA. Work in many laboratories, particularly those of Jennifer Doudna and Emmanuelle Charpentier, has produced a streamlined CRISPR/Cas9 system composed of just one protein (Cas9) and one associated RNA, consisting of gRNA and tracrRNA fused into a single guide RNA (sgRNA). The power of the system is embedded in this sgRNA, in which the guide sequence can be altered to specifically and efficiently target almost any genomic sequence (Fig. 9-21). Cas9 has two separate nuclease domains: one domain cleaves the DNA strand paired with the sgRNA, and the other cleaves the opposite DNA strand. Inactivating one domain creates an enzyme that cleaves just one strand, forming a single-strand break, or nick. The sgRNA is needed both to pair with the target sequence in the DNA and to activate the nuclease domains for cleavage.

Plasmids expressing the required protein and RNA components of CRISPR/Cas9 can be introduced into microbial cells by electroporation (p. 306). For mammalian cells, the genes encoding the CRISPR/Cas9 components can be incorporated into engineered viruses that subsequently deliver them to the cell nuclei. For many organisms, the targeted gene is inactivated in a high percentage of the treated cells. If a genomic change (mutation) rather than a simple gene inactivation is required, it can be introduced by recombination when a DNA fragment encompassing the cleavage site and including the desired change enters the cell with the CRISPR/Cas9 plasmids. This recombination is often inefficient, but success can be improved somewhat by introducing a nick rather than a double-strand break at the target site (Fig. 9-21).

CRISPR/Cas9 can be combined with other approaches to extract additional information. For example, a particular gene can be inactivated with CRISPR/Cas9. Then, the effect of that gene inactivation on the transcription of other genes can be probed with RNA-Seq at the level of tissues, cell populations, or single cells.

New applications for CRISPR/Cas9 are being developed rapidly, both for basic research and for medicine. Genetic screens based on CRISPR are described in the next section. CRISPR is being used to enhance food production, provide new approaches to combat bacterial infections, and eliminate nonnative pest species that can harbor diseases (Box 9-1). New CRISPR-based treatments for genetic diseases are being cautiously advanced to clinical trials for vision loss due to inherited retinal dystrophies, Duchenne muscular dystrophy, β-thalassaemia, and many other conditions. Uncertainties remain, particularly the potential for occasional cleavage at unintended chromosomal sites (off-target cleavage). The impact of CRISPR/Cas9 will continue to grow as problems are overcome, current applications mature, and new applications are imagined and created.