Chapter 7 – Complementary Experimental Tools  279

7.4.4  DNA-​ENCODED REPORTER TAGS

As outlined previously, several options exist for fluorescent tags to be encoded into the DNA

genetic code of an organism, either directly, in the case of fluorescent proteins, or indir­

ectly, in the case of SNAP/​CLIP-​tags. Similarly, different segment halves of a fluorescent

protein can be separately encoded next to the gene that expresses proteins that are thought

to interact, in the BiFC technique, which generates a functional fluorescent protein molecule

when two such proteins are within a few nanometers’ distance (see Chapter 3).

Most genetically encoded tags are engineered to be at one of the ends of the protein under

investigation, to minimize structural disruption of the protein molecule. Normally, a linker

sequence is used of ~10 amino acids to increase the flexibility with the protein tag and reduce

steric hindrance effects. A common linker sequence involves repeats of the amino acid

sequence “EAAAK” bounded by alanine residues, which is known to form stable but flexible

helical structures (whose structure resembles a conventional mechanical spring). The choice

of whether to use the C-​ or N-​terminus of a protein is often based on the need for binding

at or near to either terminus as part of the protein’s biological function, that is, a terminus is

selected for tagging so as to minimize any disruption to the normal binding activities of the

protein molecule. Often, there may be binding sequences at both termini, in which case the

binding ability can still be retained in the tagged sequence by copying the end DNA sequence

of the tagged terminus onto the very end of the tag itself.

Ideally, the native gene for a protein under investigation is entirely deleted and replaced

at the same location in the DNA sequence by the tagged gene. However, sometimes this

results in too significant an impairment of the biological function of the tagged protein, due

to a combination of the tag’s size and interference of native binding surfaces of the protein.

A compromise in this circumstance is to retain the native untagged gene on the cell’s genome

but then create an additional tagged copy of the gene on a separate plasmid, resulting in a

merodiploid strain (a cell strain that contains a partial copy of its genome). The disadvantage

with such techniques is that there is a mixed population of tagged and untagged protein in

the cell, whose relative proportion is often difficult to quantify accurately using biochemical

methods such as western blots (see Chapter 6).

A useful tool for researchers utilizing fluorescent proteins in live cells is the ASKA library

(Kitagawa et al., 2005), which stands for “A complete Set of E. coli K-​12 ORF Archive.” It

is a collection (or “library”) of genes fused to genetically encoded fluorescent protein tags.

Here, each open reading frame (or “ORF”), that is, the region of DNA between adjacent

start and stop codons that contains one or more genes (see Chapter 2), in the model bac­

terium E. coli, has been fused with the DNA sequence for the yellow variant of GFP, YFP.

The library is stored in the form of DNA plasmid vectors under IPTG inducer control of

the lac operon.

In principle, each protein product from all coding bacterial genes is available to study

using fluorescence microscopy. The principal weakness with the ASKA library is that the

resultant protein fusions are all expressed at cellular levels that are far more concentrated

than those found for the native nonfusion protein due to the nature of the IPTC expres­

sion system employed, which may result in nonphysiological behavior. However, plasmid

construct sequences can be spliced out from the ASKA library and used for developing

genomically tagged variants.

Optogenetics (see Pastrana, 2010; Yizhar et al., 2011) specifically describes a set of

techniques that utilize light-​sensitive proteins that are synthetically genetically coded into

nerve cells. These foreign proteins are introduced into nerve cells using the transfection

delivery methods of molecular cloning described earlier in this chapter. These optogenetics

techniques enable investigation into the behavior of nerves and nerve tissue by controlling the

ion flux into and out of a nerve cell by using localized exposure to specific wavelengths of vis­

ible light. Optogenetics can thus be used with several advanced light microscopy techniques,

especially those of relevance to deep tissue imaging such as multiphoton excitation methods

(see Chapter 4). These light-​sensitive proteins include a range of opsin proteins (referred to as

luminopsins) that are prevalent in the cell membranes of single-​celled organisms as channel

protein complexes. These can pump protons, or a variety of other ions, across the membrane