Live‐Cell Fluorescence Imaging

CHAPTER 7

Live-Cell Fluorescence Imaging Jennifer C. Waters Department of Cell Biology, Harvard Medical School Boston, Massachusetts 02115

I. Introduction II. Preparing a Specimen for Fluorescence Live-Cell Imaging A. Choice of Fluorophore B. Mounting the Specimen for Viewing C. Maintaining Temperature D. Cell Culture Media for Imaging E. Preventing Photobleaching III. Choice of Microscope A. The Microscope Stand B. Mode of Fluorescence Microscopy IV. Wide-Field Illumination of the Specimen A. Light Source B. Diaphragms C. Filter Sets D. Automation of Filter Selection V. Choosing the Best Objective Lens for Your Application A. NA: Resolution and Brightness B. Magnification C. Correction for Aberrations D. Phase and DIC Objective Lenses VI. Acquiring Digital Images Over Time A. Cameras B. Shutters C. Maintaining Focus VII. nD Imaging VIII. Verifying Cell Health and Troubleshooting Sick Cells IX. Conclusion References

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I. Introduction With the discovery of genetically encoded fluorescent proteins, such as green fluorescent protein (GFP; Chapter 6 by Straight, this volume; Shaner et al., 2005), many molecular and cell biologists who previously used microscopy only for simple localization experiments in fixed specimens now wish to use microscopy for both localization and quantitation of fluorescence in live cells over time. Accurate localization and quantification of fluorescence requires images of the highest possible quality (Chapter 17 by Wolf et al. and Chapter 22 by Murray, this volume), which is particularly diYcult to achieve in live cells. The ideal fluorescence image has a high signal-to-noise ratio and adequate spatial resolution. While spatial resolution is necessary to discern specimen detail, the most challenging and critical parameter for live-cell imaging is detection of weak fluorescent signals above background noise generated by the specimen and the imaging system. Imaging systems with high signal detection to noise ratio are essential both for detecting weak fluorescence signals and for accurate quantitation of small changes in concentration of fluorophore. Imaging live cells poses many more problems than imaging fixed specimens. For live-cell fluorescence imaging, attaining an adequate image must be balanced with minimizing illumination to protect the specimen against photodamage and photobleaching. Live cells expressing fluorescent protein fusions are usually dim compared to fixed specimens, both because the fluorescent proteins are not very bright and because there is, in most cases, only one fluorophore per protein. It is Table I Fluorescence Live-Cell Imaging Checklist To increase signal □ Choose a bright fluorophore □ Image through a clean No. 1.5 coverslip □ Mount specimen as close to coverslip as possible □ Use high NA, minimally corrected oil immersion clean objective lens with lowest acceptable magnification □ Choose fluorescence filter sets that match fluorophores □ Align arc lamp for Koehler illumination □ Use antiphotobleaching inhibitors □ Remove DIC Wollaston prism and analyzer from light path □ Use cooled CCD camera with at least 60% quantum eYciency □ Use camera binning To decrease background/noise □ Turn oV the room lights □ Close down the field diaphragm □ Use phenol red-free media □ Use cooled CCD camera with low readout noise

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also favorable to choose cells that are expressing low levels of fluorescent protein fusions to minimize the diVerence from levels of the endogenous protein in vivo. Long camera exposure times that allow accumulation of weak signals must often be avoided to reduce photobleaching and phototoxicity, and to acquire images quickly enough to capture cell dynamics. With these challenges, it is especially important that the specimen, microscope, and detector be optimized to maximize the signal while minimizing the addition of noise to the image (Table I). Live-cell imaging is further complicated by the need to keep the specimen at the appropriate temperature and pH and to maintain focus over time. In this chapter, we will begin with the specimen and work our way through the optical path to the detector, examining ways to optimize the signal-to-noise ratio while keeping the specimen healthy.

II. Preparing a Specimen for Fluorescence Live-Cell Imaging A. Choice of Fluorophore Fluorophores are molecules that absorb light of particular wavelengths, leading to the emission of longer wavelengths of light (Chapter 5 by Wolf, this volume; Herman, 1987; Tsien et al., 2006). Fluorophores vary greatly in their brightness and rate of photobleaching. Among the many fluorescent proteins that are now available, there can be brightness diVerences as much as 30-fold (Thorn, 2000). To attain the highest possible signal-to-noise and signal-to-background ratio in our final image, we want to start with the brightest possible fluorophore. Brightness is determined by the fluorophore’s extinction coeYcient and quantum yield. Extinction coeYcient is a measure of how eYciently a fluorophore absorbs light while quantum yield is the ratio of the number of photons emitted to the number of photons absorbed. These numbers are usually available from the manufacturer or in the published articles in which the proteins were introduced. Fluorescent proteins are discussed in more detail in Chapter 6 by Straight, this volume.

B. Mounting the Specimen for Viewing

1. Avoiding Spherical Aberration and Light Scattering Spherical aberration is an axial elongation and asymmetry of the image, which decreases axial resolution (resolution in ‘‘z’’; Keller, 2006; Wallace et al., 2001). Spherical aberration also greatly reduces the intensity and clarity of the image (North, 2006; Fig. 1) and should therefore be avoided as much as possible in live-cell fluorescence imaging. To avoid spherical aberration, one should consider the specifications of the objective lens (Chapter 1 by Sluder and Nordberg and Chapter 3 by Ernst Keller, this volume; Abramowitz et al., 2002; Keller, 2006) when deciding how to mount a specimen for imaging. The highest resolution

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Fig. 1 Spherical aberration can be introduced by the specimen. Fluorescence images of tissue culture cells labeled with Alexa 488 phalloidin. A diVerence in refractive index between the lens immersion medium and the specimen itself (i.e., immersion oil and an aqueous medium) causes spherical aberration. Spherical aberration was generated using an immersion oil with a higher refractive index (1.534) than that of the specimen mounting media (
1.45). Spherical aberration results in decreased intensity and resolution.

objective lenses are designed to image a specimen that is just beneath a 170-mm thick coverslip and mounted in a media with the same refractive index as the lens immersion medium. Spherical aberration is introduced when there is a mismatch in refractive index between the media surrounding the specimen and the objective lens immersion medium, as is the case when a specimen mounted in aqueous cell culture media is imaged with an oil immersion objective lens (Egner and Hell, 2006). The eVect gets worse and image intensity decreases as distance from the coverslip increases, so every eVort should be made to mount the specimen as close to the coverslip as possible. Water immersion lenses can be used to reduce spherical aberration when imaging a specimen mounted in an aqueous medium and are particularly useful when imaging deep into an aqueous solution. Water immersion lenses are used with a No. 1.5 coverslip, and should not be confused with waterdipping lenses (such as those used for electrophysiology work) which are designed to be used without a coverslip. Care must be taken when using water immersion lenses, as tilt in the coverslip relative to the lens can result in additional aberrations (Arimoto and Murray, 2004). With high numerical aperture (NA) air objectives in particular, using the wrong thickness of coverslip increases spherical aberration and decreases image intensity (Spring and Davidson, 2006). The thickness of coverslip (in millimeters) that the lens is designed to be used with is marked just after the infinity symbol (which refers to the now almost universally used infinity corrected optics) on the barrel of most modern objective lenses, and is 0.17 mm (grade No. 1.5) in most cases (Abramowitz et al., 2002). The eVect of coverslip thickness on images formed with high NA dry objectives is so detrimental to image quality that many high

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resolution air objective lenses come with a correction collar that is used to compensate for error in coverslip thickness. These collars wrap around the barrel of the objective and are adjusted by rotating the collar relative to the objective lens. If such a collar is present, it should be adjusted for each specimen every time the lens is used to minimize spherical aberration (Fig. 2A and B). It is also very important for image quality that the coverslip surface proximal to the objective lens and the top lens of the objective are clean (see Inoue´ and Spring, 1986 for thorough instructions on how to clean an objective lens). Prior to imaging, use a gentle glass cleaner such as Sparkle (www.glasscleaner.com) or a weak solution of a mild detergent to wipe any dirt and excess media from the coverslip. This is especially important when using an oil immersion objective, as residue on the coverslip will mix with the immersion oil, leaving the objective lens in need of cleaning as well. Also note that immersion oil from diVerent manufacturers does not necessarily mix, and old immersion oil accumulates dirt. Clean old immersion oil oV of the coverslip surface with Sparkle or pure ethanol before adding fresh immersion oil.

2. Chambers for Live-Cell Imaging Chambers for live-cell imaging serve multiple purposes (Dailey et al., 2006a; Swedlow and Andrews, 2005). They hold the specimen on a coverslip and allow for a volume of cell culture media necessary to keep the cells alive. Chambers designed for live-cell imaging may be open or sealed closed. Open chambers are used when the cells need to be accessed directly before or during imaging, for experiments which require drug delivery or microinjection, for example. Chambers that can be sterilized are preferable for long-term imaging experiments. And finally, specialized chambers provide temperature and CO2 control (described in Section II.C). There are several commercially available dishes for use in optical microscopy. One commonly used simple design is a round plastic tissue culture dish with a coverslip bottom (MatTek, Ashland, MA). Rectangular coverlips with plastic multiple well chambers are also available (LabTek II Chambered Coverglass, Nalgene Nunc, Rochester, NY), as are coverslip bottomed 6 through 96-well plates (MatTek; Nalgene Nunc). With all of these commercially available dishes, be sure to order No. 1.5 thickness of coverslip glass. These dishes are available sterilized and/or with coatings such as poly-L-lysine to enhance cell adherence to the glass. For a more economical solution, reusable metal chambers that hold standard coverslips can be made with easily obtainable materials and the help of a machine shop (Dailey et al., 2006a; Rieder and Cole, 1998; Sluder et al., 2005). C. Maintaining Temperature Cells must be kept at physiological temperature during imaging to ensure health and to be confident that cell behavior on the microscope is an accurate representation of behavior in vivo (Swedlow and Andrews, 2005). For the most commonly

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Collar in wrong position A

Collar aligned properly B

C o r r e c t i o n

N u C m e r i c a l

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PlanFluor, NA 0.75

40
PlanFluor, NA 1.3 D

a p e r t u r e

100
PlanApo, NA 1.4 E

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PlanApo, NA 1.4 F

M a g n i f i c a t i o n

Fig. 2 Image quality and brightness depend on the objective lens. Fluorescence images of tissue culture cells labeled with Alexa 488 phalloidin (Invitrogen, Eugene, OR). Each image pair was collected and is displayed identically to demonstrate diVerences in intensity. (A and B) High NA dry objectives

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used mammalian cell types, this requires heating the sample to 37  C. There are many commercially available solutions to heating specimens on a microscope stage. Simple and economical designs use heating elements to warm metal chambers that hold a coverslip (20/20 Technology, Inc., Wilmington, NC; Warner Instruments, Hamden, CT). These work well for shorter term experiments. However, in this design, oil objective lenses and the stage act as heat sinks, drawing heat way from the specimen. Maintaining focus during live-cell imaging can be a major source of frustration, and heating only the chamber which holds the sample introduces thermal drift as the heater cycles on and oV. One way to help alleviate this problem is to add a heated collar that is placed around the barrel of the objective lens (Bioptech, Butler, PA). However, heating the objective lens does not completely eliminate thermal drift, and cycles of heating and cooling are not particularly good for the objective lens. The most reliable way to maintain temperature over time is to build an incubator around the body of the microscope (Solent Scientific Limited, Segensworth, UK). Microscope incubators are usually made of plexiglass and are custom designed to fit around the microscope and peripheral devices. These incubators enclose the majority of the microscope and keep it at a stable temperature, which helps to eliminate thermal drift. With microscope incubation chambers, a specimen prepared on a coverslip bottomed dish (as described above) can move directly from the cell culture incubator to the microscope stage, without the need to remount the specimen in a chamber. This is not only convenient but also maintains sterility. D. Cell Culture Media for Imaging Typical media used for tissue cell culture, such as D-MEM, contains phenol red to monitor pH and is buVered by sodium bicarbonate and an atmosphere of 5% CO2. Phenol red-free versions of these media should be used for fluorescence livecell imaging, as phenol red is fluorescent and increases background in the image. A method of buVering the media while the cells are on the microscope stage is also needed. Some commercially available temperature control chambers deliver humidified 5% CO2 to the specimen, so traditional sodium bicarbonate buVered media can be used. With simple sealed chambers, media can be purged with CO2 prior to sealing the chamber. Another common approach is to change to a prewarmed media that maintains pH in atmospheric CO2 just prior to imaging such as Gibco CO2-Independent Media (Invitrogen, Eugene, OR) or Leibowitz. Supplementing sodium bicarbonate buVered media with 10–25-mM HEPES can also help to maintain pH under atmospheric CO2. It is critical that the cells be tested for viability in the media used for imaging (discussed in Section VII). (in this case, a 40 PlanApo NA 0.95) often have a correction collar for coverslip thickness. Neglecting to adjust these collars introduces spherical aberration which degrades image quality. (C and D) An image formed with a high NA oil immersion lens (NA 1.3) is much brighter than an image formed with a lower NA dry lens (NA 0.75). (E and F) Increasing magnification without increasing NA results in a dimmer image.

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Evaporation of the cell culture media can be a problem with a specimen in an open chamber, especially when heated to 37  C. We have had good results using a layer of mineral oil poured over the cell culture media to prevent evaporation. The oil allows suYcient gas exchange (to maintain pH in an atmosphere of 5% CO2 when necessary) while minimizing evaporation for, in our tests, as long as 4 days. A capillary needle can also be moved through the layer of mineral oil to reach the cells, providing a convenient way to ‘‘seal’’ a chamber during microinjection or micromanipulation experiments.

E. Preventing Photobleaching Photobleaching is the irreversible destruction of the fluorophore that can occur when the fluorophore is in an excited state, which leads to the familiar fading of fluorescence during observation (Diaspro et al., 2006). Photobleaching is best prevented by optimizing the imaging system to maximize signal and minimize noise (Table I) so that the lowest possible level of illumination can be used. A lot of photobleaching occurs when searching for the perfect cell through the eyepiece or while focusing the image of the specimen on the camera. The amount of time spent illuminating the specimen when not acquiring images should be kept at a minimum. If photobleaching is still a problem, it can be prevented by depleting oxygen from the media. One commercially available solution is Oxyrase or OxyFluor (www.oxyrase.com), which was originally made for growing anaerobes and has been successfully used to prevent photobleaching during live-cell imaging (0.5–1.0 units oxyrase per milliliter imaging media; Wittmann et al., 2003).

III. Choice of Microscope A. The Microscope Stand Inverted microscope stands are preferable for most live-cell imaging applications. With the inverted microscope design, the specimen is imaged with an objective lens from beneath the microscope stage. This allows for suYcient room on the microscope stage for cell culture dishes filled with media and heated incubation chambers, and allows easy access to the specimen for perfusion or microinjection. Highquality inverted microscope stands are heavy and stable, making them resistant to focus drift. To prevent vibration that can degrade resolution, the microscope stand should be mounted on a vibration isolation table.

B. Mode of Fluorescence Microscopy There are various modes of fluorescence imaging, including wide-field fluorescence microscopy (Herman, 1987), multipoint scanning confocal [(Toomre and Pawley, 2006); such as spinning disk (Adams et al., 2003; Maddox et al., 2003),

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swept field, or slit scanning confocal (Liebling et al., 2006)], laser scanning confocal (Chapter 25 by Bulseco and Wolf, this volume; Conchello and Lichtman, 2005), multiphoton excitation microscopy (Piston, 1999), and total internal fluorescence microscopy (TIRF; Axelrod, 2001). Confocal and TIRF are dealt with in more detail in other chapters of this volume; this section provides general guidelines for choosing the appropriate technique for a live-cell imaging experiment. Standard fluorescence microscopy (Herman, 1987) is referred to as wide-field to diVerentiate it from other fluorescence microscopy techniques. Unlike confocal techniques, wide-field uses full-field illumination, and all of the emission light that passes through the objective is sent to the detector. For live-cell imaging, widefield can be used with a high quantum eYciency, low noise cooled charge-coupled device (CCD) camera (Section V.A; Chapter 10 by Spring, this volume; Spring, 2000), helping to make it a high signal-to-noise ratio technique (Swedlow et al., 2001) and an excellent choice for live-cell imaging experiments. If a focus motor (Section V.C) is used to collect multiple focal planes at regular intervals (‘‘z-series’’), constrained iterative digital deconvolution can be used to increase the signal-to-noise ratio of wide-field images (Wallace et al., 2001). It is a common misconception that confocal microscopy is a higher resolution technique that is always preferable to wide-field when available. The benefit of confocal microscopy over wide-field is not increased resolution as much as the removal of out-of-focus fluorescence. Therefore, in specimens where there is no significant out-of-focus fluorescence, there is no benefit of using a confocal microscope. The resolution of both wide-field and confocal are limited by the NA of the objective lens (Section IV) and the wavelength of emission light (Inoue´, 2006). While there is a modest increase in axial resolution when using a confocal microscope, it is diYcult to achieve and rarely enough to uncover new information in specimens with minimal out-of-focus fluorescence. The use of the pinhole in confocal microscopy leads to a 20% loss of in focus light from the image plane (Pawley, 2006), significantly reducing the useful signal that makes it to the detector. Since wide-field fluorescence with a high-quality cooled CCD camera has been shown to be a lower noise technique than laser scanning confocal (Swedlow et al., 2001), in cases where a decrease in out-of-focus fluorescence is not needed, widefield microscopy will produce a higher signal-to-noise image than laser scanning confocal. The test that works well in most cases is as follows: if the wide-field fluorescence image looks good through the eyepiece, you do not need a confocal microscope. In live specimens that do have significant out-of-focus information that degrades the wide-field image, confocal microscopy can produce a very satisfying increase in the signal-to-noise ratio (Dailey et al., 2006b). Laser scanning confocal (also called point scanning confocal) has the benefit of variable pinhole, so its size can be matched to the objective lens and wavelength of light to provide maximum removal of out-of-focus information. For low light level live-cell applications, however, it can be diYcult to get a reasonable signal-to-noise image at laser levels that do not bleach the specimen and at speeds fast enough to capture fast cell dynamics.

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There are now several designs of faster multipoint or slit scanning confocals available (Toomre and Pawley, 2006). A commonly used model is the Yokogawa spinning disk confocal, which has been shown to be excellent for low light level live-cell imaging (Adams et al., 2003; Maddox et al., 2003). This model is designed for high resolution work, as the fixed pinhole size provides maximum reduction of out-of-focus fluorescence only when used with a 100 1.4 NA objective lens. These confocals acquire high signal-to-noise images with reduced out-of-focus fluorescence at a much faster rate than laser scanning confocals and are therefore favorable for live-cell confocal imaging. When deep penetration into biological tissue (100 mm or more) is needed, multiphoton excitation is necessary. The long wavelengths of illumination used for multiphoton penetrate deeply into biological specimens, and emission light scattered by the specimen is collected and used for image formation (Piston, 1999). Since longer wavelengths of light are less damaging to cells, multiphoton excitation can also be useful for imaging fluorophores that are normally excited by high energy shorter wavelength photons. However, multiphoton is expensive, diYcult to execute well, and a lower resolution technique, and should therefore only be used when the specimen demands it. In TIRF microscopy (Axelrod, 2001), the evanescent wave of energy that forms at the boundary between two media of diVerent refractive index (usually the coverslip and the aqueous specimen) when light totally internally reflects is used to excite fluorophores in the specimen. The evanescent wave penetrates only 50–100 nm into the specimen, so only fluorophores very close to the coverslip surface are excited, resulting in a very high signal-to-noise image with a much thinner optical section than confocal or wide-field (50–100 nm vs
700 nm). However, it is only useful for studying parts of the specimen that are close to the coverslip surface, such as membrane proteins and focal adhesions.

IV. Wide-Field Illumination of the Specimen A. Light Source The most commonly used light source for fluorescence microscopy is a mercury or xenon arc lamp (Nolte et al., 2006; Ploem, 1999). These are intense light sources that produce a large amount of heat, which can damage live specimens. A heat absorption filter (available from the microscope manufacturer) should therefore be placed in the light path, just after the lamp housing. In order to get even illumination across the entire field of view, it is essential to align the arc lamp so that it is in focus and centered at the back focal plane of the objective lens, referred to as Koehler illumination (Inoue´ and Spring, 1986; Waters, 2005). When the arc lamp is in focus at the back focal plane of the objective lens, the objective lens will project a maximally out-of-focus image of the light source onto the specimen. This provides even illumination of the specimen. To view the

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image of the arc at the back focal plane of the objective, unscrew an objective lens from the nosepiece and place a piece of paper on the stage. Use the collector lens, found at the front of the lamp housing, to focus the image of the arc onto the paper. Two translation screws on the lamp housing allow movement of the bulb in X and Y to center it in the field of view. Many lamp housings have a mirror behind the arc to reflect down the optical path light that would otherwise be lost. If a mirror is present, there will be three additional screws at the back of the lamp housing that move the mirror in Z, X, and Y so that the reflected image of the bulb can also be focused and centered at the back focal plane of the objective. When you are in the fortunate position of having a specimen that is too bright for the camera, there are often neutral density filters in the light path between the lamp housing and filter sets that can be used to attenuate illumination. These filters are most often labeled ‘‘ND’’ and then a number which is the denominator in the fraction of light that the filter allows through. For example, an ND 4 filter allows one-fourth of the illuminating light through to the specimen. B. Diaphragms There are two diaphragms present in the fluorescence light path (Inoue´ and Spring, 1986; Ploem, 1999). Both diaphragms are located in the light path between the lamp housing and the fluorescence filter sets. The aperture diaphragm (found in some but not all fluorescence microscopes) is in focus at the back focal plane of the objective. Its location means that closing down the diaphragm decreases the NA of the illumination and decreases intensity of illumination. The second diaphragm is the field diaphragm. The field diaphragm is placed in a position in the optical path such that it is in focus at the image plane. When closed down while looking at the image of the specimen, the in-focus edges of the octagonalshaped field diaphragm can be seen. It is very important in live-cell imaging to properly adjust the field diaphragm to frame the field of view. The field diaphragm should be open just enough to illuminate only the area of the sample you are interested in. Note that the area imaged by the camera is often smaller than that seen through the eyepiece, and the field diaphragm should therefore be closed down further when imaging with a camera. If the field diaphragm is open more than is needed, parts of the specimen are illuminated that you are not imaging, and light scattered from these parts of the specimen can contribute background to the image (Fig. 3A and B). C. Filter Sets A set of three filters is used to visualize fluorophores in the wide-field fluorescence microscope (Kinoshita, 2002; Ploem, 1999; Reichman, 2000). The illuminating light first encounters the excitation filter, which is chosen to allow through the wavelengths of light that excite the fluorophore. The light then hits the dichromatic mirror (or beam splitter), which is positioned at a 45  angle relative to the

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Field diaphragm open A

Field diaphragm closed B

Room lights on C

Room lights off D

Fig. 3 Stray light adds noise to fluorescence images. (A and B) Fluorescence images of live tissue culture cells expressing a GFP fusion protein. (A) With the field diaphragm all the way open, cells which are not of interest (top left corner) contribute scattered light to the image. (B) Closing down the field diaphragm so that only the cell of interest is illuminated greatly decreases the background fluorescence. (C and D) Fluorescence images of cells labeled with MitoTracker Red (Invitrogen, Eugene, OR). Light from the room is collected by the objective lens, increasing the background in the image. Images were collected and are displayed identically to demonstrate diVerences in intensity.

incident light. The excitation light is reflected 90  by the dichromatic mirror, down the optical axis of the microscope. The light is collected by the objective lens and projected onto the specimen. The fluorophores in the specimen absorb the excitation wavelengths of light and emit longer wavelengths of light. The emission light is collected by the objective, and then must pass back through the dichromatic mirror. The dichromatic mirror is therefore designed to reflect the excitation wavelengths, but transmit the emission wavelengths. The reflection of the excitation wavelengths is not 100%—a small amount of this light passes through the dichromatic mirror. However, the emission light is only 104 to 1010 as intense as the excitation light, so the excitation light that passes through the dichromatic mirror must be blocked from reaching the detector or it will contribute background to

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the image. This light is blocked by the third and final filter, the emission filter (also called the barrier filter). The emission filter can also be used to select more carefully for the emission wavelengths of light expected from the fluorophore. A ‘‘bandpass’’ filter is more selective, allowing through a narrow band of wavelengths. A ‘‘long-pass’’ filter is less selective, allowing through light above a specified wavelength. Choice of band-pass versus long-pass emission filter is an important one. The band-pass has the advantage of selectivity, making it the best choice for specimens labeled with more than one fluorophore. The lack of selectivity of the long-pass filter is beneficial if your specimen is labeled with only one fluorophore and does not have autofluorescence that needs to be filtered out. The long-pass filter will allow through all of the fluorescence from the fluorophore, resulting in the brightest possible image. It is important to accurately match the fluorophore(s) you are imaging with an appropriate filter set (Kinoshita, 2002; Ploem, 1999; Reichman, 2000). Absorption and emission spectra for fluorophores are available online from the company where they were purchased, or can be easily produced with a spectrofluorometer. Transmission spectra for filter sets are also available online from the microscope or filter manufacturer. These spectra should be compared so that the fluorophore is illuminated with wavelengths of light that it maximally absorbs, and so that all or the majority of the emission light coming from the fluorophore passes through the dichromatic mirror and emission filter. There are several websites that oVer useful tools to create overlays of fluorophore and filter spectra (e.g., the ‘‘Curv-o-matic’’ at www.omegafilters.com).

D. Automation of Filter Selection For live-cell imaging of more than one fluorophore, switching between filter sets must be automated. Motorized microscope stands are now available from all of the major microscope manufacturers. These stands include motorized fluorescence filter turrets and shutters that can be controlled by commercially available image acquisition software packages. The speed of movement varies among microscope models and manufacturers, but can take as long as 1 s or more to move between adjacent filter set positions. If rapid switching between wavelengths is required for your experiments, a faster solution is necessary. Motorized filter wheels (Dailey et al., 2006a; Salmon and Waters, 1996; Prior Scientific, Rockland, MA; Sutter Instrument, Novado, CA) can be used to switch between wavelengths at faster speeds; filter wheels switch between adjacent positions on the order of 20 ms. In this setup, excitation filters are removed from the filter cubes and placed in a motorized wheel that is positioned in front of the lamp housing. A multiple band-pass dichromatic mirror is used that reflects multiple excitation wavelengths and transmits multiple emission wavelengths. To select for emission wavelengths, a multiple band-pass emission filter can be used, but these allow more bleed through than single-pass emission filters. Single-pass emission

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filters can be used by adding a second filter wheel in front of the camera. This design is more thoroughly described in Chapter 11 by Salmon et al., this volume. For even faster switching between excitation wavelengths, a monochromator can be used (Dailey et al., 2006a). Monochromators use prisms or diVraction gratings to spatially separate colors of light. The dispersed or diVracted light is then moved mechanically to direct a narrow band of select wavelengths through an exit pinhole or slit. Switching between excitation wavelengths using these devices takes only
1 ms. Multiple band-pass emission filters are then used to select for emission wavelengths. Monochromators are commonly used for live-cell ratio imaging of calcium probes, where rapid switching between excitation wavelengths is critical (Chapter 19 by O’Connor and Silver, this volume). Some experiments call for simultaneous imaging of two or more fluorophores. This can be accomplished by simultaneously exciting the multiple fluorophores, and using a device which splits the emission light into two or four images that are focused side by side onto the same CCD camera chip (Dailey et al., 2006a; Optical Insights, Tuscan, AZ; Hamamatsu Photonics, Bridgewater, NJ). These devices are very useful for fast moving specimens or for live-cell ratio imaging experiments such as Fo¨rster resonance energy transfer (FRET; Chapter 19 by O’Connor and Silver, this volume).

V. Choosing the Best Objective Lens for Your Application In fluorescence microscopy, the objective lens is used for both illumination of the specimen and image formation. It is critical to live-cell imaging applications that an objective lens is carefully selected, with properties that will help to optimize the signal-to-noise ratio of the image. A. NA: Resolution and Brightness The NA of a lens is defined as n(siny), where n is the refractive index of the medium between the specimen and the coverslip (such as air or immersion oil) and y is the half angle of light acceptance (Keller, 2006). NA is arguably the most important property of an objective lens. In fluorescence microscopy, both the resolution of the microscope and the brightness of the image are dependent on the NA of the objective lens. The NA of the objective is marked on the barrel of the lens, just after the magnification (Abramowitz et al., 2002; Keller, 2006). A lens with an NA above the refractive index of air (
1) is designed to be used with an immersion medium of higher refractive index such as immersion oil, water, or glycerol. Use of an immersion medium decreases refraction and total internal reflection at the glass–air interface, resulting in a significantly brighter image (Fig. 2C and D). While immersion media are messy to work with, the increase in signal is well worth the eVort. The resolution of an objective lens can be defined as size of the smallest possible point source in the image, the radius of which is given by 0.61l/NA, where l is the

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wavelength of emission light (Inoue´, 2006). The axial resolution (resolution in ‘‘z’’) of the light microscope is diVerent and significantly worse. The axial resolution of the microscope is defined as (2ln)/(NA)2, where n is the refractive index of the medium the specimen is immersed in. The brightness of an objective lens is equal to 104NA4/M2, where M is the magnification (Abramowitz et al., 2002; Keller, 2006; Fig. 2E and F). High NA lenses are exquisitely sensitive to light. It is therefore important that the room lights be turned oV, especially on an inverted microscope where the objective lens points directly toward ceiling lights (Fig. 3C and D). For very low light level imaging, it can also be beneficial to turn oV bright computer monitors after acquisition has begun, or to position the monitor away from the microscope. B. Magnification If all other properties of the lens remain the same, image intensity decreases with magnification (104NA4/M2; Fig. 2E and F). The lowest possible magnification should therefore be used for fluorescence live-cell imaging. For maximum resolution, suYcient magnification is necessary to ensure that the resolution of the microscope is matched to the size of the detector elements in the camera (Chapter 10 by Spring, this volume; Spring, 2000; Waters, 2006). Sacrificing some resolution by using a high NA, lower magnification lens is one way to increase signal in low light level applications. C. Correction for Aberrations There are several aberrations that can be found in objective lenses (Keller, 2006; Chapter 3 by Ernst Keller and Chapter 18 by Goodwin and Visiting Scientist, this volume), two of which are of particular concern for fluorescence live-cell imaging—chromatic aberration and spherical aberration. Chromatic aberration causes a shift between diVerent wavelengths along the axial axis of the microscope and is, therefore, problematic when imaging more than one fluorophore (North, 2006). Chromatic aberration occurs because diVerent wavelengths of light refract at diVerent angles as they pass through the lens. Spherical aberration can be introduced by the wrong thickness of coverslip or by a change in refractive index between the coverslip and specimen, as was described earlier. Spherical aberration can also be inherent in an objective lens, since light that travels through the periphery of a spherical refractive lens comes into focus at a diVerent plane than light that travels through the center of the lens. Objective lenses with spherical aberration will produce images that are less intense and have worse axial resolution than lens that are free of spherical aberration (Figs. 1A and B and 2A and B). Objective lenses that are corrected for spherical and chromatic aberrations are called achromatic (least corrected), fluorite, or apochromatic (most highly corrected). Most research grade microscopes use fluorite or apochromatic lenses, which are marked Fluor (or Fluar) or Apo, respectively, on the objective lens

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barrel (Abramowitz et al., 2002). Correction for chromatic and spherical aberration is not perfect and the extent of correction varies with individual lenses. The best way to characterize both the resolution capabilities and the extent of aberrations in a particular lens is to collect three-dimensional images of small beads (less than 200 mm in diameter) that fluoresce in multiple wavelengths (available from Invitrogen/Molecular Probes, Eugene, OR; Chapter 17 by Wolf et al., this volume; Hiraoka et al., 1990). These images will reveal the extent of spherical aberration in the lens and any shifts between wavelengths. See exercise 8a in Chapter 17 by Wolf et al., this volume. Objectives are corrected for aberrations by adding additional elements to the lens. Unfortunately, these also decrease the transmission of light through the objective, particularly in the UV range (Keller, 2006). When imaging only one fluorophore, chromatic aberration is not a concern and a less highly corrected Fluor lens may produce a brighter image than an Apo lens. Most of the lenses we use are also marked Plan to indicate correction for an additional aberration, curvature of field (Keller, 2006). Lenses that are not Plan corrected have higher transmission in the UV range and are essential for applications such as ratio imaging of the calcium indicator dye FURA (Chapter 19 by O’Connor and Silver, this volume). D. Phase and DIC Objective Lenses When working on a fluorescence microscope that is also set up for the transmitted light methods phase or diVerential interference contrast (DIC), it is important to understand the eVect of the phase or DIC components on the fluorescence image (Inoue´ and Spring, 1986). Phase objective lenses (usually marked Ph1, Ph2, or Ph3) have a ring in the back focal plane that is necessary for generating the phase eVect (Inoue´ and Spring, 1986). The ring is designed to absorb light and decreases the transmission of light through the objective by as much 30%. Phase objective lenses are therefore not ideal for fluorescence live-cell imaging and should not be used unless the acquisition of phase images during fluorescence time-lapse recording is necessary. DIC objectives (usually marked DIC or Pol) are not modified in a way that aVects light transmission and are therefore a good choice for fluorescence live-cell imaging. However, when a microscope is configured for DIC imaging, there are two additional components in the light path that do aVect the fluorescence image (Chapter 13 by Moomaw, this volume; Inoue´ and Spring, 1986). The analyzer is a piece of polarization film that is located between the objective and the eyepieces, and when placed in the light path decreases transmission by 50%. The Wollaston prism is most often found just after the objective lens in the optical path. It shears each point source of light into two point sources separated by up to a few hundred nanometers (Inoue´ and Spring, 1986). When the Wollaston prism is in the light path, it decreases lateral resolution of the fluorescence image and decreases the

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intensity of each point source of light (and is especially harmful for confocal microscopy). Both the analyzer and Wollaston prism should therefore be removed from the light path when collecting a fluorescence image. If both fluorescence and DIC images are collected during a time-lapse experiment, the Wollaston prism is usually left in place. However, the analyzer can be placed in a motorized emission filter wheel in front of the camera so that its removal from the light path during fluorescence image acquisition can be automated (Chapter 11 by Salmon et al., this volume). Motorization of removal and insertion of the analyzer into the light path is also an option on many motorized microscope stands.

VI. Acquiring Digital Images Over Time A. Cameras Acquiring fluorescence images of live cells is especially challenging because a relatively low number of photons are emitted from the specimen. It is therefore essential that a sensitive detector is used that contributes minimal noise to the image. The CCD cameras commonly used for wide-field fluorescence imaging are covered in detail in Chapter 10 by Spring, Chapter 12 by Rasnik et al., and Chapter 13 by Moomaw, this volume (Spring, 2000; Waters, 2006). This section will review the most important considerations for acquiring high signal-to-noise ratio digital images of live fluorescent cells.

1. Quantum EYciency and Gain Quantum eYciency is the percentage of incident photons that are recorded by the CCD camera (Inoue´ and Spring, 1986; Spring, 2000; Spring and Davidson, 2004). Higher quantum eYciency means that the detector is more sensitive and will therefore require lower exposure times. Lower exposure times mean the specimen is exposed to less light, which helps to prevent photobleaching and phototoxicity. It also means that images can be collected with the higher temporal resolution necessary for capturing fast dynamic processes. The CCD cameras that are most commonly used for fluorescence microscopy have quantum eYciencies in the range of 50–70% (meaning that 30–50% of the photons coming from the specimen are disregarded; Spring and Davidson, 2004), which is tolerable for most live cell work. Back-thinned CCD chips have quantum eYciencies as high at 90% (Spring and Davidson, 2004), but they are also expensive and have fragile detectors. Color cameras should never be used for fluorescence live-cell imaging, since the filters needed to create the color image decrease the quantum eYciency substantially. Graphs of the quantum eYciency of a camera, which varies with the wavelength of light, are usually available on the manufacturer’s website. For very low-intensity fluorescence, amplification of the signal may be necessary. Such amplification is referred to as ‘‘gain.’’ This can be accomplished with an

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intensifier placed in front of a standard CCD chip (Inoue´ and Spring, 1986). While intensifiers oVer very high gain, they have low quantum eYciency. Newer electron multiplication (EM) CCDs (Basden et al., 2003) amplify the signal on the CCD chip, prior to readout (so readout noise is not amplified). EM-CCDs add more noise to an image than a standard CCD chip, so they should only be used when all other options for increasing signal have been exhausted (Chapter 12 by Rasnik et al. and Chapter 13 by Moomaw, this volume).

2. Camera Noise All CCD cameras contribute noise to each pixel in the resulting digital image (Inoue´ and Spring, 1986; Spring, 2000; Waters, 2006). High noise levels can mask weak signals coming from the specimen, eVectively decreasing the sensitivity and dynamic range of the camera. In fluorescence live-cell imaging, the signal is often weak, making it very important to choose a low noise camera. The two main types of camera noise are dark (also called thermal) noise and readout noise. Cooled CCD cameras have negligible dark noise and are therefore preferable for imaging fluorescence. Readout noise is generated as the signal is read from the CCD chip, and is significantly higher than dark noise in cooled CCD cameras. Readout noise increases with readout rate, so faster cameras generally have more readout noise. Noise levels for a camera can be found on specification sheets available from the manufacturer’s website. Noise is given in electrons to make it easy to compare one camera to another, that is for instance, the camera has a DC readout level of 100 electrons with a standard deviation of eight electrons, then eight electrons is the noise. For live-cell fluorescence imaging, readout noise of eight electrons or less is acceptable. [To make the expected noise contribution easier to appreciate, electrons can be converted into gray scale values with the following equation: gray scale value ¼ electrons/(full well capacity/maximum gray scale value).]

3. Determining Exposure Time The optimal fluorescence digital image uses the full dynamic range of the camera without saturating. A maximum gray scale value in the image which is
90% of the maximum gray scale value the camera can produce is ideal. Realistically, this is diYcult if not impossible to achieve with dim live fluorescent specimens. Even with brighter specimens, it may be preferable to underexpose the image in order to maintain cell viability and reduce photobleaching. When an image is underexposed, the maximum gray scale value decreases and gets closer to the noise level of the camera. The noise therefore becomes more noticeable, resulting in a grainy appearance to the image. More importantly, when an image is underexposed, signal noise (explained in Section V.A.4) degrades the accuracy of fluorescence intensity measurements. A common error when determining the best exposure time for a specimen is to look at the image on the monitor to judge image quality. Images from the cooled

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CCD cameras most commonly used for live-cell fluorescence imaging are 12 or 14 bit (212 or 214 gray scale values, respectively). The monitor can display only 256 gray scale values and our eyes can see only about 60. Therefore, to determine image quality, we must use the image acquisition software to measure gray scale values in the image. This can be accomplished with a histogram that displays the range of gray scale values in the image, a utility oVered in most image acquisition software packages. ‘‘Auto-expose’’ functions in image-processing software usually expose the specimen to light for an undesirable amount of time while calculating the optimal exposure time. The optimal exposure time is better determined by guessing an exposure time (which will become easier as you get to know your camera and specimen) and then looking at the gray scale values in the image to determine if the exposure time needs to be adjusted.

4. Signal (aka Shot or Poisson) Noise The intrinsic statistical variation in the detected fluorescence signal, called signal noise, is equal to the square root of the signal. Signal noise is always present and there is nothing that can be done to eliminate it from an image, so its eVect on the accuracy of fluorescence intensity measurements must be considered. The danger of underexposing a digital image is as the maximum gray scale value in the image decreases, signal noise becomes a larger percentage of the measurement, rendering the result less accurate.

5. Camera Binning Camera binning is very useful for fluorescence live-cell imaging. Binning is a function of some CCD cameras and is set in the image acquisition software. When an image is binned, the signal from subarrays of adjacent pixels (i.e., 22 pixels) is pooled together prior to readout. The outcome is a larger pixel that is four times brighter, but that has same readout noise as one single pixel. This results in a significant increase in the signal-to-noise ratio of the image (Fig. 4). The trade-oV is resolution, as the pixels in the image are now eVectively twice the size. However, since detection of weak fluorescence signals is often more important than acquiring the highest possible resolution images, the increase in signal-to-noise is often well worth the decrease in resolution. Binning has the added benefit of decreasing the image file size and transfer time, which is especially useful when faster acquisition rates are required. B. Shutters It is essential that the light source is shuttered between camera exposures to minimize illumination of the specimen. This is most often accomplished with a mechanical shutter (Dailey et al., 2006a; Prior Scientific; Sutter Instrument)

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No binning

2
2 binning

Fig. 4 Camera binning increases the signal of a digital image, without increasing read noise. Some resolution is lost, but this is often a worthwhile sacrifice in exchange for increase in signal-to-noise ratio. The loss of resolution is barely noticeable in these images of fluorescently labeled actin in tissue culture cells, while the increase in signal is dramatic.

controlled with image acquisition software. When choosing image acquisition software, it is important to be sure that the software opens the shutter just before and closes the shutter directly after the camera acquires the image. A delay (on the order of 20 ms) is sometimes needed after the shutter is opened and before the camera acquires the image to ensure vibration caused by the shutter opening has dissipated. However, software that allows the shutter to remain open significantly longer than is necessary to properly expose the image will result in unnecessary photodamage and photobleaching of the specimen. This problem can usually be detected by acquiring an image with a very short exposure time (10 ms). C. Maintaining Focus Maintaining focus during live-cell imaging can be a major source of frustration. Focus changes over time due to gravity acting on the gears within the focus mechanism, changes in temperature, and movement of the specimen. On most inverted microscopes used for live-cell imaging, the nosepiece is moved up and down by the focus mechanism. Making the nosepiece as light as possible by removing objectives that are not being used can help to reduce focus drift. For many microscope models, torque of the focus mechanism can be adjusted so that more force is needed to move the gears. The presence of a focus motor directly coupled to the fine focus mechanism of the microscope also helps to stabilize focus.

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As described earlier, microscope incubators are the best way to minimize focus drift resulting from fluctuations in temperature. After placing a dish into a microscope incubator, or after assembling a localized heated chamber, it should be expected that there will be some focus drift for 10–20 min while the temperature equilibrates. It is also important to maintain as constant an ambient room temperature as is possible. Air conditioning and heating vents should be positioned in the room so that the air does not directly blow onto the microscope. With high NA objective lenses, the smallest shifts in temperature and air currents in the room can result in loss of focus. Focus drift can also occur as a result of a specimen that is mounted in a chamber that ‘‘settles’’ over time. When placing the specimen onto the microscope stage, it is important to make sure that the specimen is held as tightly to the stage as is possible. Use stage clips or tape to hold dishes to the stage and be sure that the stage plate is locked or screwed into place. This is especially important when collecting z-series—if the specimen is at all loose it will move as the focus position is changed, rendering the z-series inaccurate. Specimens that do not naturally adhere to the glass coverslip should be immobilized during imaging to prevent movement out of the focal plane or blurring in the digital image. For example, a common way to immobilize yeast during imaging is to place them in a solidified media containing gelatin. In this design, the cells should be sandwiched between the gelatin and a coverslip so that the light between the specimen and the objective does not need to pass through the gelatin (Maddox, 2000), which would cause spherical aberration. There are a couple of ways to deal with persistent focus drift, both of which require automation of focus (Section VI). One way is to use autofocusing algorithms available with many image acquisition software packages. These can work very well, with a few caveats. The focal plane that the software chooses based on image intensity and sharpness may not be the focal plane you are interested in, and autofocusing can be slow depending on the speed of the algorithm and focus motor. Autofocusing on the fluorescence image is not a good idea, since several images need to be collected in the process resulting in additional photodamage and photobleaching. Instead, autofocusing should be performed on a transmitted light image. This requires mechanical shutters on both the fluorescence and the transmitted light path so that the software can switch between the two modes of microscopy. There are clever new commercially available technological solutions to maintaining focus that give excellent results (Hogan, 2006; Applied Scientific Instrumentation, Eugene, OR; Nikon Instruments, Melville, NY; Olympus America Inc., Melville, NY). In these systems, an infrared laser beam is sent through the back of the objective lens. The beam exits the objective at a high angle such that it totally internally reflects at the interface between the coverslip and the specimen. The light that reflects back into the objective is monitored. As the distance between the objective lens and the coverslip changes, so does the intensity and

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position of the light reflected back into the objective. A focus motor is used to continuously adjust the focus to maintain a single focal plane. These instruments are currently most commonly used in conjunction with TIRF microscopes in which focus is critical, but they work so reliably it is likely that they will soon be found on all microscopes used for live-cell imaging.

VII. nD Imaging With the development of fast and accurate motorized components of the microscope, we can now collect z-series images of multiple fluorophores at multiple stage positions over time—so-called multidimensional or nD imaging. Collecting multiple focal planes is necessary for analyzing specimens that move in three dimensions and for quantitative digital deconvolution (Wallace et al., 2001). Motorized stages can be used to collect images of multiple fields of view over time, greatly increasing the amount of data that can be collected in one experiment. Collecting multiple focal planes at precise intervals (‘‘z-series’’ optical sections or image stacks) requires automation of focus controlled by image acquisition software (Dailey et al., 2006a). This is most often accomplished with a stepper motor that drives the focus mechanism of the microscope. Stepper motors are available built within motorized microscopes, or as peripheral devices that attach to the fine focus mechanism. To increase accuracy, linear encoders can be added which monitor the distance the stepper motor moves and make adjustments as needed. Piezoelectric focus devices are alternatives to stepper motors that oVer higher accuracy and speed. Piezoelectric devices can be attached to the nosepiece to move a single objective up and down along the optical axis of the microscope. Newer piezoelectric designs move a stage plate relative to the objective lens to focus and have the advantage of being easier to use with multiple objective lenses. Piezoelectric devices are much faster and more accurate than traditional stepper motors, but only operate over a limited working range (usually between 50 and 400 mm, depending on the model). Movement between fields of view or between wells in multiwell dishes can be automated with motorized stages controlled with image-processing software (Dailey et al., 2006a). X and Y positions of the stage are memorized in the software, and the stage can be moved back to memorized positions with the click of a mouse or at each time point in a time-lapse experiment. Images from each field of view can then be collated and made into a movie. The motorized stage must return to memorized positions with high accuracy for this to work well, as inaccuracy will cause shifts between images in the movie (this can be corrected for later with software, if there is a stationary object within the field of view that can be used as a reference point). As with focus stepper motors, accuracy in motorized stage movement can be increased with the addition of linear encoders.

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VIII. Verifying Cell Health and Troubleshooting Sick Cells When imaging fluorescence in live cells, two important questions should be addressed. Is the fluorescently labeled protein behaving like the endogenous protein, and is the illuminating light damaging the cells? It must be demonstrated that the localization of the fluorescently labeled protein is the same, over time, as the endogenous protein. If the localization of the protein has not been established, a method of fluorescently labeling the endogenous protein is required. This is most commonly achieved with indirect immunofluorescence labeling of the protein. Whenever possible, fixed time point assays should be used to verify the behavior of the protein over time with live-cell imaging results. For fluorescent protein fusions, it should also be shown that the fusion protein’s expression level and stability is similar to the endogenous protein (Chapter 6 by Straight, this volume; Spector and Goldman, 2005). Optimizing the signal-to-noise ratio of your imaging setup will allow you to choose cells that are expressing lower levels of fluorescent fusion proteins to image. Light damages cells. This is especially true of shorter wavelengths such as the blue light used to image GFP (Chu et al., 2006; Gorgidze et al., 1998; Seko et al., 2001; Sparrow and Cai, 2001). Excitation of fluorophores also generates free radicals that damage cells. We must therefore always be concerned about the health of our live specimens during fluorescence imaging. It is essential that control tests are performed to ensure that the imaging conditions are not changing cell behavior. After an imaging experiment is complete, the cells that were imaged should be compared to cells outside of the imaged field of view. Using transmitted light (phase or DIC), look for the accumulation of vacuoles within cells, fragmented mitochondria, or unusually large numbers of apoptotic cells (for sample images of light-damaged cells see Dailey et al., 2006a). Many of the live-cell imaging experiments we perform are short in duration, but damage to cells may not be obvious until hours later. It is good practice to continue to image cells for a long period of time (on the order of 24 h) after the experiment, collecting transmitted light images every 20–30 min, to look for long-term eVects. Mitotic index and duration of mitosis are also good indicators of cell health. What if the cells do look sick or behave abnormally? If all of the cells in the dish appear aVected, you should check the imaging media and temperature. Ensure that the pH of the media is maintained over the course of the experiment and use a thermometer to measure the temperature of the media on the microscope stage. Try removing any antiphotobleaching reagents (e.g., Oxyrase), as they may be toxic to some cell types. On occasion, coverslips used right out of the packaging can contain contaminants that are harmful to cells. Vigorous cleaning of coverslips prior to plating cells can alleviate this problem (Canman and Salmon, 2000). A next test would be to replace the growth medium in a fresh batch of cells with the imaging media (if it is diVerent than the growth medium), and place the cells on the microscope stage in the heated chamber (if one is being used) or in a tissue

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culture incubator. Do not image the cells, but wait to see if the cells still look healthy after 24 h. If they do not, another imaging media should be tried. If the cells tolerate the imaging media, be sure that the imaging system is optimized as much as is possible for maximum signal and minimum noise, as described throughout this chapter and summarized in Table I. Use a very low level of illumination (by using a low exposure time and/or neutral density filters) to image the cells such that the fluorescent labeling is barely visible. Use camera binning whenever possible, and make sure that there is a heat absorption filter in front of the light source that cuts out infrared light. Slowly increase the exposure time and remove the neutral density filters until you determine the maximum level of light the cells will tolerate. You should then image as far away from that maximum as is possible while still attaining an adequate signal-to-noise ratio for your experiment. Experiment with illumination light level versus exposure time— some cells tolerate short exposure times at high-intensity levels while others do better with long exposure times at lower light levels. If the lowest possible light exposure still cause appreciable damage, and the imaging system has been optimized for signal-to-noise, free radical scavengers such as vitamin C or Trolox may help (Swedlow and Andrews, 2005).

IX. Conclusion Choices such as objective lens and camera determine the signal-to-noise ratio of an imaging system. Optimizing your imaging system to maximize signal and minimize noise is critical for live-cell fluorescence imaging. Imaging with high signal-to-noise ratio will allow detection of low concentrations of fluorescent fusion proteins with illumination conditions that are less likely to damage cells. Automation of an imaging system allows collection of multidimensional data while helping to maintain focus and minimize specimen exposure to light. Under all imaging conditions, maintaining and verifying cell health is essential to the validity of the experimental results. Acknowledgments Many thanks to Dr. Meg Bentley for critical reading of this chapter. Thanks also to Lara Petrak, without whom I would never find the time to write. I am also very grateful to Ted Salmon, Jason Swedlow, David Wolf, Kip Sluder, Ken Spring, John Murray, Tim Mitchison, Clare Waterman-Storer, Conly Rieder, Rich Cole, Butch Moomaw, Paul Goodwin, and Charles Felts for many helpful discussions on specimen preparation, microscopy, and digital imaging.

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