What is Photomanipulation?

Schlicker, Oliver; Laskey, Peter; Greb, Christoph

May 15, 2017

What is Photomanipulation?

The new Scientific Tool Box. Why is it important and how can you use it?

The term photomanipulation describes a wide range of techniques that enable the microscopist the transition from passive observer to instigator of events by offering a way of interacting with their sample via targeted illumination. Typically researchers are trying to observe specific processes of interest in order to understand the underlying biological process. Microscopists are often forced to hunt through large populations of cells or acquire hours of time laps footage before they’re able to observe events of interest and in many cases it’s simply not possible to observe certain processes using conventional microscopy techniques alone. Photomanipulation tools enable the microscopist to initiate biological events, precisely adjusting sample labeling, biological activity, local chemical environments and in some instances physically destroy parts of their specimen. These capabilities are hugely valuable when it comes to unpicking complex, compartmentalized biological mechanisms. Photomanipulation tools enable the users to design advanced experiments to access additional layers of information from their specimen that are not possible from observation alone.

Photobleaching and the F words

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 <link science-lab basic-principles-of-luminescence>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 [1].

Fluorescence recovery after photobleaching (<link science-lab fluorescence-recovery-after-photobleaching-frap-and-its-offspring>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 [2] (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 [3] (s. Figure 2).

Figure 1: FRAP experiment in yeast cells. The P-Bodies of these yeast cells carry a fluorescently labelled protein. In the course of a FRAP experiment a small structure can be bleached and the recovery can be observed (arrow).

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 [4] 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 (<link science-lab foerster-resonance-energy-transfer-fret>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 <link science-lab ratiometric-imaging>ratiometric imaging or via changes in fluorescence lifetime imaging of the donor (<link science-lab fret-with-flim>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.

Figure 2: Figure 1: During fluorescence recovery after photobleaching (FRAP) the fluorescence tag of a protein is bleached by powerful excitation inside a region of interest (ROI). Subsequent resettlement gives insight e.g. into the turnover of the structure of interest. For fluorescence loss in photobleaching (FLIP) a ROI within a cell is bleached repeatedly. The bleached molecules spread out and the loss of fluorescence indicates, e.g. whether cell organelles are physically connected. During inverse FRAP (iFRAP) the cell region outside the ROI is bleached. Subsequently the non-bleached molecules spread out and can be monitored.

Photochromism

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) [5], or alter their emission spectra on stimulation (photoconversion) [6] (s. Figure 3).

Figure 3: Photoactivation, photoconversion, and photoswitching can be used to track protein populations over time. Photoactivatable proteins can be “switched on” by light of a certain wavelength. Photoconvertible proteins can change their emission spectrum, whereas photoswitchable proteins can be switched on and off many times.

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 <link science-lab photoactivatable-photoconvertable-and-photoswitchable-fluorescent-proteins>Photoactivatable, photoconvertible, and photoswitchable Fluorescent Proteins.

The primary use of photochromism when used with photomanipulations is to track activated protein populations over time.

Figure 4: For this photoactivation experiment HeLa cells were transfected with pamCherry-H2B and eGFP-Actin. After photoactivation pamCherry exhibits bright fluorescence.

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 [3]. 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 [11].

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 [12] when activated, whereas some proteins like Kaede show a 2000 fold increase in brightness [13]. 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.

Optogenetics

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 <link science-lab optogenetics>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 [14]. The downstream effect of activation can be measured using electrophysiology or via optical sensors such as <link science-lab ratiometric-imaging-needs-a-specialized-setup>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 [15].

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 (<link science-lab optogenetics clap-on-clap-off-protein-on-protein-off>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.

Figure 5: Optogenetics utilizes photo sensitive protein domains to trigger cellular mechanisms. Classically photo sensitive ion channels were used to change the membrane potential. Meanwhile additional proteins have been domesticated which show conformational changes after irradiation with light. This can be used to activate or deactivate target proteins e.g. to induce protein resp. DNA binding or enzyme activation.

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 [19]. Interestingly even gene editing tools like <link science-lab crispr-cas gene-editing-with-crisprcas9-breakthrough-in-genome-engineering>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.

Cutting and ablation

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.

Figure 6: To investigate neuronal regeneration capabilities, axons can be cut with the help of a pulsed laser (arrow). Ablation and cutting experiments range from cutting whole cells or cell fragments down to single cytoskeletal elements. Courtesy of Richard Eva, Cambridge University (UK).

Uncaging

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 [23].

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 [24].

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 IR lasers to introduce targeted thermal stress to cells. Photomanipulation devices can also be used for micro pattering of substrates including protein printing and etching onto glass to produce markers for correlative microscopy. Multi photon lasers can also be used for a range of photomanipulation techniques like cutting, uncaging, photo switching and stimulations, with the main advantage of improved 3 dimensional targeting capabilities. Finally, it is also worth mentioning that it is possible to combine techniques such as both bleaching and activating of markers like mEOS to get a deep insight into some complex processes [25]. Furthermore bleaching and activation techniques can be used to investigate changes in dynamic processes that occur before and after targeted stimulation via uncaging or optogenetics actuators, or to explore cellular responses to trauma from cutting or ablation events.

Hardware for photomanipulation

There are several different types of technology that can be used for photomanipulation, all of which have their strengths and weaknesses (s. Figure 8). The simplest approach is to use a field diaphragm in the florescent light path to illuminate the center of the field of view with a specific wavelength and to then open the field diaphragm up again and switch wavelengths for imaging. This approach is slow, offers poor targeting flexibility and only provides low power illumination, so is not suitable for most experiments. However it is sufficient for basic photo activation and can be used to test the activity of constructs. As most photomanipulation experiments require more powerful illumination, lasers are typically the preferred illumination source. Laser devices such as the <link products light-microscopes life-science-research inverted-microscopes the-leica-dmi8 details product leica-wf-frap>Leica WF FRAP offer excellent tools for basic FRAP experiments. A powerful laser source is targeted to the center of the image, the profile of the beam in the sample can be modified using masks enabling users to shoot a limited range of ROI’s sizes and shapes. The limitation of these devices is that they typically offer a single laser line limiting the types of experiments they can be used for. The other restriction is that as they cannot target multiple or complex ROI’s with a single experiment, many experiments are beyond the ability of such devices.

More advanced photomanipulation experiments often require the ability to repeatedly target any area in the specimen with a range of wavelengths. There are two main technologies which offer this ability, namely digital mirror devices (DMD), and galvo scanning devices. The digital mirror device uses a chip that contains thousands of tiny mirrors that can be switched to an on or off position electronically at high speed. The entire chip is illuminated using either a laser or LED, depending of the pattern of activated mirrors the users are able to project complex illumination patterns onto their sample. DMDs enable users to simultaneously illuminate multiple regions of interest. The main weakness of DMD’s is the relatively low maximum power densities they’re able to achieve, as the illuminating light is spread out over the entire field of view. This makes DMDs good for some applications such as some optogenetics applications, but they struggle with high speed power hunger applications like bleaching and cutting and ablation. Galvo scanning devices use a high speed galvo scanner to guide a focused laser beam over a sample making them the most versatile type of photomanipulation device. Galvo scanners often come as stand-alone photomanipulation device, but many confocal scanners can also be used. Depending on the lasers coupled in, they can address almost any application. The use of a diffraction limited beam enables galvo devices to bleach and activate very efficiently. They can also be used for cutting and ablation applications. Some galvo scanners can also adjust their beam profile enabling users to have greater flexibility when designing there experiments.

Figure 8: Hardware options for photomanipulation experiments.

It is important to consider that a photomanipulation device is only part of a complete system, so the level of integration of the device into the system from both an optics and software perspective should be considered. A well-integrated device should not compromise the optical path and the software should enable seamless interlacing of imaging and manipulation tasks. Ease of operation vs the ability to design complex advanced experiment protocols should also be considered.

In summary, photomanipulation capabilities offer an extremely powerful addition to any live-cell imaging system due to the huge range of capabilities they can offer. Photomanipulation can be used to achieve a wide range of effects on the sample, with photobleaching and activation techniques enabling the users to uncover otherwise hidden dynamic process. Uncaging and the fast evolving field of optogenetics now enable users to stimulate or inhibit almost any biological pathway with subcellular resolution and ms precision. Finally, cutting and ablation techniques enable researchers to perform micro surgery on samples. These capabilities when combined with modern imaging systems offer huge potential to further our understanding in a wide number of research fields.