Live-Cell Fluorescent Speckle Microscopy of Actin Cytoskeletal Dynamics and Their Perturbation by Drug Perfusion
Fluorescent speckle microscopy (FSM) is a method used to analyze the movement, assembly, and disassembly dynamics of macromolecular structures in vivo and in vitro (Waterman-Storer et al., 1998). FSM capitalizes on the well-established method of fluorescent analog cytochemistry, in which purified protein is covalently linked to a fluorophore and microinjected or expressed as a green fluorescent protein (GFP) fusion in living cells, incorporated into cellular structures, and whose dynamics are visualized by timelapse or video wide-field epifluorescence microscopy (Wang et al., 1982; Prasher, 1995). This approach has been limited in its ability to report protein dynamics because of inherently high background fluorescence from unincorporated and out-of-focus incorporated fluorescent subunits and the difficulty in detecting movement or turnover of subunits because of their uniform labeling of fluorescent structures. In contrast, FSM offers the capability to measure variations in molecular dynamics in living cells at high spatial and temporal resolution. In addition, FSM reduces outof- focus fluorescence and improves the visibility of fluorescently labeled structures and their dynamics in three-dimensional polymer arrays such as the mitotic spindle (Waterman-Storer and Salmon, 1999; Maddox et al., 2002, 2003).
This article describes how to use FSM for measuring actin cytoskeletal dynamics in living cells using microinjected fluorescently labeled skeletal muscle actin and the manipulation of actin dynamics during high-resolution FSM imaging by drug perfusion. This allows measurement of the kinetic evolution of the response of the actin cytoskeleton to the effects of drugs with unknown or known molecular targets, including actin and its associated proteins. We have used this technique to study changes in actin dynamics in response to perfusion of cytochalasin D (an actin filament capping drug) (see Fig. 5), latrunculin A [an actin monomer sequestering drug (Ponti et al., 2002)], and ML-7 and calyculin-A [myosin II inhibitors and activators, respectively (Gupton et al., 2002)]. These experiments have provided insight into how the actin cytoskeleton is organized into functionally distinct zones in migrating cells, with an exploratory lamellipodium at the leading edge whose actin dynamics are sensitive to filament capping, and a lamellum where traction forces are generated by myosin IImediated actin dynamics. Using FSM analysis during perfusion of drugs that specifically affect signaling molecules or other structural components of the actin cytoskeleton will be a valuable approach for understanding the molecular regulation of the cytoskeleton during cell morphogenesis.
Principles of FSM Imaging
In its initial development in 1998, FSM utilized conventional wide-field epifluorescence light microscopy and digital imaging with a sensitive, low-noise cooled charge-coupled device (CCD) camera and was applied to study the assembly dynamics and movement of microtubule polymers (Waterman-Storer and Salmon, 1997). Since then, FSM has seen new applications in answering various questions about actin and microtubule cytoskeletal function in vivo during cell motility, neuronal pathfinding, and mitosis, as well as given insight into cytoskeletal dynamics in vitro. It also has been transferred from widefield epifluorescent microscopes to confocal and total internal reflection fluorescence (TIRF) microscopes, which reduce the out-offocus contribution further and thus increase the speckle contrast (Grego et al., 2001; Maddox et al., 2002; Adams et al., 2002; Adams et al., 2004). Computer-based analysis of FSM image series has begun to be developed so that the use of dynamic speckles as quantitative reporters of polymer trajectory, velocity, assembly rate, lifetime, and disassembly rate can be realized (Ponti et al., 2003).
FSM was initially discovered by accident when it was noticed in high-resolution images of cells injected with X-rhodamine-labeled tubulin that some microtubules exhibited variations in fluorescence intensity along their lattices, i.e., they looked speckled (Waterman-Storer and Salmon, 1997). These speckles were used as marks of the microtubule lattice to allow differentiation between mechanisms of microtubule movement in living cells.
To understand the origin of speckles on microtubules as an example, one must consider how the images of fluorescent microtubules are formed by the optics of the microscope. Microtubules assemble from tubulin dimers into polymers with 1625 dimers per micrometer of microtubule (Desai and Mitchison, 1997). In an image made by a microscope, a fluorescent microtubule appears as wide as the smallest region that the microscope is capable of resolving, defined by geometrical optics of light as r = 0.61λ/NAobj (Inoué and Spring, 1997), where λ is the emission wavelength of the fluorophore and NAobj is the numerical aperture of the microscope objective lens. For example, for Xrhodamine (602nm emission)-labeled microtubules imaged with the best available 1.4 NA optics, the smallest resolvable region of a microtubule is 270nm, which contains 440 tubulin dimers. A low fraction of fluorescent dimers, f = 1%, will produce a mean number n of 4 fluorescent dimers (n = 440 x f) per resolvable image region. In FSM images, the speckle pattern along the microtubule is produced by variations in the number of fluorescent dimers per resolvable image region relative to this mean. Thus, the "contrast" of the speckle pattern can be expressed as the ratio between the standard deviation and the mean of a binomial distribution with n elements:
This formula indicates (i)that the contrast c increases with lowering f and (ii) that it decreases with growing n, indicating the requirement for optics with the highest NA possible.
For example, during assembly of a microtubule with 1% fluorescent-labeled tubulin and 99% unlabeled tubulin in an image taken with 1.4 NA optics, one 270-nm segment of the microtubule may contain 1 fluorescent dimer and 339 nonfluorescent dimers, while the adjacent resolvable segment of the microtubule could contain 7 fluorescent dimers, producing a 7x difference in image brightness between adjacent resolvable regions, and thus high speckle contrast. By comparison, at f = 50%, adjacent minimally resolvable image regions may contain 212 and 234 fluorescent dimers, a 1.1x difference in image brightness and thus very low speckle contrast.
Thus, time-lapse FSM requires the ability to visualize high-resolution image regions (~0.25µm) containing few (1-10) fluorophores and the capacity to inhibit fluorescence photobleaching. This requires a very sensitive imaging system with little extraneous background fluorescence, very efficient photon collection, a camera with low noise, high quantum efficiency, high dynamic range, high resolution, and suppression of fluorescence photobleaching and photodamage in the specimen with illumination shutters and/or oxygen scavengers (Waterman-Storer et al., 1993, 1998; Mikhailov and Gundersen, 1995).
Since the original analysis of microtubules, the "speckle" method has been applied to other cytoskeletal systems. When injected with low levels of fluorescently labeled actin, actin-rich structures such as the lamellipodium of migrating cells appear speckled in high-resolution fluorescence images (see Fig. 1) (Waterman-Storer et al., 1998, 2000; Verkhovsky et al., 1999; Watanabe and Mitchison, 2002). Also, a GFP fusion of a microtubule-binding protein, ensconsin, when expressed in cells at very low levels, gave a speckled distribution along microtubules (Bulinski et al., 2001). These actin and ensconsin speckles formed in different ways from those in a single microtubule polymer. For actin, the lamellipodium is filled with a cross-linked and dense three-dimensional meshwork of actin filaments (see Fig. 1), and each filament is made up of a paired helix of 360 actin monomers per micrometer (Pollard et al., 2000; Small 1981; Svitkina et al., 1997). Here, a fluorescent speckle may arise from fluorescently labeled actin monomers within multiple different actin filaments of the meshwork falling into the same minimally resolvable image region (Fig. 2a); however, none of the filaments are detected individually. Thus, the entire meshwork appears as a relatively even distribution of fluorescent speckles in images. For ensconsin, binding of few diffusible GFP-ensconsin molecules to the microtubule lattice, together with many unlabeled ensconsin molecules, results in GFP-ensconsin speckles along microtubules.
Thus, a "speckle" in general terms is defined as a minimally resolvable image region that is significantly higher in fluorophore concentration (i.e., fluorescence intensity) than its immediately neighboring minimally resolvable image region. In addition, for a speckle to be detected, the fluorescent molecules must be immobilized for the time required for image acquisition. The rapid motion of a small number of mobile (diffusible) fluorescent molecules results in a low level of evenly distributed background signal in the many pixels the molecules visit during the 0.5- to 3-s exposure time required by the camera to acquire an image of the dim FSM specimen (e.g., for our imaging system, actin monomers move at 63 pixels/s in the image space). This concept was demonstrated nicely by Watanabe and Mitchison (2002), who showed that diffusible GFP expressed at very low levels in cells produced an even distribution of fluorescence in high-resolution images, while a similar expression level of GFP-actin produced a speckled fluorescence image.
Speckle patterns in macromolecular assemblies act as a "bar code" pattern on structures that only change with assembly or disassembly of the structure over time (Waterman-Storer and Salmon, 1998). The pattern also serves as fiducial marks on macromolecular assemblies, which in time-lapse movies can be tracked and measured, with translation of the speckle pattern encoding trajectory and velocity and appearance/disappearance of speckles relating to assembly/disassembly dynamics. FSM therefore provides an exceptional tool for studying the actin cytoskeleton during processes such as cell migration, division, morphogenesis, and neuronal pathfinding. Furthermore, FSM can serve as the quantitative readout to link pharmacological, molecular, or genetic interventions to changes in cytoskeleton dynamics to permit a systematic deciphering of molecular regulation of the actin cytoskeleton.
This chapter article gives guidelines for designing a microscope imaging system for performing time-lapse FSM and protocols for achieving time-lapse FSM imaging of the actin cytoskeleton in living tissue cells cultured on glass coverslips. Finally, it also describes a perfusion system for temporally controlled application of drugs during time-lapse FSM of actin to observe the kinetic evolution of their effects on the dynamics and organization of the actin cytoskeleton.
II. MICROSCOPY INSTRUMENTATION
As opposed to a step-by-step protocol, this section discusses the basic components needed to set up an FSM system, giving the recommendations for critical requirements in each type of component (see Fig. 2).
The following materials are needed: upright or inverted epifluorescent microscope and optics, including epi-illuminator, objective lens, excitation filter, emission filter, and dichromatic mirror; electronically controlled shutter; cooled CCD camera; and computer, digital image acquisition board, and software for control of shutter and image acquisition.
A. Upright or Inverted Epifluorescent Microscope and Optics
B. Electronically Controlled Shutter
C. Cooled CCD Camera
Choice of the camera is one of the critical, make-orbreak decisions in the design of an FSM imaging system. For imaging the low fluorescence of dim FSM specimens, the camera demands are steep: it should be highly sensitive, extremely low noise (remember that speckles look a lot like noise!), high spatial resolution, high dynamic range, and, depending on your biological application, it may need high speed as well. The camera should be a scientific grade slow-scan cooled chargecoupled device camera. To date, most intensified cooled CCD we have tested (both microchannel plate type or on-chip type) have had noise characteristics that obfuscate speckle detection, although on-nip electron multiplication LCDs appear promising. Cameras are available from several manufacturers (Hamamatsu Photonics, Roper Scientific, Cohu, Andor) at prices ranging from ~$9000 to $30,000 USD. We give an example of the specifications of the camera of choice in our laboratory, the Hamamatsu Orca II ER, as a benchmark.
D. The CCD Chip
1. Spatial Resolution
Spatial resolution is determined by the physical size of the silicon photodiodes ("pixels") that convert photons to charge on the CCD chip. These currently range in size from about 6 x 6 to 30 x 30 µm. The larger the pixel size, the more magnification from the microscope will be needed to ensure resolution-limited images. Thus, smaller pixel size is better for FSM, as it will not require photon-robbing magnification changers or optovars in the light path. The total number of pixels making up the CCD and the pixel size will determine the imaging area. For example, our Orca II ER has a 1344 x 1024 array of 6.4 x 6.4 - 41µm2 pixels, resulting in a 8.67 x 6.60-mm CCD capable of imaging an 87 x 66-µm area of the specimen at 100x magnification.
2. Pixel Well Capacity
Based on the physical composition of the silicon photodiodes, pixels will have a maximal number of photoelectrons that it can "hold" before it is saturated with charge, which will correspond to white saturation in the image. The greater this "full well capacity," the greater the potential for a high dynamic range, after taking noise into consideration (see later). The full well capacity is also a function of the pixel size, so for FSM imaging one should consider the best full well capacity per micrometer of pixel area. The Orca II ER has a full well capacity of 18,500 electrons/41 µm2 = 45 electrons per µm2.
3. Illumination Geometry
CCDs can be illuminated from their front or back sides. Front-side illumination requires that the light pass through substrate materials to reach the photosensitive area, reducing quantum efficiency (QE). This is the configuration of our Orca II ER. Back-illuminated CCDs are physically thinned (also called "backthinned") to allow illumination directly on the photosensitive surface, making them much more sensitive (and much more fragile and expensive!). However, because of the thinning process, there are limits to the size of the pixels, with the smallest currently available at 13 x 13µm. Thus, one has to weigh whether the increased sensitivity is worth the photon loss in having an optavar in the image path, as well as whether one can afford the expense. For FSM applications, sensitive front illuminated CCDs, such as the Orca II ER, work quite well.
4. Spectral Sensitivity
Different types of CCDs have specific probabilities at any given wavelength of converting a photon striking the pixel to an electron that is counted as signal by the camera (quantum efficiency). Manufacturers supply graphs of the wavelength vs QE for their available CCDs. A CCD should be chosen that has a high (>50%) QE in the wavelength range of your fluorophore of choice. The Orca II ER has ~70% QE between 450nm (blue-green) and 600nm (orange-red). Some back-illuminated cameras achieve ~90% QE throughout the visible spectrum, but have excessively large pixels and terribly slow readout (see later).
5. Readout Geometry
Once photons are converted to charge in the array of pixels, the charges must be read out to an image acquisition board so that the image can be reconstructed in the computer by assigning a gray value to the relative charge at each pixel position. Charges are transferred out of the CCD in three basic ways. Fullframe readout occurs as each row of pixel charges is transferred serially out of the CCD one row after another. This type is the slowest and introduces the most noise (nonphoton-associated charge) into the image, although can still acceptable for FSM if other camera electronics do not introduce sources of noise. In contrast, in frame transfer and interline transfer CCDs, either the entire pixel charge array or whole rows of pixels are transferred simultaneously to an array of pixels that are masked from light. The charges are then read out from the masked area while the imaging area is being exposed to light again. These types are much faster and less noisy than the fullframe readout type. All three geometries are acceptable for FSM, although attention should be given to the manufacturer's specifications for the noise introduced during readout ("readout noise") as this will deter mine the dynamic range of the camera (see later). Our Orca II ER is the interline transfer type, giving a good balance of higher speed and low readout noise (three to five electrons).
E. Camera Electronics
Heat on the CCD can cause nonphoton-associated charge to build up in pixels, thus contributing to image noise. The coldest camera possible within a reasonable budget should be chosen. Manufacturers will house the same CCD in cameras with different degrees of cooling, ranging from 20°C below ambient temperature to -50 or-60°C. Do not try to go the inexpensive route here because heat is an avoidable source for noise that can easily mask your very faint FSM signal. The Orca II ER is cooled to -60°C, contributing to its exceptionally low readout noise (see earlier discussion).
2. Readout Speed
In general, the faster the readout speed, the more error is introduced during charge transfer, which translates to noise in your image. Speeds in modern cameras range from 100kHz/pixel in some low-noise back-illuminated cameras to 14-15 MHz/pixel in interline and frame transfer cameras. For quantitative FSM imaging of cytoskeletal dynamics, image acquisition rates of 1-2 images/s may be required, which cannot be accomplished by the slower cameras. Here, a reasonable compromise of speed and low noise must be sought, but it is recommended not to buy a camera much slower than 1MHz. The Orca II ER has a choice of readout speeds, a higher noise, fast readout at 10mHz/pixel or a slower low-noise (quoted earlier) 1.25-mHz/pixel readout rate.
3. Dynamic Range
For FSM imaging of very dim specimens, it is important to have the biggest dynamic range possible (again, within a reasonable budget). Although pixel full-well capacity is set for a given CCD, the number of gray levels this amount of charge is divided up into is not fixed. It can be encoded by 8, 10, 12, 14, or 16 bits of information per pixel, corresponding to 256, 1024, 4096, 16,384, or 65,536 (28 , 210 , 212 , 214 , 216 ) gray levels. However, statistically, it is not possible to distinguish between two gray levels that differ by less than the noise level. Thus the actual dynamic range is determined by the pixel full-well capacity divided by the readout noise. For example, our Orca II ER camera advertises 14 bit dynamic range (16,384 gray levels) and has a full-well capacity of 18,500 photoelectrons. Thus, it must have a noise level of 18,500/16,384 = 1.1 electrons per pixel or less to make use of the full 14 bit range. Because the Orca II ER has a minimum of three electrons noise, the actual dynamic range is 18,500/3 = 6166 discernible gray levels. For FSM imaging, a bigger dynamic range (at least 12 bit) is required so that differences between the intensity of two or three fluorophores can be detected quantitatively.
4. Subarraying and Binning
Being able to read out only a specified portion of the CCD (subarraying) can increase image acquisition speed for imaging small areas of a cell, but it is not necessary. Binning, in which the charges in a group of pixels are combined and read out as a single pixel to increase sensitivity, should not be done in FSM, as this effectively increases pixel size and decreases CCD resolution.
F. Computer, Digital Image Acquisition Board, and Software for Control of Shutter and Image Acquisition
A computer with the fastest processor and most random access memory (RAM) affordable should be used. Currently, computers with ~2-GHz processors and 2-GB RAM can be had for ~$3000 USD. Time-lapse FSM image series are large files often on the order of 500 MB or more and computer "horsepower" is necessary to view and manipulate these. A large hard drive (100 GB) is useful for temporary file storage.
2. File Storage
A DVD R/W device is the most economical choice recommended to archive the large files generated by time-lapse FSM. The fastest write speed available within budget should be chosen. A portable USBbased hard drives allow rapid transfer between computers. However, in the best case, a large networked file server is preferred.
3. Image Acquisition Board
Use the board recommended by your camera and software manufacturer, making sure that the board can handle the bit depth of the camera. Many cameras come with their own boards.
Software should be capable of time-lapse digital image acquisition and triggering the shutter during camera exposure. The software should allow easy viewing of time-lapse series as movies, with control of play-back rate and adjustment of brightness and contrast in the entire image series. Basic image processing, including the ability to perform low-pass filtering and image arithmetic (subtraction, multiplication, etc.), is required. The software should provide the ability to perform quantitative analysis of intensity, position, and distance. We have used Metamorph (Universal Imaging) with outstanding success. However, NIH-Image freeware (available at http://rsb.info.nih.gov/nih-image/) is also very versatile and many free macros are available.
G. Matching Microscope and Detector Resolution
A critical key to obtaining resolution-limited images of fluorescent speckles is matching microscope and camera resolution. The resolution-limited image region has to be magnified to an area on the CCD large enough to achieve a sampling frequency that is high enough to be able to digitally resolve structures at the resolution limit (Stelzer, 1998). As a rule of thumb, magnifying the resolution limited image region to the size of 3 x 3 pixels on the CCD is sufficient so that the CCD does not limit imaging system resolution or produce aliasing between pixel rows. This is referred to as the Nyquist sampling criterion. Any magnification over this value does not contain significantly more information and simply reduces the area of the specimen that is imaged. The magnification (M) required to achieve this is given by
where Pwidth is the width of a pixel and r is the size of the resolution-limited image region. Thus, for red fluorescence with a resolution limit of 0.27µm and a camera with 6.7 µm pixels and a 1.4 NA objective lens, the magnification required to satisfy the Nyquist criterion is 75x. Thus either a 100x objective or a 60x with 1.25x intermediate magnification should be used, whichever transmits more light.
III. MATERIALS AND INSTRUMENTATION FOR CELL PREPARATION, DRUG PERFUSION, AND FSM IMAGING
Successful time-lapse FSM imaging of actin dynamics is highly dependent on the choice of cell type. Cells should grow in tissue culture well adhered to glass coverslips and should be relatively large (>50µm diameter), well spread, flat, and thin (<1.0µm). We routinely use PtK1 cells, an epithelial line from ratkangaroo kidney (American Type Culture Collection, Manassas, VA, ccl-35). These cells are optimal as they are large, thin cells and are relatively easy to microinject. PtK1 cells are maintained in F-12 Hams media (Sigma, St. Louis, MO, Cat. No. N-4388) and plated on 22x 22-mm #1.5 cover glasses (Corning, Kennebunk, ME, Cat. No. 2870-22) in 35-mm tissue culture dishes and maintained in a humidified incubator at 37°C supplemented with 5% CO2. Use of this type of coverslips is critical, as high-resolution optics are specifically corrected for this thickness (#1.5 = 0.17mm. thick), and our custom-made perfusion chamber described in Fig. 2 is specifically designed for them. Prior to plating cells, the coverslips are cleaned by sequential washes and sonications in detergent, water, and ethanol, as described in detail in Waterman-Storer (1998). This level of coverslip cleanliness is of paramount importance, as coverslip dirt results in background fluorescence that degrades speckle contrast.
Fluorescently labeled skeletal muscle actin can be bought (Cytoskeleton, Denver, CO, Cat. No. APHR-A) or made using the detailed protocol published elsewhere (Waterman-Storer, 2001). A labeling ratio of 0.3-1.0 fluorescent dye molecules per protein molecule is acceptable. We recommend using longer wavelength fluorophores, such as Texas red (ex 596 nm, em 615 nm), X-rhodamine (ex 575nm, em 602nm), Alexa 568 (ex 578 nm, em 603 nm), or tetramethyl rhodamine (ex 541nm, em 572nm), as cellular autofluorescence that degrades speckle contrast tends to be at shorter emission wavelengths (~500-550nm). Very short wavelength excitation fluorophores (i.e., <400nm) should not be used, as the UV light required for their excitation is very damaging to living cells. In addition, we have found no qualitative difference in FSM of actin dynamics using actin coupled to fluorescent dyes via either lysine residues (i.e., succimidyl ester fluorophore derivates) or cysteine residues (malemide derivates).
Labeled actin should be stored long term (up to 2 years) in flash-frozen aliquots at higher protein concentrations (5-10mg/ml) at -70°C and can be diluted to the working concentration of 1 mg/ml in G buffer (2mM Tris-Cl, pH 8.0, and 0.2mM CaCl2; just before use add 0.2mM ATP and 0.5 mM 2-mercaptoethanol; store up to 1 day at 4°C), refrozen in 5-µl aliquots, and stored for several months before use. To prevent problems of clogged microinjection needles, prior to microinjection into cells, fluorescent actin must be clarified by centrifugation in a swinging bucket rotor (Sorvall, Newtown, CT, Cat. No. S55-S or Beckman Cat. No. TLS-55) in a microultracentrifuge (Sorval, Cat. No. RCM120 or Beckman Cat. No. TL-100). If possible, keep the rotor stored at 4°C for convenience. Microinjection needles can be bought from Eppendorf (Femtotip II, Brinkman, Westbury, NY, Cat. No. 930000043), although we pull our own from thin wall borosilicate glass capillaries with an outer diameter of 1.00mm, an inner diameter of 0.78 mm, and a length of 10cm (Sutter, Novato, CA, Cat. No. BF100-78-10) using a Sutter Instruments P-87 needle puller with a 2.5 x 4.5- mm box filament (Sutter, Cat. No. FB245B). Needles are treated further with hexamethyldisilazane (HMDS, Pierce, Rockford, IL, Cat. No. 999-97-3) to prevent clogging caused by protein adhering to the insides of the needle. A 10-µl Hamilton syringe (Hamilton, Reno, NV, Cat. No. 80008) is used to back load needles with fluorescent actin.
Coverslips of cells are injected in 35-mm tissue culture dishes using an Eppendorf Injectman (Cat. No. 5179 000.018), microneedle holder and grip heads (Cat. No. 5176 190.002, Cat. No. 5176 210.003), and transinjector (Cat. No. 5246 000.010) (Brinkman, Westbury, NY) system mounted stably on a microscope. Although many fine-control microinjectors are acceptable, be sure that they are designed for mammalian cell injection, as those designed for in vitro fertilization of larger eggs may not have fine enough positional control. For back pressure in the needle, systems capable of repeatable femtoliter injection volumes are needed; other systems including those made by Picospritzer are acceptable. The microscope on which the microinjector is mounted must be a stable platforminverted configuration. Although we use a Nikon TE-2000-S, all those mentioned previously are suitable, as well as less expensive "clinical" models. The microscope must be equipped with a quartz-halogen transilluminator and a long working distance phase-contrast condenser lens system that allows room for an open tissue culture dish on the stage (we use a Nikon 0.35 NA extra-long working distance condenser lens). A long working distance dry (nonimmersed) 40x phase contrast objective lens is required to image cells through the thickness of the plastic dish and coverslip during microinjection (we use a Nikon 40x 0.55 NA objective with a 2.1-mm working distance).
In order to keep cells happy and healthy during FSM imaging over time, it is necessary to keep the cells in an optimum environment on the microscope stage. This includes proper temperature and pH regulation and minimizing photodamage and photobleaching caused by exposure to excitation light. Because suppression of fluorescence photobleaching is one of the most critical aspects of successful FSM imaging, when filming PtK1 cells, we supplement their F-12 Hams media with Oxyrase (Oxyrase Inc., Mansfield, OH, Cat. No. ES-50), an oxygen-scavenging enzyme purified from Escherichia coli, which decreases photobleaching and photodamage to the cell. To make this reagent effective, cells and their surrounding media must be sealed from exposure to air. In order to minimize deleterious effects of pH, one might consider adding 5 mM HEPES buffer (pH 7.0) to the medium. However, because we use F-12 Ham's media for PtK1 cells, which is already buffered, this does not require extra HEPES. For temperature control, we find that an airstream stage incubator, such as those produced by Nevtek instruments (ASI-400), is the best bet. One should experiment ahead of time to determine the setting on the thermostat that gives a sample temperature (with the objective lens coupled by immersion oil!) of 37°C and keeps cells viable over several hours.
For drug treatment studies we have custom designed a live-cell perfusion chamber that allows use of high-resolution oil-immersion objective and condenser lenses and gives near laminar flow for complete exchange of media in a minimal volume while remaining sealed from the environment (Fig. 3). The chamber portion comes in direct contact with media that washes over cells, and thus must be made of inert stainless steel. The chamber top that holds down the coverslip of cells on the chamber has no contact with media, and thus can be made of less inert aluminum, while the coverslip gasket is made of 0.005-in. thick Teflon sheet, for flexibility. A 24 x 60-mm coverslip (Corning, Cat. No. 2940-246) must be sealed to the bottom of the chamber with hot VALAP (a 1 : 1 : 1 mixture of Vaseline, paraffin, and lanolin prepared in the laboratory) to complete the condenser lens-facing side of the imaging chamber. Any good machine shop should be capable of reproducing this chamber based on the diagram in Fig. 3.
A. Microinjection of Fluorescent Actin
Microinjection of 1 mg/ml fluorescently labeled actin allows later visualization of protein dynamics by time-lapse FSM.
B. Live-Cell Imaging and Drug Perfusion
C. Interpreting Speckle Dynamics
Interpretation of actin speckle dynamics provides information about actin turnover and actin flow in a cell and how this relates to cell behavior. Using image analysis software such as Metamorph, images can be processed and analyzed.
Successful FSM imaging of live cells requires several critical components, such as proper fluorescent protein concentration, successful suppression of photobleaching, and high-quality, well-labeled, fully functional fluorescent protein. Fluorescent actin speckles cannot be detected if the concentration of fluorescent protein in the cell is too high, which can occur if the microneedle concentration is >1mg/ml. Even at proper needle concentrations, too much fluorescent actin can get into the cell if the microinjection technique is not optimized. Conversely, if there is not enough fluorescent protein injected into cells, the camera exposures required to acquire a decent image will be excessively long and may result in motion artifacts within the image. Photobleaching is also a major pitfall in FSM imaging. This can occur if the imaging chamber is not fully sealed, if there are air bubbles present in the chamber, or if the Oxyrase has begun to lose its potency. Ensuring an airtight chamber with fresh Oxyrase will reduce photobleaching greatly. Finally, ensuring that the actin remains functional during the labeling process is an extremely important consideration for quality FSM imaging. If you are making your own fluorescent actin, be sure to use high-quality acetone powder that is finely powdered or "fluffy" and that the dye is fresh and has been stored in dessicant at -20°C. Old dye will render actin nonpolymerizable, so avoid using dye that is not fresh.
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