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The first step in the detection of light by vertebrate photoreceptors is the photoisomerization of the retinyl chromophore of their visual pigment from 11-cis to the all-trans configuration. This initial reaction leads not only to an activated form of the visual pigment, Meta II, that initiates reactions of the visual transduction cascade, but also to the photochemical destruction of the visual pigment. By a series of reactions termed the visual cycle, native visual pigment is regenerated. These coordinated reactions take place in the photoreceptors themselves as well as the adjacent pigment epithelium and Müller cells. The critical initial steps in the visual cycle are the release of all-trans retinal from the photoactivated pigment and its reduction to all-trans retinol. The goal of this monograph is to describe methods of fluorescence imaging that allow the measurement of changes in the concentration of all-trans retinol as it is reduced from all-trans retinal in isolated intact salamander and mouse photoreceptors. The kinetics of all-trans retinol formation depend on cellular factors that include the visual pigment and photoreceptor cell type, as well as the cytoarchitecture of outer segments. In general, all-trans retinol forms much faster in cone cells than in rods.
Absorption of incoming light by the visual pigment of vertebrate photoreceptors isomerizes its retinyl chromophore from 11-cis to all-trans. This photoisomerization results in the formation of Meta II, an enzymatically active visual pigment conformation, and begins the transduction of light to an electrical signal that can be transmitted to the brain (see (1) for review). The all-trans chromophore is then removed and recycled to reform 11-cis retinal that can be used to regenerate the pigment (see (2-5) for reviews). The first steps of this process take place in the photoreceptor outer segment and culminate in the generation of all-trans retinol, which is formed through the reduction of the all-trans retinal released from the photoactivated visual pigment. The conversion of all-trans retinal to all-trans retinol is catalyzed by the enzyme retinol dehydrogenase (RDH) and requires metabolic input in the form of NADPH. In the case of rods, all-trans retinol is transferred to neighboring cells of the retinal pigment epithelium (RPE) where it is esterified by lecithin retinyl acyltransferase. The resultant ester is the substrate for the Rpe65 isomerohydrolase that generates 11-cis retinol, which is then oxidized to 11-cis retinal and translocated to the photoreceptor outer segments; there, it condenses with opsin left behind following the release of all-trans retinal to regenerate the visual pigment. In the case of cones, an additional pathway involving the Müller cells can generate 11-cis retinal and regenerate the visual pigment independently of the RPE (6, 7). The transfer of all-trans retinol from the outer segments appears to occur through mass action (8-11), and allows for the recycling of the chromophore to make fresh 11-cis retinal. In the rest of the chapter, where unspecified, retinal and retinol refer to their all-trans isomers.
Several steps comprise the pathway that forms all-trans retinol in the outer segment after the photoisomerization of the visual pigment’s 11-cis retinyl chromophore. The initial step is the hydrolysis of the Schiff base bond between the chromophore and the visual pigment protein and the release of all-trans retinal. The released all-trans retinal may be sequestered inside the disks in the form of a Schiff base with phosphatidylethanolamine; it can then become available for reduction after it has been transferred to the cytosol by the ABCA4 transporter (12). The final step is the reduction of all-trans retinal by RDH, a reaction that uses NADPH as a co-factor. Because of the substantial amount of chromophore present in photoreceptor outer segments (3-4 mmol/L (13)), the kinetics of RDH and the availability of NADPH may have a significant impact on the overall kinetics of retinol formation. In summary, the kinetics of retinol formation depend on the rate of retinal release, local NADPH availability, retinal access to RDH, RDH kinetics, and retinol elimination rate (that depends on outer segment morphology).
The kinetics of the formation of retinol after light excitation can be monitored in the outer segments of single photoreceptors from its intrinsic fluorescence, which has been shown to provide a good measure of its concentration (9, 14). Such single cell fluorescence measurements with their high sensitivity and time resolution provide a unique window for examining the different steps in the formation of retinol. They have been used to study the formation of retinol in the rod and cone photoreceptors from several vertebrate species, including tiger salamander (Ambystoma tigrinum), grass frog (Rana pipiens), lizard (Gecko gecko), and mouse (Mus musculus) (8-10, 14-16). From among these species, tiger salamanders and mice offer several distinct advantages as model systems for the study of retinol formation. From the salamander retina, one can obtain different types of rods and cones that include two types of rods, green- and blue-sensitive, and three types of cones, red-, blue- and UV-sensitive (10, 17). These salamander cells are large, robust, and can survive for several hours after isolation from the retina, facilitating the experimental manipulations for single cell imaging. Their large size also allows measurement of the time course and kinetics of retinol formation in local regions of the outer segments. Salamander photoreceptors utilize different pigment types as well as the same pigment in different cell types (10, 18). Thus, they allow the comprehensive examination of the dependence of retinol formation on cell type, visual pigment type, and outer segment architecture. The mouse retina offers complementary advantages. It is dominated by a single cell type, the rods, allowing comparisons with biochemical measurements from isolated retinas (14) and the eyes of whole animals. Furthermore, the availability of genetically modified animals offers the opportunity to probe specific enzyme involvement and disease relevance.
Figure 1 shows a diagram of the setup that is used for such measurements, and includes an inverted microscope, fitted with a near UV fluorescence excitation light source and a high sensitivity camera. The orientation and function of these separate components is described below (Methods) and in the figure legend. An experiment begins with placement of dark-adapted photoreceptor cells in the experimental chamber. After an initial fluorescence measurement in their dark-adapted condition, the cells are illuminated to ensure that virtually all of the visual pigment chromophore has been isomerized to the all-trans conformation. The reason behind this is that the UV light used to excite the fluorescence of retinol is also absorbed by the visual pigment and thus isomerizes the 11-cis chromophore. So, to avoid having each measurement of fluorescence initiating the reactions that lead to additional retinol formation, each experiment begins with photoactivation (bleaching) of virtually all of the visual pigment. Figure 2 shows an experiment performed on an isolated salamander green-sensitive rod photoreceptor in this way. Here it can be seen that, following quantitative bleaching of the visual pigment, retinol fluorescence increases within the outer segment, assuming a maximal value at about 30 min, and declining slowly thereafter. The data presented in Figure 3 illustrate a similar experiment performed on an isolated salamander red-sensitive cone cell. In this case, retinol fluorescence increases rapidly after bleaching, reaching a maximal value after ca. 1 min. Subsequently, it declines at a much faster rate than in the rod. Finally, Figure 4 presents an experiment with an isolated mouse rod, performed at 37 °C.
A dark room is necessary for the fluorescence imaging setup. The same room can be used for dark-adapting animals and for dissection. An area of ~100-150 sq.ft. is sufficient. A revolving door for entering is convenient, but a thick black curtain is also adequate.
1It is critical that the buffer composition is accurate within a few percent. The osmolarity of the solution affects the function and viability of the cells, especially the murine ones.
2We have obtained the same results using a Ringer’s solution with a slightly different composition: 110 mM NaCl, 2.5 mM KCl, 1.6 mM MgCl2, 1.0 mM CaCl2, 10 mM glucose, 10 mM HEPES, pH 7.8.
3The addition of bovine serum albumin is not strictly necessary, but we have found that it appears to improve the viability of the cells. At this concentration, albumin does not affect the removal of retinol from the outer segment.
4This long pass filter can be used for bleaching the visual pigments of the salamander green-sensitive rods and red-sensitive cones, and mouse rods. These three cell types comprise the overwhelming majority of the cells isolated from the retinas of these two species. For experiments with the salamander blue-sensitive rods and cones and UV-sensitive cones, a much more demanding technical approach is required, including different filters for bleaching the visual pigments of these cells (see section 3.2 paragraph 5).
5You can place solutions for perfusion on the top of the cage (instead of inside). In that case, the top plate should have a 2-3 inch diameter hole to allow the tubes carrying the solutions to enter the cage.
6You can improve the light-tightness of the enclosure by attaching a 5-inch skirt made of black cloth around the bottom edges of the plates.
7The width of the inside curtain should be the same as the width of the frame; its top side should be permanently attached to the frame. The horizontal rod at the bottom is used to roll the curtain up or down as needed.
8The outside curtain must fully cover the inside one, so its width is longer than the frame. It is not permanently attached to the frame, and is kept in place during experiments with Velcro strips at the top, and the right and left sides. Place the Velcro on the frame and the curtain, so that the curtain is securely held but is also loose at the bottom. This is necessary to allow the hands of the experimenter to go under the outside curtain and through the slits of the inside one to reach the microscope.
9A ½-inch thick piece of wood that is about 5 inches deep and runs the length of the front side of the cage is adequate.
10The flow rate can be controlled by adjusting the height of the reservoir containing the solution or by compressing the plastic tubing that brings the solution from the reservoir to the chamber.
11Placement of the needle is critical for avoiding complete removal of the solution in the chamber and drying the cells. A disadvantage of the suction system is that it is difficult to avoid pulsation, because the height of the solution in the chamber tends to fluctuate. Because the suction generates a sound however, it allows monitoring of perfusion from the outside of the light-tight cage. Cessation of the sound is an important warning sign, indicating potential flooding on the microscope stage.
12The high numerical aperture is necessary to collect as much fluorescence signal as possible. A 100× lens might be somewhat more helpful especially for visualizing the small mouse photoreceptor cells, but because of the smaller field of view it makes it more difficult to find a cell. In our experience, an oil-immersion 40× lens with 1.3 numerical aperture (for example the Zeiss Plan Neofluar or even the Fluar) is perfectly adequate.
13The objective lens is a major heat sink and heating it helps to stabilize the temperature in the experimental chamber. Temperature fluctuations in the lens change the temperature of the immersion oil, resulting in changes in its refractive index and focus drift.
14The shutter is essential to avoid photobleaching of retinol. The cells should be exposed to the excitation light only for image capture.
15Do not crowd or try to cover the Xenon light source to reduce light leak. The lamp generates a lot of heat and good ventilation is necessary to avoid overheating.
16The absorption maximum of retinol is 325 nm, and its fluorescence emission maximum is ca. 480 nm (15). The glass optics used (lenses and light guide) transmit poorly <350 nm, hence the use of the particular excitation filter. Use of very expensive quartz optics should allow efficient excitation with 325 nm light, but we have no direct experience with such a system. The dichroic mirror and the longpass emission filter are selected to collect most of the fluorescence emission.
17Do your best to ensure the health of the animals, because the health of the cells depends on it.
18It is not necessary to use filter paper, but we have found that it helps for handling the small salamander and mouse eyes by keeping them in place.
19Sometimes it is difficult to separate the retina from the pigment epithelium. Slowly peel off starting from the periphery. If you still cannot separate the retina, a likely possibility is incomplete dark adaptation, which could be caused by too bright a red light. Ensure that the animal is dark-adapted properly, and dim the red light.
20It is a good idea to cut a small piece of retina and transfer it to a separate Petri dish. Under room lights this piece of retina should be a bright red color (due to rhodopsin) that fades rapidly. The red color indicates the presence of rod outer segments and confirms that you have obtained a healthy retina. Lack of red color indicates either an unhealthy retina or a failure to separate the rod outer segments from the retinal pigment epithelium. In such case, you should ensure the health of the animals and proper dark adaptation.
21Before you embark on actual experiments, you need to ensure that the chopping and the cell density have been optimized. A very high cell density will result in cells settling on top of each other, disallowing an experiment. A very low density might result in failure to find a cell for experiment. Check your isolated cell preparations under the infrared illumination of the microscope, and adjust chopping and density until you can regularly obtain isolated intact cells, that have settled without cells above or below them.
22At the end of the day, remove any residual oil on the lens with lens paper. With use, oil seeps under the lens. Remove and clean the lens regularly with cleaning fluid and lens paper. You can use cotton-tipped applicators to clean the back surface of the lens.
23After the first experiment of the day, the equipment has been switched on. If you need to switch off the Xenon lamp to place a second dish on the microscope stage, then you will need to switch off all pieces of equipment (see Note 24 below). However, with the Xenon lamp on the floor, the light leaking from it should not reach the microscope stage inside the cage. If this is indeed the case, switch off only the computer monitor, and open the light-tight box containing the chamber with the cells inside the cage. Continue as with the first dish of the day.
24When the Xenon lamp is turned on, it could generate a voltage spike that might damage other electronic equipment.
25Blue-sensitive salamander rods and blue- and UV-sensitive cones are each morphologically distinct and can be distinguished from the green-sensitive rods and red-sensitive cones (17). For experiments with these types of cells however, it is best to independently corroborate their identity by measuring their spectral sensitivity. Such measurements require additional equipment (10).
26An important additional parameter for image acquisition is image binning. Binning improves the signal-to-noise ratio at the expense of resolution by grouping the output of camera pixels. With the specified hardware components, and exposure times 100-1000 ms, we find that 2×2 and 4×4 binning provide images with adequate resolution and good signal-to-noise ratio.
27In cone outer segments, retinol fluorescence increases rapidly after bleaching, so it is necessary to employ a time delay image acquisition routine to capture a series of images in rapid succession. This image acquisition routine is an option provided by the software and is necessary for capturing images with time delays less than 1 min. For longer time delays between images, single image capture is generally better. Using a time delay routine for an experiment lasting for more than 5 min, although not impossible, is usually unsuccessful, because the focus drifts and the cell might move. Refocusing with the IR before each image capture is necessary for these longer experiments.
Yiannis Koutalos, Departments of Ophthalmology and Neurosciences Medical University of South Carolina 167 Ashley Avenue Charleston, SC 29425 Tel: (843)-792-9180 FAX: (843)-792-1723 ; Email: ude.csum@olatuok..
M. Carter Cornwall, Department of Physiology and Biophysics Boston University School of Medicine 715 Albany Street, L 714 Boston, MA 02118 Tel: (617)-638-4256 FAX: (617)-638-4273 ; Email: ude.ub@llawnroc.