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Imaging & Photomanipulation - Approaches

The Imaging and Photomanipulation group is developing and expanding a variety of different kinds of imaging technologies for application to migration research both in vitro and in vivo.



Photoactivation can provide instantaneous in vivo activation of various physiological modulators of adhesion and migration that are introduced to the cell prior to the experiment. These molecules are originally inactive, due to a caging moiety covalently attached to or near their active site. Upon brief (5 - 500 ms) irradiation with light, typically UV, the caging moiety is photolytically released, thus activating the molecule of interest (Figure 1). It is also possible to use epifluorescence excitation simultaneously with photoactivation, in order to monitor indicator fluorescence of some other molecule that is known or suspected to be involved in a particular signaling pathway or cell behavior, e.g., cell locomotion (Ishihara et al., 1997). A mercury arc lamp (365 nm), a He-Cd (354 nm) or an argon ion multiline laser can be used as UV light sources.

Figure 1. Illustration of the basic principle of photoactivation; the caged protein or peptide of interest is present (purple) but inactive; the UV light severs the bond between the protective caging group and the protein, and activates the protein (red); the uncaging by-product can be neutralized by a scavenger molecule (green). >

Caged molecule synthesis and characterization: The caging moiety is usually a nitrobenzyl-based group (e.g., for thymosin 4) or carboxylic acid (e.g., for cofillin), chosen so as to form a bond that is stable under physiological conditions and that cleaves only due to photoisomerization (Marriott & Walker, 1999). In some cases the scavenger molecules also have to be added in order to neutralize the by-products of photoactivation. One of the considerable difficulties in using the caged proteins and peptides is that their design and synthesis involve complex procedures that are still being perfected. Some caged proteins are available commercially, but in majority of studies the caged reagents are custom-synthesized in biochemistry labs. In one group of procedures, developed for the studies of actin dynamics and related signaling pathways, the reactive amino acid residues are chemically modified within the key recognition sequences, yielding a caged peptide (Marriott, 1994; Wood et al., 1998; Pan & Bayley, 1997). Another, considerably challenging method that yields only small quantities of caged protein, is based on introduction of photolabile amino acids at a desired site on the chain, using the nonsense codon suppression technique (Noren et al., 1998; Bain et al., 1998). The already caged amino acids can also be introduced into polypeptides and small proteins using solid-phase peptide synthesis technique.

Characterization of the caged protein or peptide involves estimating the caging efficiency and comparison of molecule activities before and after uncaging. It is also important to confirm that the by-products of photolysis do not react with other proteins in the cell or that they are neutralized by scavengers. The quantum yield for uncaging is used as a measure of the photoactivation efficiency of a caged molecule of interest, relative to a known caged molecule (e.g., caged Pi) (Ellis-Davies & Kaplan, 1994). A higher irradiation power results in shorter uncaging times, thus increasing the photoactivation efficiency, but it may also cause considerable cell damage. Optimal photoactivation requires the highest irradiation power and the shortest uncaging times at which there is no discernable cell damage. Finally, the final local concentration of the uncaged molecule depends on the size of the caged molecule: small molecules diffuse rapidly into the cytoplasm and outside the irradiated region, so higher concentration of caged molecule is needed to cause the desired perturbation.

Introducing the caged molecules in vivo: The caged protein or peptide is introduced into the cell by microinjection or bead loading. Microinjection is used to introduce the caged molecules into specific cells, by use of commercially available pressure-driven precision pipette system (Eppendorf FemtoJet Microinjector) capable of delivering as little as several hundred femtoliters per cell. An inexpensive alternative to microinjection is bead loading, which is also employed for cell types that are too sensitive to damage due to microinjection. When sprinkled over the cells, the beads create transient pores in cell membranes allowing for the molecules in the culture medium to enter the cell. The loading efficiency of this nonselective technique is very low compared to microinjection and much larger quantities of caged molecule are needed to achieve the same results.

Some applications to the studies of cell migration: A study of migrating eosinophils (Walker et al., 1998) employed two different peptides targeted against calmodulin and myosin light chain kinase. The peptides were caged with 9-fluorenylmethoxycarbonyl amino acid moieties, and photoactivated globally with a series of 5 ms pulses of an argon ion laser (near-UV), resulting in rapid (within 10 s) inhibition of lamelipod protrusion and cell net movement.

In a more recent study, local photoactivation of caged thymosin 4 (actin sequestering protein) was shown to cause turning of fast-moving keratocytes (Roy et al., 2001). The activated thymosin 4 binds to and sequesters G-actin, reducing the actin polymerization rate and, consequently, arresting the movement within irradiated region, thus creating a pivot point around which the cell turns. In this case, the ([n-nitroveratryl]oxy)chlorocarbamate caging group was released by a 100 ms burst of UV irradiation produced by a He-Cd laser (354 nm), resulting in a visible effect on migration within tens of seconds after uncaging. A slight delay is believed to be due to the 'inertia' in signaling.

In summary, unlike conventional genetic and biochemical methods, photoactivation allows high spatiotemporal control of molecular perturbations in the cell. The ability to regulate the activity of a molecule or a signaling pathway has been and will be particularly useful for studies of cell migration.

Caged phosphopeptides and phosphoproteins: Photoactivation can also be used for the generation of active phosphopeptides and phosphoproteins from caged and inert precursors. When appropriately caged precursors are available, uncaging via UV light mimics the biological process of kinase-mediated phosphorylation without a dependence upon the upstream pathway. The spatial and temporal control afforded by photochemical activation also enables new approaches for understanding complex signaling pathways.

The Imperiali group has recently developed the synthetic methodology for the chemical synthesis of caged phosphoserine, phosphothreonine and phosphotyrosine building block that are suitable for solid phase peptide synthesis (Rothman et al., 2002; Rothman et al., 2003) The methodology is general and enables for the solid phase synthesis of caged phosphopeptides including phospho-serine, threonine and tyrosine. For example, the synthesis of caged phosphoserine is illustrated below. Peptides up to 35 amino acid residues have been prepared using the caged phosphoamino acid building blocks.

Recent publications illustrating biological applications of the caged phosphopeptide methodology have recently appeared in the literature (Vazquez et al., 2003; Nguyen et al., 2004).

This is summarized in the following animation;



The Imperiali group is currently focusing research efforts upon the integration of caged phosphoamino acids into native protein structures. Two approaches are being adopted;

I. Protein total synthesis or semisynthesis; This procedure involves large segment coupling via native protein ligation or expressed native ligation using intein chemistry. The methodology is practically limited to the incorporation of unnatural amino acid residues within 40 amino acids of the C- or N-terminus of the proteins. These methods have been pioneering Muir, Dawson, Kent. For a review on the semisynthesis of proteins see Dawson & Kent, 2000.

II. Hijacking mRNA translation; Native proteins with caging at internal phosphorylation sites will be accessed via chemically mischarged aminoacyl tRNAs as developed by Schultz, Chamberlain, Hecht, and Dougherty. (Hohsaka & Sisid, 2002; Hendrickson et al., 2004)

Chromophore-Assisted Laser Inactivation (CALI)

This technique uses a 620nm pulsed laser to photoactivate malachite green (MG) dye molecules, causing local damage to a protein of interest by the subsequent generation of hydroxyl radicals (Jay, 1988). An antibody against the protein of interest is labeled with this chromophore (6-10 dyes per antibody is optimal (Beerman & Jay, 1994), which needs to be positioned within 15 Å of the protein of interest in order for the radicals to reach it (Liao et al., 1994) due to their short lifetimes (10-9 to 10-12s). Thus any unbound MG-labelled antibodies cause no damage, due to the short penetration distance of the radicals, and only the bound proteins which fall within the diameter of the laser spot are inactivated. In this way a non-function-blocking antibody is converted into an inactivating reagent with high spatial and temporal resolution.

Figure 2. Antibody binds specifically to protein A and is armed with the covalently bound dye (green circles) which when irradiated by the laser light (red arrows) causes the dye to generate reactive hydroxyl radicals. The proximity of the antibody to its binding partner enables the short half-life, highly reactive radicals to damage protein A while leaving other proteins in the mixture (e.g. B) unaffected. (adapted from Jay & Sakurai, 1999)

CALI therefore allows the inactivation (within minutes) of a specific protein in a living cell. Recovery requires new synthesis of the protein, and so can take hours or days. Thus the dependence of cellular processes on this protein can be investigated in situ, within this window of time.

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Micro-CALI: A modification of CALI, called micro-CALI, uses microscope optics to focus the laser beam such that only a 10 µm spot in a cell is photoactivated. This acute inactivation of the protein of interest means that CALI-treated regions of the cell can be compared with other non-irradiated areas as an internal control, and that statistical analysis can be performed on the quantitative data acquired. Also, effects of protein activity in a particular region of the cell can be determined during processes such as motility and cell shape change, e.g. of Myosin V and ezrin respectively (Wang et al., 1996; Lamb et al., 1997).

CALI in practice: This technique has been used successfully on enzymes, cytoskeletal proteins, membrane receptors, signal transduction molecules and transcription factors (see table 1, Jay & Sakurai, 1999); in processes such as neuronal growth cone guidance (Chang et al. 1995; Buchstaller & Jay, 2000) and cell fate switching (Schmucker et al., 1994) and has been shown to phenocopy Drosophila genetic loss of function mutations precisely, in vivo (Schroder et al., 1999). In some cases it has failed to inactivate the protein of interest (e.g. hexokinase, actin) and there could be several contributing factors to this failure: a) the geometry of the binding site is such that the MG is more than 15 Å from the functional domain(s) of the protein relevant to the process being studied, b) that domain is not sensitive enough to damage by hydroxl radicals, and c) the protein is so abundant that residual activity is enough for normal functioning of the cellular process to continue. These must be kept in mind as possible causes of a null-result when interpreting the results of a CALI experiment.

Similarly, if the protein of interest is part of a complex, there is the possibility that functional domains of neighboring proteins will fall within this 15 Å radius of the MG label, depending on the exact antibody binding site and geometry, although this has not been observed to date.

To check that the experimental set-up is functional, one must first of all test that the antibody is specific and recognizes the protein of interest. For intracellular antigens, the loading of the MG-tagged antibody can be accomplished by several methods, depending on the number of cells required. Electroporation, scrapeloading and trituration are used for bulk loading, (Sydor et al., 1996) and microinjection can be used for experiments needing less than 100 cells, (Wang et al., 1996) although the success of the loading method also depends on the cell type. Lastly, in vitro assays using purified protein with the MG-labelled antibody should be performed, to show that laser irradiation leads to loss of function in this simplified environment.

Although CALI can show that acute loss of function of a specific protein has a particular effect on a cellular process, it proves nothing about whether the link is direct or not. There is also no information on the molecular mechanism by which the protein of interest is ultimately inactivated. e.g. inactivation of kinesin in a motility assay can lead to either immobilization of microtubules (through irreversible attachment) or detachment from microtubules (through loss of motor activity), depending on whether the photoactivation takes place in the presence of or prior to the addition of microtubules (Surrey et al., 1998). So CALI may be arresting the protein in a particular state of its biochemical cycle, or may be destroying its activity totally.

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Advantages of CALI are a) high spatial and temporal resolution for deactivation of a protein, b) it can be used to study the function of proteins whose absence causes embryonic lethality, since knockout methods are of no use for investigating processes involving these proteins at later stages of development. c) there is less likelihood of genetic compensation occurring (Wang & Jay, 1996). d) CALI's photochemical mechanism of protein inactivation is much simpler to perform than other target validation strategies for functional genomics, such as mouse or invertebrate knockouts and genetic screening. Antibody library screening for proteins which are involved in particular cellular processes could also be greatly enhanced by the function-perturbing effect of CALI.

Alternative chromophores for CALI: 620 nm light is not damaging to cellular components, hence the use of MG as the CALI chromophore. However, fluorescein is more soluble than MG, and requires orders of magnitude less energy to inactivate the protein having a 50-fold higher inactivation efficiency than MG, (in fact in neuronal growth cones, the inactivating effect of fluorescein after 30 s of illumination matched that of MG after 5 minutes of photoactivation (Nakai & Kamiguchi, 2002). It has also been shown that pulsed illumination is not necessary, continuous-wave light achieving the same or higher levels of specific inactivation (Surrey et al., 1998). Thus with fluorescein as the chromophore, a microscope lamp can be used for CALI experiments, rather than a high-energy pulsed laser. Two disadvantages of using fluorescein rather than MG are that the spatial resolution of the CALI effect is reduced to a radius of 30nm, (although higher resolution than this is often unnecessary, especially for in vitro experiments) and that the use of 488 nm light for the photoactivation may be problematic since other proteins may be present in the cell which have chromophores that naturally absorb blue light.

Most recently, EGFP has been used instead of MG as the chromophore for in vivo micro-CALI, to inactivate specific focal adhesion components using a 488 nm laser beam focused to a 2.2 µm diameter spot (Rajfur et al., 2002). This is a less invasive method than the antibody-based MG one, but since EGFP is used for normal confocal imaging because it is relatively non-phototoxic, a light dose of the order of 106 times greater than that used in typical imaging is needed to inactivate the protein. So EGFP has a poor efficiency as a CALI dye, but on the plus side, it allows the possibility of observing the cellular locations of the protein of interest whilst investigating its function, and also avoids the difficulties involved in finding a non-inhibitory antibody against the target protein and introducing the dye into living cells, being attached by gene fusion instead.

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Other developing methods for dye attachment include 1) His-tag-binding dyes, that are more easy to introduce into cells due to their smaller size, 2) an epitope tag genetically inserted close to the active site of the protein (eg., for HAkinesin, used to avoid inhibiting functionality of the motor domain) which provides a general method for chromophore attachment by eliminating the need to find a non-inhibitory antibody against each new target protein, and 3) for proteins that undergo biotinylation, chromophore-labeled streptavidin provides an indirect inactivator, although generally more energy is needed to inactivate the protein with this method. (Surrey et al., 1998)

Recently, the use of FLASH (4',5'-bis(1,3,2-dithioarsolan-2-yl)fluorescein), a membrane permeable arseno fluorescein derivative, has been introduced as a CALI/FALI reagent to inactivate synaptogamin I (Marek & Davis, 2002). In this technique, a tetracysteine motif is incorporated into the target protein so that protein will bind FLASH. This variant has the advantage of employing fluorescein, a more effective CALI reagent. The disadvantage is that FLASH may non-specifically bind to some cells (Stroffekova et al., 2001). This means that the amount of externally introduced FLASH must be carefully titrated to obtain specific binding but avoid non-specific binding. Other FLASH binding motifs are being developed to reduce non-specific binding.

The choice of chromophore thus determines the spatial resolution of CALI from the lifetime of the radicals produced, the energy needed for generation of these radicals, and the possible methods by which the chromophore can be bound to the protein of interest. Investigation into this area also provides a systematic study of photodamage, which is a factor that needs to be taken into account in all fluorescent microscopy techniques.

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Fluorescence Recovery After Photobleaching (FRAP)

Cell migration is only possible because of the highly coordinated intracellular movement of many proteins in response to various signals. There are many processes that contribute to the mobility of proteins and macromolecules within the cell, e.g. diffusion, binding to other molecules, active transport and contractile flow, to name but a few. The intracellular environment, which varies greatly between cellular compartments such as the nucleoplasm, Golgi, cell membranes and the cytoplasm, is another important factor that affects the mobility of the molecule. In combination with a mathematical model that suitably describes this environment, quantitative analysis of the mobility data gathered in a FRAP experiment allows the extraction of parameters which characterize the different processes involved in protein kinetics, leading to a determination of their relative contributions to the in vivo motion of the molecule being examined.

FRAP technique: Based on the phenomenon of photobleaching, FRAP is a particularly powerful technique for evaluating the distribution of molecules within or on living cells. Cells are loaded with the molecule of interest fluorescently labeled with tags such as fluorescein or GFP (Initial condition in figure below). A laser beam directed at a defined region of the cell (circled region in figure below + or - 0.5 µm) destroys the fluorescence emission from that region (dark spots in center of circle). The exchange of bleached fluorophores for unbleached fluorophores (blue spots) leads to the subsequent recovery of fluorescence into the region and reflects the type of transport processes occurring. Recording the changes in fluorescence intensity over time within the region yields measurements of protein mobility, as an average for the population. Two of the parameters that can be extracted are the rate of mobility and the mobile fraction of the population.

Figure 3. Simple diagram demonstrating FRAP within a cell and a typical recovery curve with Fi - initial fluorescence, F0 - fluorescence immediately after bleaching and Finfinity - fluorescence after complete recovery. (Adapted from Roy et al., 2002.)

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Rate of mobility: In the absence of active transport and unidirectional flow, the spatiotemporal dynamics is assumed to be due to diffusion, thus the protein mobility is expressed in terms of the diffusion coefficient (D). D is related to the characteristic diffusion time D and there are different mathematical models of varying complexity for describing this Brownian motion, both in two dimensions (e.g. within membranes or thin films of liquid, (Axelrod et al., 1976) and taking into account the membrane-bound topology of three dimensional compartments, (e.g. the cytoplasm or nucleoplasm, where recovery can be due to molecules from outside the focal plane). They also take into account such factors as viscosity and the hydrodynamic radius of the particle.

In practice there are many barriers to completely random motion that are not accounted for by the simpler diffusion models, so the measured parameter is usually termed the effective diffusion coefficient. For example, according to calculations based on molecular weight alone, most proteins move slower than expected; and biologically inactive molecules move 10-200 times as fast as similar sized proteins that are active (Phair & Misteli, 2000). Recently, more complex models have been developed which do allow quantification of the effects of some of these influences, such as binding and diffusion events, on in vivo movement. (Table 2. Carrero et al., 2003 summarizes mathematical models for nuclear protein diffusion).

Mobile fraction: As the name suggests, this expresses the ratio between the molecules that eventually move out of the region of interest (ROI) and the total photobleached sub-population. It is defined as R = (Finfinity - F0)/(Fi - F0) (see figure 3 above) (Reits & Neefjes, 2001) and depends strongly on the environment. Any barriers to free diffusion, such as membranes, the presence of microdomains or binding to other molecules have a strong effect on the mobile fraction.

Association/dissociation rates, and binding constants can also be found from fitting equations to the recovery curve, and information has been extracted about the dynamics of many proteins crucial to cell motility. Example FRAP studies include: cytoplasmic actin dynamics, to look at both the diffusion kinetics of G-actin (short term recovery) and also the movement of actin monomers between the two pools/subpopulations of G- and F-actin within the cytoplasm (long term decay constant) (Tardy et al., 1995; McGrath et al., 1998). Microtubule turnover in the leading edge of migrating cells, to show its dependence on F-actin convergence speed during contractile flow (Gupton et al., 2002). Plasma membrane microviscosity, to show how this and membrane composition physiologically regulate motility of endothelial cells (Ghosh et al., 2002).

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Determining experimental parameters: For quantitative analysis, time lapse images of this influx of fluorescent molecules into the bleached region must have low background noise and a high dynamic range. Thus the optimum photobleaching conditions for this need to be ascertained. Since most FRAP experiments last only minutes, phototoxicity is not usually a problem, experiments having been performed over time courses of more than 24 hours without any phototoxicity being observed (Ellenberg et al., 1997). Whether there is damage occurring during photobleaching has been addressed in a number of studies (Jacobson et al., 1978; Wolf et al., 1980).

Sampling about 50 data points during the recovery period is sufficient to identify the different processes contributing to mobility and also minimizes photobleaching of the molecule during imaging of the recovery. But if a longer time-span needs to be observed, e.g. for diffusion of integral membrane proteins (Klonis et al., 2003), the time interval between acquisitions can be increased after the initial 5-10 s of recovery, since this is usually the crucial period for collection of free-diffusion data. Pilot experiments should be conducted to determine the optimum interval between measurements and the total observation time, by plotting the raw intensity data from a trial run over a couple of minutes, with a sampling frequency of a few seconds.

The duration of the final experiment will depend on how long it takes for the recovered intensity to reach a plateau, this being the minimum observation time needed. The sampling frequency should depend on the extent of recovery within the first 5 s, as explained above. In order for more quantitative analysis to be performed, the raw data needs to be normalized to take the following phenomena into account:

  1. There is always some loss of intensity from low-power photobleaching during imaging.
  2. There will be a decrease in total cellular fluorescence due to the movement of proteins into and out of the ROI during the initial photobleaching process.
  3. Background noise is always present.

Therefore, during the experiment, the whole cell/cellular compartment under investigation and also an area free from fluorescence should be included in the image region, to allow these corrections to the intensity data to be made.

Usually a confocal set-up is used, but FRAP experiments can be done with any imaging system with a detector such as a CCD camera and a stable light source that can rapidly bleach a small region. Most early experiments, with the exception of video FRAP, were performed with a widefield microscope and a focused Argon or Argon/Krypton laser (Kapitza et al., 1984) and photomultiplier tubes as detectors.

The initial FRAP experiments used hydrophilic or lipophillic fluorophores such as fluorescein (Edidin et al., 1976) but now the GFP family are the favored fluorescent reporter proteins used since they:

  1. are very photostable and can therefore be imaged over a long time period to monitor recovery of fluorescence in the bleached ROI,
  2. rarely block the functionality or localization of the protein being labeled,
  3. can be expressed internally, allowing experiments to be performed on living cells that have not been disturbed by microinjection.

Although the technique of FRAP has been around for a while, current advances in molecular biology, technology and mathematical modelling are giving it new importance as a tool in studying protein dynamics in living, moving cells.

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Text References

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