Photobleaching is a photochemical process whereby powerful excitation of a fluorophore renders it permanently unable to fluoresce. The protein to which the fluorophore is attached remains biological active, effectively enabling users to generate 2 distinct populations of protein (fluorescent/non fluorescent). The F words describe a range of techniques that involve bleaching away the fluorophore using a short burst of targeted illumination and then recording redistribution of the remaining labeled population. These techniques enable users to quantitatively studying compartmentalization and turnover of labeled protein in sub cellular structures .
Fluorescence recovery after photobleaching (FRAP) involves bleaching away parts of the labeling and recording the recovery of the fluorescent signal in the target region through migration of unbleached protein into the bleached zones  (s. Figure 1). The plotting of recovery curves enables users to quantify recover rates and differentiate between mono and multi phasic recovery profiles. The technique enables the user to explore turn over and compartmentalization of structures of interest. Fluorescent labeling with fluorescent proteins only lets you observe where a protein is located; it does not allow you to peer inside seemingly solid labeled objects (vesicle/ filament) to understand how molecules move around and turn over within the structure. In addition to this, internal dynamics of objects may vary in different regions of the cell or in response to external stimuli like cytokines or pathogens. In short FRAP is still an extremely useful technique that can be employed to study the plasticity and turnover, and inter connectivity of structures within cells  (s. Figure 2).
Fluorescence loss in photobleaching (FLIP) is another bleaching technique that can be thought of as reverse FRAP. A target region is bleached repeatedly over time and an image acquired after each bleaching cycle. Unlike FRAP which looks for the recovery of fluorescence within the target region, FLIP looks at the steady loss of signal throughout the rest of the cell. The technique reveals the molecules’ mobility throughout the whole cell  and enables the user to identify regions of high and low turnover in multiple cellular compartments. This technique can be used to observe complex turnover pathways of a protein such as cytokeratin that otherwise may not be apparent. Whereas FRAP enables you to study the turnover within a specific region, FLIP enables you to observe turnover and compartmentalization of a molecule throughout the entire cell.
Förster resonance energy transfer (FRET), also known as fluorescence resonance energy transfer, is a process where the energy is transferred from one fluorophore to another and can be used to measure inter and intra molecular distances. FRET only occurs over extremely small distances (typically 1-10 nm) and so can be used to quantify and map out binding and protein interactions inside cells. Once excited, the donor fluorophore transfers energy to the acceptor fluorophore, which typically then emits at a longer wavelength than the donor. The efficiency of FRET is typically measured using ratiometric imaging or via changes in fluorescence lifetime imaging of the donor (FLIM). When FRET occurs, the donor emission is quenched and the acceptor emission channel increases. In FRET acceptor photobleaching the acceptor is bleached in a target region, so FRET can no longer occur, de-quenching the donor emission. The increase in the donor signal (de-quenched) can be used to directly measure the FRET efficiency.
As opposed to destroying the fluorescent properties of dyes via photo bleaching, photomanipulation can also be used to activate many photo sensitive fluorescent proteins. Many fluorophores have the ability to switch between different molecular states upon illumination at specific wavelengths, a property known as photochromism. Photochromism enables fluorophores to switch between low and high fluorescent states (photoactivation) , or alter their emission spectra on stimulation (photoconversion)  (s. Figure 3).
A well-known example of photoconversion is mEOS which exhibits a green to red switch upon activation with 405 nm, shifting its emission peak from 516 nm to 581 nm. Photoactivatable proteins enable users to track two distinct populations (activated/non activated) of one protein, whereas photoconvertible proteins can be useful when trying to look at multiple tagged proteins. Photoswitchable fluorescent proteins exhibit reversible photochromism and are able to cycle repeatedly between different states enabling users to switch them on and off (example Dronpa [6,7]). For more information see Photoactivatable, photoconvertible, and photoswitchable Fluorescent Proteins.
The primary use of photochromism when used with photomanipulations is to track activated protein populations over time.
It is worth noting that fluorophores exhibiting photochromism can also be used for single molecule localization super-resolution techniques such as in sptPALM and fPALM. Once a photoswitchable fusion protein has been produced it can often be used for multiple complimentary techniques [8,9].
Photochromic proteins are particularly useful when trying to observe the behaviors of abundantly expressed proteins. For example photoactivatable fluorophores enable users to easily track specific objects in otherwise crowded environments. This approach can be useful when trying to track individual objects, using long time-lapse intervals for reduced photo stress. Photoactive fluorophores also enable users to visualize diffusion between different cellular compartments, or examine plasticity and turn over inside seeming solid structures such as cytoskeleton filaments. The technique enables users to observe whether subunit exchange occurs between adjacent objects. Furthermore the technique enables users to study the lineage of objects from vesical fusion and splitting, to the location and identity of daughter cells . Finally photochromic proteins can also be used for pulse chase experiments whereby all the proteins in a cell are converted enabling users to differentiate between proteins made before and after the photomanipulation event. Finally some photoswitchable proteins can also be used as optogenetics actuators whereby in addition to switching the fluorescent properties of the dye, the conformational change that occurs is used to switch the activities of other conjugated biomolecules .
It is important to consider your application carefully and to select the right fluorescent protein (FP) for your requirements. Some photoactivatable FPs are significantly brighter than others, for example paGFP shows ~100 fold increase in brightness  when activated, whereas some proteins like Kaede show a 2000 fold increase in brightness . Brighter FPs enable users to track small quantities of proteins or allow users to achieve comparable signal to noise levels with much lower excitation settings, minimizing phototoxicity. It’s also worth noting that photoconvertible proteins can be easier to work with than photoactivatable proteins, as they are inherently visible prior to activation. Conversely they require using two imaging channels for one protein population, so users planning to image multiple proteins in a single experiment may be better off working with photoactivatable proteins.
In addition to controlling the properties of fluorescent labels, it’s also possible to use photomanipulation to selectively activate biological pathways, using photo sensitive protein domains via a process commonly referred to as optogenetics. Optogenetics tools enable users to stimulate cells on command in a highly targeted manner. Classic optogenetics involves targeted activation of photo sensitive ion channels to change the membrane voltage potential. The most well-known optogenetics effectors belong to the rhodopsin family. Rhodopsins are nonspecific cation channels, however anion conducting rhodopsins as well as ion specific channels have since been discovered or engineered . The downstream effect of activation can be measured using electrophysiology or via optical sensors such as ratiometric calcium dyes. Photo sensitive ion channels are extremely valuable tools as they permit high spatial and temporal resolution stimulation of neurons and other cell types. It is also worth noting that optical control of G protein mediated signaling is also possible .
In addition to the classic receptor based optogenetics actuators, a whole optogenetics tool kit is emerging with discovery or many other photo sensitive protein domains. Photo sensitive domains such as light, oxygen, voltage (LOV) domains, phytochromes or cryptochromes from plants, can be engineered into effector proteins enabling researchers to build in photo sensitive optogenetics actuators into almost any protein they want to study [14,16]. A LOV domains’ exposure to blue light introduces a conformational change. With some thoughtful protein engineering the change can be used to activate or deactivate target proteins of interest [17,18]. The LOV domain can be used to induce targeted DNA binding, protein dimerization and enzyme activation or deactivation.
One example of the use of LOV domain is to use it to produce a photo sensitive RAC Kinase. Activation enables the group to effectively control cell motility . Interestingly even gene editing tools like CRISPR/Cas9 have been bought under optical control, by the addition of photoinducible dimerization domains named Magnets, enabling optically targetable gene editing. The emerging optogenetics tool kit offers huge potential for new avenues of research.
As well as being used to adjust protein activity, or the fluorescent sates of dyes, photomanipulation can be used to manipulate physical and chemical states of the sample itself. By replacing continuous wave lasers commonly used for imaging with high power pulsed lasers, it becomes possible to physically destroy parts of the sample or induce photolysis of chemical agents in and around the sample. With cutting and ablation, depending on the power applied to the sample, effects can vary from cutting individual cytoskeletal fibers or amputations of parts of a cell, through to total ablation of cell clusters (s. Figure 6). This ability can be used for a number of different applications enabling users to study how organisms react to trauma and to investing the downstream biological pathways involved in repair. These capabilities have a wide range of applications from exploring the role of single microtubules in mitosis [21,22] through to the mechanics behind wound healing, inflammation and metastatic processes. Ablation can be used to make wounds in tissues, enabling researchers to explore signaling pathways, cell migration and the kinetics involved in complex multi cellular processes. In developmental biology ablation can be used to knock out parts of an embryo to see the impact on development and organization of a whole organism.
Furthermore, photomanipulation techniques can be used to introduce biologically active compounds to a sample in a highly targeted manner. Many organic compounds undergo photolysis when exposed to UV light. Using organic chemical synthases it’s possible to design chemically encapsulated compounds that are biological inert when intact, but which degrade into biologically active compounds when exposed to UV light. Such compounds are referred to as caged compounds and targeted photolysis of these compounds is referred to as uncaging. A common use for uncaging is in neurobiology where it can be employed to introduce active neuro transmitters into single synaptic junctions. To date a wide range of caged compounds is commercially available and includes neuro transmitters such as glutamate and GABA, nucleotides like ATP and cAMP, ions such as calcium, and even some macromolecules (s. Figure 7). Molecules such as proteins, peptides, DNA and RNA can also be caged with commercially available reagents that covalently modify specific residues .
In addition to using photomanipulation to physically destroy the material in the sample, it is also possible to inflict very specific types of damage to a cell. One such example is using a UV laser to introduce targeted breaks into the DNA. This technique can be used to trigger and study cellular DNA repair mechanisms. Many of the proteins involved in recognizing and responding to DNA damage play a key role in both the development and treatment of cancers and so the ability to activate these pathways on demand offer an extremely valuable cancer research tool .
Photomanipulation techniques offer an extremely versatile set of research tools, as well as the applications already described, the photomanipulation tool box also offers several other less well known capabilities. These include the use of