Two-Photon Microscopy for 4D Imaging of Living Neurons
    Steve M. Potter
    Neuroengineering Laboratory, Coulter Department of Biomedical Engineering, Georgia Institute of Technology and Emory University, Atlanta, Georgia 30332
    
    T
    
    wo-photon laser-scanning fluorescence microscopy (Denk et al. 1990) has made it possible to image neurons over 600 pm deep within a living slice or organism, with submicrometer resolution and in three dimensions (3D) for many hours without photodamage (Potter et al. 1996a).With true 4D microscopy (i.e., 3D with time), it is now feasible to capture neural development and synaptic plasticity in the act of happening, eliminating many uncertainties associated with between-animal comparisons of fixed-tissue specimens. By using pulsed IR laser light to excite fluorescent labels (e.g., dyes, fluorescent proteins, or endogenous fluorophores) that are normally excited by visible light, excitation is restricted to the focal plane, greatly reducing photobleaching and phototoxicity (see Chapter 7). The IR illumination is scattered less than visible light, allowing imaging 2-3 times deeper than with standard confocal microscopy. In addition, because the fluorescent signal emanates only from the focus of the scanning IR laser beam within the specimen, no confocal aperture is necessary to remove the out-of-focus signal. This means that two-photon microscopy has an inherently higher signal-to-noise ratio (SNR) compared with confocal microscopy. Excellent references for two-photon microscopy, and for labeling and imaging living specimens, include Denk et al. (1995) and Terasaki and Dailey (1995). In this chapter, I describe two-photon imaging hardware, pointing out potential pitfalls in microscope construction and operation. I also describe technical considerations for successful two-photon microscopy in two experimental systems: (1) 4D imaging of living neurons transplanted to cultured hippocampal slices from neonatal rats and (2) 4D imaging of dendritic spines within acute hippocampal slices from adult rats. Figure 1 shows the components of the imaging setup.
    
    TWO-PHOTON HARDWARE
    Although both confocal and two-photon microscopes scan a laser beam in the focal plane within the specimen, a two-photon microscope can actually be a much simpler device than a confocal microscope. There is no need to focus or even descan the emitted fluorescence to create an image. In most cases, however, it is probably easier to convert a confocal microscope to a two-photon microscope than to build a two-photon system from the ground up. I converted a Molecular Dynamics Sarastro 2000 confocal with an upright Nikon Optiphot I1 microscope, preserving its ability to be used in standard confocal mode (see Fig. 2) (Potter et al. 1996~). every case where Fraser and I imaged the same In
    
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    Sarastro optics @*
    
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    mp laser
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    Mobile Slice Unit
    
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    FIGURE 1. The imaging setup. In the foreground is the Mobile Slice Unit, with oxygenated ACSF tank and peristaltic and syringe pumps. In the center is the Molecular Dynamics Sarastro 2000 confocal microscope (converted for two-photon microscopy), enclosed in insulation and warmed by an egg incubator (upper left). Behind the microscope are the argon-ion pump laser and Ti:sapphire IR laser. The argon-ion laser has been replaced with a shoe-box-sized,all-solid-stateVerdi pumplaser (not shown).
    
    \
    
    specimen using confocal microscopy with visible excitation (either the Sarastro or the Bio-Rad 600) and two-photon microscopy (converted Sarastro), we got better resolution, deeper penetration, and less photodamage with two-photon microscopy. The confocal capabilities of the converted Sarastro are therefore seldom used.
    
    Lasers
    To produce adequate two-photon excitation, an IR laser that compresses all its output into very short (-100 fsec) pulses is necessary. Although the peak power of these pulses is enormous, the pulses are widely spaced, so the mean power is not enough to cause any heating of the specimen. (For a detailed discussion on lasers in multiphoton microscopy, see Chapter 79.) We use the Coherent Mira900 titanium:sapphire laser, which is tunable from approximately 700 to 1000 nm. The Ti:sapphire laser converts green light from a pump laser into pulsed IR light. The Coherent Verdi pump laser is superior to the Coherent Innova310 8-W argon-ion laser. The Verdi is an all-solid-statelaser that can be powered from a normal 110-V outlet and requires no cooling water, unlike the Innova310. With only 5.5 W of green light, it produces more mode-locked (pulsed) IR output from the Mira900 compared with the 8-W argon laser, due to improved beam quality. The IR beam is brought into the microscope with four dichroic mirrors, optimized for broadband IR reflection at a 45' angle (Newport BD.2). Between two of the mirrors is a beam expander (Newport T81-3x), which is used to focus the beam so that it is slightly larger than the back aperture of the objective lens. Without it, most of the large beam from the trinocular (scanner) eyepiece does not enter the lens, and two-photon excitation is greatly reduced. All beams are covered with tubes connected to sealed mirror boxes. This allows safe use, even by biologists, and protects the mirrors from dust buildup and accidental bumping.
    
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    Laser for 1-photon
    
    I
    
    FIGURE 2. Optics of Molecular Dynamics Sarastro 2000, modified for two-photon microscopy. The pulsed IR laser beam is brought into the Sarastro by a periscope to the left. Before the periscope is a 3x "beam expander" (not shown) used to match the IR laser beam width to the back aperture of the objective lens. To switch from two-photon to confocal imaging, one need only block the IR beam, turn on the on-board laser (upper right), rotate the primary dichroic beam-splitter wheel to a long-pass mirror, and rotate the pinhole aperture wheel from the open hole to one of the three pinhole settings.
    
    Conversion of the Sarastro 2 0 0 0 Confocal to a Two-Photon System
    The Sarastro 2000 has a very simple light path compared with most available confocals, which is desirable because valuable photons are inevitably lost at every mirror, lens, and filter. To convert it to a twophoton microscope, I replaced mirror 2 (see Fig. 2) with a 680-nm long-pass (680 LP) beam splitter that allows us to use either the IR laser or the on-board visible laser. We use a 680-nm short-pass (680 SP) dichroic mirror to separate excitation from emission in the primary beam-splitter wheel. Mirror 3 and the two scanning mirrors were coated with silver and a protective dielectric coating (Ventura Optical Industries). Silver reflects both visible and IR light well, unlike standard aluminum mirrors, which reflect IR poorly. The dielectric is crucial to prevent the silver from tarnishing. No changes were
    
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    made to the OptiphotII, which has a motorized z-stage with 0.1-pm accuracy. The stage and scanner are controlled by an SGI Indigo workstation, running Molecular Dynamics Imagespace software. No changes to the software were necessary for two-photon imaging.
    
    Non-descanned Detection: Every Photon Is Sacred
    To get images with the best SNR, it is important to collect as many photons as possible. Our microscope sends the emitted light through a trinocular eyepiece (beneath the scanner), two scanning mirrors, and an achromat lens before it gets to the on-board detectors. To avoid the losses associated with these optics, and to collect more of the photons that are scattered on their way out of the specimen, it is best to detect the emitted light as close to the specimen as possible (see Fig. 3). We replaced one
    
    + To on-board
    'descanned path" detectors Scanner box
    
    Trinocular eyepiece
    
    Lost
    
    Epifluorescence attachment with custom filter cube (670LP)
    
    Excitation blocker (BG39) Objective lens sealed to membrane-insert weight with latex barrier Petri dish with specimen on membrane insert
    
    FIGURE 3. Non-descanned detection. By placing a detector close to the specimen, it is possible to collect more scattered photons and avoid losses associated with optics in the descanned path. For low background, this "non-descanned" detector must be well shielded from room lights and scattered IR excitation. To avoid damage to the sensitive photomultiplier tube, it should be automatically shut off when the transilluminator, the epi-illuminator, or the on-board laser is turned on. The Y-scanner reflects the IR beam down through a trinocular eyepiece and custom filter cube to the objective lens. Emitted fluorescence is reflected by the cube into a custom detector mounted directly in front of it, or if the cube is gulled out, it is descanned and sent through a 680-nm short-pass beam splitter and achromat lens to the on-board dual-channel detector (descanned path).
    
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    of the OptiphotIIYs epifluorescence filter cubes with a custom cube (Omega Optical) that transmits IR from above and reflects visible fluorescence forward to a photomultiplier tube (PMT; Hamamatsu R928) that was installed just below the binocular eyepiece. One antireflection-coated lens in the filter cube allows scattered photons that would not have made it to the on-board ("descanned path") detectors to enter this "non-descanned path" detector. It is crucial to protect the PMT from scattered and reflected excitation light by sealing a BG39 colored-glass filter to the PMT housing. (A different filter may be more appropriate for red-emitting fluorophores.) I installed circuitry to turn off the PMT when any of the microscope's other light sources are turned on (substage tungsten lamp, epifluorescence lamp, or on-board confocal laser), to prevent accidental damage to the PMT. Although the original detectors are much less sensitive than the nondescanned detector, they are still used for double-labeling experiments.
    
    Imaging Neurons Transplanted to Cultured Slices
    Survival of transplanted neurons is often disappointingly low (Shetty and Turner 1995). Caltech professor Jerry Pine and I developed a model system for neuronal transplants that allows continuous imaging of transplant integration into the host tissue (Potter et al. 1996b; Fraser et al. 1997) to shed light on the dynamics of transplant migration and integration success or failure. The idea was to label a suspension of neurons from embryonic rat hippocampus with fluorescent membrane dye, wash away all free dye and labeled debris, and seed the cells onto cultured hippocampal slices from neonatal rats (see Fig. 4). 4D two-photon imaging showed that after one day in culture, the transplanted cells migrated throughout the slice and began to extend axons and dendrites. For an example of such a 4D movie, see the following WWW site: http://www.neuro.gatech.edu/groups/potter/movies.html. At 17 days, transplanted neurons had developed pyramidal morphologies, with processes extending for several hundreds of microns (Potter et al. 19960).
    
    Cell Labeling
    Lipophilic dyes, such as DiI or DiO (generic acronyms for dialkylindocarbocyanines and dialkyloxacarbocyanines, respectively), are often used to trace living neuronal processes. Small structures such as axons, filipodia, and dendritic spines contain little cytosol and are better visualized with a membrane
    
    El 8 hippocampus
    
    <=^
    -
    
    Dissociatecells, stain supensionwith Dil or DiO, wash by spinning thru BSA solution
    
    -a
    300 pm transverse hippocampal slice
    
    Seed washed, stained cell suspension onto cultured slice Incubate at 35OC, 5% C02
    
    P9-PI3hippocampus
    
    Slice using tissue chopper or vibratome
    
    Grow
    
    slices on porous tissue culture membrane
    
    /
    
    Image on 2-Photon microscope
    
    FIGURE 4. "In Slico'' transplant model. Cells from an embryonic rat hippocampus are labeled and seeded onto cultured neonatal hippocampal slices. After some time in culture, transplant integration is observed in living slices using two-photon microscopy.
    
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    dye than a cytosolic dye. The problem is that these dyes tend to crystallize in aqueous media. Dye crystals are difficult or impossible to remove from a cell suspension, and will label host neurons if transplanted with the labeled cells. To surmount this problem, I used the more soluble 12-carbon form of DiI, along with a nonionic macromolecular surfactant Pluronic F127 (Lojewska and Loew 1986,1987), to ensure that the dye remains completely dissolved and to aid the transfer of dye to the cell membrane. The 16-carbon form of DiO, which is slightly less soluble, was also used. Normally excited by blue light (485 nm), DiO is easier to excite with the Tizsapphire laser than DiI (normally excited by green light around 548 nm). The following methods are from Potter et al. (1996b),which has further details.
    
    PROCEDURE A Preparation of Clean Cell Suspensions
    Cut brain tissue into 1-mm pieces. Digest with 0.25% trypsin , 0.02 mglml DNase (Sigma) in Hanks' balanced salt solution (no calcium or magnesium [GIBCO]) 37OC for 15 min. at Gently wash pieces in plating medium. Repeat wash. Note: For hippocampus, use Neurobasal with B27 supplement (GIBCO)and 500 (IM glutamine,25 p~
    glutamate.
    
    Gently triturate in 1 ml of plating medium with five passes through the 0.78-mm opening of a blue tip of a P- 1000 pipetman. Note: This is much more reproducible than with the ubiquitous "fire-polished Pasteur pipette." Decant suspended cells and triturate the remaining pieces once more (see step 3, above). Gravity-filter the cell suspension through a 40-pm nylon mesh (Falcon) to remove large debris. To remove smaller debris, layer 0.5 ml of 5% bovine serum albumin in IXphosphate-buffered saline (BSAIPBS) under cell suspension in a 15-ml tube. Centrifuge the cell suspension through the BSAIPBS layer in a swinging-bucket rotor at 160g for 2 min. Discard supernatant and resuspend cell pellet in plating medium by gentle trituration (three passes) as described in step 3 (above).
    Caution:See Appendix 3 for appropriate handling of materials marked with .
    
    PROCEDURE B Dil Labeling of Neurons in Suspension
    Prepare a 40 mglml stock solution of DiICn  (Molecular Probes) by dissolving the tar-like dye in DMF containing 2.5% (wlv) Pluronic F127 (BASF). Note: This solution can be stored at -20° and dissolved easily upon warming to room temperature. Prepare staining solution: Add 2 pl of dye stock to 2 ml of plating medium in a 15-ml tube (final concentration: 40 pglml DiIC,,, 0.0025% Pluronic F127,0.1% DMF). Note: This solution should easily pass through a 0.2-pm sterile filter with no diminution in color, indicating that it is completely dissolved.
    
    Warm staining solution to 37OC in a water bath. Add approximately 0.5 ml of cell suspension (from step 7, above) with gentle mixing to the staining solution. Note: ~ a b e l i will be incomplete a t room temperature due to decreased membrane turnover. n~ Incubate cells at 37OC for 15 min. Layer 0.5 ml of 5% BSAIPBS under the cells in staining solution using a long Pasteur pipette. Centrifuge in a swinging-bucket rotor at 160g for 6 min. Carefully decant the staining solution floating above the BSAIPBS, as well as most of the BSAIPBS.
    
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    7. To resuspend the stained cells, triturate (three passes) in 1 ml of plating medium using the
    P-1000 as described in step 3 of Procedure A.
    
    8. Repeat steps 5-7 (above) to remove any remaining dissolved dye. Caution: See Appendix 3 for appropriate handling of materials marked with .
    
    PROCEDURE C
    
    Labeled Transplants to Slice Cultures
    These static (membrane) cultures are prepared according to the method developed by Stoppini and coworkers (Stoppini et al. 1991; Buchs et al. 1993).Routinely grown over 1 month in culture, they are several cell layers thick (-150 pm) and retain more of the original architechtonics than do rollertube cultures.
    
    1. Cut hippocampal (or other brain tissue) slices (300 pm) from postnatal day 7-1 1 rat pups using a tissue chopper or vibratome, keeping the slices submerged in Hanks' solution (GIBCO).
    2. Add 1 ml of Organotypic Slice Culture Medium (Vanderklish et al. 1992) to a 35-mm petri dish and place a Millicell-CM membrane insert upon the medium. (Organotypic Slice Culture Medium: minimal essential medium [MEM] with Hanks' salts, no glutamine [GIBCO], 25% horse serum [Hyclone], 30 m glucose, 5 m~ NaHCO,, 30 mM HEPES, 2.5 mM MgSO, , 2 M m CaC12, 3 m L-glutamine,1 mglliter insulin [Sigma], 10 mllliter penicillin/streptomycin M M [Sigma P07811. Adjust pH to 7.2 at 35OC.) Note: The membrane becomes clear when wet, facilitating imaging from below. The plastic ring on
    standard inserts gets in the way of petri dish lids, electrodes,and large objective lenses (upright scope), so this was initially cut off with a hot nichrome wire. I later persuaded Millipore to produce "Low height" inserts (#PICMORG50) to avoid this hassle.
    
    3. Transfer 1-3 slices to the membrane using a small spatula whose edges have been rounded with a grindstone. Note: The culture medium soaks through the membrane from below, and the moist slice is well oxygenated from above.
    
    4. Incubate slices at 35OC in humidified 5% carbon dioxide atmosphere. Replace half the medium under the membrane with fresh medium weekly. Note: Slices will adhere well to the membranes in a day or two and flatten out to about 150 pm. For
    submerged imaging sooner than this, the slices must be glued to the membrane using a- clot of fibrin (Brown et al. 1993).
    
    5. Dilute freshly labeled cells (from step 6 , p. 64) to 50,000 cells/ml in slice culture medium.
    6. Place a drop (-5 pl) of cell suspension on each slice, at least one day after preparing slice cultures. Notes: In a few minutes after addition of the cell suspension, excess medium will soak through the membrane, and the cells will settle on and adhere to the slice. I have found that within a day, embryonic hippocampal neurons have migrated throughout the 150-pm thickness of a cultured slice. This
    method provides 20-50 labeled cells per slice.
    
    Caution: See Appendix 3 for appropriate handling of materials marked with .
    
    Environmental Control during Imaging
    I enclose the microscope body within Reflectix Mylar bubble-plastic insulation (http://www.reflectixinc.com/) (see Fig. I), connected to a chicken egg incubator (Marsh Automatic Incubator, Lyon Electric) to warm the air to rat body temperature. Using this low-wattage heater (100 W), there are no thermostat-related temperature fluctuations. It takes about 12 hours for the microscope stage, objec-
    
    tive, and other components of the setup to warm up to S Â ° C everything remains thermally stable but during the imaging. This means there is no focus drift during protracted imaging sessions, even if the room temperature changes. The opaque enclosure also serves to block ambient light from entering the objective and causing unwanted background signal. This is especially important when using the sensitive non-descanned detector. For slices that have adhered to the membranes, I usually flood the slice with slice culture medium during imaging, and 40~10.75-NANikon or 63~11.2-NAZeiss waterimmersion objective is used. Nonadhered slices must be glued to the membrane using a fibrin clot (Brown et al. 1993). The membrane insert is weighted down by a custom-made, stainless steel ring. To prevent evaporation of medium and pH drift, a condom with the tip cut off (Trojan nonlubricated, Carter-Wallace) seals the ring to the objective lens (see Fig. 3). Because the objective lens is warm, it is also possible to image a moist nonflooded slice using an air objective, without condensation fogging the lens.
    
    Data Acquisition
    The two-photon absorption maximum for DiI is approximately 1030 nm (Xu et al. 1996), and the power of the Ti:sapphire laser falls off from 900 to 1000 nm; thus, an empirically determined excitation wavelength of 960 nm seems to give the best images of DiI-labeled specimens. At 900 nm, where the laser is more powerful, background autofluorescence increases more than DiI fluorescence. For imaging neurons labeled with DiO, two-photon excitation anywhere from 850 to 900 nm produces excellent signal-to-background ratios. Although photodamage is tens or hundreds of times less with two-photon microscopy than with standard wide-field fluorescence or confocal microscopy, it is possible to use too much laser power, excite already-excited fluorophores, and cause excessive dye bleaching. For this reason, I always set the PMT at 80-100% of its maximum voltage, and reduce the laser power to the minimum necessary to provide a good fluorescence signal at this gain, usually 10-30% transmission. Each time point of a 4D time-lapse movie consists of a series of 20-50 z-sections, taken at 0.3- to 2-pm increments. With a 4 0 objective, a volume as large as 250 x 250 x 100 pm can be imaged every ~ 5 minutes for over 8 hours. As long as the above guidelines are adhered to, there is no sign of photobleaching or phototoxicity of labeled neurons. To save on data storage and post-processing time, one can collect widely spaced (2 pm) z-sections and average two or three scans per section to reduce shot noise. However, better noise reduction is obtained by taking more single-scanned sections that are more closely spaced (0.5 pm or less), and then subjecting them to a 3 x 3 x 3 median filter using Imagespace (or equivalent) software. Using 3D filters is only valid when each section is very similar to the two adjacent sections.
    
    TWO-PHOTON, 4D TIME-LAPSE IMAGING OF DENDRITIC SPINES
    In addition to its usefulness for revealing the dynamics of developing neural systems, two-photon microscopy is useful for revealing potential morphological correlates of synaptic plasticity in adult brain tissue. Dendritic spines have been imaged in acute hippocampal slices from juvenile rats, to study structural changes associated with synaptic facilitation (E. Schuman et al., unpubl.).
    
    Maintenance of Healthy Slices
    Maintaining acute slices poses a number of technical difficulties compared with organotypic cultured slices. The adult hippocampus is much more susceptible to excitotoxic injury from anoxia than the neonatal hippocampus. The Mobile Slice Unit (see Fig. l), an instrument designed to keep acute slices healthy for up to 24 hours during imaging, was developed (A. Mamelak and S. Potter, unpubl.). This is a portable heart-lung machine that bubbles blood gas (95% oxygen/5% carbon dioxide) through artificial cerebrospinal fluid (ACSF) that is perfused continuously across the slice (150 mllhour) by a peristaltic pump.
    
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    Medium Preparation
    ACSF is equilibrated with blood gas several hours before use to prevent foaming on addition of B27 and then mixed with B27 nutrient supplement (GIBCO) using a syringe pump at 150 volume ratio (ACSF: 25 M M glucose [Edwards 19951,126r t NaCl, 2.5 mM KC1 , 1 mM MgCl,  , 2 m~ CaC12 ~ , 1.25 mM sodium phosphate , 26 m NaHCOJ. The pH and osmolarity are adjusted to 7.3 M and 295 mOsm, respectively.
    Note: Phenol Red pH indicator is routinely included in the ACSF to ensure that the gassing is effective, because this is a carbonate-buffered solution. Ideally, an in-line pH sensor and oximeter would be included.
    
    Temperature Control
    As with cultured slices, we control the temperature of the microscope to prevent thermal focus drift, but imaging is usually conducted at 25OC, because acute slices kept at 37OC survive for only 6 hours or so.
    
    Measurements
    We routinely check the field potential of slices before and aftei imaging and can induce long-term potentiation by tetanic stimulation after 20 hours of imaging, as long as the osmolarity, pH, oxygenation, and temperature are carefully controlled.
    
    Slice Movement during Imaging
    Acute slices are not adhered to a membrane as the cultured slices are. To prevent movement during imaging, I use slice weights that are a modification of those described in Edwards et al. (1989). A 1-mm-diameter silver wire is shaped into a "U" and pressed in a vise until it is 50 pm thinner than the acute slice, which is usually cut at 500 pm. This weight is then glued to a Millicell-CM insert using cyanoacrylate glue. The weight is cut out of the insert, leaving membrane in the middle of the U, and a small hole is then cut in the membrane to allow access to the slice by electrodes or micropipettes. The porous membrane slice weights are transparent when wet and do not cut into the slice as the nylon fibers of the weights used by Edwards et al. (1989) do. To allow several different sites to be imaged during one time-lapse session, we attach the slice chamber to a micromanipulator (e.g., Sutter MP-285). The manipulator controller can remember the location of several interesting regions and return to them in sequence under computer control. As long as the slice is well weighted down, the positional relocation is accurate to approximately 1 pm. This device also makes hunting around the slice for labeled cells much easier than using the microscope's mechanical stage translation knob, especially when imaging at high magnification.
    
    Acute-Slice Labeling
    A variety of approaches for labeling living slices have been developed. They include the following:
    
    Biolistics and GFP
    McAllister et al. (1995) had great success using biolistics to shoot gold particles coated with GFPencoding DNA into slices of developing ferret cortex. However, it takes 12-24 hours for GFP to be expressed and oxidized, which is not acceptable for acute hippocampal slices.
    
    Rusty-nail Approach
    Slices can be labeled by drying a solution of DiI or DiO on a glass pipette to form small crystals and poking the slice with the pipette to deposit the crystals within it (Sorra and Harris 1997; S.M. Potter, unpubl.). Although quicker than the biolistics approach, this also takes several hours for maximal labeling of dendritic arbors, presumably due to the small contact between dye crystals and cell mem-
    
    brane. One can also combine the biolistics and rusty-nail approaches, using "DiOlistics," in which combinations of dyes are dried onto gold particles that are shot into brain tissue to provide rapid multicolor labeling of a random subset of neurons (Gan et al. 2000).
    
    Oil-drop Approach
    Hosokawa et al. (1992) and Dailey (Dailey and Smith 1993) successfully labeled acute and cultured slices by dissolving DiI or DiO in fish oil and applying a small drop of this solution to the surface of the slice. Unfortunately, the cells best labeled by this technique are the unhealthy ones near the cut surface. In addition, the oil drop must be removed by microaspiration before imaging, because it acts like a lens to defocus the excitation beam, reducing two-photon excitation.
    
    Intracellular Dye Injection
    We have produced some nice two-photon images of single neurons filled with fluorescein-dextran by intracellular iontophoretic injection. However, seldom do neurons survive for more than an hour after the trauma of impalement and electrode removal. Also, as mentioned on p. 63, membrane dyes produce clearer images of small structures such as dendritic spines.
    
    Picospritzing a DMF Solution
    We have had great success using a solution of DiO (16 mglml) in DMF. A very small amount can be applied by extracellular pressure injection into the middle of the slice. By varying pressure and pulse duration, any number from two to many neurons can be labeled. Because the carrier is miscible with water, the dye surrounds the cells and they become maximally labeled within 30 minutes. No toxicity has been observed from either the carrier or the dye.
    
    Imaging and Post-processing
    The demands and parameters for imaging spines in acute slices are similar to those used on cultured slices. However, to observe submicrometer changes in the structure of spines, which are typically 1-3 pm long and less than 1 pm wide, the smallest pixel size available should be used. Using a 63~11.2-NA water-immersion objective lens, this is 0.08 pm. Thus, a 512 x 512 scan is only 40 pm wide. This exacerbates any drift problems and makes control of temperature and osmolarity (which affects swelling or shrinking of tissue) even more critical. Inevitably, some drift or movement must be removed during post-processing, for example, by using the "register" function in NIH Image. This Macintosh software is available free at the following WWW site: http://rsb.info.nih.gov/nih-image. To virtually eliminate noise and greatly increase the z-resolution of the images, we have found it helpful to perform image deconvolution of the raw data (see Fig. 5). A model of how the microscope optics blurs the image is created by imaging subresolution fluorescent spheres (Molecular Probes MultiSpeck kit) under conditions as similar as possible to the imaging of the dendrites. The Huygens System is presently the only deconvolution software that deals appropriately with multiphoton images (Vankempen et al. 1997). It takes several hours on an SGI 0 2 workstation (Silicon Graphics) to deconvolve a single z-series, but the improvement is dramatic. As with the 3D median filter, slices must be collected at very close intervals (0.5 pm or less) for effective deconvolution.
    
    -
    
    IMPROVEMENTS FOR THE FUTURE
    The benefits of two-photon microscopy for imaging living specimens are clear. However, the technology will not be widely implemented until the lasers needed become much cheaper. Many labs that are not mechanically or optically inclined are waiting for a true "turnkey" two-photon system, one that is designed from the ground up for optimum IR throughput and efficient, non-descanned detection.
    
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    Blurring due
    * -
    
    to optics
    
    4
    
    -.
    
    Deconvolution
    
    Sub-resolution point
    
    Point-Spread Function
    
    Deconvolved Point
    
    Real dendrite, actual shape unknown
    
    Raw data
    
    'Maximum Likelihood'' model of dendrite
    
    FIGURE 5. Image deconvolution. A 0.1-pm fluorescent sphere is imaged to obtain the microscope's pointspread function, which is then used to numerically deblur (deconvolve) a raw set of 3D-image data. We use the maximum likelihood estimation algorithm of the Huygens software suite.
    
    Although most labels that are in current use with standard fluorescence microscopy also work with multiphoton microscopy, the potential exists for dyes optimized for multiphoton imaging (Albota et al. 1998). Fluorescent proteins provide relatively noninvasive labeling for a variety of specimens. In collaborations with Paul Garrity and Peter Mombaerts, I have had great success using two-photon microscopy to image GFP-labeled neurons in living fruitfly optic lobes and mouse olfactory bulb (Potter et al. 1996c; Mombaerts et al. 1996, respectively).Transgenic mice expressing membrane-associated GFP in neuronal subpopulations would eliminate the need for tedious labeling procedures (Feng et al. 2000; Hadjantonakis et al. 2003).
    
    ACKNOWLEDGMENTS
    I thank Professors Jerry Pine and Scott Fraser for their continued support and guidance. I thank Arno Klein for technical assistance with the detector installation and a critical reading of this chapter. This work was supported by the Beckrnan Foundation and National Research Service Award F32NS1025702 from the National Institute of Neurological Disorders and Stroke.
    
    REFERENCES
    Albota M., Beljonne D., Bredas J.L.,Ehrlich J.E., Fu J.Y., Heikal A.A., Hess S.E., Kogej T., Levin M.D., Marder S.R., McCord-Maughon D., Perry J.W., Rockel H., Rumi M., Subramaniam G., Webb W.W., Wu X.L., and Xu C. 1998. Design of organic molecules with large two-photon absorption cross sections. Science 281: 1653-1656. Brown L.F., Lanir N., McDonagh J., Tognazzi K., Dvorak A.M., and Dvorak H.F. 1993. Fibroblast migration in fibrin gel matrices. Am. J. Pathol. 142: 273-283. Buchs P.A., Stoppini L., and Muller D. 1993. Structural modifications associated with synaptic development in area CAI of rat hippocampal organotypic cultures. Dev. Brain Res. 71: 81-91. Dailey M.E. and Smith S.J. 1993. Confocal imaging of mossy fiber growth in live hippocampal slices. Jpn. J. Physiol. 43: S183-S192.
    
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