Maintaining Live Cells on the Microscope Stage
An increasing number of investigations are using live-cell imaging techniques to provide critical insight into the fundamental nature of cellular and tissue function, especially due to the rapid advances that are currently being witnessed in fluorescent protein and synthetic fluorophore technology. Because of these advances, live-cell imaging has become a requisite analytical tool in most cell biology laboratories, as well as a routine methodology that is practiced in the wide ranging fields of neurobiology, developmental biology, pharmacology, and many other related biomedical research disciplines. Among the most significant technical challenges for performing successful live-cell imaging experiments is to maintain the cells in a healthy state and functioning normally on the microscope stage while being illuminated in the presence of synthetic fluorophores and/or fluorescent proteins.
Tight control of the environment is one of the most critical factors in successful live-cell imaging experiments. In particular, the conditions under which cells are maintained on the microscope stage, although widely variable in many requirements depending upon the organism, often dictate the success or failure of an experiment. Aspects of the environment that are readily manipulated include the physical parameters of the chamber in which the cells are grown and imaged, the localized degree of temperature control, atmospheric conditions (gas mixture and humidity), nutritional supplements, growth medium buffering (pH), and osmolarity of the culture medium.
Illustrated in Figure 1 are a series of images captured from several unrelated cell lines, each labeled with a different combination of synthetic fluorophores and/or fluorescent proteins. The rabbit kidney epithelial cells (RK-13 line) in Figure 1(a) were transfected with a fusion plasmid of enhanced yellow fluorescent protein (EYFP) and a nuclear targeting signal peptide to localize a greenish-yellow label in the nucleus. The cells were subsequently treated with MitoTracker Red CMXRos to stain the mitochondria. In Figure 1(b), opossum kidney proximal tubule epithelial cells (OK line) were transfected with an EYFP-actin subcellular localization vector to label the filamentous actin cytoskeletal network. Mitochondria were targeted with a DsRed2 fusion vector in the Indian Muntjac cells presented in Figure 1(c), while an EGFP-peroxisomal chimeric plasmid highlights peroxisomes in the human cervical carcinoma (HeLa line) cells in Figure 1(d). The adherent culture of normal Syrian golden hamster kidney fibroblast cells (BHK-21 line) featured Figure 1(e) was transfected with a mixture of DsRed2 FP-endoplasmic reticulum and EGFP-nucleus subcellular localization vectors, thus localizing a green fluorescent protein tag to the nucleus and an orange-red probe to the endoplasmic reticulum. Finally, the human bone osteosarcoma cells (U2OS line) illustrated in Figure 1(f) was transfected with Cerulean fluorescent protein fused to a mitochondrial targeting sequence to label the mitochondria. For each of the images in Figure 1, a separate channel was recorded using differential interference contrast and overlaid on the fluorescence channel(s) to identify cell boundaries and other common structural features, such as the nucleus.
During the course of formulating plans for live-cell observations and long-term imaging experiments, a number of important factors are worthy of serious consideration. The specimen should be accurately labeled with the fluorescent protein or synthetic fluorophore(s) of interest in order to clearly visualize the target biological components. Perhaps even more important, the cell culture must be maintained in a condition that promotes growth and normal function in order to avoid potential artifacts in the interpretation of experimental results. In addition, the cells should be imaged with sufficient spatial and temporal resolution in a manner that does not induce phototoxicity or perturb localization of the fluorescent probes. Among the most important routine considerations for live-cell imaging that must be addressed (and discussed in detail below; see Table 1) are temperature, oxygenation, humidity, osmolarity, pH (medium buffering), phototoxicity, the laboratory environment, microscope focus drift, fluorescence signal strength, bleed-through, and resolution.
Maintaining living cells in a healthy state on the microscope stage is undoubtedly the most critical aspect for any live-cell imaging investigation and generally requires a combination of mechanical ingenuity along with keen insight into the biology of the cell or tissue being studied. Although many laboratories are adept at growing cultured cells in temperature controlled carbon dioxide incubators, the task of maintaining cells on the microscope stage for long-term imaging experiments is far more demanding. The imaging chamber must keep the cells (or tissues) functioning normally for the duration of the experiment, while allowing unrestricted access by the microscope objective. This feat can become particularly difficult when high numerical aperture oil or water immersion objectives are being used. In many cases, the investigator must be able to introduce a reagent while imaging (to perturb a particular cellular process) without disturbing a time-lapse sequence by shifting focus or stage position. Other important factors are simplicity, reliability, and reasonable cost. The discussion that follows focuses primarily on mammalian cells, but the techniques for a variety of other organisms differ only slightly and are usually not as stringent. For example, cultures of yeast, insect, and plant cells do not have the strict temperature requirement, whereas the composition of media is not as demanding for bacterial cells.
Culture Media for Mammalian Cell Lines
Although the media for early attempts at cell and tissue culture consisted of mixtures containing embryo extracts, serum, protein hydrosylates and a host of other body fluids, propagation of established lines quickly required the transition into defined media based on biochemical requirements. Of these, the most popular are Eagle's Basal (EBM) and Minimal Essential (MEM) culture media, Dulbecco's modification of MEM (DMEM), Ham's media (F-10 and F-12), and two highly refined media formulations designed at the Roswell Park Memorial Institute (RPMI 1640 and RPMI 199). In addition, a medium designed for culturing cells in the absence of bicarbonate buffer (and carbon dioxide), Leibovitz L-15, has been widely employed. All of these culture media require the addition of serum (usually derived from fetal and newborn calves or horses) to a final volume fraction ranging between 5 and 20 percent. Serum-free media have also been developed for culturing highly specialized cells under strictly defined conditions that benefit applications in the biopharmaceutical industry. Many laboratories involved in culturing a wide variety of cell types often compromise on the rigorous requirements of an exact formulation by using a mixture of a complex medium, such as Ham's F-12, with a second medium (DMEM, for example) containing higher amino acid and vitamin concentrations.
The composition of cell culture media varies widely, but most recipes include amino acids, vitamins, inorganic salts (minerals), trace elements, nucleic acid constituents (bases and nucleosides), sugars, co-enzymes, lipids, tricarboxylic acid cycle intermediates, and a variety of other biochemicals. Simple media, such as MEM, contain only the essential amino acids, vitamins, and salts, whereas more complex formulations (RPMI and serum-free media) have hundreds of components. Cell culture media are usually designed for specific purposes, including routine growth of normal and immortalized (transformed) cell lines, primary culture initiation, virus propagation, pharmaceutical preparation, and defined growth conditions for genetic variants. Among the variables that are controlled in all tissue culture media formulations are pH, buffering capacity, oxygen concentration, osmolarity, viscosity, and surface tension. When cells are imaged in the microscope, even for short periods of time, these same medium conditions must be carefully reproduced in the live-cell imaging system.
Environmental Variables for Mammalian Cell Lines
A majority of the popular cell lines used in live-cell imaging experiments grow very well in a narrow range of pH between 7.2 and 7.4, although some normal fibroblasts perform better at slightly higher pH values (up to 7.7), while many transformed cell lines grow faster in more acidic media (down to pH 7.0). In cases where pH is critical for experimental accuracy, a plating efficiency assay should be performed at the target pH to ensure the selected cell line will perform satisfactorily. Most commercially available media formulations contain an indicator dye (usually Phenol red) for visual determination of the approximate pH value. In solution, Phenol red produces a bright red hue at pH 7.4, becomes orange at pH 7.0, and yellow at pH 6.5, a shift in color that is often seen when the medium becomes more acidic as cultures form confluent monolayers. At higher pH values, Phenol red is pink at pH 7.6 and purple at pH 7.8 and above. Many tissue culture laboratories find it useful to construct a set of pH standards using Phenol red in a balanced salt solution at varying pH values for comparison to culture media. Although the indicator dye is essential for routine cell culture, due to the high visible light absorption extinction coefficient, its use should be avoided in live-cell fluorescence imaging experiments in order to reduce the level of background noise and to prevent phototoxicity. Recognizing this fact, manufacturers provide most of the common media formulations in sterilized liquid or powder form without Phenol red.
Virtually all cell lines require a carbon dioxide and bicarbonate buffer system to regulate pH and must be cultured in an atmosphere containing a small percentage of carbon dioxide (usually 5 to 7 percent, depending upon the bicarbonate concentration) in specialized incubators to strictly control the concentration of dissolved gas. For live-cell imaging on the microscope, producing a suitable atmosphere with carbon dioxide can be difficult and usually requires culture chambers that are specifically designed for a regulated atmosphere. The introduction of synthetic biological buffers, such as TRIS and HEPES, has been of dubious value in eliminating the carbon dioxide requirement, as many cell lines will not tolerate a lack of carbon dioxide, especially at low cell concentrations. In general, a concentration of 10 to 20 millimolar HEPES buffer can control pH within the physiological range in the absence of a carbon dioxide atmosphere, but the culture medium should still be supplemented with sodium bicarbonate for optimum cell growth.
Attempting to grow cells on the microscope using HEPES alone usually results in dramatically reduced growth rates (especially for long-term experiments), and each cell line should be carefully scrutinized for its ability to grow and function in media without the carbon dioxide buffer system. Note that supplementing the bicarbonate buffer system in cell culture media with HEPES only reduces the rate of pH drift, and does not eliminate the progressive increase in alkalinity that occurs when the culture is exposed to the atmosphere. In addition, numerous reports have surfaced of HEPES toxicity during live-cell imaging experiments, presumably due to increased free radical formation by the synthetic buffer under the illumination conditions required to visualize fluorescent probes. A specialized formulation, Leibovitz L-15 medium, is designed to eliminate carbon dioxide through the use of sodium pyruvate and buffering with high amino acid concentrations. Including sodium pyruvate in the culture medium allows cells to increase their endogenous production of carbon dioxide, theoretically rendering them independent of the gas (as well as bicarbonate). However, many cell lines do not adapt well to L-15 medium and should be thoroughly examined for several passages before configuring live-cell imaging experiments based on this formulation.
Cell lines can vary widely in their oxygen requirement, although normal atmospheric oxygen tension levels will meet the needs of most cultures. Mammalian cells usually require oxygen for respiration in vivo, but can often successfully substitute glycolysis (an anaerobic process) when grown in culture vessels as primary lines or after immortalization. The depth of the culture medium above the cells can influence the rate of oxygen diffusion to adherent cells growing on a glass or plastic surface and should be kept below 5 millimeters. In most cases for live-cell imaging, strict oxygen regulation is not necessary and, conversely, depletion of oxygen is often used as a strategy to reduce the photodamage during fluorescence illumination that can occur through reactions with oxygen free radicals. However, it should be noted that lowering oxygen tension can be just as deleterious to cells if they begin to suffer hypoxic stress. The most common method of reducing oxygen levels involves the application of a commercial oxygen depletion system, such as Oxyrase. Alternative techniques to limit free radical damage due to molecular oxygen include supplementing the medium with scavengers such as ascorbate (ascorbic acid; vitamin C) or Trolox (a derivative of vitamin E), but the strategy of reducing illumination intensity coupled with highly sensitive camera systems should also be considered.
Most of the popular cell lines have a fairly wide tolerance for osmotic pressure and will grow well at osmolarities between 260 and 320 milliosmolar. In cases where cells are routinely grown in Petri dishes or open-plate cultures, hypotonic medium can be substituted to compensate for evaporation. It is important to monitor osmolarity of the culture medium when altering the constitution by the addition of organic buffers or plasmid selection drugs, such as HEPES and G-418, respectively. The concentration of ions and organic nutrients in live-cell imaging will initially be set by the medium chosen for the experiment, but the small volumes of media accommodated by most imaging chambers are subject to changes in osmolarity due to evaporation (this problem is usually more severe when the medium is heated to 37 degrees Celsius). Therefore, special care must be taken when assembling cells into chambers and when changing culture medium if any evaporation has occurred (cells are very sensitive to rapid changes in osmolarity). In addition, during the imaging experiment, evaporation should be minimized either by using a sealed system, by covering the medium in an open chamber with an oil that has a lower density than water (usually mineral oil), or by humidifying the chamber during imaging. Note that, in general, the micro-environment provided by the small volume in a live-cell imaging chamber is inherently less stable than a carbon dioxide incubator and requires considerably more attention in every detail.
Choosing Cell Lines for Live-Cell Imaging
The choice of cell line used for live-cell imaging experiments is often dictated (and limited) by a number of factors, including the target biological observations of the investigation, the ability of the cells to be labeled with synthetic fluorophores, transfection efficiency, and the tolerance of a particular cell line to the rigorous culture chamber environmental and illumination conditions. All too often, a cell line that displays excellent properties in one of more of these categories either performs marginally or fails completely in another. For example, normal bovine pulmonary artery endothelial cells (BPAE line) can be fixed and stained using synthetic fluorophores to reveal intricate cellular structural details with exquisite clarity, but the line can only be transfected with fluorescent protein vectors at low efficiency (less than 5 percent) and is relatively intolerant to long-term illumination at low light levels that do not affect many other cell lines. Alternatively, rabbit kidney epithelial cells (RK-13 line) can be transfected at high efficiency with a variety of plasmids and are very tolerant to high illumination levels (including laser light) during time-lapse sequences extending for several days, but are not adequately stained with many of the common synthetic fluorophores (such as MitoTrackers) designed for live-cell imaging.
An overriding factor in the successful observation of biological phenomena in living cells is that the particular line chosen for study and imaging must display the necessary morphological and physiological properties to clearly demonstrate the concept of interest. In studies targeting mitosis, for example, many cell lines are less than adequate for imaging due to the fact that dividing cells become spherical and may detach from the substrate. Instead, cell lines that remain relatively flat and attached to the substrate during mitosis are far superior in revealing the fine details of the mitotic spindle during cell division. Among the most useful lines for mitosis studies are rat kangaroo kidney cells (PtK1 and PtK2 lines; see Table 2), which can only be transfected at relatively low efficiency, but contain a small number of chromosomes that are easier to distinguish in the microscope. Several other kidney cell lines, including one from the pig (LLC-PK1) and another from the African green monkey (BS-C-1) also remain attached during mitosis and are far more susceptible to transfection. Both the pig and monkey cells contain more chromosomes than the rat kangaroo, but their ease of transfection and imaging make them excellent alternatives for investigations of mitosis. In addition to being useful for observations of the mitotic spindle, cells that remain flattened on the culture chamber glass during cell division can also reveal the distribution of other cellular components, such as the Golgi apparatus, cytoskeletal elements, endoplasmic reticulum, and mitochondria.
Useful Mammalian Cell Lines for Live-Cell Imaging Experiments
Investigations of the cytoskeleton should be conducted with cells that exhibit high levels of expression and specific localization of the protein(s) that are being studied. Filamentous actin stress fibers are usually much more clearly defined in fibroblast cells than epithelial cells, but there are many exceptions. Cytokeratin intermediate filaments form extensive networks throughout the cytoplasm, which are readily visualized with immunofluorescence or fluorescent proteins in several epithelial cell lines (although not universally). Unfortunately, cytokeratin networks are either poorly defined or virtually absent in fibroblast cells, as well as many varieties of epithelial and endothelial cells. Likewise, vimentin, desmin, peripherin, neurofilaments, lamins, and glial fibrillary acidic protein (GFAP) intermediate filaments form prominent structural networks in many cell types, but are difficult to detect in others. In almost all cases, the target cell line should first be fixed and tested with synthetic fluorophores and/or antibody labeling prior to attempting to localize fluorescent proteins for live-cell imaging.
A variety of applications coupling fluorescent proteins to live-cell imaging have opened many new avenues in the quest for information concerning dynamic processes in cell biology. The advanced fluorescence microscopy techniques of recovery after photobleaching (FRAP), resonance energy transfer (FRET), correlation spectroscopy (FCS), and speckle microscopy (FSM) have benefited significantly in their development from the use of fluorescent proteins. These ubiquitous molecules have also been genetically modified to produce a new generation of optical highlighters that can be photoactivated to specifically label individual members of a larger molecular population. Going a step farther, the coupling of Förster resonance energy transfer techniques to fluorescent proteins has yielded a new class of physiological biosensor probes that are useful for reporting various ions, such as calcium, sodium, potassium, chloride, and pH, in addition to a plethora of cellular events including enzymatic activity, changes in membrane potential, neurotransmitter release, and oxidation-reduction. The foundation for all of these powerful new techniques is built upon the imaging of living cells that are expressing genetically-encoded fluorescent probes.
Presented in Table 2 is a listing of several mammalian cell lines that have been of significant service to many of the live-cell imaging experiments reported in the scientific literature. The well-studied human cervical carcinoma (HeLa) line is an immortalized epithelial cell from which a wealth of information has been gathered. This hearty cell line can be transfected at high efficiency with most of the fluorescent protein vectors to produce high levels of expression and defined localization. Transformed African green monkey kidney cells (COS-7 line) have been used to explore protein dynamics in the Golgi apparatus and endoplasmic reticulum, whereas the hamster cell lines (BHK-21 and CHO-K1) are favorites for investigations involving the molecular and cellular biology of viruses, intracellular enzymatic activity, receptors, promoter function, and transport mechanisms. The other cells listed in Table 2 are also responsive to transfection, microinjection techniques, and labeling with synthetic fluorophores for long-term imaging experiments in widefield and confocal fluorescence microscopy.
Brief Overview of Live-Cell Imaging Chambers
Specimen chambers are an integral branch in the history of microscopy and a number of designs have been published over the years describing systems that offer excellent optical properties while allowing specimens to be maintained for varying amounts of time. Short term imaging experiments (20 to 30 minutes or less) can be conducted simply by attaching a coverslip containing adherent cells onto a microscope slide using spacers to keep the cells from being damaged (physical stress can induce autofluorescence in some cell lines). The coverslip can be secured with any one of a number of sealants, including molten agarose, rubber cement, vacuum grease, or a useful preparation known as VALAP (a 1:1:1 mixture of Vaseline, lanolin, and paraffin), to provide a watertight seal and eliminate evaporation of the culture medium. Thin gaskets cut from silicone rubber (also commercially available) or broken pieces of coverslip can be used as spacers to keep the cells from coming in direct contact with the microscope slide. Make certain the coverslip surface containing the cells is placed face down on the spacer, and fill the void between the coverslip and slide with a physiological buffer (such as phosphate buffered saline; PBS). Seal the edges around the coverslip using the reagent of choice and place the microscope slide on the stage for imaging. Without growth medium and temperature control, the cells will function normally for only a few minutes, but this is often enough time to obtain the necessary images.
For longer term experiments, specially designed environmental chambers provide a mechanism for viewing and imaging living cells on the microscope stage, as well as keeping the culture very close to the optimum growth conditions for extended periods of time. In general, imaging chambers include a glass window, usually the thickness of a coverslip (approximately 170 micrometers), through which the cells can be readily viewed with objectives operating at high numerical aperture. Temperature control, a critical parameter for most cells, is often achieved using peripheral sources of infrared radiation or heated air (such as a hair dryer or egg warmer), a metal heating plate under thermistor control coupled directly to the chamber, or with optically transparent thin coatings of electrically conductive metal oxides applied by evaporation onto the coverslip surface to provide a more efficient conductive heat transfer to the chamber.
The wide variety of commercially available chambers that can be purchased (or, alternatively, easily constructed in-house) for imaging living cells generally fall into two basic functional categories: open chambers, similar to Petri dishes, which have free access to the atmosphere; and closed chambers that are sealed to protect cells from evaporation of the culture medium. An open chamber system will usually allow quick access to the growing cells, thus readily permitting microinjection, addition of drugs, changing of the culture medium, or other manipulations to the cells. In contrast, closed chambers provide better insulation from the external environment, but make access to the cells more difficult. Most closed chamber designs include ports that permit the addition of fresh medium and drugs during the experiment without interrupting an imaging sequence. In these systems, perfusion is regulated by either a peristaltic pump, motor-driven syringe, or through a gravity-controlled manifold. When new solutions are added to a closed chamber, it is critical that before addition they are equilibrated to the same temperature as the cells. Furthermore, many cells are sensitive to shear, so perfusion of adherent cells attached to a coverslip should be performed at very low flow rates. Several of the more advanced closed chamber systems are designed to offer control of shear forces.
Many of the simplest commercial open chamber imaging systems are constructed by mounting a coverslip onto the bottom of a ordinary tissue culture vessel or Petri dish. Standard 35 and 60 millimeter sterile Petri dishes are available that have a small hole (approximately one centimeter in diameter) drilled in the dish with a 170 micrometer coverslip fused to the plastic to enable high-resolution imaging. Rectangular coverslips and microscope slides with a small single or multiple well plastic imaging chamber sealed to the glass are also commercially available, but are quite expensive. Both of these chamber designs are relatively simple to use, but they are not tightly sealed, so the amount of culture medium that evaporates over the course of an experiment must be carefully monitored. In addition, most of the simple imaging chambers do not include any heating system, and must be mounted on a microscope stage equipped with an auxiliary heating unit designed specifically to house the chamber. Without temperature control, simple open chamber system performance is only marginally better than using the sealed coverslip method described above.
Sealed closed chambers similar to the Bioptechs FCS2 live-cell imaging chamber illustrated in Figure 2(a) are more expensive than most of the simple open chamber systems, but they offer a far more controlled environment and can maintain cells in a healthy state for many hours (and even days or weeks). A typical closed chamber system provides two optical surfaces separated by a perfusion ring sealed with gaskets. This sandwich is then clamped together with a metal or composite housing that is designed to provide temperature control and secure adaptation to the microscope stage. With such an enclosure, the perfusion rate, media volume, temperature, atmosphere, flow geometry, and optical stability of the imaging chamber are controlled to a relatively high degree compared to open system chambers. Advanced closed chamber systems (Figure 2(a)) reduce the fluid exchange time, provide flow control of the perfusion medium in order to avoid disturbing adherent cells, offer superior temperature regulation, and maintain close proximity of the optical surfaces for observation using high numerical aperture microscope objectives. In addition, the user can define culture medium flow conditions across the cell surface to meet the experimental requirements. A wide variety of closed chamber live-cell imaging systems are commercially available.
The pinnacle of live cell imaging chambers effectively combines a cell culture incubator with an inverted microscope to provide almost total control of the environment, an example of which is presented in Figure 2(b). The incubator enclosure is most often constructed of Plexiglas and surrounds the microscope stage, objectives, fluorescence filters, and transmitted light condenser. These chambers can be used with a variety of culture vessels, including standard culture bottles, Petri dishes, microscope slides with mounted coverslips, and many of the other open and closed systems discussed above. Temperature is maintained with an external heating unit (usually forced air) and the carbon dioxide concentration is controlled with a sensing unit coupled to a regulator that is fed by a cylinder of pure gas. These units can also be equipped with humidity control and several designs provide rubber glove access to avoid disturbing the environmental equilibrium when manipulating the cells during imaging. In order to maintain a high degree of temperature control, several of the more sophisticated incubator chambers enclose virtually the entire microscope with the exception of the eyepieces, camera, and lamphouses. On the downside, environmental chambers can impede rapid access to the specimen and are cumbersome when repeated manipulation is necessary. In addition, the high humidity level inside the chamber can add to the expense of maintaining the instrument due to premature degradation of gear lubricants and the oxidation of metal surfaces and lens coatings. Most of the microscope manufacturers offer a custom incubator option for their inverted microscopes, while aftermarket suppliers fill the gaps with both simpler and more advanced models, as well as a host of useful accessories.
As discussed above, a wide spectrum of open and closed live-cell imaging chamber designs are commercially available, many of which are intended for quite specific applications. It is worthwhile to investigate the various options available when embarking down the long road to successful live-cell imaging. Most investigators have favored techniques that reflect their experience using a particular system, and these preferences span the gamut of chamber designs. In fact, several functional systems have been reported that were constructed with readily available home repair insulation sheeting, duct tape, and low-cost humidifiers connected to the chamber using clothes dryer exhaust tubing. The key point is that in a given experiment, the chamber must maintain an optimal environment for cellular function and provide a clear optical window in which to capture the events occurring within the chamber. Because the experimental variables differ from one investigation to another, design preferences can change and the best approach is to test a variety of systems in order to identify the one that is best suited to the cell line and the experimental conditions.
With all live-cell imaging configurations, difficulties can arise when experiments are performed on preparations that require temperatures significantly different from that of the surrounding laboratory environment (note that temperature fluctuations in live-cell imaging are usually the rule rather than the exception). Cellular function is extremely sensitive to temperature variations, with changes of even a couple of degrees having profound effects on cell physiology. A variety of methods are available for controlling the temperature of cells on the microscope stage, and many of the commercial systems described above include heating elements directly coupled to the chamber. Although this strategy provides a simple integrated solution, temperature control is too often limited to the chamber itself and ignores associated components that might exert negative influences on maintaining a steady temperature. Among the most significant factors associated with temperature fluctuations is that the microscope stage, frame, and objectives can act as heat sinks and counteract the efforts of the specimen heating system. This problem is compounded when immersion objectives are used because the optical coupling medium, which can be oil, glycerin, or water, has a much higher thermal conductivity than air. Coupled with the close proximity of high numerical aperture objectives to the specimen, as well as the thermal load of the objective itself, the entire system can be rapidly deprived of heat if the objective is not thermally controlled. In the case of the static chambers described above, the area directly under the objective is often up to 5 degrees (Celsius) cooler than the remainder of the specimen chamber.
Depending upon the configuration, the entire microscope can be enclosed and heated, but several critical temperature control issues can be avoided simply by using commercially available objective heaters (see Figure 3) that employ circulating heated water or resistive heating elements. Objective heaters, when combined with a suitable specimen-heating system, can partially offset the temperature gradient between the specimen and the front lens elements. Note, however, that even with an objective heater, there can still be a temperature gradient along the objective barrel or between the objective and the microscope itself. The adjustable heating blanket and circulating water jacket objective heaters illustrated in Figures 3(a) and 3(b) are essentially the same from a thermodynamic point of view. The only difference is that heat is electrically produced in the blanket (Figure 3(a)), while heat is externally generated and transferred by a fluid into the jacketed objective (Figure 3(b)). In both cases, heat transfer is relatively inefficient with much of the heat being radiated away from the objective to the region directly beneath the specimen instead of being transferred into the objective. The unfortunate result is excessive external heat being convectively channeled upwards in the vicinity of the specimen to produce large temperature variations and excessive thermal cycling in the chamber control system.
Microscope objectives have thermal profiles that vary as a function of their physical parameters. Most objectives are designed and sold for the purpose of fixed-cell microscopy (conducted at room temperature), so when selecting an objective to be employed in live-cell imaging, care should be taken to consider only those objectives with the ability to be efficiently heated. Aside from objective heating issues, cycling of the heating system can alter the coverslip position and cause the specimen to drift out of focus. In addition, the investigator should be aware that repeated heating and cooling of the objective has been incorrectly reported to considerably shorten the lifespan, especially in terms of the strain-free character of internal lens elements. In fact, there is little reliable evidence that temperature cycles affect the strain characteristics in specialized objectives, and most can withstand temperatures up to 50 degrees Celsius without damage. The only negative effect of heating microscope objectives is that the retraction stopper barrel lubricant can increase in viscosity (achieving the pliability of gum) over a shorter period of time than with objectives that are not heated. However, in the case of heated immersion objectives, the tendency of immersion oil to creep into the barrel often prolongs the lifetime of the original lubricants.
For permanent installations, a large box can be built around the microscope and heated with warm air. In this case, most of the microscope can be equilibrated to a single temperature to provide the advantage of eliminating any movement resulting from thermal expansion of the microscope components. However, air currents surrounding the specimen chamber itself must also be minimized. When constructing an enclosure, access to the microscope and its adjustable components may be limited, so it may be worthwhile to construct the box from relatively common and inexpensive components in case significant modifications are required.
A final consideration is tight control of temperature not only for the microscope, but also the entire laboratory. Modern microscopes are fabricated with a wide spectrum of materials, including aluminum, plastic, composites, glass, brass, and steel, all of which have different thermal expansion coefficients. Even a change of a single degree Celsius can produce unwanted movements in the microscope optical train, resulting in focus or alignment shifts. Air conditioning or heating ducts that are in close proximity to the microscope will often produce localized temperature fluctuations that ultimately result in focus problems. For long-term observation, many investigators build a large thermostatically controlled box around the stage (see the discussion above) or even place the entire microscope in a room that is maintained at 37 degrees Celsius (a relatively uncomfortable working situation). The exact strategy that is employed ultimately depends on the specific application, but it is absolutely critical to consider these issues when designing a live-cell imaging system and the laboratory in which it will be housed.
Laboratory Environmental Considerations
When choosing a room that will be utilized for live-cell imaging experiments, it is necessary to ensure that adequate ventilation is available to dissipate the ozone released by mercury and xenon arc-discharge lamps, as well as the fumes from organic solvents used to clean optical surfaces and disinfect the microscope stage. Allow sufficient space around the microscope system for proper ventilation as well as cleaning of the floors, benches, tables, and experimental apparatus. Equipment failures can often be traced to air intakes that are clogged, located close to the floor, or placed in an inaccessible location. The laboratory should be kept meticulously clean and maintained in an orderly fashion to reduce the levels of dust, smoke, and other damaging vapors that can diminish optical as well as electronic performance. In order to reduce the incidence of live-cell culture contamination by microorganisms, the microscope stage and surrounding area should be periodically wiped with 70-percent ethanol or commercial antiseptic towels. Culture media spills, an unavoidable factor in live-cell chamber manipulation, should be cleaned immediately and the surrounding area thoroughly disinfected.
Among the mechanical vibration sources that can affect microscope performance are central heating and cooling units (or air handlers) in the attic or on the roof of the building, refrigerators, low-temperature incubators (even in adjacent rooms), and traffic through nearby hallways. Vibrations from refrigerators and other sources that may not be immediately obvious can have a severe impact on microscope stability. A variety of techniques can be applied to reduce room and building low-frequency vibrations, including feedback-controlled isolation tables that are gas-filled (the most expensive option) and relatively low-cost flexible synthetic polymer vibration isolation pads (see Figure 4). The latter are marketed in a variety of geometries and damping levels to suit a wide spectrum of configurations. A combination of the vibration pads and a heavy sheet of half-inch aluminum or a pre-drilled isolation platform often will reduce vibrations to an unnoticeable level. Although inverted tissue culture microscope frames are much heavier and usually less sensitive to vibration compared with upright microscopes, they still benefit from isolation. High-frequency vibrations can, in many cases, be substantially reduced by loading the table top with additional mass, such as lead bricks. Finally, when very small displacements of sub-resolution features are the subject of investigation, additional steps may be required, such as working very late at night or early in the morning, when the environment is quieter.
In addition to vibrations conducted through the floor, possible airborne sources must also be considered. Equipment cooling fans or air-conditioning fans within the laboratory, and even exhaust noise from nearby vehicles (conducted through laboratory windowpanes), have been reported to produce troublesome vibrations for high-resolution microscopy on otherwise stable equipment. The vibration of cooling fans for lasers and other major equipment can also affect live-cell imaging stations. A section of flexible duct tubing placed between the cooling fan and the laser head can be effective in decoupling fan vibrations. In perfusion and gravity-fed media systems, vibrations due to gas bubbling through the feeding solution can couple through the tubing to the specimen chamber and produce periodic mechanical displacements. Acoustic noise can also be a source of vibration and reduced microscope stability. Noisy fan units should be replaced and other equipment that tends to produce excessive amounts of noise (and possibly vibration) should be relocated, if feasible, to another room. Microscopes are often equipped with built-in light sources and power transformers that can gradually heat the microscope frame and produce a slow focal drift of the specimen.
Control of room temperature for an otherwise stable microscope live-cell imaging system is another major concern and can be one of the most significant problems encountered when recording time-lapse sequences. A microscope that is confined to a small and poorly ventilated room can produce a considerable increase in ambient temperature. The heat load resulting from light sources, temperature controllers, shutters, cameras, computers, and other equipment may exceed the nominal capacity of the room, requiring the services of an auxiliary cooling system. Additional cooling improves the operation of all electronic equipment provided it does not introduce additional vibration and dust. In this regard, the cooling-fan capacity of microscope system computers and high-speed disk drives may become insufficient as the chassis is filled with heat-producing cards, such as those used for camera controllers, image processing software, and additional memory. Problems are manifested as system crashes, increased noise, and, in the worst extreme, lost data. Addition of a low-noise box fan to the computer housing can often alleviate this difficulty.
Although modern inverted tissue culture microscopes (those most favored for live-cell imaging experiments) are designed to be solid stand-alone instruments through a considerable amount of engineering effort to minimize vibration sources, aftermarket auxiliary components can often compromise this inherent stability. Fast shutters and optical filter changers (filter wheels) produce significant levels of vibration during operation and, when attached directly to the microscope frame, often induce perturbations that can last for tens to hundreds of milliseconds. In addition, many research-level microscopes often contain built-in motorized components (nosepieces, focus controls, etc.) that can be sources of vibration unless the manufacturers limit the speed of such devices and/or introduce delays between motorized functions and image acquisition. Vibration from the auxiliary components can be dramatically reduced or even eliminated by mounting these devices independently on separate stands adjacent to the microscope. Rigid aluminum optical breadboards containing pre-drilled mounting holes are commercially available, and are ideal for housing both the microscope and accessory components.
Fluctuations in the axial position of the microscope focal plane during the collection of sequential images from living cells is one of the most serious and frequently encountered problems in time-lapse microscopy. Often termed focus drift, changes to the microscope focal plane usually occur due to temperature variations in the imaging chamber or within the room in which the instrument is housed. Generalized qualitative analysis of the relationship between the thermal state of a microscope and the focal plane position indicates that a one-degree (Celsius) increase or decrease in temperature can shift the focus by approximately one-half micrometer (500 nanometers). Irregularity of the surface in glass coverslips, as well as mechanical instability arising from gear slippage, compression of lubricating grease layers, and settling between the moving components of the microscope, although not usually a major source of concern, can also contribute to focus drift. Without a feedback device to continuously monitor and correct focus, one of the best remedies for drift is to employ a thermostatically controlled enclosure that fully envelops the microscope and associated components. Although it is possible to obtain image sequences that remain in focus for short periods of time without auxiliary equipment, a high percentage of longer term experiments fail due to focus drift. In almost all cases, thermal instability leading to flexing of the coverslip or temperature gradients in the microscope optical train can usually be traced as the source of the failure.
Defeating focus drift should be a principal consideration during configuration of a microscope for time-lapse imaging, especially when using high numerical aperture objectives where the narrow depth of focus (approximately 300 nanometers) requires a focal position that is held to within a 100 nanometers of the initial plane. The entire system, including the microscope, camera, shutters, filter wheels, illuminators, live-cell chamber assembly, and host computer should be brought to operating temperature for at least 24 to 48 hours prior to initiating time-lapse imaging sequences. When assembling the imaging chamber, ensure that coverslips containing adherent cells are mounted securely in their housings and that the chamber itself is positioned on the stage in a manner that does not allow movement either in the lateral or axial directions. At high imaging resolutions, oil immersion objectives can be the source of focus drift as the oil spreads across the chamber coverslip and the front lens element (dry objectives do not have this problem). An objective heater, discussed above, should be used for all configurations that employ immersion objectives. Once the microscope is at the correct operating temperature and all other equipment is staged for the experiment, monitoring focus with a preliminary time-lapse sequence over a period of 12 to 24 hours using a fixed and permanently mounted specimen will provide an excellent indication of the system stability.
A variety of commercially available software and hardware solutions have been introduced by microscope and aftermarket manufacturers to contend with focus drift. Several of the hardware devices are autofocus systems that measure the distance between the objective front lens and the specimen by sensing light or sound reflected from the lower surface of coverslip (closest surface to the objective). This approach can be hampered, however, when high resolution oil immersion objectives are used, due to loss of contrast and reflectivity as the sensing light passes through the oil. The most advanced autofocus systems use low intensity near-infrared laser or light emitting diode (LED) sources to reflect a beam of illumination through the objective and onto the upper surface of the coverslip (supporting the cells and bathed by the culture medium), subsequently recovering the reflected light with the objective and passing it on to a detector that controls a feedback circuit to adjust the position of the objective in relation to the coverslip-culture medium interface.
Instantaneous feedback of specimen position enables the distance between the coverslip and objective to be detected in millisecond intervals during observation, thus automatically correcting for focus drift in real time. These systems are especially useful in terminating focus drift resulting from a temperature drop that occurs when changing and perfusing culture medium or adding reagents (such as drugs) to the culture. In addition, examining the cells to determine candidates for imaging is facilitated by constant focus corrections while translating the stage during observation and setting locations for multipoint image collection. Software accompanying the autofocus systems allows freely selectable focal planes through adjustment of an offset control. Due to the long wavelengths used by the laser or LED light source, the near-infrared detection system does not intrude on wavelengths used for observation and should be invisible to the fluorescence detector.
Illustrated in Figure 5 is the schematic diagram of a typical automatic focus drift correction mechanism for an inverted tissue culture microscope (Figure 5(a)) along with a spectral window comparing fluorescence emission profiles of common fluorescent proteins with the wavelength of the autofocus laser spectral line (Figure 5(b)). The laser (or light emitting diode) is focused into the rear focal plane of the objective and introduced parallel to its centerline axis with a specified offset. The positioning feedback loop corrects for minute geometrical shifts due to focus drift in the position of the beam that is reflected from the interface between the upper coverslip surface and the culture medium. The spectral emission profiles outlined in Figure 5(b) represent several of the most useful fluorescent proteins covering the wavelength region between 450 and 700 nanometers, including enhanced cyan (ECFP), green (EGFP), and yellow (EYFP) fluorescent proteins, as well as reef coral red-shifted variants, monomeric Kusabira Orange (mKO) and mCherry. The 770-nanometer laser line used by the focus drift correction system does not interfere with the fluorescent protein signals.
In laser scanning confocal microscopy focus drift correction during time-lapse investigations is often accomplished by obtaining multiple axial images at each time point with subsequent analysis of every image stack to identify a common focal plane that corresponds to a pre-selected reference point. Other approaches employ software algorithms that determine the focal position between time points using contrast functions that record images in successive upward and downward steps and compare the results until the highest level of contrast is obtained (at which point the focus is set for that interval). Software techniques rely on a relatively constant level of specimen contrast, however, which is often not the case with live-cell imaging where debris and other artifacts can randomly float into the viewfield and alter the apparent optimum focal plane.
Phototoxicity and Photodamage
Aside from the toxicity that occurs due to excessive concentrations of synthetic fluorophores and over-expression of fluorescent proteins, the health and longevity of optimally labeled cells in microscope imaging chambers can also suffer from a number of other deleterious factors. Foremost among these is the light-induced damage (phototoxicity) that occurs upon repeated exposure of fluorescently labeled cells to illumination from lasers and high-intensity arc-discharge lamps. In their excited state, fluorescent molecules tend to react with molecular oxygen to produce free radicals that can damage subcellular components and compromise the entire cell. In addition, several reports have suggested that particular constituents of standard culture media, including the vitamin riboflavin and the amino acid tryptophan, may also contribute to adverse light-induced effects on cultured cells. Fluorescent proteins, due to the fact that their fluorophores are buried deep within a protective polypeptide envelope, are generally not phototoxic to cells. However, many of the synthetic fluorophores, such as the MitoTracker and nuclear stains (Hoechst, SYTO cyanine dyes, and DRAQ5), can be highly toxic to cells when illuminated for even relatively short periods of time. In designing experiments, fluorophores that exhibit the longest excitation wavelengths possible should be chosen in order to minimize damage to cells by short wavelength illumination.
Presented in Figure 6 are three examples of phototoxicity induced by the illumination of cells labeled with synthetic fluorophores or expressing fluorescent fusion proteins. The normal rabbit kidney cells (RK-13) shown in Figure 6(a) were first transfected with a fusion vector of EGFP and the SV40 virus T-antigen nuclear targeting sequence to localize green fluorescence within the nucleus. The cells were subsequently treated for 20 minutes with MitoTracker Red CMXRos and imaged sequentially in 30-second intervals for 24 hours. After several hours, vacuoles began to form in the region containing mitochondria surrounding the nucleus. Illustrated in Figure 6(b) is a single human glioblastoma cell (U-251 line) expressing mCherry fluorescent protein fused to human beta-actin after several hours of observation. Note the significant deterioration of cellular structure that has occurred as the cell appears to enter necrosis. Detachment of Swiss albino mouse embryo cells (3T3 line) after an hour of imaging following treatment with the DNA-binding nuclear stain Hoechst 33342 is depicted in Figure 6(c). Ultraviolet-absorbing nuclear dyes (such as Hoechst) usually demonstrate phototoxic effects more quickly than do probes that are excited at longer wavelengths in the visible spectrum.
As discussed above, the synthetic biological buffer HEPES has been reported to produce a phototoxic effect on cells in some circumstances (although in many cases this might be attributed more directly to inadequate levels of bicarbonate). The investigator should keep in mind that all cells are intrinsically photosensitive, and adding fluorophores or aromatic components to the culture medium only compounds this sensitivity. Damage by the free radicals generated through excited state fluorophores can only be limited, not prevented. However, healthy cells have inherent enzymatic mechanisms for converting free radicals to less harmful compounds and can tolerate fluorescence excitation provided their enzyme systems are not saturated. Reducing the level of oxygen in the medium, provided the cells can survive oxygen withdrawal, can serve the dual purpose of limiting the degree of photobleaching and free radical production.
Regardless of the potential for cells to enzymatically deal with toxicity arising from fluorophores and culture medium components, the exposure of cells to light in the imaging setup should be reduced to the lowest possible level in order to limit other sources of cell damage and to minimize the potential for artifacts in the experiment. The illumination dose per image can be limited by the judicious application of neutral density filters (for arc-discharge lamps), lowering the output power of the illumination source (for lasers), and programming shutter assemblies to restrict exposure to fluorescence excitation light only during image capture. In addition, fluorescence filter bandwidths should be carefully chosen to reduce the number and intensity of unnecessary wavelengths that will illuminate the cells with light that is not useful for imaging. The ability to successfully decrease illumination levels in live-cell imaging is aided by using high numerical aperture objectives that feature the best light throughput and the fewest optical elements.
Even in the absence of fluorophores, the sensitivity of mammalian cells to ultraviolet light exposure has been well documented (a phenomenon known as photodamage), and many cell lines are at least equally sensitive to blue light in the wavelength regions used to excite cyan and green fluorescent proteins. To minimize photodamage and phototoxicity when setting up an imaging experiment, visualization of the living cells during microscope configuration must be performed at the lowest light levels under which the cells can be observed, and this should be conducted as quickly as possible. Neutral density filters (or very low laser power) should be used to attenuate the illumination source and visualization of the cells is best accomplished with the digital imaging system. Observing the cells through the microscope eyepieces takes several seconds, which is at least tenfold longer than is often required to obtain an image of sufficient quality for cell selection and focusing. Alternatively, candidate cells for imaging can be located using brightfield, phase contrast, or differential interference contrast (DIC) imaging modes. In all cases, a green or red interference filter should be used during setup to improve contrast and minimize exposure of the cells to blue light.
The widespread application of 546-nanometer (green) filters for live-cell imaging studies originated because this wavelength region matches one of the major spectral lines emitted by mercury arc-discharge lamps. However, achieving adequate levels of illumination intensity is rarely a problem with live-cell investigations and modern objectives are corrected to such a degree that they do not require the use of green light for high-resolution imaging. Thus, the choice of illumination wavelengths should be limited to those in the regions best tolerated by the cells. Immediately prior to cell division (prophase), most mammalian cell lines are very sensitive to ultraviolet and infrared light, and are least sensitive to red, green, and blue light, in descending order. Thus, in order to minimize photodamage, red interference filters with a bandpass region between 600 and 650 nanometers are the ideal choice for live-cell observations. Autofluorescence in mammalian cells and tissues is also reduced in the longer wavelengths of the visible light spectrum. Note that optical resolution is dependent upon wavelength and red light yields the lowest theoretical values for resolution due to the longer wavelengths involved. In most cases, however, the use of red light is not the limiting factor because the degree of resolution achieved in live-cell imaging is often compromised by internal cellular motion, temperature drifts, imperfections in the optical system, and illumination fluctuations.
In choosing filters for live-cell imaging experiments, bandwidths should be carefully selected so that even trace levels of infrared and ultraviolet light are eliminated. Even though modern bandpass filter designs perform well in the central regions of the visible light spectrum, they often pass radiation at very low and very high wavelengths. It is therefore advisable to install specialized glass filters near the illumination source(s) to block damaging ultraviolet and infrared wavelengths. Mercury and, to a lesser extent, xenon lamps produce high levels of ultraviolet light, while tungsten-halogen (transmitted) lamps emit significant amounts of infrared light. As a final step, electronic shutters should be installed (for both tungsten-halogen and arc-discharge lamps) to limit exposure of the cells to damaging radiation during periods when images are not being captured. Judicious shuttering of the illumination source is one of the most important factors in successful live-cell imaging experiments.
Advances in detector technology over the past few years have made it feasible to further reduce illumination levels in live-cell imaging experiments. Increasingly sensitive photomultiplier tube cathodes for confocal microscopy and advanced charge-coupled device (CCD) camera systems for widefield microscopy are continually being introduced. The intensified and electron multiplying camera systems now available are capable of imaging living cells with high sensitivity at light levels that are exceedingly low. Many of these cameras employ back-thinned CCDs, which are usually cooled and feature high quantum efficiency across the visible and near-infrared spectral regions to further increase sensitivity. If the best cameras are not available, sensitivity can be increased by combining the signal from multiple pixels (a process known as binning) at the expense of spatial resolution. In confocal microscopy, maintaining low zoom ratios will reduce the level of phototoxicity to cells. Increasing the confocal zoom factor causes the total amount of laser light to be scanned over a smaller region of the specimen, thus exposing the cells to more intense illumination.
Choosing the exact level of light attenuation and the correct exposure time is almost always an empirical exercise. For a new cell line with unknown parameters, the best strategy is to attenuate the light as much as possible and apply very short exposure times so that subcellular structures are just barely visible in the acquired image. A good place to start is a neutral density filter with an optical density of 1.0 and an exposure time of 100 milliseconds or less. For laser scanning confocal microscopes, start with a laser power of approximately 1 percent and increase the voltage (and gain, if necessary) of the photomultiplier. Use pixel dwell times about 50 percent shorter than those that usually produce adequate signal levels. If the cells are able to tolerate this light level through a long-term time-lapse experiment, then the illumination intensity and exposure time can slowly be increased in subsequent experiments until a workable compromise is achieved between signal-to-noise and cell viability. It should be noted that a nonlinear relationship often exists between the total amount of light exposure a cell can tolerate and the length of individual exposure times. In general, cells appear to be the healthiest when exposed to very brief pulses of light, since extended exposures (greater than one-half second) are often lethal over long periods of time.
Monitoring Cell Viability and Variability
After the live-cell imaging chamber has been loaded with fresh cells, assembled, and mounted on the microscope stage, the next step is to visualize the cells to establish their overall condition and morphology, and to identify candidates that are appropriate for imaging. In a vast majority of experiments, especially when imaging cells that have recently been transiently transfected with fluorescent proteins, there will be a significant amount of morphological variability in the cell population. It is not uncommon to observe cells that were either not transfected or exhibit poor localization of the probe. In addition, a percentage of the transfected cells will over-express the fluorescent protein, often to the point of creating a potentially toxic effect. Other cells may exhibit common health problems (as illustrated in Figure 7) manifested in the form of detachment from the substrate, excessive vacuole formation, swollen mitochondria, and cytoplasmic blebs. The same cell line in different cultures can demonstrate varying patterns of beading and blebbing that may be a symptom of dissimilar stress factors. Declining health often affects the rate of cell growth and motility, usually resulting in a general decrease in activity. However, poor health is not always indicated by a decline in cellular activity as compromised cells can exhibit a marked increase in Brownian-like organelle motions. Cells showing even a slight deviation from a normal and healthy appearance should not be pursued for imaging and data collection. Furthermore, if more than 50 percent of the population is judged as unhealthy, the entire culture should be discarded and exchanged for one that is in better physiological condition.
Many live-cell imaging experiments are performed with only a single or, at most, a few cells. The investigator should keep in mind that the morphology of cells in culture can be quite varied with regard to the apparent phenotypes present. This heterogeneity can be the result of cells in different phases of the growth cycle or possibly intrinsic differences among individual members of the population (the latter is more apparent in primary cultures). For this reason, it is often necessary to record data from a number of individual cells in order to gain a statistically significant sampling of cellular behavior and dynamics. As discussed above, the investigator should bear in mind that the imaging conditions in live-cell observations must be minimally perturbing to the cells in order not to significantly affect the experimental result. Among the most critical aspects of live-cell imaging experiments is establishing a criterion for judging the health of the cells under study so that the success of the experiment can be evaluated objectively. The exact criteria will vary depending upon the experiment, but one of the most important factors to be considered is whether the expected result was achieved without damage to the cells incurred by the imaging process. In some cases, the phenomenon under study can be matched to results obtained with fixed cells, but this is often not possible.
Imaging experiments should be monitored to ensure that the experimental conditions (culture medium, buffering strategy, atmosphere, chamber configuration) do not substantially alter the growth rate, mitotic index, or apoptosis properties of the cells. If possible, the potential negative effects of each experimental component should be separately assessed. For example, cells should be grown in the culture medium used for the imaging experiment in a standard carbon dioxide incubator and judged for viability, mitotic index, and general morphological variations. As a final check, the cells can then be installed in the environmental imaging chamber (without illumination) for a number of hours and then similarly assessed. This strategy isolates the various contributions to general cell health and readily identifies any procedures that require modification.
As discussed above, cells are exposed to high doses of illumination during live-cell imaging experiments and those that are labeled with fluorophores are potentially venerable to the generation of reactive molecular species that can seriously affect cellular function. It is therefore safe (and prudent) to assume that at least some level of damage has occurred during the imaging experiment and steps should be taken to determine whether it is significant enough to have produced detrimental effects. This is best accomplished by leaving the cells in the imaging chamber on the microscope stage after completion of the experiment with subsequent periodic examination to determine whether the cells initiate apoptosis or enter and complete mitosis. During the post-experiment evaluation process, time-lapse images can be recorded at widely spaced time intervals (10 to 30 minutes) over a number of hours.
The criteria for determining cell viability depend on the appropriate pathways present in the cells under examination. Some transformed mammalian cell lines have defunct cell cycle checkpoints that render determinations of mitotic and apoptotic progression useless. Each cell type should be carefully evaluated to determine the criteria that can be used for viability assessment following imaging sequences with the microscope. Among the simplest assays following an imaging experiment is to compare the apparent health and morphology of the cells that have been exposed to light during the investigation with neighboring cells in the same chamber that haven't been subjected to illumination. Similar features in the two populations, when observed with phase contrast or DIC, are a good indication that general health has not been compromised. This observation should be followed with fluorescence imaging to ascertain whether fluorophore localization has changed during the course of the experiment. Finally, both populations of cells should be intermittently observed for several hours (or days) following the experiment to see if the cells subjected to imaging behave similarly to the non-imaged cells. Comparisons of this type often reveal whether significant damage has occurred to the cells under study.
Examination of the mitotic pathway during live-cell imaging is an excellent mechanism with which to monitor cells for photodamage. Cells that are just beginning the mitotic cycle by initiating chromosome condensation (prophase) often fail to enter prometaphase and subsequently complete cell division when damaged by excessive illumination. Scanning the culture with the microscope operating in differential interference contrast mode can reveal those cells that are commencing mitosis, and a suitable candidate can be isolated for closer examination under the actual experimental imaging conditions. If the chromosomes undergo decondensation during the observation period and the cell fails to re-enter mitosis within several hours, there is a good likelihood that photodamage has occurred. Note that changing the culture medium will also initiate decondensation of prophase chromosomes in many cells, so this analysis should be performed several hours after the cells are placed on the microscope stage. Cell lines differ widely in their ability to tolerate light during mitosis. Established lines and primary cultures of human and animal cells can be extremely sensitive to light, whereas cells derived from embryos (such as fruit flies and nematodes) often lack pathways to arrest the division cycle in response to DNA damage and are more tolerant to photodamage.
Aside from the numerous problems associated with photodamage and phototoxicity, as well as the rigorous maintenance requirements incumbent on live-cell imaging, the investigator must also be alert to the possibility of microbial contamination during the course of the experiment. The most common infections are those that occur due to bacteria, fungi, mycoplasma, yeasts, molds, and in rare circumstances, protozoa. Unless it becomes a frequent event, the nature of the infection or species involved is not as important as determining where the contamination originated. In general, rapidly growing microorganisms are less problematic due to the fact that they are usually readily detected and the culture can be quickly discarded. More important are those infections whose presence is cryptic, unable to be visualized during routine examination of the culture because of the physical size, or enabling the invader to grow at a level that escapes detection. Overuse of antibiotics in the culture medium is a common problem that often results in a low-level contamination, which can remain undetected for long periods of time and may ultimately interfere with normal mammalian cellular function.
Presented in Figure 8 are micrographs illustrating three of the most common visible sources of microbial contamination in live-cell cultures. Unchecked bacterial growth (Figure 8(a)) is readily detected at high magnification (often accompanied by a rapid drop in culture medium pH) and can even be visualized as turbidity in the culture medium with the unaided eye when the organism numbers begin to reach saturation. Yeast (Figure 8(b)) grow far more slowly than bacteria, but have a distinct colonial motif that reveals their presence. Mold contamination (Figure 8(c)) usually overtakes a culture within 24 to 48 hours, and can often be identified as a bushy, fibrous intrusion. Perhaps the most serious form of contamination, however, is not obvious using routine contrast-enhancing microscope techniques (phase contrast and DIC). Mycoplasma (not illustrated) can seriously alter cell behavior and metabolism, but are not easily identified in cell cultures other than through often subtle signs of gradual deterioration. Assays for mycoplasma should be routinely conducted to ensure cultures are free of this serious artifact in live-cell imaging. A variety of detection methods are useful for revealing the presence of mycoplasma, including fluorescent staining (using the nuclear dye Hoechst), polymerase chain reaction, and autoradiography.
Dynamic imaging of biological activity was introduced in 1909 by French doctorial student Jean Comandon, who presented the earliest reported time-lapse cinema films of syphilis-producing spirochaetes, 5 years before Charlie Chaplin made his first movie. Comandon's technique, which he called microcinematography, enabled the production of movies capturing events in the microscopic world that could be recorded using an enormous cinema camera bolted onto a fragile darkfield microscope. These films proved instrumental in teaching physicians how to distinguish disease-causing spirochaetes from those that are harmless, and demonstrated how time-lapse observations can be employed to gain important biological information without recourse to image analysis, processing, or even empirical quantitative measurements. For the next 75 years, many microscopists adapted increasingly advanced cinema film cameras to the microscope in order to generate progressively better films at much higher resolution.
As tube-based video cameras became affordable in the late 1970s and early 1980s researchers began to couple these devices with optical microscopes to produce analog time-lapse image sequences and real-time videos. The tube camera ultimately gave way to the area array CCD in the early 1990s, heralding a new era in photomicrography and signaling the ultimate demise of film. With the advanced digital camera systems available in the 21st century, the increasingly popular technique of time-lapse cinemicrography is becoming broadly applied to capturing events that occur in living cells over periods ranging from a few seconds to several weeks (or even months). The technique involves repeated imaging of a cell culture at defined time points, thereby providing information on myriad dynamic processes that often occur with a wide distribution of time scales. When time-lapse investigations are coupled to labeling cells with synthetic fluorophores and genetically encoded fluorescent proteins, events at the subcellular and molecular levels can be investigated.
Time-lapse imaging can be performed in two spatial dimensions using widefield techniques and extended even further to three-dimensional imaging with confocal microscopy. In addition, modern confocal microscopes are equipped with line-scanning software for rapid and repeated imaging of single scan lines. Two-dimensional time-lapse imaging involves sequential capture of single focal planes (x-y in widefield microscopy and x-y, x-z, and y-z in confocal microscopy), whereas three-dimensional imaging produces optical stacks from multiple focal planes in a variety of dimensional formats with thick specimens. When a single focal point in the lateral plane (x and y) is combined with z-stack imaging as a function of time, the technique is referred to a 4-D time-lapse imaging. Likewise, adding a fifth dimension (wavelength) yields 5-D imaging whereas adding either multiple wavelengths or multiple lateral regions to the 4-D stack is referred to as 6-D time-lapse imaging. As described above, the time intervals for sequential image gathering using these techniques can range from milliseconds to days or even months.
A growing number of small molecule, vital synthetic fluorescent probes that yield highly specific cellular or subcellular labeling patterns are now commercially available. In addition, the huge effort to produce useful fluorescent proteins having emission colors spanning the visible and near-infrared spectrum is beginning to produce encouraging results. Green fluorescent protein and related spectral variants are now routinely being fused to other proteins of interest to reveal details concerning protein geography, movement, lineage, and biochemistry in living cells. In this regard, these biological probes have provided an important new approach to understanding protein function, which is the next logical step for investigations of cellular processes now that the genome sequence of many organisms has been determined. The inherent brightness and photostability of many fluorescent proteins render them well-suited for the repeated imaging required in time-lapse studies. Together, the synthetic and genetically encoded fluorescent probes are affording a seemingly endless array of possibilities for imaging molecular components in living cells.
Illustrated in Figure 9 are several images from a 24-hour time-lapse sequence of rabbit kidney epithelial (RK-13) cells that were captured using a combination of fluorescence and differential interference contrast. The cells were transfected with a chimera of mCherry fluorescent protein fused to human beta-actin to reveal the distribution of this cytoskeletal component in the main cell body as well as the lamellapodia. The white arrow in Figure 9(a) indicates the initiation of a cytoplasmic ruffle near the large pool of actin in the central region of the cell. As the sequence progresses, the small ruffle begins to swell and extrude towards the left-hand side of the image, in a wave-like motion, concentrating the brightly labeled actin fusion protein into the leading edge as it grows (indicated by the arrows in Figure 9(b) through 9(d)). As the lamellapodium spreads to cover a large area, small clusters of labeled actin form behind the leading edge (arrows in Figures 9(e) and 9(f)) in structural elements that may be podosomes involved in the process of cellular adhesion to the glass substrate.
Time-lapse imaging techniques are significantly aided by the application of critical microscope auxiliary components, such as electronic light shutters, filter wheels, motorized stages, and focus drift correction mechanisms, which can be coordinated and controlled by a host computer using commercially available image acquisition software. Electronic shutters are necessary to block illumination of the specimen between camera exposures when imaging fluorescently labeled cells in order to minimize photodamage, phototoxicity, and photobleaching, thus dramatically extending the quality of images and cell viability over the long periods of time often used in time-lapse experiments. Simultaneous imaging of multiple fluorophores, as well as combined fluorescence and DIC imaging, requires motorized filter wheels that can rapidly switch between several fluorescence filters and/or a polarizer. In practice, excitation illumination is controlled by a filter wheel that contains two or more bandpass excitation filters, while fluorescence emission is gathered through a second filter wheel that houses either bandpass or longpass barrier (emission) filters. Beneath the objective, a multiple bandpass dichromatic mirror is installed to coordinate blocking and reflecting of excitation wavelengths and simultaneous passing of emission wavelengths through a single, stationary optical element. Advanced filter wheels can switch between adjacent filters in 30 to 50 milliseconds with electronic shutters having operating specifications in the same range, which is the limiting factor in determining the shortest possible time interval between successive images in time-lapse sequences.
In confocal microscopy, time-lapse image collection is limited by the speed of the scanning mirrors. The highest scanning rates are achieved by reducing the image pixel dimensions and employing the fastest raster speed available on the instrument, but this is usually restricted to approximately 8 to 10 frames per second. Collecting image sequences over shorter time intervals requires single-line scanning, swept-field instruments, or spinning disk microscopes. Modern spinning disk and swept-field microscopes can routinely capture image sequences at high rates, ranging from one to several hundred frames per second, and are usually only limited by the speed of the digital camera system or photomultiplier. These instruments are designed to accommodate a host of experimental variables in high-speed time-lapse imaging at very low light levels.
Live-cell imaging experiments can be extraordinarily powerful, but they can also significantly benefit from complementary investigations using fixed cell assays to assist in the validation of phenomena that are observed in the living cells. Although this may seem counterintuitive due to the fact that live-cell assays are often held as the standard for cellular and molecular dynamics, the technical difficulties and risk of damaging cells during investigations can be substantially overcome if similar results in terms of kinetics and time point events can be verified with fixed cells. In many cases, comparing fixed cells to live-cell imaging is simply not possible, either because the kinetics of an event are too fast or because the nature of the experiment (for example, dynamics) cannot be meaningfully performed within the fixed cell. Moreover, the point of the live-cell experiment is to reveal events or properties that are not observed or easily interpreted in fixed cells. Nonetheless, it is worthwhile to consider the use of this approach as a technique for monitoring events seen in longer time-lapse experiments to confirm the absence of deleterious effects by extended illumination.
Modern instrumentation enables the imaging of living specimens at high signal-to-noise ratios using extremely low levels in incident illumination and now stands as a powerful technique for the analysis of molecular dynamics within cells. Continued advances in imaging techniques and fluorescent probe design enhance the power of this approach and ensure its future as an important tool in modern biology. Technological and conceptual advances in instrumentation are also likely to push spatial and temporal resolution to new limits, as well as perfecting the modes of fluorescence microscopy currently in use. Several of the recently introduced approaches, such as stimulated emission depletion and 4Pi microscopy, appear to be promising techniques for live-cell imaging, but their potential for widespread use in biological applications has yet to be established, and there are limitations on the thickness of acceptable specimens. In the final analysis, however, the technical care and expertise required to conduct a successful live-cell imaging experiment with currently available instrumentation is considerable, and even with likely advances there remain numerous obstacles.
Michael E. Dailey - Department of Biological Sciences and Neuroscience Program, 369 Biology Building, University of Iowa, Iowa City, Iowa, 52242.
Daniel C. Focht - Bioptechs Inc., 3560 Beck Road, Butler, Pennsylvania, 16002.
Alexey Khodjakov and Conly L. Rieder - Wadsworth Center, New York State Department of Health, Albany, New York, 12201, and Marine Biological Laboratory, Woods Hole, Massachusetts, 02543.
Kenneth R. Spring - Scientific Consultant, Lusby, Maryland, 20657.
Nathan S. Claxton, Scott G. Olenych, John D. Griffin, and Michael W. Davidson - National High Magnetic Field Laboratory, 1800 East Paul Dirac Dr., The Florida State University, Tallahassee, Florida, 32310.