"Confocal Microscopy". In: Current Protocols Essential

Confocal Microscopy David Carter 1

UNIT 9.4

1

Center for Plant Cell Biology, University of California at Riverside, Riverside, California

ABSTRACT This overview covers the two main types of confocal scanner. Point scanners are more flexible and give the best resolution, while camera-based systems can be more sensitive, faster, and more automated. The procedure for setting up and using these instruments is described in sufficient depth for basic operation. Beyond collecting single images, a Z-series, which captures a volume of data, a kinetics series, collecting a sequence of images, and generation of a large montage image are all described. Confocal is an extension of regular fluorescence microscopy, with the ability to handle thicker and more heavily stained specimens and to work more precisely with physiological markers in live samples and organelle stains. Such techniques are highly divergent, depending on specimen type, so they cannot be covered in any detail here, but a general idea will give the user confidence to explore the literature and collect good data on their own systems. C 2013 by John Wiley & Sons, Inc. Curr. Protoc. Essential Lab. Tech. 7:9.4.1-9.4.36. ⃝

Keywords: confocal ! imaging ! laser scanning ! Z-series ! kinetics ! montage

OVERVIEW AND PRINCIPLES A confocal microscope is a high-performance fluorescence microscope that uses scanning optics to see only one very thin focal plane within the sample (Fig. 9.4.1). Everything above and below this optical section is hidden from the detector to give better resolution and a prettier picture. Thick samples can be imaged without physical cutting, and there are a myriad of vital stains available for labeling features of living samples. Readers should familiarize themselves with the microscopy and fluorescence chapters, since these form the foundation of any confocal work.

Types of confocal microscope There are two main types of confocal microscope: high-resolution point-scanning systems and high-sensitivity camera-based systems. Point scanners are more versatile and give sharper images, while camera systems are faster, more sensitive, and easier to automate. Confocal is also seen in a wide range of highly specialized instruments, including endoscopes for looking inside a live animal, ophthalmoscopes for studying the back of the eye, and high-content screening systems that automatically collect vast amounts of data from multi-well plates. Point scanners and PMT detectors The most common type of confocal microscope uses lasers as the light source and photomultiplier tubes (PMTs) as detectors. Lasers are ideal because the parallel light coming out of them can be focused to a precise point, and they emit pure colors which can easily be separated from the returning fluorescence. PMTs have a very wide sensitivity range, capable of handling any brightness from sunlight to starlight. Very dim signals can be massively amplified inside the PMT, depending on what voltage is applied. The area being scanned can be zoomed in to increase magnification, then panned around to exactly frame the most interesting part of the sample. Lasers and detectors can be used Microscopy Current Protocols Essential Laboratory Techniques 9.4.1-9.4.36, October 2013 Published online October 2013 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470089941.et0904s07 C 2013 John Wiley & Sons, Inc. Copyright ⃝

9.4.1 Supplement 7

sample

XY scan

objective lens

dichroic mirror

beam expander

confocal aperture

prism

PMT detector

lasers AOTF

Objective Lens

Figure 9.4.1 The main components of a confocal scanner. Laser light is launched through an acousto-optic tunable filter (AOTF) and an optical fiber, and then bounced off a dichroic mirror. The scanning mirrors sweep the beam through the objective lens. Returning fluorescence light is descanned by the mirrors and returned via a pinhole and spectral filter to the detectors.

simultaneously to look at multi-colored samples, and each image is watched as it forms from top to bottom as each scan progresses. A selection of overlaid laser beams is launched through an acousto-optic tunable filter (AOTF) into an optical fiber for delivery to the scanner. The AOTF is a solid-state device that uses a standing wave of sound as a diffraction grating to bend specific wavelengths so they exactly hit the core of the optical fiber. Up to eight notes can be played on an AOTF at once, and the loudness exactly controls how much light is launched. The output is expanded out and bounced off a dichroic mirror, which reflects only the pure colors of the laser lines and transmits all the other colors. Up to eight different dichroics may be available with the most popular permutations of colors, plus a semi-silvered mirror for reflection imaging. Always make sure the needed colors will bounce off the selected dichroic. The illuminating light goes through a scanning system to move the beam around, and is focused through the objective lens and into the sample. All the light hits one point in the sample at a time. This gives better contrast and higher resolution than conventional fluorescence because only one X-Y position is lit up at a time, so there is no stray light from other locations to blur the image. Fluorescence signal is collected by the objective lens and descanned by the scanning mirror back through the dichroic mirror. Then it is focused through a pinhole before being detected. Light from above or below the plane of focus is either underfocused or overfocused so that it hits the pinhole as a disc instead of a point, and is blocked. A filter wheel may be used to select a range of colors for detection. More versatile spectral systems disperse the colors with a prism or grating, whereupon any exact color range can be picked off for detection.

Confocal Microscopy

Full-field fluorescence and confocal are compared in Figure 9.4.2, which shows a mammalian cell with fluorescently stained vesicles. With full-field, an average view of the whole thickness is seen and it is impossible to tell whether the solid volume of the vesicles are fluorescently stained, or just the membrane. However, confocal shows only

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Current Protocols Essential Laboratory Techniques

Figure 9.4.2 Full field fluorescence (A) and confocal image (B) of a thin mammalian cell. The confocal view clearly shows that the vesicles are membrane labeled only, while in the full-field view they look solid.

one height and clearly shows the vesicles as circles rather than solid disks. Therefore, it is the membrane that is labeled and not the solutes inside those vesicles. Seeing only one very narrow thin layer of the sample is the advantage of confocal, but also the disadvantage. If the image plane is set a little too high or low, the screen is completely black. If the sample did not stain, or the scanner is not correctly configured, again nothing is seen and it is hard to tell the difference between those possibilities. With full-field fluorescence, even when tens of micrometers out of focus, there is something to see, but it is dim and blurred. By moving the focus knob, it either gets sharper and brighter or dimmer and fuzzier, so the direction in which to go can be determined easily.

Parallel detection camera-based systems Rather than looking at just one location in a sample at a time, camera systems collect light in parallel from all over the field of view, to give much faster imaging. A confocal module fits between a camera port on the microscope and a good grayscale CCD camera. The module launches laser light through a disc with spiral patterns of pinholes in it. As the disc spins, points of light from the pinholes sweep across the specimen (Fig. 9.4.3). Returning light is filtered so that only one plane at a time is imaged. Over a thousand pinholes can be viewed at a time, so the amount of light being detected can be orders of magnitude higher than for point scanning. This makes images clearer, or enables them to be collected extremely quickly. Bleach rate at the sample is determined by the local intensity experienced by the fluorophores, so paradoxically even with the higher light throughput, the spinning disc confocal is much gentler on the sample, which can then be imaged for longer, with less photo-damage. The illuminating light is distributed over the entire imaged area, with the pinholes of the disc blocking most of it to generate the optical sections. Yokogawa (http://www.yokogawa.com/) makes laser-based confocal scanners that have two spinning discs instead of one. The first disc is an array of lenses, which focuses most of the light through the second disc containing the pinholes. Returning light is picked off by a dichroic mirror between the two discs and sent to the camera.

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Figure 9.4.3 (A,B) Spinning-disc confocals make fast, high-quality images for kinetic studies with minimal laser damage. Single discs are bright enough to allow direct viewing, while double-disc scanners view against a darker background for finding dimmer signals with a camera. (C) Four 100-msec images were averaged every 2 sec on an EMCCD camera to collect small stacks of images of GFP-labeled microtubules in live Arabidopsis leaf epidermis.

This double-disc format has two major benefits. First, lasers of moderate power (20 to 30 mW per color) are sufficient to image the whole field of view, and second, the camera is looking at returning light from the pinholes against a completely dark background. In single-disc systems, the detection optics have to cope with back-reflected illumination light from the whole face of the disc, except for the 7% or so that consists of open pinholes. This limits the peak sensitivity of the whole system, because even if the emission filters are designed to let through only one in 108 excitation photons, there will still be more stray light in the image. The benefit of having the most sensitive, most expensive cameras is only realized with these dual-disc systems, but single-disc scanners with less expensive cameras are an excellent choice for more routine imaging, and are often bright enough for confocal viewing directly to the eye. Cameras have a higher quantum efficiency than PMTs, and have less internal noise, so images are smoother, can discriminate more levels of intensity (bit depth), and are easier to quantify. Back-thinned CCD cameras boast a quantum efficiency up to 95%, and deeply cooled cameras have extremely low noise characteristics. The camera can easily be the most expensive part of the whole system, so sometimes a front-illuminated progressive scan camera with relatively modest cooling is sufficient. Parallel detection across the whole field of view enables automation of focus and brightness adjustments. The field of view is fixed, so zooming in on a small area is not possible. Camera systems are fast, but there is nothing to see while the camera is collecting light, so patience is needed when finding initial focus. Wait for the image to refresh, then quickly move the sample position a small distance until the best location is found. If multiple colors are used, they either have to be collected in sequence, or a second camera with precisely matched pixel positions must be set up for each additional color. If the camera has enough pixels, an image-splitting eyepiece is a third option (for example the DV2 from Photometrics). This sends greener light to one side of the camera, and red to the other, to make two half-sized images which are overlaid with software to make a two-color image. Confocal Microscopy

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Any defects in the optical system, such as brightness variation in the light source, internal reflections, or dust in the detection light path are very apparent in a camera system, but can sometimes be hidden with software correction. A slightly frosted plate placed between the specimen and the condenser scrambles optical artifacts outside the plane of focus, and improves the transmission image, where artifacts are most intrusive.

SAMPLE PREPARATION Mounting the sample Confocal microscopes are quite forgiving of sample type, so almost anything that can be made fluorescent can be imaged. If the sample is too thick to be seen by conventional microscopy, navigation becomes more difficult, and first finding the region of interest is often the most challenging step of the whole experiment. Each field of study has its own tricks for optimal staining procedures and sample preparation (Matsumoto, 2002), but this general overview helps describe what is possible, and how it all works. On an upright microscope, the stage can be dropped to accommodate bulky samples, or a ceramic dipping objective can be lowered into a dish of culture medium to look directly at the sample without even needing a coverglass. With an inverted microscope, the sample can be placed on a coverglass-bottomed dish to be looked at from below. Alternatively, a slide and large coverslip can be used, with the slide flipped so that the coverslip faces down. Figure 9.4.4 shows the marking of slides for easy reference. Multiwell plates with coverglass or plastic-membrane bottoms are also an option for handling many samples at once. Time saved and ease of use soon compensate for the higher cost of consumables, but

Figure 9.4.4 (A) Delicate samples can be protected from compression with coverslip bridges, which are held in place by surface tension. A sharpie mark or diamond scribe helps find the plane of the specimen. (B) A slide and no. 1.5 coverslip are sufficient for most samples. Use LabTek or Willco dishes for wet samples if a coverglass base is needed. Inverted microscopes can also handle multiwell plates, including membrane-bottomed clear plates, coverslip 96- and 384-well plates, or custom designs such as this 48-well gasket set inside a universal plate lid.

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a plain slide and a number 1.5 coverglass are usually adequate. High-numerical-aperture lenses are needed for good optical sectioning, so polystyrene flasks, dishes, or plates that have 1-mm-thick walls are not suitable for confocal work.

Fixed samples Samples can be live or fixed. Fixed samples can be sectioned, stained and mounted in the same way as for fluorescence microscopy, but much thicker sections can be prepared, and higher levels of staining can be tolerated. Clearing of thicker samples requires more care, so look for protocols appropriate for your model organism, and try different variants to find the one that gives the best clarity and preservation of fluorophore (Hama et al., 2011). Mounting media Samples can be quickly mounted in a mixture of up to 75% glycerol:buffer, where the glycerol reduces the cloudiness of the sample and the buffer maintains the fluorophores at a stable pH. Do not allow stained sample to dry out, since exposure to air can oxidize the dyes. Make a reasonable effort to keep them away from bright light, for example by wrapping clear plastic slide boxes in aluminum foil. Keeping them cool in a refrigerator will slow any chemical degradation, but will also introduce condensation when the samples are returned to a warm, humid room. Water-compatible commercial mountants give similar clearing, but contain a setting agent for more durable preparations. Products are available that boast better preservation and slower bleach rate than lab recipes, such as Vectashield (Vector Labs) and Prolong Gold (Life Technologies), which are worth trying for difficult samples, or when the sample needs to be preserved for many weeks instead of imaged quickly and discarded. Mountants that set hard protect the sample from accidental shearing force when cleaning the coverglass. Those that do not set should be handled with more care, and can be sealed with a ring of nail varnish or cyanoacrylate adhesive to achieve the same end. Most fixed samples are scanned within a week of preparation, then discarded and replaced with new preparations. Many commercial mounting media and lab recipes contain free-radical scavengers like n-propyl-gallate or 1, 4-diazobicyclo-2,2,2-octane (DABCO; Air Products & Chemicals, Inc., http://www.airproducts.com/), which protect the fluorophores from oxidation, but may slightly increase background signal. Newer fluorescent dyes are much more robust, so these additives may still be helpful but are not always needed. Mounting media are listed in Table 9.4.1.

Live samples Live samples are observed in a minimal physiological buffer, or distilled water in the case of plant tissues. Some microscopes have incubators to keep animal cells warm and at high carbon dioxide levels. These either cover the entire microscope, or are small inserts that fit on the stage. The objective lens will draw heat from the sample if it is not inside the enclosure, so use a heating collar to warm it and prevent this intrusion (e.g., Bioptechs, http://www.bioptechs.com/).

Confocal Microscopy

Mounting methods Pieces of tissue can be mounted in buffer under a 24 × 50–mm no. 1.5 coverglass. If there is any air under the coverglass, surface tension will hold it in place even on an inverted stage, but applying too much liquid will cause the coverslip to float and let the

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Table 9.4.1 Mounting Media

Mountant

Refractive index

Sets

Ethanol

Notes

Water/agar/buffer

1.33

No

No

Can be live

75% glycerol

1.44

No

No

Can be thick

DPX/DEPEX

1.51

Sets

Yes

Traditional histology

Vectashield

1.51

No

No

DAPI optional

ShurMountAq

1.44

Sets

No (Yes)

(Toluene version)

Prolong Gold

1.44

Sets

No

DAPI optional, 24 hr set

BABB

1.44

No

Yes

Very thick; shrinkage; quenches GFP

Scale (4 M urea)

1.38

No

No

Extremely thick tissue

sample move. To protect from compression under the objective, a bridge can be made with smaller coverslips or pieces cut with a diamond scribe. Cultured plant cells, pollen, and protoplasts are more delicate and can be made to tumble by the focusing movements of the objective. Use a smaller bridge to give the span more support, and reduce its tendency to flex. Touching the carrying liquid with a Kimwipe draws off any excess and prevents the coverslips from floating. Water objectives should be used with watery samples such as plant tissue for optimal imaging, and glycerol is a better match for animal tissues. If a water lens is moved to the edge of the coverslip, the immersion fluid may mix suddenly with the sample water and cause a sudden flow of fluid in the sample. The viscosity of immersion oil or glycerol will tend to move a small coverglass as the stage or focus is adjusted, so it will need to be secured more firmly. If cyanoacrylate (Krazy Glue) is used to seal coverglasses, it must be completely set before use or the vapors can fog nearby optics. A quick dip in water will ensure complete polymerization. If nail varnish is used, the acetone solvent needs to evaporate; it will not set in a sealed container.

TYPES OF FLUOROPHORE Any fluorescent staining techniques for conventional microscopy can be applied equally well to confocal. However, the right colors of laser must be available for a particular stain to work well. For example, Texas Red works very well with an arc lamp, but the less common orange or yellow lasers (561 nm to 594 nm) are needed for good confocal work. Many thousands of dyes can be used for confocal imaging, but there are a few common colors that closely resemble the fluorescence spectra of some of the earliest fluorescent dyes. Fluorescein isothiocyanate (FITC) excites with the 488-nm blue laser line and emits green. All dyes with similar spectra are imaged in the “FITC channel” and not listed separately in setup software. The many chemical variants of rhodamine cover a wider range, but tetramethyl rhodamine isothiocyanate or TRITC has given its name to an optical layout for collecting green-to-orange fluorescent signal. Cy5 was the first popular red to deep-red dye, and Texas Red the most popular orange to red dye. A UV layout is commonly referred to as the DAPI channel, after the nuclear stain diamidino2-phenylindole. Quantum dots or nanoparticles, which are electron dense and can thus be seen with a transmission electron microscope (TEM), are fashionable alternatives to fluorescent dyes.

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A quantum dot is excited by any color of shorter wavelength than its narrow emission band, so one laser can excite many shades of dot at once (Deerinck, 2008).

Green fluorescent proteins Molecular biology has revolutionized fluorescence microscopy by making available perfectly stringent fluorophores. Rather than having to raise an antibody against a purified protein and immunostain tissue sections, it is possible to genetically modify the whole organism so that every time the protein of interest is made by a cell, it adds its own fluorescent tag to it (Avila et al., 2003). Extensive collections of recombinant model organisms are available, with a range of fluorescent protein colors labeling a wide range of marker genes to highlight specific organelles or signal proteins. GFPs work in cultured cells too, but poorly expressing cells can replicate faster than their brighter siblings, leading to a steady loss of label intensity over time. Observation is therefore best shortly after transformation. Variable expression also happens in organisms, so check a few nematodes or Arabidopsis seedlings before settling on an organism to study in detail. The sequence for GFP protein tags can be spliced into the C-terminal or N-terminal end of the protein of interest, with an aggressive promoter in front of both, to amplify the amount made. This gives less control of the final brightness than is possible with fluorophores, but there is no solvent or chemical to cause toxic effects. If too much tag is made, the localization may be affected by the overabundance of the modified protein. Using the native promoter gives a more accurate picture of the localization and amount of the protein of interest, but fluorescence may be too dim to see easily. For efficient production of the protein tag, the codons need to work well in the species being studied, so there is often a lag between a fluorescent protein becoming popular in one model (e.g., mammals) and being ported over to other model species (e.g., Arabidopsis). The most popular fluorescent proteins are cyan (CFP; 442 nm or 458 nm excitation, 480 emission), green (GFP; 488 excitation, 515 emission), and yellow (YFP or citrine; 514 excitation, 530 emission), with a protein from a different species, dsRed, giving a rhodamine-like color (543 nm excitation, 560 nm emission). Double protein constructs have been made where the proximity of the two halves is changed by a physiological property like calcium level, to cause a color change (Chameleon). Half-protein constructs can show if two separate proteins come close enough together in vivo for the two halves to snap together and produce a YFP signal by biomolecular fluorescence complementation (BiFC, Kwaaitaal et al., 2010). There are switchable GFPs that can be switched on or changed in color by appropriate light stimulation. This is a very active field of development, with new versions gaining popularity every year.

Organelle stains Fluorescent dyes that readily enter cells and target just one organelle are invaluable for live work on animal models (Fig. 9.4.5). Loading may rely on esterase activity, where a less polar dye precursor is loaded, then the acetate groups are cleaved to release a polar dye which is concentrated and trapped inside the cell. Enzyme activity also exists in cell walls, so dye precursors get stranded on the outside of plant plasma membranes, making this approach less useful in plants or fungi, but still possible in protoplasts.

Confocal Microscopy

Rhodamine 123 is concentrated by the very high potential difference across live mitochondria, and a range of Mitotracker dyes of different color are also available, some of which can then be fixed in place by formaldehyde fixation. BODIPY ceramide is a fluorescent membrane lipid that concentrates so much in Golgi that it changes color and fluoresces red instead of green. Nuclei are stained with many dyes including DAPI, Hoechst 33342 which quickly penetrates live cells, and propidium iodide, which does

9.4.8 Supplement 7


A

B

GFP

RH414 RH414

hexyl rhodamine

DAPI

phalloidin

DAPI TO-PRO3

FM4-64

Y

RH414

PI

immunostaining

ER Tracker CFDA

mitotracker

rhodamine 123

CF

WGA BODIPY ceramide

Nile Red

FISH WGA

Figure 9.4.5 Examples of popular stains for live (A) and fixed (B) cells. Some stains like DAPI and hexyl rhodamine work well on both. Some rely on the live physiology for loading, such as Rhodamine 123 and CFDA. Others need more open-access post-fixation, such as immunostaining, FISH, phalloidin and propidium iodide (PI).

not, and thus is used as a cell-death indicator. Carboxyfluorescein diacetate (CFDA) is cleaved inside live cells to render them green, and makes a good partner to PI for livedead assays. TO-PRO-3 has been used with great success as a deep-red DAPI substitute that does not require a UV laser, but it bleaches quickly under 633 nm excitation. Rhodamine B hexyl ester (or hexyl rhodamine) is a nonfluorescent TRITC-like dye that becomes brightly fluorescent when dissolved in any organelle membrane. Most hexyl rhodamine is in the aqueous nonfluorescent phase which does not bleach and which rapidly interchanges with the membrane labeling, so preventing that phase from bleaching significantly either. Fluorescently labeled lectins, like wheat germ agglutinin (WGA), bind to sugar residues on glycoproteins in Golgi, plasma membranes, and nuclear envelopes to give similar results to immunostaining, but in a single stain/rinse cycle. Fluorescent in situ hybridization (FISH) targets specific nucleic acid sequences in fixed cells. Most dyes are soluble in dimethylsulfoxide (DMSO) and can be kept in the freezer as 1 mg/ml stock. Break the purchased dye into aliquots so that most of it is not frozen and thawed or opened and closed too often. 5 µl of 1 mg/ml stock added to a ml of buffer gives a 5 µg/ml staining solution.

LASERS The workhorse of confocal microscopy remains the argon ion laser. This is a relatively inexpensive, high-power laser which provides the most useful line for fluorescent imaging, the 488-nm fluorescein line. It also delivers 514-nm light, which is ideal for exciting yellow fluorescent protein, and 458 nm, which is good for cyan fluorescent protein. Two relatively inexpensive helium-neon lasers, the 633-nm HeNe (as seen in supermarket checkout lines), which works for deep red dyes like Cy5, and the relatively dim, 543-nm green HeNe or GreNe laser, which is a good color for rhodamine-type dyes.

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A violet solid-state 405-nm laser enables DAPI staining of DNA to be visualized, and is used to switch on photoactivatable GFPs or toggle photoconvertable proteins between colors (Ando et al., 2002; Rizzo et al., 2010) for pulse-chase experiments. For true UV excitation, a whole separate light path is needed within the scanner, because UV focuses to a different height below the objective, and corrective optics are needed to precompensate for this error. The krypton ion laser provides 561-nm and 647-nm yellow and red light, and used to be commonly paired with the visible argon. However, it has a shorter lifespan and is noisy compared to more expensive solid-state alternatives. It will also heat up a room quickly if its cooling system is not vented through a wall or ceiling exhaust.

SAFETY CONSIDERATIONS Under normal use, the laser is completely contained within the system and offers no hazard to the user. It does of course exit through the objective lens, which is pressed against a sample at a safe distance from the eye of the user. The high-numerical-aperture objective lenses necessary for confocal imaging ensure that the light is diverging rapidly as soon as it leaves the lens, so after a few centimeters of travel, the light has dissipated to very safe levels. Some confocal systems are also equipped with very powerful pulsed infrared lasers for multiphoton microscopy. The beam is invisible, but still dangerous to the human eye. Power levels are high enough to burn skin and present a fire hazard if allowed to play on a flammable surface. As with confocal, the beam is fully contained in normal operation, but the output should be treated with respect, and never be allowed to escape the system, for example through an unpopulated objective port in the nosepiece turret. Many fluorescent microscopes have amber screens to protect the user’s eye from being dazzled by the much higher intensity of the conventional fluorescence light source. These are more for the comfort of the user than for safety. Our irises respond to the brightest feature in our field of view, which may be the point of illumination, or the glow from the monitor. The human eye takes 10 min in a darkened room to gain full sensitivity, and on a confocal system with its computer monitors and bright illumination, such levels of sensitivity are never achieved in practice. This can make the first finding of a sample very challenging, even though the scanner itself has plenty of sensitivity once it has something to look at. A greater danger comes from the slide, coverglass, and sample itself. Fluorophores are potentially toxic, and are mostly supplied dissolved in dimethyl sulfoxide, which will carry its solutes through exposed skin, and leave a sulfur taste on the tongue. All DNA stains are possible mutagens. Quantum dots may be made from heavy metal salts, and animal materials are potential biohazards. Even genetically modified plant material must be disposed of carefully to comply with legal regulations. A medical-grade sharps box that is autoclaved before disposal is suitable for controlling this waste stream. It is very rare for a mercury arc lamp to burst in operation, but hot mercury vapor is to be avoided. If it happens, switch off, then leave the room for 30 min while the lamp cools down. Major lab supply companies sell small mercury cleanup kits.

OBJECTIVE LENSES

Confocal Microscopy

Confocal is an expensive imaging methodology that requires the very best quality objective lenses (Fig. 9.4.6). Lenses are either dry or immersion. Immersion lenses are designed for oil, water, or glycerol to be sandwiched between the top of the slide and the front of the objective lens. These fluids increase the numerical aperture, or NA,

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Figure 9.4.6 A good set of confocal objective lenses. New designs are improving working distance, field of view, color correction, resolution, and brightness. The water and silicone objectives have collars to correct for coverslip thickness and sample density. The 30×/1.05 NA silicone objective has a working distance of 800 µm, with a moat to protect against drips entering the microscope. It even has a second scale for optimization at 37◦ C.

which is a measure of the angle of collection of the lens. NA = nsinθ, where n is the refractive index of the immersion fluid (or air) and θ is the half-angle of collection at the objective. Some immersion lenses have a correction collar that allows them to be set for more than one type of immersion fluid (e.g., glycerol or water) or to be adjusted for the thickness of the coverglass or even the imaging temperature. The correction collar can also correct for spherical aberration, where the incoming light is focused to a line instead of a spherical dot, preventing a rapid loss of both brightness and resolution as the scan goes deeper into the sample. Always use a no. 1.5 thickness coverglass (170 µm thick) and have the collar correctly set. These collars affect the focal plane, so to adjust them by eye, move the collar and the focus knob simultaneously to keep the image as sharp as possible while searching for the best value. Oil objectives do not need to correct for glass thickness because the refractive index of the oil is identical to that of the coverglass, such that the top surface of the coverglass optically disappears. Oil lenses do sometimes have collars, but only to restrict the NA of the lens for use in dark-field applications, to prevent illumination light from being seen directly. Leave them fully open. Effective numerical aperture is the most important property of lens quality, and a high NA is essential for confocal imaging. NA usually increases with magnification, but the scanner’s unlimited ability to zoom in means that higher-magnification lenses merely have a more restricted field of view. A 40×/1.2 NA water lens for example outperforms its 63×/1.2-W sibling in all respects, but may be considerably more expensive. An exciting new trend in microscopy is the development of ultrahigh-performance objective lenses with extremely high NAs, long working distances, and low magnification for unprecedented light collection efficiency. Just one such lens on an upright multiphoton microscope may replace the entire turret, and afford the microscopist access to a very large volume of the sample to pan and zoom around in. Resolution in depth is always worse than X-Y resolution. It is easier to see between two particles that are side by side than if they are sitting on top of each other. When imaging

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Table 9.4.2 Objective Lenses

Resolution Working distance

Field of view

x-y

z

Air

3100

1275

0.8

9.5

20×/0.75

Air

600

630

0.4

2.4

30×/1.05

Silicone oil

800

425

0.3

1.6

40×/1.3

Oil

180

320

0.2

1.1

60×/1.42

Oil

150

210

0.2

0.8

60×/1.2

Water

240

210

0.2

1.1

Lens

Fluid

10×/0.4

deeper into the specimen, it will get worse still, because now all the laser light has to fight through a chaotic sample and the fluorescence light has to fight its way back. Typically, most tissues can be imaged to about 50 µm in depth for confocal, but this is very sample dependent, and depths of over 2,000 µm have been achieved in perfectly cleared and stained samples (Hama et al., 2011). Objectives are listed in Table 9.4.2.

PROTOCOLS Basic Protocol 1: Use of a Confocal Microscope Basic use of a confocal microscope involves scheduling the use of the instrument (especially if using one in a core facility), turning on the microscope, lasers, and electronics, and allowing for proper warm-up and calibration. Once a region of interest on the slide is found by simple ocular viewing, the scanning optics are set to optimally collect images. Slow, careful collection of high-quality data, with subsequent data removal and shut-down, completes the procedure. The goal is to collect sets of images as described in Basic Protocols 5, 6, and 7: Z-series (collecting a stack of images at slightly different depth), T-Series (collecting the same data repeatedly, to track changes over time), photobleaching or photoconversion (using high-power light to change the fluorophores in specific small regions, as part of a T-series), and montage (collecting a series of adjacent regions to stitch together a larger image).

Reserving time Instruments in a private lab are available whenever needed, but core-facility instruments get very busy and need to be booked by the hour in advance. (for an example of a typical scheduling system, go to http://faces.ccrc.uga.edu). If there is heavy demand, fees may be higher at peak times, or users may have allocation limits, such as a rolling maximum of 10 hr booked. Once some of those reserved hours have passed, they can be reused to reserve more future hours, so long as the maximum does not exceed the personal limit. This is a popular and flexible system, which tends to space bookings evenly, and prevents one user from monopolizing the system.

Confocal Microscopy

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Turning on the microscope In a busy core facility, turning on and shutting down the microscope are frequently omitted during the day as users take over from each other, and it is only necessary to change to the lasers that are needed and clean the objective lenses. If time is limited, one should show up early to give the previous user some time to complete the last scan and remove the data. Current Protocols Essential Laboratory Techniques

1. Turn on the microscope body, so its internal processor is ready to communicate. 2. Turn on the system electronics and wait for the firmware to fully initialize. 3. Start the software last, so all electronic components are ready to communicate. 4. Arc lamps now have good shielding, so they no longer have to be started first and shut down last. They should be run long enough to fully warm up (30 min), or the arc will drift and cause flickering 5. Turn on all needed lasers. Solid-state and HeNe lasers use a simple key to switch them on. Argon and krypton lasers have a main power button, then Off, On, and Start key positions. Like starting a car, turn the key from Off to Start, then allow it to spring back to the On position for normal lasing. Do not use Standby mode for imaging, as intensity is too unsteady when the laser is running on minimum power. 6. Most lasers can be powered down immediately when done, but argon and krypton have to be left in the Off position for at least 5 min while the cooling fan removes the intense heat from the plasma tube, before the power switch is shut off. Failure to do so will shorten the life of the laser. 7. Some systems have integrated software control of the lasers, so start up and shutdown are handled automatically.

Ocular viewing Biological samples are inherently complex, so finding the most interesting parts to look at is a challenge. Even finding the focal plane of the specimen can be a chore, especially if there are a lot of users with a wide range of different specimen types. Anything that can be done to make the initial find easier is well worth the effort. 8. Start by looking at the sample with a low-magnification dry objective, find the best plane of focus, and center on a good region of interest. Try to prepare samples with a higher density of features of interest. Locate them more precisely under the coverslip for an easier find. A scratch with a diamond scribe, or a small cross made with an indelible marker, can help identify the surface nearest to the sample. The top of the slide, the bottom of the coverslip, and the top of the coverslip can otherwise be confused. If no reference mark has been placed, use the edge of the coverslip as a reference. A Pap pen or grease pencil can limit the flow of staining reagents to a small circle, then act as a reference feature in the finished slide. As a last resort, while looking directly at the sample, traverse across the width of the coverslip, then down a field of view, then back across, to completely scan the sample area until a flash is seen on the edges of the coverslip. Slight cloudiness of the sample scatters some light perpendicular to the slide, which is funneled to the very edge of the coverslip to light it up in a full-field flash.

9. Look down the eyepieces and adjust focus to make the fluorescence brighter, then sharper as it comes into focus. The human eye is exquisitely sensitive to movement, so gently rocking the stage from side to side can help in spotting exceedingly dim samples. It takes 10 min in a completely dark room for the eye to reach full sensitivity, so dimming the lights and shading the eyes from bright task lights and monitors will also aid in detection. Counter-staining with DAPI is useful even if not required for the study, because DAPI is a bright and robust stain that lights up all nuclei and makes finding the sample easier. Some commercial mounting media (e.g., Vectashield) are available with DAPI mixed in, so that DNA staining is ensured at the final step of sample preparation.

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10. Switch to white light, DIC, or dark-field to locate the sample. Confocals do not have phase contrast because phase objectives have reduced light transmission. Chlorophyll is an excellent fluorophore that absorbs at nearly all wavelengths and emits deep red fluorescence, which can obscure GFP signal in green plant tissue and make selection of a good region of interest difficult. If there is a filter slot between fluorescence cubes and eyepieces, a ∼630 short-pass filter can be inserted to hide the chlorophyll glare, then pulled out of the way for scanning.

11. Lower the stage, add appropriate immersion fluid, and set the higher-magnification lens in place. The stage will rise to the chosen focal plane without risk to the objective. Re-center the region of interest if necessary, then switch to scanning mode. If a motorized stage is used, survey the sample at low power and mark regions of interest in the software. Switch to high power and jump between locations, to avoid a great many lens changes. If lost, switch back to ocular viewing and low power, to re-find a good region. By starting with a long working distance lens, the risk of bumping into the sample is minimized, but once the optimal height has been found, similar samples can be swapped in at high power.

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Figure 9.4.7 Simple user interface (beam path interface from Leica SP2) for controlling the optics. (1) Double click on a factory preset. (2) Use the laser brightness slider panel to adjust intensity. (3) The right choice of dichroic mirror launches needed laser lines towards the sample. (4) An exact spectral range is defined by moving windows, which can be dragged to different colors and resized. (5) Each active channel has a check mark, a display color, and a gray trapezoid showing which range of color it is detecting. (6) Visible light going all the way through the sample is collected by an additional non-confocal transmission detector. (7) User-defined presets can be saved, and any sequence of optical layouts can be listed for sequential scanning.

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Setting up the optics Pick from a list of preset optical configurations to correctly set up a workable imaging layout (Fig. 9.4.7). If a modification is made, it can be saved as a user-defined layout for future use. Alternatively, each image is saved with the information of how it was collected, and this can be accessed and reused later. Each dye in the sample needs to be excited by the best available laser, which has to reflect off the delivery dichroic in order to reach the sample. Not every permutation will be there. For example, the 488- and 514-nm laser lines are so close together that only one can be used at a time. A 70/30 neutral density mirror works for unsupported combinations, and for reflection scanning. The laser can be turned brighter to compensate for only 30% reflection, and 70% the transmission is adequate for normal signal strengths. Backreflected laser light can be problematic though, since the lasers are being used at higher power, and more is bouncing back towards the detectors. Stray laser light may appear as a diffuse ring pattern, especially when imaging close to the coverglass. On the detection side, the ideal pinhole aperture is one Airy unit (UNIT 9.1) in diameter, to collect the first diffraction fringe of each point of light. Then select a detector with the right filter in front of it for collecting the emission peak of the fluorophore. Fluorescein-like dyes are imaged through a 530/30 BP (band-pass) filter, which lets through a 30-nm band of wavelengths, centered on 530-nm green. If no other dyes are present, use a 505 LP (Long Pass) filter, which lets through all colors longer than 505 nm, including the whole 530/30 band, and so gives a brighter signal. The emission filter must block all laser colors with a stringency of at least 107 , or backscattered laser light will show up alongside the relatively very dim fluorescence signal. Many systems now have either a prism or a diffraction grating to spread the rainbow of emitted light and pick off an exact range for each channel of interest. This removes the limitation of having a filter wheel with only eight choices in front of each detector, since any wavelength range can now be specified. The detection window still needs to be at least 5 nm away from the laser lines on either side.

Basic Protocol 2: Raster Scanning A point scanner illuminates only one point in the sample at a time. To generate an image, the scan mirrors sweep along a line, then laser delivery is turned off while the mirrors swing back to the beginning and a second line is swept in a raster pattern. This continues until a 512 × 512–pixel preview image is generated. Then the pattern repeats. Each line is broken into a series of time-resolved chunks that are quantified then rendered on the screen as colored squares or pixels (Fig. 9.4.8). A microscope equipped with a fast Z-drive can just as easily raster-scan in Z-depth as in Y, where each subsequent sweep is recorded slightly higher up inside the sample. This optically cuts the sample to reveal a profile through the side of the sample. To do this, take a preview X-Y image in the usual way, then select X-Z profile mode and draw the line along which the profile will be collected. Each channel of data can be displayed in any of a wide range of color palettes, including pure red, green, blue, magenta, cyan, yellow, or grayscale. There are also false color palettes that use the whole rainbow to exaggerate more subtle brightness differences within a single channel, and are often used for physiology studies. If there are two channels of data, the shorter-wavelength dye is best shown in green and the longer one in red. Magenta and green are another popular pair, which looks white instead of yellow where both channels are equally bright and works well for color-blind users. If a third color is needed, blue gives unique colors for any mix with red and green, but the human

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Figure 9.4.8 Raster scanning. The laser is swept along a series of lines, turning off at the end of each. These lines are quantified as a series of time-resolved fragments, which are rendered on the screen as colored squares according to brightness. Get higher resolution by scanning more lines and breaking them into smaller pieces. Reduce signal noise by rescanning each line many times and reporting the mean.

eye is not very sensitive to blue. Instead, cyan is typically substituted, and the channel with the brightest or most morphologically distinct structure is shown in this color. The most useful palette shows the whole image in grayscale or a pretty glow orange palette, then starkly highlights saturated and zero-value pixels with a bright prime color. This makes it easy to optimize the sensitivity of the detector so that the maximum amount of information is present in each image. While preview scanning, increase the gain of each channel in turn until the saturation color is seen, than back down until only a few pixels are saturated. At the dim end, an image looks better without any loss of data when the darkest regions are as close to black as possible. This is done with an offset control. Lower the offset until the dimmest part of the image starts to change to zero, then raise it slightly to avoid losing any data in the shadows. Images can be viewed as a single overlay of all data, or as a set of panels, one for each channel and one for the overlay. Use separate panels to set up brightnesses and a large overlay to confirm final quality. Single channels can be fluttered on and off to see if any signal from one channel is hiding under that from the others. A scale bar can be added to the image, but is rarely in a suitable format for a journal figure. Instead, verify the size of the whole image, or the size of each pixel, then use this information to draw a solid bar scale bar on the finished figure.

Mark and revisit with motorized stage Inverted systems with motorized stages can screen thin-bottomed 96- or 384-well plates. Use the Navigate mode to visit different wells and preview the contents. Then use the Select mode to choose which wells are of interest for automatic scanning with a macro program (Fig. 9.4.9). On a regular slide, the sample can be surveyed with a low-power lens to mark locations of interest, before switching to high power and auto-visiting each flagged position. Dry or water objectives are best for this, since oil and glycerol stick to the glass and get left behind.

Confocal Microscopy

Zoom, pan, and crop While preview scanning, the scanned area can be compressed as much as necessary to zoom in, then the scanned area can be nudged around the field of view to neatly frame the area of interest. Alternatively, place a target square over a finished image to exactly position the next scan (Fig. 9.4.10). The software may allow unlimited zoom, but be aware that the lens itself has limited resolution, so beyond a few-fold magnification, the scanned area is shrinking without revealing more detail.

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Figure 9.4.9 Plate Map dialog from the Pathway HT. Multiwell plates can be screened on an inverted confocal. Use Select mode to highlight the populated wells, and Navigate to preview them manually. A macro program can auto-visit each well once to screen the whole plate, or repeatedly with autofocus to generate very long time series experiments.

Figure 9.4.10 Screen shots from the Zeiss 510. Navigation controls within a field of view. The swept area can be compressed to magnify, and scan mirrors can sweep at a diagonal to frame the image at a different angle. Alternatively, A target can be placed directly on a preview image (A), resized and rotated (B). The next scan will exactly image the enclosed square (C). Microscopy


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Optimizing channels 1. Begin a continuous preview scan at 512 × 512 pixels and about 1 frame per second.

2. Start with the easiest, strongest channel and adjust the gain until the brightest features of interest just begin to saturate. 3. Use a range indicator palette that clearly shows the saturated and zero-value pixels. Only the leading edge of the raster scan will show any changes in setting, so in a sparse field of view, wait for a whole scan to complete before settling on a final gain value. 4. For a multi-channel image, adjust the brightest channel for full range signal. 5. Check the focus to verify that the best depth within the specimen is in view. 6. Adjust the other channels to bring them all on scale. 7. Laser light going all the way through the sample can be collected by a transmission detector and displayed as a grayscale bright-field or differential interference contrast (DIC or Nomarski) image. This transmission image uses any of the visible lasers but is not confocal, since there is no filtering pinhole. 8. UV is not transmitted well by condenser optics, so if transmission is needed in combination with DAPI, add a little red laser light. 9. It is best not to optimize the transmission channel, but leave it a dull gray so it does not detract from the more important fluorescent confocal data when all colors are overlaid.

10. PMTs leak a small amount of current even when there is no light present, so adjust the offset to bring the darkest parts of the image close to zero. 11. Offset is largely a property of the detector circuit, so does not vary much from one region to another, or even from one lens to another. Once the offset is chosen, most adjustment will be just the PMT gains. 12. Move the focus to confirm that the depth being scanned is optimal, then continue with the other channels until all colors are nicely balanced. This simple procedure is the essential skill for setting up any point-scanning confocal. Different brands have their adjustment controls in different places, and offer different range indicator palettes, but the procedure is identical. With practice, you can quickly find saturation then back off, and find zero offset then back off using a suitable palate (Fig. 9.4.11). Auto-adjust controls may be available too, but do not work perfectly since the gain range is so large and only one point is in view at a time. If there are highlights in the image that are not of interest, allowing them to saturate makes the rest of the image brighter. However, large areas of saturation look fake in the final image, and will deplete the detector so that it is less sensitive for fewer pixels after crossing this area. Images look better against a very dark, almost black background, so setting the offset carefully not only increases the amount of data by using more gray levels, but also gives a more visually attractive result.

Confocal Microscopy

For very dim, poorly stained channels, the PMT settings may reach the maximum voltage and still not give a good image. In this case, reconsider whether the staining protocol could be improved. Also, be sure to compare with unstained controls to verify that the observed signal is real, and not nonspecific autofluorescence. Insect cuticle and trachea and plant xylem vessels and stomata are all autofluorescent, and this extends into all colors, being slightly stronger at shorter wavelengths.

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Figure 9.4.11 Screen shots from the Zeiss 510. Balancing channels. When the gain is too high, bright parts of the image are flat and featureless (A). If the offset is too low, all detail is lost from the dim parts of the image (B). A properly adjusted image has very few saturated or zero-value pixels (C).

As the voltage on the PMT increases, it gives more random flashes which are unconnected to the underlying signal. These can be averaged down for the final image, but will severely limit what can be seen in the dimmest of samples. Allow more pixels to saturate or hit zero on these dim channels, then make adjustments if the final image is too bright or has too low a background. If needed, increase the pinhole size from Airy 1 to Airy 2 to give many times more signal. This makes a thicker optical section, but with a minimal loss of lateral resolution. Increasing laser power improves signal, but causes a much higher bleach rate, which then limits how often the dim signal can be averaged. Balancing a multichannel image takes time, during which a fragile sample is being bleached slightly. Make adjustments on a representative part of the specimen before finding the most critical location, so less time is expended and hence there is less bleach damage before the image parameters are optimized. If wide-angle and zoomed-in images are needed in the same field of view, scan the wide images first, or else the tiny bit of bleaching might be apparent as a sharp-edged region of slightly lower brightness. The human eye is very good at spotting artificial sharp boundaries, so even if the bleach damage is only a few percent, the image will be spoiled.

Brightness comparison Quantitation of confocal data is fraught with pitfalls, the first of which is saturation. Once a pixel hits saturation, it is not known whether it would have been two percent brighter or two-fold brighter had it been on scale. Therefore, even a few saturated pixels are enough to make any attempt at quantitation suspect. Comparative brightness of images rather than numerical quantitation does not pretend to be so precise, and is easy and popular. Start with the brightest sample, and set scan parameters so that the brightest regions are comfortably below saturation. Then go on to scan the dimmer samples with exactly the same scan parameters. PMTs have an exponential gain capability, so a few volts increase effectively doubles the signal brightness. Comparing two images is not possible if PMT settings were not exactly the same. Camera systems are much better for quantitation because they boast a much better dynamic range and their response is linear to within less than a percentage point. They are also gentler on the sample, so loss of signal due to bleaching during the scan is not such a problem. Microscopy


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Basic Protocol 3: Careful Collection of Quality Data Averaging There is little benefit in collecting higher numbers of pixels per image, if the noise level in the image is so high that they are mostly recording random intensity changes. To suppress noise, each location can be scanned multiple times, and the mean value saved as data. This is essential for dim samples where the detectors are running at a high voltage. PMT flashes are random, but real signal is always there and is not changed by averaging. Noise is reduced by the square root of n, so a 4 average cuts noise in half, while a 16 average is needed to cut it in half again. Multiple readings can be taken at each pixel by slowing the scan speed and measuring each pixel several times consecutively. Alternatively, a whole frame can be scanned and rescanned until the desired quality is obtained. Sweeping the laser slowly exposes the fluorophore to a higher intensity and duration of excitation, which can increase bleach damage, so it is better to scan fast and average more, than just scan more slowly. In the time it takes to scan and rescan the whole frame, parts of a live sample may have moved, to produce multiple ghosts of themselves at each subsequent scan. Line averaging is an attractive compromise, since scan rate is so fast that a vesicle moving under Brownian motion will still be captured as a single particle, but the fluorophores get a brief rest between each line sweep, which reduces their bleach rate.

Pixel number Once all the channels of data look good at preview speeds, a higher-quality image can be made by averaging and scanning more pixels. Increase the pixel number to 1024 × 1024 pixels, and select Line Average 4 (Fig. 9.4.12). This will scan twice as many lines of data, breaking each line into twice as many pixels to double the resolution in both directions. Modern systems can scan at 2048, 4096, or even 8192 pixels square, and can also zoom in 32 times. It is fun to explore the different settings to see what works best. Bear in mind, though, that resolution is ultimately set by the laws of physics, and a high-magnification

Confocal Microscopy

Figure 9.4.12 Scan parameter settings from Zeiss 510. X and Y pixel number can be selected from factory preferred presets, or entered directly. It is better to scan fast and average. The smaller the swing angle of the scan mirror, the faster it can scan.

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lens can only deliver a certain density of information, equivalent to roughly half the wavelength of the excitation laser line. A 63× objective has a field of view a little over 200 µm across, with a resolution just less than 200 nm. Hence, just over a thousand resolvable features can be fitted across the field of view. A two-fold over-sampling will just capture all this information, at either 2048 × 2048 or 1024 × 1024 and zoom 2. Lower-power lenses do not bump into the resolution limit quite so quickly, so a 40962 image taken with a 10×/0.4 NA objective is only over-sampling by a factor of 3 and thus will capture all the information present with minimal empty magnification. To optimize the collection of one-dimensional data, sampling speed should be at least twice as frequent as the peaks and valleys of the underlying data. This is called the Nyquist criterion, and is often applied to 2-D data collection on a scanning microscope. If the pixel density is twice the resolution, a data-rich, visually appealing image with no wasted magnification will be generated. Experience shows that exceeding Nyquist reveals a bit more information in a 2-D or 3-D confocal scan and gives a smoother, more convincing image of important fine features, even if the total detail per image is reduced.

Cross-talk or bleedthrough Fluorescent dyes absorb the pure color of the laser, then emit a range of colors, centered on a sharp peak, but spreading to a gentle tail which extends into other colors. On the excitation side, the spectrum is a mirror image, with a leading tail extending to bluer colors. Knowing the peaks’ locations enables a best guess of what laser and filter range to select, but the shape of the whole curve is especially important when several colors are present. Tails overlap with other detectors and lasers to cause spurious signals in the wrong channel, called cross-talk or bleed-through. If we fully understand the behavior of this stray signal, we could simply subtract it away. Suppose the FITC channel has a 20% bleedthrough into the TRITC channel for a given laser power and detector voltage. Subtracting 20% of the green image from the red image corrects this error and gives cleaner, brighter colors in the final image. With three or four colors, a matrix of corrections can be filled out. Do not be tempted to subtract too much between channels, as this will give the final image brighter, cleaner colors, but on close inspection can hollow out the real data and lead to clearly inaccurate artifacts. If a complete set of immunofluorescence controls is prepared, run the multicolor slide to optimize settings. Then, use each single-channel control in turn with the exact same settings to determine the level of cross-talk between channels. Unstained controls will give the level of autofluorescence, which is generally very broad and a little higher at bluer wavelengths. Draw regions of interest around features which are known to be stained with each of the dyes, then look at the statistics for those regions to determine the percentage cross-talk. New spectral systems can capture the whole curve of emission colors for each dye and save them in a database as a hyperspectral stack. Any mixture of these dyes can then be deconvolved automatically to be rendered in corrected colors corresponding to the level of dye present, not just the level of signal detected. Such software makes it possible to pack many more fluorophores into a single sample for investigating the inter-relationship of more components at a time.

Sequential scanning A simple solution to reduce the amount of cross-talk between channels is to scan the channels in sequence rather than simultaneously. If the DAPI laser is not on when the

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Figure 9.4.13 Sequential scanning (Leica SP2). When all lasers and all detectors are on together, emission from one dye can show up in the wrong detection window. Sequential scanning avoids this by cycling laser and detector line by line. The total scan speed slows accordingly, but the instrument otherwise behaves the same and gives cleaner colors.

FITC channel is observed, the very bright cross-talk signal vanishes and now FITC signal is visible inside the cell nucleus. This too can be done automatically, because the instrument has instantaneous control of laser power and which PMT signal is being collected (Fig. 9.4.13). Line-by-line sequential scanning takes a bit longer as the lasers and detectors are cycled, but the instrument behaves the same in preview and quality image-collection modes. Bleach rate is unchanged because the net amount of light delivered is identical. If moving a physical optic is beneficial between sequential scans, the scan can be done frame by frame, or Z-stack by Z-stack, where a whole set of data is collected, then the optics reconfigured and the data set collected again. A live sample would have time to change between frames, spoiling the alignment, but such scans can be used for collecting more channels of data in fixed samples than there are detectors in the system.

Time considerations for point scanning How long it takes to get a good image entirely depends on the quality of the sample, especially how bright it is, since PMTs are much less noisy at lower gain and less averaging is therefore needed. But it is also important to know how robust the sample is, since this limits how many times it can be scanned, especially for a Z-series or live kinetics series.

Confocal Microscopy

Generally, preview scans take a second each, and since the update scrolls down the screen, plenty of adjustment can be made during each second. High-quality scans take about 30 sec, and Z-stacks anything from 5 min to half an hour if the sample can stand it. Sequential scanning slows everything two-fold or three-fold for two or three fluorescence channels, respectively, with transmission usually being collected alongside the FITC channel.

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Basic Protocol 4: Imaging with a Camera Camera-based imaging is completely different from raster scanning with laser and PMT. Images are collected faster, and have greater dynamic range (Fig. 9.4.14). The field of view is fixed, so the flexibility to zoom and pan around is lost. Also, parallel detection with a spinning-disc module gives thicker optical sectioning compared to sweeping a single point through the sample. Light going in through one pinhole to the wrong depth can cause fluorescence to exit through neighboring pinholes, which introduces a slightly higher background and makes a thicker optical section. Image brightness on a camera system is exactly proportional to exposure time. Look at the histogram of pixel values to see what the brightest values are. If the image is too dim, increase the exposure proportionately. For example, a 12-bit camera can report gray levels from zero to 4095, so if the lowest 800 gray levels are used on a 0.1-sec exposure, increase to 0.5 sec for a full range image, or to 0.4 for a comfortably bright image with no saturation. This process can be automated, so an autoexposure can be requested in the same way as with a retail-grade pocket camera. The system takes an extremely short exposure, looks at the values, then takes a second exposure of exactly the right duration.

Figure 9.4.14 Histogram controls on the BD Pathway HT high-content screening system. Cameras have a large dynamic range, and image brightness can be scaled to always look good on the screen. Specify a brightness range to exaggerate structures of interest, or to let brightness vary as the sample is explored.

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Binning If it takes 2 sec to generate an image with 800 gray levels, quadrupling the exposure time makes the system difficult to use because the wait between images gets too long. Instead, binning can be used, where 2 × 2 blocks of pixels are all added together to give a final image that is four times brighter, but with half the resolution. A 1300 × 1024–pixel camera may give perfectly adequate images at bin 2 (650 × 512) for routine work. Then, bin 1 is available for the occasional high-resolution final image. Binning is handled right on the camera chip, so data-transfer rates are also proportionately faster. EMCCD cameras If extremely dim samples need to be imaged extremely quickly, a detector needs to capture and report every photon that reaches, to produce a grainy image. This is possible with intensified and EMCCD cameras when they are used with Yokogawa scanners. Electron-multiplied charge-coupled device (EMCCD) cameras can operate like regular cameras for bright samples, but have an optional trick for data read-out. When a photon strikes the back-thinned CCD sensor, it has up to a 95% chance of kicking loose an electron from the silicon substrate, which is held in place within the pixel. Rows of pixels are transferred line at a time to a read-out register, then quantified one at a time by the analog to digital (A-to-D) converter. EMCCD cameras have a hairpin readout column, where each transfer from pixel to pixel is done with a stronger than usual voltage. As electrons are moved, they kick loose additional electrons to increase the pile in each pixel. By doing this hundreds of times, a thousand-fold amplification of the captured signal can be achieved. This is different from simply multiplying the output number because it happens between the sensor and the analog-to-digital converter. A-to-D converters have inherent noise or error, which increases as the speed of data transfer increases. At a clock speed of 10 MHz, this error could be 35 electrons or more. In ordinary CCD mode, at least 35 photons per pixel must be collected to be sure the signal is real. But in EMCCD mode, the massive amplification means that every captured photon counts and can be seen above the noise floor of the A-to-D converter. The camera has single-photon sensitivity. EMCCD cameras are delicate and can be damaged by too much signal going through them. They will also pick up the tiniest leak or internal reflection within the optical light path. Do not be tempted to use them at maximum settings, as this will shorten the life of the camera without contributing to final image quality. Instead, start with a very low gain setting, then only rise to mid-range for the dimmest of samples. Make a note of optimal scan settings for future use, and do not tamper with the alignment of scanner and camera once the system is working well.

Other camera types An image intensifier is a plate full of narrow channels with a phosphor screen on one side and an accelerating voltage along the bore. This can be coupled to the front end of a CCD camera for extremely fast imaging at high sensitivity. Quantum efficiency is lower than for the best CCD cameras, and cameras set up this way are more fragile, but they benefit from massive amplification of signal and the ability to be shuttered on and off very quickly. Synchronized to the frame rate of a spinning disc scan head, they produce the very fastest low-light images. Accidentally exposing an active intensified camera to a bright image can burn it into the phosphor screen as a permanent shadow, so great care is needed. Confocal Microscopy

CMOS technology is beginning to be used for high-performance imaging (Andor, http://www.andor.com). Rather than having an array of wells that pass charge from one to

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the other, the CMOS chip uses the charge generated by light capture to turn on flow in a transistor circuit in each pixel location. This arrangement enables random-address access to anywhere on the chip, much higher speeds, lower power requirements, and smaller, more tightly packed pixels. CMOS do not have on-chip binning capability, however, so any summing and averaging is handled after collection.

Time considerations for camera imaging The speed limit on camera imaging is the refresh rate of the spinning-disc pattern, since the pinholes have to smoothly cover the whole field of view without leaving gaps or narrow bright stripes. This can be as short as a 10-msec exposure, plus the read-out time of the camera. A 20-MHz camera clock speed allows 20 million pixels to be transferred to the computer per second, which could be 20 whole 1,000 × 1,000 frames or eighty 512 × 512 frames at bin 2. But this is transfer only, so if the exposure time is 100 msec, six full frames is the theoretical maximum. Binning makes the images four times brighter, so the 512 images can either be four times brighter, or collected at 25 msec for a maximum 20 frames per second at the lower quality. Use a subregion of the camera to push the speed faster. Generally, the cameras are run at bin 2 and bright samples generate good images in 0.4 sec on a single disc scanner, or 100 msec on a Yokogawa dual-disc system. A piezo stage or lens holder allows very fast cycling of focal plane, to allow a ten-slice Z-series to be collected every second for four-dimensional data collection. Changing color drastically slows data rate on a camera system, since filter wheels need to be moved, and the colors collected in sequence. Spinning disc systems lose their speed advantage when multiple colors are needed, although they are still more gentle on the sample.

Data removal and shutdown Most companies now use their own data format, rather than standard TIFF or lossy JPEG, so they can save all the information of how each image is scanned. Data then has to be exported before use in other programs. A free “lite” version of the scanning software is available to make it easy to look over collected data and export to other programs. Commercial 3-D software like Imaris (http://www.bitplane.com/) or Volocity (http://www.perkinelmer.com/) can input data from any confocal scanner for analysis and display, but popular image archiving software like Picasa does not support exotic formats. ImageJ is an open-source image analysis program with plug-ins that enable data entry from all popular brands of microscope. Save data to a named subfolder, to keep the host computer tidy and prevent accidental loss or overwriting of similarly named experiments. Remove the day’s work from the microscope to back it up and free up space for future users. Burning CDs and DVDs may be less convenient than a removable hard drive, but can act as an inexpensive secondary back-up, which will not be overwritten by accident.

Keeping the confocal microscope clean A compound microscope is cunningly designed so that any dust that gets inside and settles on an optical element is out of focus and not noticed. But it is still reducing the light throughput of the system, and introducing stray scattered signal, which will affect the resolution. When any microscope is not in use, it must be under a dust cover to prevent this degradation (Fig. 9.4.15). When the dust cover is not being used, it should not be allowed to fall on a dusty floor.

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Figure 9.4.15 Dust covers are an essential part of any microscope, since any dust that gets inside cannot be removed. Your work session is complete when the system is shut down and covered, or when it has been handed over to the next user.

Dust accumulates under gravity, so if a convoluted air path is sufficient to keep a Petri dish sterile, it is also sufficient to keep a microscope dust-free. If a room has a particularly bad dust burden, a whole-room HEPA filtration system is a sound investment to constantly pull dust out of the air before it can contaminate the optics (e.g., IQ Air, http://www.iqair.com). The same approach is used inside some scanners and lasers, which often have a small air-filtration system to keep the interior under positive pressure with clean air. These filters can become clogged over time, so they should be replaced or cleaned at the time of a preventive maintenance inspection (PMI). Dry objective lenses never touch the sample, and thus do not need any special cleaning. On an inverted stand, dust may accumulate, and can be blown off with a rubber bulb or carefully with canned air. Test the can trigger before use, to avoid hitting the lens with a blast of liquid refrigerant, which can contaminate the lens surface and give the assembly a risky thermal shock. Water objectives are also easy to clean with a single sheet of lens tissue or a folded Kimwipe moistened with a drop of either Sparkle or Windex. Assume aseptic technique and do not use the parts of the tissue that have been handled, or finger-grease and flecks of skin will transfer into the solvent on the wipe. Use a new wipe each time, and for each lens. Stretch the wet area of the tissue across the lens, then sweep sideways so the cleaning fluid is all drawn into the dry part of the tissue. Glycerol can be cleaned off in the same way. A fully dry wipe is a potential scratch hazard, so follow wet with dry in quick succession. Confocal Microscopy

Immersion oil is much thicker and has a stronger affinity for the glass and metal of the lens surface than water. Repeat the wipe three or four times until the lens is clean and the paper dries with no sign of contamination. Methanol or ethanol can also be used, but

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care must be taken not to allow the solvent container to pick up dissolved grease from any surfaces that have been touched. Otherwise, when they dry, a milky residue or oily smear is left behind. Xylene is out of favor because it is a suspected carcinogen. If solvent is allowed to dry on the lens, rather than being sopped up, it will redeposit any grease that was dissolved in it, requiring additional cleaning steps. Bubbles broken from a block of expanded polystyrene (e.g., Styrofoam) can be used as a gentle final wipe. The interior surfaces of this material are free from dirt, and are soft and lipophilic. They work well to remove that last sheen of grease from the front of an objective lens. Dipping objectives have ceramic and Teflon exteriors that are resistant to physiological buffers and dissolved fluorescent stains. Some of them have a concave front face though, which makes wiping less efficient. Instead, make a pool of cleaner in a dish lid and lift it to fully immerse the lens. Hold for a few seconds, then remove and wipe in the usual way. Fold a wipe to make a clean point, which can be lightly pushed against the concave surface. No hard pressure should ever be applied to an objective, as there may be some grit that would scratch the face of the lens.

Figure 9.4.16 Scanning artifacts. If slices in a Z-series are too far apart, stripes will appear between slices (A). Air bubbles are strong negative lenses that block one side of the lens (B). Bilateral scanning can cause zippering artifacts where left and right scans do not exactly line up (C, far left and far right). EMCCDs are so sensitive that internal reflections can show up in the image (D). Camera systems show dust in the light path as a rough transmitted-light backdrop that does not move with the specimen (E). These are hidden by a frosted beam diffuser in the condenser (F). If the field of view is too small, a montage of any size can be collected on a motorized stage (G). Microscopy


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Larger lenses, like those in the condenser, are harder to clean to visual perfection because of their size and often highly curved shape. It is better to avoid contamination in the first place by not using too much immersion fluid and by addressing any spills or breakages promptly. Do not worry about the last interference fringe of oil left in the periphery of the condenser lens or polarizer filter as its effects will be trivial. Most immersion lenses have a spring-loaded front face to protect against bumping and breaking the slide. If a lens is driven through the sample by mistake, it should be inspected for damage and all shards removed from the condenser or inverted turret. The condenser may have to be removed from the microscope and partially disassembled to get out every crumb of broken glass. To inspect a lens, remove it from the microscope, then cover the opening or move the turret to protect against dust. Remove an eyepiece in the same way and hold the eye side of the eyepiece close to the lens. Turn the lens to catch a reflection off a bright light source, and any problems on the front face can be seen clearly through the back of the eyepiece. A stereo microscope can also be used.

Troubleshooting The most frustrating aspect of microscopy is when a sample is presumed to be properly prepared, and a scanner properly configured, but there is no signal on the screen. It takes a bit of patience to plod through each step of the setup procedure to find any mistakes. Have a well stained test slide available to aid in this troubleshooting, and check settings in sequence until the error is detected. Go back to ocular viewing in fluorescence, or even bright field, to be sure a good location is chosen. Know the sample well enough—e.g., dim signal from autofluorescence can act as an internal control of imager performance. Insect trachea and cuticle are slightly autofluorescent at most wavelengths, and thick cell walls in plant trichomes, stomata, and xylem sieve elements are easy to recognize. Problems most often arise after turning on the system for the first time, but software can freeze or drop communication to the hardware at any time. Save frequently to minimize the risk of lost data in the event of a crash. Examples of image quality problems are shown in Figure 9.4.16.

Basic Protocol 5: Collecting a Z-Stack A volume of the sample can be captured by taking a series of equally spaced images or Z-stack. The spacing between slices can be matched to the resolution of the microscope, to avoid leaving gaps in the data set. But if a thick Z-series is needed, 1- or 2-µm sections are preferred, to reduce the time needed to collect the data, and hence the resultant bleach damage. 1. While preview scanning, change the focus to one extreme of the depth range of interest, usually where the structure of interest begins to completely fade into the background. 2. Mark it in the software as the Start position, then focus to the other side and mark the End (Fig. 9.4.17). It does not matter if top or bottom are marked first, as the system will move against gravity to collect the data. 3. Stop the preview, and see how thick a range has been marked. Then enter a number of slices to be fit within this range and/or the spacing of the sections. If the Z-range is 19 µm, request 20 slices so they are exactly 1 µm apart. Confocal Microscopy

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4. Change resolution and averaging if needed, but use the smallest effective average and lowest pixel number, or the scan will take too long and the lower levels will be bleached out. Current Protocols Essential Laboratory Techniques

Figure 9.4.17 Z-stack controls on the Pathway HT. Collect a Z-stack by marking both extremes of the thickness to be sampled, then entering the number of slices to fit between those extremes. Use a round-number thickness, so each slice is a meaningful distance from its neighbors.

5. Press Go to collect the whole Z-series. If too deep a Z-stack is requested, it can always be stopped early, but if the tolerances are set too tightly, the top may be missing and the whole stack spoiled.

3-D rendering of finished data The finished 3-D volume is typically 512 or 1024 pixels square and 20 to 50 layers deep. Depth resolution is so much worse than lateral resolution, so a rendering of the whole stack will always look better if it is top-down or almost top-down (Fig. 9.4.18). The closer to perpendicular, the more the lower resolution of the Z-dimension intrudes into the image quality. 6. Use an orthogonal view mode to evaluate the quality and content of the stack. This shows the X-Y image in one panel, the X-Z image in an adjacent panel, and the Y-Z image in a third adjacent panel. The intercept of all three can be moved around to peruse the whole stack at any depth and along any line of X and Y. 7. To see the whole volume in a single image, use a maximum intensity projection. This looks through all levels and displays the brightest pixel at each position of X and Y, producing a bold, attractive image. Any pixel noise will be very visible in dark areas, and this view is of no value for quantitation. 8. Save the projection for future reference. Some software applications autosave all data management steps, but be careful not to lose any work. 9. Request an Average projection to simply add up all the sections and divide by the number of slices. This produces an arithmetic mean image through the whole depth, giving a smooth, soft image, which is quantitatively honest because each pixel contributes equally to the final image. 10. Movie loops showing the volume being rocked from side to side, or spun around, can be made by rendering a projection from a series of vantage points and reassembling them as a movie. This is easy to do and takes relatively little processing power. 11. Commercial 3-D software applications hold all the data in memory as a cloud of voxels, which can be manipulated in real time, making it possible to spin it

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Figure 9.4.18 3-D visualization controls on the Zeiss 510. Orthogonal planes (A) show an XY, XZ, and YZ composite view, where the lines of intersection can be moved interactively to explore the sample. A series of vantage points for reconstruction can be specified as a start angle, number of views, and increment between views (B) to make simple projection movies. A maximum intensity projection shows the brightest pixel along each line of sight for a more visually striking image set (C,D,E) while an average projection (F,G,H) is more realistic.

around, zoom in and out, or even fly through the volume. Isosurface planes can be drawn inside the volume to highlight structures of interest, or particles can be found automatically, and their brightness, shape, and position tracked over time

Confocal Microscopy

Basic Protocol 6: Kinetics Imaging By repeatedly collecting data at different time points, the temporal behavior of a live sample can be monitored. With a fast Z-drive and high-speed scanners, a whole Z-series can be collected several times a second to monitor the behavior of features in three dimensions. The difficulty with such experiments is they require so many scans in the same location, so bleach damage becomes a serious concern. It takes practice to balance image quality with collection speed, so that enough scans can be taken before too much of the fluorophore is bleached away, or laser toxicity starts changing the behavior of a living system. Camera-based spinning-disc confocal microscopes are better suited to this

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kind of experiment because they are much gentler on the sample, have a higher quantum efficiency, and collect more light per second of exposure time. Higher speeds can be achieved with the simple expedient of collecting fewer lines of data. Scanning in the X direction is fast, but accumulating lines of data into the Y direction necessarily takes longer. If the distance traveled in Y is reduced to a quarter, the scan time is correspondingly reduced. This is often a better solution than cutting pixel number in both directions, since it scans a larger area per second than a square scan. The letterbox profile can be rotated to capture the feature of interest. The logical extreme of this is where just a single line of data is collected repeatedly to visualize events happening in the millisecond range, for example to track blood cells flowing through a capillary. An X-T image can be rendered from the results, where the Y direction is replaced by time in a two-dimensional rendering of the data set. If the laser is parked at a single location and not scanned at all, microsecond-scale phenomena could be captured, subject to the mobility of fluorophores and their stability. Fluorescence correlation spectroscopy (FCS) relies on this imaging mode to see single fluorophores drifting through the illumination/detection window. When the concentration of fluorophore is very low, and the detection volume is known, this can be used to precisely quantify the concentration of fluorophores in the surrounding medium and the speed at which they can move around. Scan speed can be doubled by scanning in both directions instead of raster scanning. The two sweep directions must exactly line up, or there will be a zippering effect visible on sharp edges where the edge is encountered sooner in one direction than the other. Careful adjustment of “phase” corrects for this empirically by adjusting the sweep controls until they both appear to mesh together exactly. Some point scanners have custom hardware for extremely fast point scanning. For example, the Leica SP5 has an optional resonant scanner that can be moved in place of the regular X-axis mirror. Resonant-scanning galvanometers have a more limited swing angle and work at a very high fixed speed. The time spent sampling each pixel is then much shorter, so there is more random shot noise in the image, but this can be averaged down to whatever trade-off works best for speed and image quality.

Holding focus Confocal optical sections are so thin that any drift in depth during a time series will lead to a change in the area being sampled. If the sample is stuck to the coverglass, a focus-lock infra-red laser inside the microscope can be used to track the position of the glass/water interface and keep the offset constant (Fig. 9.4.19). If the specimen is nonadherent, use an image-based autofocus that quickly takes a series of images at different heights, then chooses the one with best contrast as the reference height. This is slower, even with camera systems, so some trial and error is needed to optimize how many depths to sample at what image quality. But once calculated, extremely long time series can be generated with no visible drift. A third option is to just over-sample with a Z-series, then either use the whole stack, or pick out which levels match as the time series progresses.

Photobleaching and photoconversion Deliberately bleaching a small region of the sample is a powerful technique for monitoring the behavior of the fluorescently tagged molecule. Fluorescence recovery after

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Figure 9.4.19 Autofocus controls on the Pathway HT. Image-based autofocus and laser focus. If the sample is nonadherent, a series of images is taken to pick the best height. If the coverslip is an effective reference, an infrared laser bouncing off it can tell the difference between the top surface of the coverglass and the more reflective underside, so the microscope can keep a steady focus at any given depth.

bleach

Intensity

diffusion rate

bound in place freely diffusing

Time

Figure 9.4.20 FRAP measures the proportion of a molecule that is bound verses freely diffusing and also the diffusion rate can be calculated from the speed of recovery after a bleach event.

photobleaching (FRAP) can determine the mobility of a fluorescently tagged protein by bleaching then measuring the rate and extent of recovery (Fig. 9.4.20). Bleached molecules diffuse away from the region and are replaced by unbleached molecules from the surroundings. If recovery is not complete, that suggests there is a subpopulation of the molecule that is bound in place (Axelrod et al., 1976). 1. To do a FRAP experiment, define a small region within the field of view using the Bleach module of the software. Confocal Microscopy

2. Scan a series of five images, then trigger the bleach

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3. Take an extended sequence of kinetics images to record any recovery of signal in those areas to complete the experiment. 4. If the amount of fluorophore is limited, recovery of signal in the bleached zone will correspond to a decrease in signal in the unbleached zone due to diffusion mixing. This can be used in Gap-FRAP to determine the gap-junction connectivity of cells that have been labeled with a bleachable dye.

Basic Protocol 7: Montage Imaging When collecting images on any microscope, start with the lowest magnification and work up until the necessary resolution is achieved. However, what if the feature of interest is too big to fit the field of view? If the microscope has a motorized stage, a montage image can be requested, where a series of adjacent images are scanned and then stitched together to make one extra-large image. Montage can be performed with Z-series, to stitch together whole volumes of image data, or with slow kinetics experiments, where a whole population of cells is being monitored and the extra image size is needed to monitor a statistically significant number of targets. The montage images may exactly abut against each other to make a continuous field of pixels, or an overlap of 10% is automatically included, and the data in the overlap used for exact registration and blending of adjacent images, much like the panorama feature in a pocket camera. For very large scans, be aware of the data size limit of the instrument, so the software does not seize up, and consider how long the whole image will take to scan. 1. To prepare a montage, specify the area to be scanned by driving to each extreme of the area of interest and noting its position. The software may give guidance, asking to mark the top-left and bottom right of the rectangle to be scanned, but be aware that those corner positions may be in open space, and that the edges of the tissue section may be ragged and have several local extremes. 2. If one axis of the XY stage controls can be locked, it is easy to drive along the presumed perimeter to confirm that nothing is being cropped. Otherwise, watch the stage’s X-Y position closely, to avoid drifting on one axis while traveling a long distance in the other. 3. Subtract X coordinates from the extreme left and right, then divide by the field size to calculate how many fields of view are needed. Then navigate to the center. 4. Do the same for the Y axis. 5. Enter the number of fields needed in X and Y, then start the montage scan. Some systems allow a single row to be added in case of an error, while in others the whole area must be re-scanned.

6. When moving very large distances to collect a big area at high resolution, the focal plane may be different from one side to the other. If part of a Z-stack, the whole thickness would have to be collected to avoid cropping out one side. But if only a single depth is needed, use an autofocusing routine, as with kinetics scanning. Decide how frequently an autofocus scan is needed to save a little time, but be aware that it cannot find image-based focus in empty fields of view, and therefore may have difficulty in the corners. 7. Some software applications allow irregular shapes to be specified and scanned, or offer the option of deliberately leaving a gap between scanned areas, to statistically sample a very large specimen without collecting all the data.

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Table 9.4.3 Troubleshooting Guide for Confocal Microscopy

Problem

Possible cause

Solution

System failed to initialize

Software started too soon after Turn off hardware and program. Restart hardware and hardware wait 30 sec before restarting software.

Field of view fuzzy to eye on one side only

Air bubble in immersion fluid

Lower and raise stage to pop bubble. Clean and reapply fluid.

Field of view to eye flickers or Arc lamp bulb old or is uneven misaligned

Switch to test slide and adjust arc lamp for flat, bright illumination. Replace bulb after 200 hr, or if it flickers.

No image in any scanned channel

Wrong focal plane

Turn on transmission channel to find focus.

Pinhole too small

Check software knows which lens is active, and open to Airy 1.

Zoom too high

Zoom needs to be reset to 1 whenever lens is changed.

Sample absent from field of view

Switch back to ocular viewing and confirm position of sample, and settings of microscope for scanning.

Laser off

Check laser power supply for status/warning lights and confirm keys and switches are set properly.

Wrong dichroic selected

Turn down working lasers to verify missing channel laser is visible at the sample, and check optical layout.

Gain too low

Increase gain until random PMT snow starts showing at high voltage.

No image in one channel

Look-up table (LUT) invisible Switch to quantitative LUT, which shows zero and max; avoid pure blue in a dim channel Transmission image saturated

Tungsten lamp still on

Switch off lamp, select detector in transmission path,

Transmission image blank

Condenser misaligned

Adjust condenser for good Kohler illumination.

Optic blocking light path

Adjust microscope settings until white light is seen at the slide.

Transmission software off

Check optical layout and make sure transmission is on.

Gain too low

Increase gain. If each channel has separate pinhole, increase pinhole in dim channel.

Laser not selected

Signal is from cross-talk only if correct laser is off. Briefly turn off other lasers to see correct color at sample.

Focus always different between scanner and eye

Diopter adjustment set low

Each eyepiece has a correction collar which should be set to zero, not screwed all the way to one extreme: Leave eyeglasses on for normal infinity vision: Some systems have a custom scan tube lens that must be selected.

Error message, image too big

Memory capacity of display exceeded

Scan fewer channels, or lower pixel density on all channels. Save and close old images.

Images fade out during kinetics scan

Laser power too high

Lower power, and increase averaging if needed to maintain image quality; reduce pixel density; increase time interval.

Stripes appear in Z-series reconstruction

Step size is greater than resolution

If sample can stand additional scans, reduce step size to below lens resolution; if not, increase pinhole size and lower laser power to thicken sections and reduce bleach damage.

Image too dim

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CONCLUDING REMARKS Confocal images have become the standard for published optical microscopy figures. Even when the third dimension is not required for the science in question, the sharper, brighter images add impact to an article. Modern confocal systems are still expensive, but are easy to set up and use, so should not be avoided out of fear of their complexity. Optical sections reveal more detail within a sample, and digital imaging gives more control of the data collection process. It does take several hours of practice to get to know a new type of sample in three dimensions, and to get comfortable with taking high-quality images (see Table 9.4.3 for common troubleshooting issues and resolutions). New users should allow enough time to work on each sample, and only hope to image three or four samples per hour on a point scanner. If a particularly good region is found, spend the time to frame a really pretty shot, in case it will be useful for a publication. Book the first few sessions during the day when there are staff available to help. After that, book quieter times when there are fewer distractions.

LITERATURE CITED Ando, R., Hama, H., Yamamoto-Hino, M., Mizuno, H., and Miyawaki, A. 2002. An optical marker based on the UV-induced green-to-red photoconversion of a fluorescent protein. PNAS 99:12651-12656. Avila, E., Zouhar, J., Agee, A., Carter, D., Chary S.N., and Raikhel, N.V. 2003. Tools to study organelle biogenesis: Point mutation lines with disrupted vacuoles and high speed confocal screening of green fluorescent protein tagged organelles. Plant Physiol. 133:1-4. Axelrod, D., Koppel, D.E., Schlessinger, J., Elson, E., and Webb, W.W. 1976. Mobility measurement by analysis of fluorescence photobleaching recovery kinetics. Biopys. J. 16:1055-1069. Deerinck, T.J. 2008. The application of fluorescent quantum dots to confocal, multiphoton, and electron microscopic imaging. Toxicol. Pathol. 36:112-116. Hama, H., Kurokawa, H., Kawano, H., Ando, R., Shimogori, T., Noda, H., Fukami, K., Sakaue-Sawano, A., and Miyawak, A. 2011. Scale: A chemical approach for fluorescence imaging and reconstruction of transparent mouse brain. Nat. Neurosci. 14:1481-1488. Kwaaitaal, M., Keinath, N.F., Pajonk, S., Biskup, C., and Panstruga, R. 2010. Combined bimolecular fluorescence complementation and F¨orster resonance energy transfer reveals ternary SNARE complex formation in living plant cells. Plant Physiol. 152:1135-1147. Matsumoto, B. (ed.) 2002. Cell Biological Applications of Confocal Microscopy. In Methods in Cell Biology, Volume 70, 2nd ed. (, 70) Academic Press, San Diego. Rizzo, M.A., Davidson, M.W., and Piston, D.W. 2010. Fluorescent protein tracking and detection: Fluorescent protein structure and color variants. Cold Spring Harb Protoc. 2009 Dec;2009(12):pdb.top63. doi: 10.1101/pdb.top63.

INTERNET RESOURCES http://faces.ccrc.uga.edu/ A free calendar reservation system which is widely used for core resources such as seminar rooms and shared instrumentation. http://www.emsdiasum.com/ The Electron Microscopy Sciences online catalog contains most supplies needed for optical microscopy too, including all types of mounting media, slides, and coverslips. http://www.bioptechs.com/Products/OBJ HTR/obj htr.html Describes a well designed objective lens heater for use with a heated stage insert. http://www.vectorlabs.com/catalog.aspx?prodID=428 Non-setting mounting media with DAPI for nuclear counter-staining. Vector labs sells many kits and reagents for all kinds of staining protocols, including secondary antibodies for a dozen species for immunostaining, and kits for fluorescent in-situ hybridization (FISH).

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http://www.iqair.com/ An air-filtration system that reduces the dust burden in a microscopy lab and optionally can remove chemical contaminants, such as DMSO, from a robotics room. http://www.macbiophotonics.ca/downloads.htm A version of ImageJ that is a good starting point for microscopy analysis of images. ImageJ is free and has a large user base who constantly add to its functionality. http://www.carolina.com/category/life+science/microscope+slides/histology+slides.do This school supply house has a large collection of histological slides, some of which are fluorescent (from the eosin in H & E), and can be used as practice pieces. http://wardsci.com/ Another online school supply house with an extensive collection of histological slides. http://www.tedpella.com/histo html/histolgy.htm Fine science tools supplier with fixatives, stains and equipment for sample preparation. http://www.invitrogen.com/site/us/en/home/brands/Molecular-Probes.html The most comprehensive source for fluorescent reagents is Molecular Probes, now part of Invitrogen. http://micro.magnet.fsu.edu/primer/techniques/confocal/index.html Titled “Molecular Expressions,” this Florida State University site is an exhaustive resource for all things microscopy and includes many interactive Java applets for hands-on demonstration of various aspects of instrument performance. http://www.microscopyu.com/articles/confocal/ This Nikon-sponsored extension to “Molecular Expressions” has many tutorials on various aspects of confocal imaging and sample preparation. http://www.andor.com/ This camera company is pioneering scientific CMOS cameras and offers integrated spinning disc confocal systems. http://www.olympusamerica.com/seg section/seg microscopes.asp?section=confocal Olympus makes relatively affordable confocals with the most extensive analysis and quantitation software. Their FluoView 1000 can be configured with a whole second scanner for photobleaching and photoconversion applications. http://www.leica-microsystems.com/products/confocal-microscopes/ Leica makes premium confocal systems with excellent sensitivity and ease of use. They were the first to market with a real-time super-resolution system. http://www.zeiss.de/c12567be0045acf1/Contents-Frame/3d7a40584d71593841256a71004cb2d4 Zeiss makes premium confocal systems with excellent image quality and high performance objectives.

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